Progressive abnormalities in skeletal muscle and ... · Progressive abnormalities in skeletal muscle and neuromuscular junctions of transgenic mice expressing the Huntington’s disease

Progressive abnormalities in skeletal muscle andneuromuscular junctions of transgenic mice expressingthe Huntington’s disease mutation

Richard R. Ribchester,1 Derek Thomson,1 Nigel I. Wood,2 Tim Hinks,2 Thomas H. Gillingwater,1 Thomas M. Wishart,1

Felipe A. Court1 and A. Jennifer Morton2

1Division of Neuroscience, University of Edinburgh, George Square, Edinburgh EH8 9JZ, UK2Department of Pharmacology, University of Cambridge, Tennis Court Road, Cambridge CB2 1PD, UK

Keywords: atrophy, electrophysiology, Huntington’s disease, neuromuscular junction, R6 ⁄ 2 mice, skeletal muscle

Abstract

Huntington’s disease (HD) is a neurodegenerative disorder with complex symptoms dominated by progressive motor dysfunction.Skeletal muscle atrophy is common in HD patients. Because the HD mutation is expressed in skeletal muscle as well as brain, wewondered whether the muscle changes arise from primary pathology. We used R6 ⁄ 2 transgenic mice for our studies. Unlikedenervation atrophy, skeletal muscle atrophy in R6 ⁄ 2 mice occurs uniformly. Paradoxically however, skeletal muscles show age-dependent denervation-like abnormalities, including supersensitivity to acetylcholine, decreased sensitivity to l-conotoxin, andanode-break action potentials. Morphological abnormalities of neuromuscular junctions are also present, particularly in older R6 ⁄ 2mice. Severely affected R6 ⁄ 2 mice show a progressive increase in the number of motor endplates that fail to respond to nervestimulation. Surprisingly, there was no constitutive sprouting of motor neurons in R6 ⁄ 2 muscles, even in severely atrophic musclesthat showed other denervation-like characteristics. In fact, there was an age-dependent loss of regenerative capacity of motorneurons in R6 ⁄ 2 mice. Because muscle fibers appear to be released from the activity-dependent cues that regulate membraneproperties and muscle size, and motor axons and nerve terminals become impaired in their capacity to release neurotransmitter andto respond to stimuli that normally evoke sprouting and adaptive reinnervation, we speculate that in these mice there is a progressivedissociation of trophic signalling between motor neurons and skeletal muscle. However, irrespective of the cause, the abnormalitiesat neuromuscular junctions we report here are likely to contribute to the pathological phenotype in R6 ⁄ 2 mice, particularly in latestages of the disease.

Introduction

Huntington’s disease (HD) is a fatal neurodegenerative disordercharacterized by progressive decline in motor and cognitive functionwith insidious onset in the third to fifth decade. In adults, the first motorsymptom of HD is usually chorea, but the motor deficits progress inadvanced disease to rigidity, bradykinesia and dystonia. By the endstages, theHDpatient is usually bedriddenwith very limited capacity forvoluntary movement (for references, see Bates et al., 2002).Although the genetic mutation causing HD has been identified

(as an expanded CAG repeat that is translated into a polyglutaminerepeat in the protein huntingtin), the mechanism underlying thepathology is unknown. The pathological changes in the brains ofHD patients have been well described, and the striking neurode-generation seen in the caudate and putamen has, for many years,focused attention on the mechanisms underlying pathology in theseregions. However, with the cloning of the gene came the discoverythat the expression of the HD gene and its protein producthuntingtin is not restricted to the brain (The Huntington’s DiseaseCollaborative Research Group, 1993). Indeed, huntingtin expression

is not even neuron-specific, but is found in many tissues includingheart and skeletal muscle. The role of huntingtin in these tissues isunknown, although many patients exhibit signs of peripheral motorpathology, including abnormal eye movements, difficulty swallow-ing, gait abnormality, dysarthria and skeletal muscle wasting (forreferences, see Bates et al., 2002). Further, skeletal muscle atrophy(which undoubtedly contributes to weight loss) is observed in manyHD patients despite an adequate diet and feeding (Sanberg et al.,1981). Because the general health of HD patients declines as theirdisorder progresses, it has generally been assumed that changes inbody mass are secondary to other symptoms. However, significantbody weight changes are measurable in early (Djousse et al., 2002)as well as late stage HD patients. This suggests that weight lossdue to atrophy is a significant pathological component of HD.Nevertheless, the cause of the atrophy remains unknown and,because motor function is under the control of the brain, thepossibility that a peripheral pathology contributes directly to thesesymptoms has not been widely considered.The dominant nature of the HD mutation has allowed the

development of a number of mouse models (Hickey & Chesselet,2003). The best characterized is the R6 ⁄ 2 line which is transgenic forthe human HD mutation (Mangiarini et al., 1996). R6 ⁄ 2 mice exhibita progressive neurological phenotype that includes abnormal

Correspondence: Dr Jenny Morton, as above.E-mail: [email protected]

Received 16 June 2004, revised 10 September 2004, accepted 27 September 2004

European Journal of Neuroscience, Vol. 20, pp. 3092–3114, 2004 ª Federation of European Neuroscience Societies

doi:10.1111/j.1460-9568.2004.03783.x

involuntary movements, tremor and progressive deterioration of motorand cognitive function (Mangiarini et al., 1996; Carter et al., 1999;Lione et al., 1999). As well, they exhibit a number of peripheralsymptoms which replicate those seen in HD, in particular, wasting ofskeletal muscle (Sathasivam et al., 1999).

The cause of skeletal muscle atrophy in R6 ⁄ 2 mice and HD patientsis unknown. However, skeletal muscle atrophy is an indicator of nerveand ⁄ or muscle pathology in a wide variety of diseases, and very subtleatrophies can be strongly indicative of pathology. In this studytherefore we sought to establish whether the muscle atrophy observedin the R6 ⁄ 2 mouse is part of the primary pathology associated with theHD mutation or whether it is secondary to other previouslyunrecognized features of the disorder, for example, impairment ofneuromuscular transmission. Our data suggest that both motor neuronsand muscle fibers are independently affected by the HD mutation andthat neuromuscular changes may contribute to the pathologicalphenotype in HD, particularly in late stages of the disease.

Materials and methods

Animals

All studies were carried out in accordance with the UK Animals(Scientific Procedures) Act 1986. R6 ⁄ 2 mice (Mangiarini et al., 1996)and their wild-type (WT) littermates were taken from a colonyestablished in the Department of Pharmacology, University ofCambridge, and maintained by back-crossing transgenic males ontofemale CBA · C57Bl ⁄ 6 F1 mice. Some experiments were carried outusing mice that were transferred to Edinburgh. In all cases, dry chowwas supplemented by twice daily moist chow (mash) from weaningonwards, and lowered waterspouts were also provided. We haveshown previously that this regime improved health and survival ofR6 ⁄ 2 mice (for further details of husbandry see Carter et al., 2000).Genotyping was carried out using a polymerase chain reaction basedon a modification of the method described by Mangiarini et al. (1996).Tissues from 50 WT and 42 R6 ⁄ 2 adult mice of both sexes were usedin the study.

Surgery

R6 ⁄ 2 (N ¼ 18) and WT (N ¼ 16) mice were used for experiments tostudy the neuromuscular response to denervation or reinnervation.Reinnervation experiments were done after crushing the entire sciaticnerve. Partial denervation experiments were done by cutting the tibialnerve, and examining the sprouting response of endings of the intact(sural) nerve in the partially denervated fourth deep lumbrical muscles.The mice were anaesthetized by inhalation of halothane. The sciatic ortibial nerve was exposed and a 1–2-mm section was either cut (but notremoved), or crushed using fine forceps. Mice were then kept for1–40 days before being killed by cervical dislocation.

A number of different muscles from the hind limb and the chestwere used in this study. The hind limb muscles were: flexor digitorumbrevis (FDB), the peroneal, soleus and extensor digitorum longus(EDL) muscles from the anterior calf, and the deep lumbrical musclesof the hind foot. Different muscles were used as appropriate. Forexample, FDB is a superficial muscle from the foot which is wellsuited to location and electrophysiological recording of synapticresponses. (This is on account of the short length of the muscle fibers;< 500 lm in length. As a consequence, they are ‘isopotential’ withrespect to membrane potential changes induced locally at any pointalong their length. Thus, it does not matter where the intracellularmicropipette tip is located with respect to longitudinal distance from

the motor endplate, or where the synaptic currents originate; synapticpotentials of virtually identical magnitude and time course arerecorded at all points along the length of the fiber.) The deeplumbrical muscles of the hind foot were used because they areparticularly amenable to fluorescence immunocytochemistry (becausethey are very thin, and therefore there are minimal problems withpenetration and diffusion of fixative and antibodies for immunostain-ing). Nerve terminals and endplates are readily observable in whole-mount preparations, either in regular fluorescence or confocalmicroscopes. Finally, the fourth deep lumbrical muscle has a dualnerve supply in rodents (Nakanishi & Norris, 1970; Betz et al., 1979;Taxt, 1983; Costanzo et al., 2000), one via an anastomosis of a branchof the sural nerve with the lateral plantar nerve, the other supplied bythe same nerve (tibial nerve) as the FDB muscle. Section of the tibialnerve in anaesthetized animals thus renders FDB and the mediallumbrical muscles completely denervated, but the fourth deeplumbrical muscle is subject to a controlled partial denervation, makingthe muscle ideal for studies of motor nerve sprouting. We also madewhole mounts of the triangularis sterni muscle, an exceptionally thinrespiratory muscle, well suited to immunocytochemistry.

Histology

Peroneal muscles of the hind limb were dissected and snap-frozen inliquid isopentane cooled over liquid nitrogen. Serial transversesections (20 lm) were cut from the belly of each muscle using acryostat. For fiber typing, sections were stained histochemically forenzyme activity of NADH diaphorase or succinate dehydrogenase(SDH), both standard markers for the oxidative enzymes thatcharacterize type I fibers (Filipe & Lake, 1983). Parallel sectionswere prepared for standard histological analysis by staining withHarris’s haematoxylin and eosin (stains cytoplasm and nuclei), CresylViolet (stains Nissl substance), Weigert’s iron haematoxylin (stainsnuclei) and van Geison’s stain (stains connective tissue). Selectedsections were also stained with Oil Red O (stains lipid) and theperiodic acid–Schiff reaction (stains glycogen). For detailed methods,see Culling et al. (1985). Unless otherwise stated, all reagents forhistological staining came from Sigma, UK.Fiber diameters were measured in peroneal muscle using three

sections from each of three WT and four R6 ⁄ 2 mice. Within eachsection, fibers were classified into type I or type II fibers according totheir NADH diaphorase reaction, and the diameters of at least 75 type Iand 75 type II fibers were measured and recorded using the methodo-logy described by Patel et al. (1969). Proportions of fiber type wereassessed in three sections from each muscle. All the fibers within three500 · 500-lm square fields were classified according to their NADHdiaphorase reaction, and the number of each type was counted using anoverlaid grid. To visualize intranuclear inclusions, cryosections of fresh-frozen tissues were cut onto gelatinized slides, fixed for 30 min in 4%paraformaldehyde and processed for immunocytochemistry. Sectionswere incubatedwith a rabbit antiubiquitin antibody (DAKO) at 1 : 2000dilution. A horseradish peroxidase-conjugated second antibody(1 : 1000 dilution, DAKO) was used and immunoreactive componentswere visualized using diaminobenzidine (Sigma, UK).

Neuromuscular junction staining

FDB, lumbrical and triangularis sterni muscles were prepared forimmunocytochemistry by fixing in 0.1 m PBS containing 4% parafor-maldehyde for 30–40 min. Acetylcholine (ACh) receptorswere labelledby incubation for up to 30 min in 5 lg ⁄mL TRITC-conjugateda-bungarotoxin (a-BTX: Molecular Probes, USA). Muscles were

Abnormal neuromuscular junction function in HD mice 3093

ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114

washed in PBS and blocked in 4%BSAand 0.5%Triton-X in 0.1 mPBSfor 30 min before incubation overnight in primary antibodies directedagainst either the 165-kDa neurofilament proteins (mouse monoclonal2H3), the synaptic vesicle protein SV2 (mouse monoclonal; both1 : 200 dilution; from the Developmental Studies Hybridoma Bank,Iowa, USA) or S100 antibody (rabbit polyclonal; Diagnostics Scotland,UK) or ubiquitin (rabbit polyclonal, 1 : 1000, DAKO). After washingfor 30 min in blocking solution (see above), muscles were incubated for4 h in a 1 : 200 dilution of appropriate (antimouse IgG or antirabbit)secondary antibodies conjugated to either FITC or TRITC (DiagnosticsScotland or DAKO), as indicated in the relevant figures. Muscles werethen whole-mounted in Vectashield (Vector Laboratories, Burlingame,CA, USA). Some preparations were stained for cholinesterase by amodified Karnovsky–Roots method, described in Harris & Ribchester(1979). Fiber diameter and endplate area analysis were made using thepublic domain NIH Image program (developed at the US NationalInstitutes of Health and available on the Internet at http://rsb.info.nih.-gov/nih-image/). For endplate area analysis, the region covered by theendplate was outlined and measured. This procedure overestimates thearea of motor nerve terminal contact, but it is sufficiently robust to testthe correlation between endplate size and muscle fiber size (Harris &Ribchester, 1979). Fiber diameter was measured within 100 lm of theendplate.Some preparations were ‘vital’ stained with the dye FM1-43, a

marker of endocytosis (Molecular Probes), a marker of endocytosis infunctioning nerve terminals (Betz et al., 1992; Ribchester et al., 1994;Barry & Ribchester, 1995). Preparations were incubated in oxygenateddepolarizing saline (K+ increased to 50 mm, Na+ decreased by 45 mm;concentrations of the other ions in the bathing medium were as givenunder Electrophysiology, below) containing 4 lm FM1-43, for5–15 min, washed and then imaged in situ. Immunofluorescent andvitally stained preparations were imaged using either a standardfluorescence microscope (Olympus) or a laser scanning confocalmicroscope (Biorad Radiance 2000). Confocal z–series were mergedusing Lasersharp (Biorad) software.

Electron microscopy

FDB muscles were fixed in ice-cold 0.1 m phosphate buffer contain-ing 4% paraformaldehyde and 2.5% glutaraldehyde for 4 h. Prepara-tions were then washed in 0.1 m phosphate buffer, postfixed in 1%osmium tetroxide solution for 45 min and dehydrated through anascending series of ethanol solutions before embedding in Durcupanresin (Sigma, UK) and sectioning at 75–90 nm. Sections werecollected on Formvar-coated grids (Agar Scientific, UK), stained withuranyl acetate and lead citrate and viewed in a Philips CM12 TEM.EM negatives taken between 2000· and 60000· were scanned at600 dpi using a Linoscan 1200 (Heidelberg Instruments, Heidelberg,Germany) equipped with a transparency adaptor, before importing intoAdobe Photoshop for analysis and presentation.

Electrophysiology

Intracellular recordings were made from isolated FDB nerve–musclepreparations. In initial experiments, FDB muscles were isolated inCambridge early in the morning, bathed in oxygenated HEPES-buffered physiological saline and couriered the same day toEdinburgh, where they were then transferred to bicarbonate-bufferedsaline (below) for electrophysiological recording the same afternoon,as described previously (Mack et al., 2001). The isolated muscles werepinned out in a Sylgard (VWR International, Poole, UK)-lined bath

and perfused with normal mammalian physiological saline (in mm:NaCl, 120; KCl, 5; CaCl2, 2; MgCl2, 1; NaH2PO4, 0.4; NaHCO3,23.8; d-glucose, 5.6) bubbled to equilibrium with a 5% CO2)95% O2

mixture. In later experiments, batches of R6 ⁄ 2 mice were transferredfrom Cambridge to Edinburgh and killed, and muscles were directlydissected into the bicarbonate-buffered medium. Muscle contractionswere reduced or eliminated by bathing them in 2.5 lm l-conotoxin(l-CTX) GIIIB (Scientific Marketing Associates, UK) for 30–45 min.In some experiments, l-CTX was retained in the bathing mediumthroughout subsequent recordings. Normally, up to 30 muscle fibersper muscle were sampled using microelectrodes filled with 4 m

potassium acetate (impedance � 40 MW), according to standardtechniques. Spontaneous and evoked endplate potentials were recor-ded using either WPI M707 (WP Instruments Inc.) or Axoclamp 2Bamplifiers (Axon Instruments) and stored and analysed on a PC usingeither WinWCP v3.0.8 software (developed and distributed by DrJohn Dempster, Strathclyde University), Spike-2 (Cambridge Elec-tronic Designs, Cambridge, UK), or MiniAnalysis (Synaptosoft,Atlanta, USA). In some experiments, input resistance was measuredby applying current to the recording microelectrode via a Wheatstone-bridge circuit built into the recording amplifier. Constant current wasapplied to set the resting membrane potential at )80 mV. Theadditional steady-state membrane hyperpolarization evoked by super-imposed weak hyperpolarizing current pulses (1–5 nA, 30–100 msduration) was then measured. In some fibers, stronger current pulseswere injected to test for ‘anode-break’ excitation of the muscle fiberson termination of the current pulse (Marshall & Ward, 1974). In otherexperiments, iontophoretic responses to ACh were tested usingmicropipettes filled with 1 m ACh (> 100 MW input impedance). Aholding current of 1–3 nA was applied to the pipettes to preventleakage of ACh. Carefully reducing the holding current produced slowdepolarization of the muscle fiber impaled with the recordingmicroelectrode when the tip of the iontophoretic pipette was in thevicinity of its endplate. After restoring the holding current, theelectrode was displaced � 100 lm to place its tip over the extrajunc-tional membrane and brief outward current pulses (1–10 nA, 1 ms)were applied to the iontophoretic pipette. In other experiments,extracellular recordings of muscle fiber action potentials were madeusing blunt (1–2 MW) pipettes filled with 1 m NaCl. The pipette tipwas inserted into the belly of the FDB muscles and electromyographic(EMG) responses were evoked by suprathreshold stimulation of themuscle nerve (nominally 10–30 V, 0.1–0.5 ms duration). In someexperiments, muscle tension recordings were made by pinning theproximal tendon of FDB muscles to the Sylgard-lined base of therecording chamber. The distal tendons were gathered and tied with ashort silk suture then attached to a sensitive force transducer (Akers,Olso, Norway) connected to a custom Wheatstone-bridge circuit andpreamplifier. The output was filtered and further amplified usingNeurolog equipment (Digitimer, UK). Resting muscle length wasadjusted to maximize the twitch tension and ⁄ or tetanic (50 Hz)tension responses evoked by nerve stimulation. ACh chloride (10 lm)and l-CTX (2.0 lm) were added directly to the bathing mediumduring recording of either evoked EMG or muscle tension responses.

Statistical analysis

Data are presented as mean ± SEM in each group, when n ¼ numberof muscle fibers and N ¼ number of muscles or animals used, or asmedian, interquartile range and 5–95% outliers. Where appropriate,data were analysed either by unpaired Student’s t-test, Mann–Whitneytest or one-way anova. Dunnett’s post hoc test was used to determinelevels of significance following anova.

3094 R. R. Ribchester et al.


Results

Experiments were performed over a period of 44 months, from 2000to 2004. Initially, muscles for physiological recording were dissectedin Cambridge and couriered to Edinburgh in physiological saline foranalysis and further processing the same day. In later experiments,surgery, physiological measurements, immunocytochemistry, histo-chemistry and electron microscopy were made on muscles isolatedfrom R6 ⁄ 2 mice and their WT littermates following their delivery toEdinburgh. Over the course of the study, it became evident that thedelay to onset of discernible weight loss and muscle atrophy increased,from � 9 weeks of age at the start of the study to > 11 weeks of age bythe end. Mice did not normally live beyond 16 weeks at the start of thestudy, but routinely survived to 18 weeks or older by the end. Thisimproved longevity occurred in spite of a spontaneous and progressiveincrease in the CAG repeat length in the colony over the period of thestudy (data not shown) and is attributed primarily to improvements inhusbandry (Carter et al., 2000). None the less, the data are presentedhere in relation to the chronological age of the mice, with ‘severelyaffected’ mice in all cases being older than 15 weeks.

Clinical presentation of R6 ⁄ 2 mice

R6 ⁄ 2 mice show no overt behavioural phenotype until they are� 8 weeks of age (Carter et al., 1999). Thereafter, growth of R6 ⁄ 2mice begins to slow and by 12 weeks they begin to lose weight (seebelow). At the same time they show an increasing number of abnormalmotor signs, such as excessive hind limb grooming and tremor. Likemany other mutant mice with neurological dysfunction, R6 ⁄ 2 micealso exhibit abnormal hind-limb reflex responses to tail lifting: hipflexion and toe clasping rather than extension. By 12 weeks, R6 ⁄ 2mice are noticeably weaker and less active than their WT littermatesand they perform poorly in behavioural tasks requiring strength. Forexample, they have difficulty swimming, performing on the rotorod orlifting their own body weight out of water onto an elevated platform,tasks that pose no difficulty to age-matched WT mice (Carter et al.,1999; also A.J. Morton, unpublished observations). However, in theirhome cage they appear generally healthy, and are not overtly differentfrom their WT littermates.

In the present study, body weights of R6 ⁄ 2 mice at 12 weeks were21.9 ± 0.4 g (N ¼ 17) compared to 23.6 ± 0.6 g (N ¼ 16 WT mice).By 16 weeks their body weight dropped significantly (17.5 ± 0.5 gcompared to 24.5 ± 1.2 g for R6 ⁄ 2 and WT mice, respectively). Thus,the R6 ⁄ 2 mice lost � 20% of their body weight while WT micecontinue to grow. By 15 weeks, the mice show a marked lordoky-phosis (hunchedback spine; Fig. 1a and b), poor mobility andpronounced muscle weakness. However, flexion reflex responses totail lifting persisted. Overt motor symptoms such as spontaneoushindlimb grooming are present by 12 weeks. In R6 ⁄ 2 mice, 1.2 ± 0.6instances of hindlimb grooming were recorded (in a 10-min obser-vation period in the open field; N ¼ 17) compared with 0.3 ± 0.3instances recorded in WT mice (N ¼ 16). By 15 weeks, 7.7 ± 2.7instances of hindlimb grooming were recorded in R6 ⁄ 2 mice (in10 min) compared with 0.2 ± 0.2 instances in WT mice. The motorsymptoms progress until the premature death of the mice, usuallybetween 16 and 20 weeks of age.

Skeletal muscle atrophy

By 12 weeks of age, R6 ⁄ 2 mice show visible atrophy of hind limbskeletal muscles compared with WT littermates and, by 15 weeks, this

muscle atrophy is very severe (Fig. 1c and d). To examine thepathology of the skeletal muscle in more detail, cryosections ofgluteus maximus and peroni from 15-week-old mice were processedand examined using a number of histological stains for myopathies(Anderson, 1985). Apart from the reduced diameter of fibers,microscopic examination of sections from R6 ⁄ 2 mouse showednormal morphology with haematoxylin and eosin, Cresyl Violet, VanGeison’s, Oil Red O and periodic acid–Schiff stains. The frequencyand prominence of nuclei was normal, and there were no centrallylocated nuclei. Other pathological signs that were sought, but notfound, included central cores, nemaline rods, granular cytoplasm (inhaematoxylin and eosin staining), basophilic rims, target fibers, ringfibers, cytoplasmic vacuoles, basophilic blebs, concentric lamellae,fibrosis and thickened endomysium (data not shown). Althoughatrophic, the muscle fibers looked healthy and there was no evidenceof cell death, such as inflammatory cell infiltrates, fibro-fatty change,or regenerating fibers. Oil Red O revealed no evidence of lipidaccumulation. Apart from the diffuse atrophy and fiber type changes(see below), there was no evidence of any of the other changestypically associated with muscle pathology.The extent of muscle atrophy was measured in both transverse

sections (Fig. 1e and f) and teasedmuscle fiber preparations of peroneal,SOL and EDL muscles (Fig. 2). Some transverse sections were stainedhistochemically for NADHdiaphorase to distinguish type I from type IImuscle fibers (Fig. 1c and d). By 8 weeks, muscle fiber diameter wasslightly, though significantly, smaller in R6 ⁄ 2 mice than WT mice(Fig. 1i and j). However, muscle fiber diameter collapsed between 8 and12 weeks. By 16 weeks, there was a severe, generalized atrophy of allmuscle fibers in R6 ⁄ 2 mice that we examined and diameters of musclefibers from R6 ⁄ 2 mice were half the size of age-matched WT muscles.For instance, in teased muscle fiber preparations of EDL muscles, fiberdiameter of muscles from R6 ⁄ 2 mice was 28.32 ± 0.78 lm (mean ±SEM; n ¼ 84 fibers, N ¼ 2 muscles), compared with 45.33 ± 1.06 lm(n ¼ 80 fibers, N ¼ 2 muscles) from WT muscle. Notably, both type Iand type II muscle fibers were atrophic. In transverse sections of theperoneal muscles, the diameter of type I and II muscle fibers in WTmuscles were 25.4 ± 0.3 and 41.3 ± 0.3 lm, respectively, comparedwith 15.9 ± 0.2 and 17.4 ± 0.2 lm in R6 ⁄ 2 (n ¼ 675 muscle fibers ineach case). Although atrophy occurred in both fiber types (see alsoSathasivam et al., 1999), the proportion of type I and II fibers wasaltered. InWTperoneal muscles, 51 ± 2%of fiberswere type I comparewith 81.8 ± 2.1% in the R6 ⁄ 2 mouse. However, there were no signs ofdegeneration, nor was there any marked change in grouping of musclefibers within the muscle. Thus it appears that type II fibers wereconverted to type I during the process of muscle atrophy rather than thechange occurring by another process (e.g. fiber splitting).

Fiber diameter and motor endplate area

Muscle atrophy was measured in teased fiber preparations of R6 ⁄ 2mouse EDL, soleus and FDB muscles, after staining histochemicallyfor cholinesterase at motor endplates (Harris & Ribchester, 1979). Thismethod typically stains the rims of junctions, delineating the extent ofsecondary folds. There were no discernible differences in the pattern ordensity of cholinesterase staining in R6 ⁄ 2 muscles (Figs 2a and b, and3). Every fiber showed a single motor endplate, and the ‘pretzel’-likestructure of the endplates were similar to those seen in WT muscle.Further, there was no overt fragmentation or expansion of endplateareas as observed in other neurological mutants with muscle atrophy(Harris & Ribchester, 1979; Blanco et al., 2001; Lin et al., 2001).Muscle fiber diameters declined uniformly as the animals deteriorated.



By 16 weeks, R6 ⁄ 2 muscle fiber diameters were 20–40% less than WTmuscle diameters. This atrophy was evident in all three muscles butgreatest in EDL (Fig. 2c). Interestingly, endplate areas also shrank inR6 ⁄ 2 muscles as muscle atrophy progressed (Fig. 2d), but the ratio ofendplate area to fiber diameter was unaltered, as evident from plots ofendplate area against muscle fiber diameter (Fig. 2e and f).

Ubiquitinated inclusions in myonuclei

One of the hallmarks of pathology in HD brain and brains of mousemodels of HD is the presence of abnormal aggregates of ubiquitinatedprotein (Davies et al., 1997). Inclusions have been observed inquadriceps muscle (Sathasivam et al., 1999). To check for thepresence of inclusions in the muscles we immunostained for ubiquitinin both whole mounts and transverse muscle sections. WT musclesshowed no evidence of ubiquitin accumulation. Some myonuclei inR6 ⁄ 2 EDL and soleus muscles stained immunopositive for intranu-clear inclusions. The morphological appearance of these inclusionswas similar to those seen in the brains of R6 ⁄ 2 mice (Morton et al.,2000). They were infrequent up to 12 weeks of age but by 16 weeksthey were present in � 20% of EDL muscle fibers (Fig. 1g and h). Asimilar percentage (� 20%) of myonuclei was immunopositive forubiquitin in whole-mount immunofluorescent specimens of thetriangularis sterni muscle. This included nuclei in the motor endplateregion of the muscle fibers (Fig. 3b).

Morphology of neuromuscular junctions

ACh receptors at motor endplates of R6 ⁄ 2 muscles were stained withTRITC-a-BTX. Co-immunostaining for NF ⁄ SV2 showed that, at allages and in all muscles examined, the endplates in R6 ⁄ 2 muscles werepretzel-shaped and mononeuronally innervated by single axon collat-erals in alignment with postsynaptic junctional folds, as in WT muscles(Fig. 3c–h). This appearance suggests that the adult neuromuscularinnervation pattern develops normally, and this pattern is sustainedthroughout the life of R6 ⁄ 2 transgenic mice. Myelinating and perisy-naptic terminal Schwann cells were also of normal appearance and therewas no evidence of any constitutive Schwann cell or nerve terminalsprouting (see section on sprouting in response to nerve injury, below).Several distinctive abnormalities became apparent in R6 ⁄ 2 mice

from the age of � 12 weeks. Most were not quantified because theirincidence was generally very low (estimated at fewer than 5% ofendplates in all muscles). The abnormalities included: longpreterminal axon branches (Fig. 4a); thin, wispy or ‘untwisted’neurofilaments (Fig. 4b); poor penetration of neurofilaments intonerve terminals (Fig. 4c); and accumulations of neurofilaments inpreterminal axonal swellings (Fig. 4d). Similar abnormalities havebeen reported in mouse models of dying-back neuropathy (Miuraet al., 1993) and motor neuron disease (Frey et al., 2000; Fischeret al., 2004). Motor nerve terminal abnormalities included partiallyoccupied motor endplates, with exposed regions of ACh receptorstaining (Fig. 4e). Although this was observed rarely (in three out

of > 600 neuromuscular junctions in three lumbrical musclesexamined; 0.5%), this feature is notable because it has also beenreported in mouse models of motor neuron disease (Cifuentes-Diazet al., 2002; Ferri et al., 2003).A remarkable feature of immunostained triangularis sterni muscles

observed in three out of six severely atrophic R6 ⁄ 2 mice was asignificant reduction in or absence of endplate staining of AChreceptors (Fig. 4f). This abnormal feature was not observed inlumbrical muscles of R6 ⁄ 2 mice at any stage although it is consistentwith physiological failure of synaptic transmission observed inrecordings from FDB muscles (see below). Absence of receptorstaining could not be explained by failure of the TRITC-a-BTX toxinto penetrate the muscle because endplates in the same confocal planewere clearly stained (Fig. 4f).Some lumbrical muscles were vitally stained with FM1-43, a dye

which highlights regions of nerve terminals that contain recyclingsynaptic vesicles (Ribchester et al., 1994; Gillingwater et al., 2002).Almost all neuromuscular junctions in R6 ⁄ 2 mice took up FM1-43 inresponse to depolarizing stimuli, indicating they were able to recyclesynaptic vesicles. As in the immunostained material, we sawoccasional instances of partial occupancy of endplates by FM1-43-loaded terminals (data not shown).

Electron microscopy

We examined endplates in FDB muscles from two 12-week R6 ⁄ 2mice, seeking evidence for early changes in nerve terminal ultrastruc-ture. Of 44 endplates examined in detail, 33 appeared normal and 11showed abnormalities, including membrane inclusions or vacuoles,pre- or postsynaptic detachment, and insinuation of terminal Schwanncell processes into synaptic cleft (arrows, Fig. 5a). Two endplates werefound where junctional folds appeared abnormally wide (Fig. 5a andb). Synaptic vesicles appeared normal but there were vacuolatedinclusions in the presynaptic nerve terminals and the density ofmitochondria appeared abnormally high (Fig. 5a and b).Myelinated axons in intramuscular nerves appeared normal

(Fig. 5b, inset), confirming the normal appearance of axons inwhole-mounts immunostained and studied using confocal micr-oscopy.

Nerve regeneration and sprouting

Motor nerve endings sprout when muscles atrophy as a result ofmuscle paralysis or disuse (Betz et al., 1980; Brown et al., 1981; Punet al., 2002). We were therefore surprised by the normal appearance ofmost R6 ⁄ 2 neuromuscular junctions in spite of the significant, uniformand progressive atrophy of the muscle. We hypothesized that atrophicR6 ⁄ 2 muscles might be incapable of generating nerve regeneration orsprouting signals, or that motoneurons in these muscles wereincapable of responding to such stimuli. To test this, we firstexamined the ability of injured axons in R6 ⁄ 2 mice to regenerate

Fig. 1. R6 ⁄ 2 mutant mice have abnormal skeletal muscles. (a) A WT mouse aged 15 weeks and (b) its R6 ⁄ 2 littermate. Note wasting of hindlimbs andlordokyphosis in the R6 ⁄ 2 mouse. The muscle atrophy of R6 ⁄ 2 mice presents as a pronounced generalized wasting of all muscles, as can be seen in thigh groupsfrom (c) WT and (d) R6 ⁄ 2 15-week-old mice stained with NADH diaphorase. Note that these two photomicrographs were taken through the same level of themuscles and are at the same magnification. (f) The uniform wasting of muscle fibers can be seen clearly in the transverse sections of a 12-week-old symptomaticR6 ⁄ 2 limb muscle. (g) There were no ubiquitinated deposits in the myonuclei of WT muscles. (h) About 20% of R6 ⁄ 2 myonuclei contain ubiquitinatedintranuclear aggregates (arrows) similar to those seen in the brains of R6 ⁄ 2 mice. (i and j) Muscle fiber diameters in the EDL muscle (i) did not change significantlyin WT mice between 9 and 16 weeks, but (j) declined steeply between 9 and 16 weeks in symptomatic R6 ⁄ 2 mice, becoming uniformly� 30% less than in the age-matched WT fibers (P < 0.05, anova). Calibration bar in (h), 3 mm (c and d), 75 lm (e and f), 25 lm (g and h).





c

a WT b R6/2

Wt R6/2 Wt R6/2 Wt R6/2WT R6/2 WT WT R6/2R6/2

FDB EDL SOL

0

10

20

30

40

50

* ****

fiber

diam

eter

(µm

)

d

Wt R6/2 Wt R6/2 Wt R6/20

250

500

750

FDB EDL SOLWTWTWT R6/2 R6/2 R6/2

FDB SOLEDL

*** **

endp

late

area

(µm

2 )

e

endp

late

area

(µm

2 )

0 10 20 30 40 50 600

250

500

750

1000

1250

fiber diameter (µm)

1

1f

endp

late

area

(µm

2 )

0 10 20 30 40 50 600

250

500

750

000

250

fiber diameter (µm)

Fig. 2. Cholinesterase staining in atrophic R6 ⁄ 2 muscle appears normal. Examples of teased EDL muscle preparations from (a) 16-week-old WT and(b) R6 ⁄ 2 littermate stained histochemically for cholinesterase. There were no discernible differences in the staining pattern, although muscle fiber diameter andendplate area were both reduced in the R6 ⁄ 2 muscle. (c) Muscle fiber diameter and (d) end-plate area in FDB, EDL and soleus muscles were all significantlyreduced (P < 0.05; t-tests) comparing R6 ⁄ 2 with WT in each case. The correlation between endplate area and muscle fiber diameter was sustained during progressionof muscle atrophy. (e and f) Examples of WT (s) and R6 ⁄ 2 (d) from littermates aged (e) 13 weeks and (f) 16 weeks. Calibration bar in (b) 30 lm (for a and b).

Fig. 3. Most immunostained neuromuscular junctions appeared normal. Examples of confocal projections from whole-mount preparations of (a,c,e,g) WT and (b,d, f and h) R6 ⁄ 2 mice aged 12–15 weeks. (a–d) Triangularis sterni and (e–h) lumbrical muscles were costained with TRITC-a-BTX to label ACh receptors (red). Inpanels a and b, nuclei were stained using ToPro (blue) and immunostained for ubiquitin (green). (a) Ubiquitin staining was completely absent from WT muscles.(b) About 20% of myonuclei stained positive for ubiquinated inclusions (small arrows), similar to that seen in transverse sections of R6 ⁄ 2 mouse muscle (Fig. 1) orbrain. Inset in (b) is a higher magnification of the nucleus indicated by the arrowhead in the main panel showing an inclusion in the same confocal plane. Axons andsynaptic terminals in R6 ⁄ 2 muscles also showed diffuse immunostaining for ubiquitin. (c and d) Immunostaining for S100 (green) shows no discernible abnormalityin the organization of either myelinating or terminal Schwann cells. (e–h) Neurofilament–SV2 staining (green) shows near-normal appearance of axons, motor nerveterminals and motor endplates in liminally (12 weeks old; e and f) and overtly (16 week; g and h) atrophic muscles of R6 ⁄ 2 mice. Calibration bar in (h), 30 lm (a,b, e, f, g and h), 50 lm (c and d).





by crushing the tibial nerve in 6-week-old animals, and studied theinnervation pattern of muscles physiologically using FM1-43staining and microelectrode recording. In six R6 ⁄ 2 mice studied5–6 weeks after the nerve crush was done, flexion reflexes hadreturned and, in isolated preparations, FM1-43 stained regeneratedmotor nerve terminals (Fig. 5c and d). Intracellular recordings fromtwo of the muscles showed spontaneous miniature endplate

potentials (mEPPs), evoked EPPs and ⁄ or action potentials in15 ⁄ 20 and 19 ⁄ 20 fibers, respectively. Thus, motor neurons inyoung R6 ⁄ 2 mice are capable of axon regeneration and neuromus-cular synapse repair.Reinnervation studies were not possible in the oldest, most severely

affected R6 ⁄ 2 mice because they did not live long enough for axonregeneration to occur after complete nerve section. However, we

Fig. 4. By 16 weeks, some R6 ⁄ 2 motor endplates showed distinctive diverse abnormalities. Endplates were immunostained for (a–c) neurofilaments or (d–g)neurofilaments and SV2. All preparations costained with TRITC-a-BTX. Examples of abnormalities observed include (a) a neuromuscular junction in a triangularissterni muscle whose motor nerve terminal is supplied by a long preterminal axon branch (arrow); (b) neurofilaments that are ‘untwisted’ in this triangularis sterniterminal (arrow), a feature of � 5% of terminals; (c) thinning of neurofilaments with poor penetration into a lumbrical nerve terminal; (d) a lumbrical terminalsupplied by an axon showing swollen accumulations of neurofilaments (arrows), with an ultra-fine preterminal branch supplying the motor endplate; (e) a partiallyoccupied endplate (arrow) (the inset shows a partially occupied lumbrical neuromuscular junction contacted by only two synaptic boutons, leaving the remainder ofthe endplate vacant; arrow); (f) a triangularis sterni neuromuscular junction with intact nerve terminal but complete absence of junctional ACh receptors (inset); (g)a neuromuscular junction with absent ACh receptors in a small fraction of the endplate (inset, arrow). Calibration bar in g, 20 lm (a and c–f), 10 lm (b and g).



examined whether motor axons in R6 ⁄ 2 muscles older than 12 weekswere able to generate compensatory nerve sprouts in response topartial denervation. Sprouting of motor axons is normally preceded bysprouting of terminal Schwann cells (Reynolds & Woolf, 1992; Sonet al., 1996) and also, in mice, by down-regulation of the intensity ofS100 immunostaining (W.J. Thompson, personal communication).The fourth deep lumbrical (4DL) muscles in rodents receive motorinnervation by axons running in both the tibial–lateral plantar andsural nerves (Nakanishi & Norris, 1970; Betz et al., 1979; Taxt, 1983).Section of the tibial nerve in anaesthetized animals thus renders FDBand the medial lumbrical muscles completely denervated but the 4DLmuscle is subject to a controlled partial denervation, making themuscle ideal for studies of motor nerve sprouting of the sural nerve.

Tibial nerves were cut under anaesthesia in nine R6 ⁄ 2 mice (four at9–14 weeks and five at 15–18 weeks) and eight age-matched WTmice. 4DL muscles were immunostained 7 days later and examinedfor sprouting of sural nerve motor axons.The results showed that axons in partially denervated R6 ⁄ 2

muscles from 9–14-week-old mice retained their competence tosprout as their muscle became atrophic. Figure 6a–d shows imagestaken from 4DL muscles 7 days after partially denervation, stainedfor S100 protein. Taking account of the settings of laser power andthe gain and offset on the photomultiplier tubes in the confocalmicroscope, the attenuation of the S100 immunostaining signal wassimilar in WT and R6 ⁄ 2 mice. All muscles showed strongindications of Schwann cell and axonal sprouting (Fig. 6e–h). To

Fig. 5. Ultrustructural abnormalities were evident in some R6 ⁄ 2 presynaptic terminals but synaptic vesicle recycling appeared largely normal. Electronmicrographs in (a) and (b) show motor nerve terminals in a 12-week-old R6 ⁄ 2 FDB muscle. There are expanded postsynaptic junctional folds and pockets andinfiltration of Schwann cell processes (arrows) into the synaptic cleft, and the presynaptic terminals also contain vacuolated structures. The intramuscular nervesappear normal in structure and myelination (inset in b). (c,d) In a reinnervated R6 ⁄ 2 muscle, 6 weeks after sciatic nerve crush, (c) FM1-43 and (d) TRITC-a-BTX staining show that motor axons are competent to regenerate functional synapses. This was confirmed by electrophysiological recording and recycled synapticvesicles (data not shown), as in unoperated R6 ⁄ 2 muscles (see Fig. 4e, inset). Calibration bar in (d), 2 lm (a and b), 5 lm (inset in b), 10 lm (c and d).



quantify the sprouting of axons, we scored the number of sproutsgrowing either into or out of a-BTX-stained ACh receptor pretzels.Collateral sprouting from nodes of Ranvier of intact intramuscularnerve axons was observed in all 9–14 week muscles and in four outof five muscles from severely affected mice older than 15 weeks(Fig. 6e–h). We pseudorandomly selected confocal microscope fieldsstraddled by an intramuscular nerve branch. In the four muscles fromthe 9–14-week-old group of R6 ⁄ 2 mice, 17 endplates out of 71sampled (24%) received collateral axonal sprouts from intact suralnerve motor axons, compared with 12 out of 53 (23%) of endplatesin WT littermates. In the muscles from R6 ⁄ 2 mice older than15 weeks, we observed that 13 endplates out of 50 analysed (26%)received collateral axonal sprouts. However, the sprouts werediscernibly finer than in WT muscles, suggesting a qualitativeimpairment. Interestingly, in the WT muscles we also saw severalinstances of terminal sprouting (outgrowths from nerve terminalsextending to nearby denervated endplates; arrows in Fig. 6f).However, we did not observe any of this form of sprouting in any offour partially denervated muscles from two severely affected R6 ⁄ 2mice older than 16 weeks.Together, the morphological data suggest that the innervation

pattern and morphological appearance of neuromuscular junctions in avariety of R6 ⁄ 2 muscles is quite normal between 8 and 12 weeks ofage, despite the decline in muscle mass during this period. However,significant and increasing instances of abnormal nerve terminals weredetected in coincidence with increasing severity of the diseaseand there was a weakened sprouting response following partialdenervation in muscles from mice older than 12 weeks. Most notably,whereas a compensatory sprouting reaction in motor nerve terminalsmight have been expected in response to the pronounced atrophy seenin mice older than 12 weeks, none was observed.

Synaptic transmission at R6 ⁄ 2 mouse neuromuscular junctions

Previous physiological studies of motor function in R6 ⁄ 2 mice havefocused on central nervous system defects in synaptic transmission, asmeasured behaviourally and electrophysiologically. Because R6 ⁄ 2mice show deteriorating cognitive and behavioural function yet remainmobile despite their severe muscle wasting and weight loss, it isunclear whether muscle atrophy is a consequence of central orperipheral neural defects or whether it is caused by intrinsic changes inskeletal muscle properties. In preliminary tests we measured forelimbgrip strength. These showed that both WT and severely affected(15–17-week-old) R6 ⁄ 2 mice were motivated to exert consistentvoluntary force, but R6 ⁄ 2 mice showed some fatigue on repeatedtesting (data not shown). However, such tests do not distinguishneuropathic changes from myopathic ones. We therefore studiedpassive membrane properties of R6 ⁄ 2 muscle fibers (resting mem-brane potential, input resistance and time constant) as well as synapticproperties (mEPPs and EPPs). We also examined the sensitivity ofmuscle fibers to bath-applied and iontophoretically applied ACh, andthe sensitivity of action potentials to the selective muscle NaV1.4sodium channel blocker l-CTX.

Resting membrane potentials

The mean resting membrane potential of FDB muscle fibers fromWT mice was )72.2 ± 3.2 mV (SEM, N ¼ 6 muscles). This wassignificantly different from muscle fibers of presymptomatic R6 ⁄ 2mice aged < 14 weeks ()62.8 ± 1.52; P < 0.001; N ¼ 10 muscles).There was a further deterioration of resting membrane potential inR6 ⁄ 2 muscle fibers as the animals became overtly symptomatic andthe muscle fibers more atrophic (Fig. 7a). By 15–18 weeks, musclefiber resting potentials were )55.2 ± 4.3 mV (N ¼ 10 muscles).This decline was not an artefact of impalement of the atrophicmuscle fibers because we made the recordings using high imped-ance microelectrodes filled with potassium acetate (> 40 MW). Lowresting membrane potentials were also evident in R6 ⁄ 2 muscleswhen using ultra-sharp, very high impedance (� 100 MW) micro-electrodes, which also registered normal resting potentials in WTmuscles.

Input resistance and time constant

The small diameter of atrophic muscle fibers would be expected toconfer a high input resistance and prolonged membrane time constant,as measured by the ratio and time course of the steady-state change inmembrane potential in response to injection of rectangular constant-current pulses. Thus, input resistance is proportional to d)3 ⁄ 2, where dis the muscle fiber diameter (Katz & Thesleff, 1957). FDB musclefibers in 15–18-week R6 ⁄ 2 mice are � 40% narrower than in WTmice (see above). As a consequence, input resistance should be abouttwice as great. Input resistance and membrane time constant wereestimated using a single microelectrode and Wheatsone bridge method(Fig. 7b–e). Though imperfect, the single-electrode and bridge methodminimizes damage to muscle fibers and allows effective comparisonsbetween experimental and control groups of muscles (Barry &Ribchester, 1995). There was no signficant difference between inputresistances of presymptomatic, nonatrophic R6 ⁄ 2 muscles and thoseof WT (data not shown). However, by 15–18 weeks, as expected fromthe degree of muscle atrophy, the input resistance of R6 ⁄ 2 fibers was� 4.38 ± 0.36 MW (SEM; n ¼ 21 fibers, N ¼ 3 muscles and threemice) compared with WT (2.34 ± 0.27 MW; n ¼ 27 fibers, N ¼ 3muscles and three mice; P < 0.001, t-test).The membrane time constant in atrophic R6 ⁄ 2 muscle fibers was

extended to more than twice that of WT fibers (Fig. 7b–e). FDBmuscle fibers are equivalent to short cables with open-circuited ends,and are isopotential along their length in response to focal constantcurrent injection (e.g. by synaptic currents, or via intracellularmicropipettes; Bekoff & Betz, 1977). Biophysically, they are similarin their overall passive membrane behaviour to spherical cells. Themembrane time constant was therefore estimated from the time takento reach 63% of the steady-state membrane potential displacementduring the same constant current injections (Jack et al. 1974). Inpresymptomatic R6 ⁄ 2 muscles the membrane time constant was notsignificantly different from WT (data not shown). However, inatrophic muscle fibers from 15–18-week R6 ⁄ 2 mice, with higher inputresistances than normal, the membrane time constant was about

Fig. 6. Motor axons in R6 ⁄ 2 mice are competent to sprout. Immunostaining for (a–d) S100 protein in Schwann cells (green) or (e–h) neurofilaments and SV2(green), in partially denervated 4th deep lumbrical muscles, suggest that sprouting capacity is retained following partial denervation even as the disease becomesadvanced. Defined settings of the confocal laser power, gain and offset demonstrated that Schwann cells down-regulate S100 protein in response to section of thetibial nerve 7 days earlier in both WT (a, unoperated; c, 7-day partially denervated) and R6 ⁄ 2 mice (b, unoperated; d, 7-day partially denervated). Neurofilamentstaining (green, e–h) shows that axons sprout (arrows), even in liminally affected (12 weeks old; f) and severely affected (17 weeks; h) R6 ⁄ 2 mice, compared withage-matched WT mice (12 weeks old, e; 17 weeks old, g). Calibration bar in h, 60 lm (a–d), 50 lm (e and f), 40 lm (g and h).



WT R6/2

S10

0N

F/S

V2

dc

ba

hg

fe

Power = 4.7%Gain = 4.9

Offset = -3.9


Offset = -3.9


Offset = 1.9


Offset = 1.9



threefold greater: 6.53 ± 0.72 ms (SEM, n ¼ 10 fibers) comparedwith 2.22 ± 0.19 ms (n ¼ 17) in WT muscles.

Spontaneous transmitter release

Measurements of spontaneous mEPPs were made in 137 muscle fibersfrom 12 muscles of R6 ⁄ 2 mice aged 8–18 weeks. In the youngermice, mEPP frequencies were in the normal range (mean ± SEM

0.23 ± 0.2 ⁄ s at 18 �C, n ¼ 40 fibers from two muscles; comparedwith 0.16 ± 0.04 ⁄ s, 25 fibers from two age-matched WT muscles).mEPP amplitude, rise time and decay time constants were also withinthe normal range.Analysis of mEPP frequency, amplitude and time course was

performed on a subset of fibers in older, atrophic R6 ⁄ 2 muscles.Examples of averaged mEPPs are shown in Fig. 8a and b. Between-event interval histograms of mEPPs in 18-week-old mice showedan exponential time course, indistinguishable from the Poisson

Fig. 7. Passive membrane properties of R6 ⁄ 2 muscle fibers change as they atrophy. (a) There was a decline in resting potential as the mice became more severelyaffected. Each point is the mean of 7–30 muscle fibers recorded from one muscle. The box-whisker plot shows the median and range of values from WT muscles,which did not change with age. (b and c) Typical records showing responses to constant-current injections via the recording microelectrode used to measuremembrane time constant and input resistance in (b) R6 ⁄ 2 and (c) WT muscles. (d and e) Both properties were two to three times greater in R6 ⁄ 2 mice (P < 0.01;Mann–Whitney test in each case).



distribution that characterizes normal mEPP interval histograms(Fig. 8c and d). However, though highly variable between fibers,mEPP frequencies were reduced overall by about half: medianfrequency was 0.27 ⁄ s (interquartile range 0.17–0.5 ⁄ s) compared with0.60 ⁄ s overall in WT (interquartile range 0.47–0.72 ⁄ s; P < 0.001,Mann–Whitney). Overall, these data suggest that the stochastic

determinants of synaptic vesicle exocytosis are not altered in R6 ⁄ 2mouse neuromuscular junctions. However, further analysis of thecharacteristics of spontaneous transmitter release frequency may bewarranted, especially in the light of the finding that at least one synapse-specific protein thought to modulate exocytosis (complexin II) is down-regulated in the R6 ⁄ 2 mouse brain (Morton & Edwardson, 2001).

Fig. 8. Miniature EPP amplitudes and time courses increase as R6 ⁄ 2 muscle fibers atrophy. (a and b) Examples of intracellular recording of mEPPs from (left) WTand (right) 18-week R6 ⁄ 2 muscle fibers are shown. mEPP amplitudes were more variable in R6 ⁄ 2 muscle fibers. Normalized average mEPPs (20–60 events)showing prolonged decay time constant (c, 5.84 ms; d, 16.08 ms). (e and f) Histograms of mEPP intervals in (e) WT and (f) R6 ⁄ 2 muscles (129 and 211 events,respectively) also showed variability in mean frequency but approximately exponential distribution of inter-mEPP intervals in both groups. (g and h) Box-whiskerplots summarizing overall median, interquartile range and 95% outliers of mEPP amplitude and frequency in (h) severely affected R6 ⁄ 2 mice and (g) their WTlittermates. Overall, mEPP amplitude was significantly increased about two-fold, as expected from the difference in muscle fiber input resistance in atrophic FDBmuscle fibers in these mice (see Fig. 7), and mean mEPP frequency was significantly reduced in R6 ⁄ 2 mice (P < 0.001; Mann–Whitney in both cases).



As expected from the measurements of input resistance, mEPPamplitudes were markedly increased as the phenotype of the R6 ⁄ 2mice deteriorated. In R6 ⁄ 2 mice older than 15 weeks, mEPPs weretypically 2–4 times greater in amplitude than in WT mice (Fig. 8e).Overall, the median mEPP amplitude in R6 ⁄ 2 fibers was 1.96 mV(interquartile range 1.58–2.16 mV, n ¼ 21 fibers from six muscles)compared with 0.54 mV (interquartile range 0.49.0.74 mVn ¼ 11 fibers) from two age-matched WT muscles (P < 0.001,Mann–Whitney). The coefficient of variation of mEPP amplitude insome R6 ⁄ 2 muscle fibers was numerically higher (mean ± SEM0.47 ± 0.27) but not statistically significantly different from WT(0.34 ± 0.06; P > 0.1, t-test). The instances where the variabilityseemed higher may thus have been merely a consequence of the bettersignal-to-noise ratio for detection of mEPPs in R6 ⁄ 2 muscle fibers onaccount of their high input resistance. However, we cannot rule out atthis stage a real difference in the distribution of quantal sizes.Also as expected from the increase in membrane time constant,

time-to-half-decay of mEPPs were longer in R6 ⁄ 2 than in WT fibers(Fig. 8f). The decay time constants of exponential curve fits to therepolarization phase of the normalized average mEPP were consis-tently 2–3 times longer in R6 ⁄ 2 than WT.

Evoked EPP characteristics

Muscles were pretreated with l-CTX GIIIB (2.5 lm), with theintention of blocking muscle action potentials and thereby exposingthe full amplitude and time course of nerve-evoked EPPs. The l-CTXwas very effective in WT muscles, but we experienced considerabledifficulties with older R6 ⁄ 2 muscles because they were resistant to theblocking effects of the toxin (see below). Movement artefacts wereevident in many of the recordings, and action potentials rather thanEPPs were frequently registered. However, continuous incubation inl-CTX eventually produced sufficient block to enable recording ofEPPs (Fig. 9). In most fibers, robust EPPs with a low coefficient ofvariation were recorded, indicating a high quantal content (Fig. 9b).We found no instances of random ‘failures’ of synaptic transmission inany of the trains of EPPs recorded from single muscle fibers, at anystage in the deterioration of R6 ⁄ 2 mice. For instance, EPP trains weresustained during repetitive high-frequency (> 30Hz) stimulation(Fig. 9c) as in WT muscles and there was no evidence for anysystematic change in mean quantal content. Highly variable responses(including random occurrences of ‘failures’ in response to nervestimulation) would have indicated a low quantal content and weaksynaptic transmission. This was never observed in any of therecordings from > 500 muscle fibers recorded during the course ofthis study. Analysis of EPP amplitudes using the coefficient ofvariation method indicated that the overall mean quantal contentof EPPs in atrophic muscle fibers was not statistically significantlydifferent from WT (data not shown). Voltage-clamp analysis ofsynaptic currents would perhaps clarify whether there are any subtledifferences in quantal content of EPPs in R6 ⁄ 2 muscle fibers but,based on the voltage recordings, any such differences are unlikely tobe physiologically significant.Despite the absence of significant progressive deterioration in EPP

quantal content, there was an increase in the number of muscle fibersthat did not respond at all to nerve stimulation in severely affectedR6 ⁄ 2 mice (older than 14 weeks). Only two out of 206 fibers in 16WT muscles did not show evoked EPPs or action potentials when themuscle nerve was stimulated supramaximally. However, in 269recordings from 18 muscles in R6 ⁄ 2 mice aged 15–18 weeks, 52 ofthe fibers (i.e. � 20% overall) were consistently unresponsive to

repeated stimulation and failed to show evoked EPP responses. Thiswas not a consistent feature between mice or muscles because in eightout of 18 of the muscles all fibers impaled gave evoked responses.Nevertheless, complete failure of evoked synaptic transmission wasevident in muscles from half of the severely affected R6 ⁄ 2 mice. Itremains to be established whether this was due to abnormal nerveconduction or complete pre- or postsynaptic failure of neurotransmitterrelease or action.In spite of the high input resistance of atrophic R6 ⁄ 2 muscle

fibers and the concomitant increase in mEPP amplitudes, theamplitudes of evoked EPPs were not significantly different fromthose in WT muscles (Fig. 10a). However, resting potentials of theR6 ⁄ 2 muscles were reduced (see above, Fig. 8), and differences inEPP amplitude could have been obscured by the nonlinearsummation of synaptic potentials as the membrane potentialapproached the transmitter null (i.e. reversal) potential (McLachlan& Martin, 1981). We therefore measured the membrane potential atthe peak of the EPP response. A nonlinear polynomial curve fitted tothe data showed a trend towards increased synaptic efficacy in theageing R6 ⁄ 2 mice that did not reach statistical significance (EPPsdepolarized muscle fibers to )40 ± 5 mV (N ¼ 10 muscles) in olderR6 ⁄ 2 mice compared with )48 ± 4 mV in WT mice (N ¼ 4muscles; P > 0.05; t-test). There were similar trends in EPP latencyand time-to-peak (Fig. 10c and d), although only the time-to-half-decay of EPPs in atrophic R6 ⁄ 2 muscles was statistically signifi-cantly different: about three times longer in R6 ⁄ 2 mice older than15 weeks compared with WT.

Effect of anticholinesterase

The prolongation of EPPs as R6 ⁄ 2 mice age is not a consequence ofany functional reduction in cholinesterase activity. Histochemically,cholinesterase distribution at endplates was unaffected (see Fig. 2).Further, the addition of the cholinesterase inhibitor neostigmine

200 ms

10 mV

dc

ba10 mV

10 ms

10 mV

10 ms

10 mV

10 ms

Fig. 9. Evoked EPPs in R6 ⁄ 2 muscles are robust. Typical superimposed serialtraces of EPPs from different FDB fibers are shown from (a) WT and(b) severely atrophic 16-week-old R6 ⁄ 2 muscle. Both records show a lowcoefficient of variation in amplitude (high quantal content), but note the longerdecay time course in the R6 ⁄ 2 muscle fiber. (c) R6 ⁄ 2 fibers showed robustresponses to high frequency (50 Hz) repetitive stimulation. (d) The prolongeddecay time constant of R6 ⁄ 2 EPPs could not be explained by inactivecholinesterase because, after a 10-min incubation in the anticholinesteraseneostigmine, all EPPs in R6 ⁄ 2 muscles were significantly prolonged. Thisrecording was made from a different fiber but the same muscle as in (b).



(10 lm) to the bathing medium prolonged the decay time of EPPs inR6 ⁄ 2 mice about seven-fold, showing that the cholinesterase inunblocked muscles was functional. The mean ± SD half-decay timewas 22.95 ± 7.06 ms (n ¼ 4 fibers) in one 16-week R6 ⁄ 2 muscleafter neostigmine compared with 3.34 ± 1.54 ms (n ¼ 7) in the samemuscle before neostigmine (see Fig. 8d).

Resistance of muscle action potentials to l-CTX

Voltage-gated sodium channels in mammalian skeletal muscle(NaV1.4 channels) are normally blocked by l-CTX GIIIB atconcentrations of 1–3 lm (Cummins et al., 2002; Li et al., 2003).At this concentration, the toxin is ineffective on axons and motornerve terminals, which therefore release physiologically normalamounts of neurotransmitter in response to nerve stimulation(Wood & Slater, 1997; Costanzo et al., 1999; but see Bragaet al., 1992).

Incubation of WT muscles in l-CTX routinely and visibly blockednerve-evoked muscle contractions within 5–30 min and this blocknormally persisted for 1–4 h after returning muscles to normalphysiological saline. As described above, robust endplate potentialswere recorded from virtually all WT muscle fibers during the period ofmuscle paralysis induced by the toxin. Muscles from R6 ⁄ 2 mice aged< 11 weeks were also sensitive to l-CTX; muscle fiber actionpotentials were blocked, and robust nerve-evoked EPPs remained.However, isolated FDB muscles from R6 ⁄ 2 mice older than 12 weekswere consistently resistant to the blocking effect of l-CTX, showingvisible, moderate-to-strong contractions in response to nerve stimula-tion, even 1–2 h after continuous incubation in 2.5 lm l-CTX.To quantify the sensitivity of WT muscles and the resistance of

R6 ⁄ 2 muscles to bath application of l-CTX, we measured muscletension and EMG currents from FDB muscles. Twitches and tetanideclined following l-CTX administration. In two WT muscles,tension declined to � 10% of the initial response (Fig. 11a). In one16-week R6 ⁄ 2 muscle, tension declined to only 35% of the initial

6 9 12 15 18

0.0

0.5

1.0

1.5

2.0

WT

0.0 2.5 5.0 7.5 10.00.0

0.5

1.0

Half-Decay Time (ms)6 9 12 15 18

0.0

2.5

5.0

7.5

W T

WT6 9 12 15 18

0

1

2

3

4

Age (weeks) WT

6 9 12 15 18

-60

-50

-40

-30

-20

-10

0

Age (weeks)

6 9 12 15 18

0

10

20

30

40

Age (weeks) WT

WTR6/2

c d

a b

e f

R6/2 age (weeks)

R6/2 age (weeks)

R6/2 age (weeks)

R6/2 age (weeks)

R6/2 age (weeks)

EP

P a

mpl

itude

(m

V)

EP

P h

alf d

ecay

(m

s)la

tenc

y (m

s)

time

to p

eatk

(m

s)E

m a

t pe

atk

(m

s)

WT

WT

WT

WThalf-decay time (ms)

freq

uenc

y

Fig. 10. Synaptic strength is not impaired in R6 ⁄ 2 muscles. A summary of raw data on (a) peak EPP amplitudes and (b) membrane potential at the peak of theEPP after allowing for the depolarized resting membrane potential of R6 ⁄ 2 muscle fibers (see Fig. 7), showing that evoked synaptic strength did not changesubstantively in most muscle fibers as R6 ⁄ 2 mice deteriorated, compared with WT (data are mean ± SEM, N ¼ 7 muscles). There was a no statistically significantchange in (c) EPP latency and (d) time-to-peak, but (e) time-to-half-decay increased significantly from � 13 weeks of age (P < 0.01, Mann–Whitney). Lines ina–e are best-fit second-order polynomials. (f) Cumulative distributions of relative frequencies of EPP half-decay times from all muscle fibers recorded in musclesfrom R6 ⁄ 2 mice (solid line) aged 15–18 weeks were significantly different from WT (dotted line; P < 0.01, Kolmagorov–Smirnov test).



response (Fig. 11b), still producing visible contractions that appearedquite strong under the dissecting microscope.Next, we recorded extracellular action potentials using 1 m NaCl-

filled blunt-tipped (� 2–20 lm) glass micropipettes inserted into thebelly of the FDB muscles. In three R6 ⁄ 2 preparations recorded in thisfashion, l-CTX depressed nerve-evoked muscle fiber inward currentsby < 5%, 28% (Fig. 12a) and 48%, whereas extracellular responseswere reduced by 54% and 76% of initial current amplitude in two WT

muscles (Fig. 11c and d). No contractions were visible in the case ofthese two WT muscles. The residual responses were therefore causedby endplate currents. As expected, adding 10 lm a-BTX to thebathing medium completely blocked junctional ACh receptors andabolished all evoked responses and contractions in both R6 ⁄ 2 and WTmuscles (data not shown).Lastly, we measured the amount of l-CTX resistance in intracellular

recordings. In 15–18-week-old R6 ⁄ 2 muscles, some fibers showed

Fig. 11. R6 ⁄ 2 muscles were relatively resistant to l-CTX. l-CTX almost completely blocked FDB muscle contractions and action potentials in WT muscle fibersleaving robust EPPs (see Fig. 9a). (a and b) A significant fraction of nerve-evoked muscle tension and action potentials remained following addition of 2 lm l-CTXto the medium bathing (b) isolated R6 ⁄ 2 FDB muscles compared to (a) that in the WT muscle. In both a and b, the upper traces show recordings made beforeadding the toxin while the lower traces show recordings made � 15 min after toxin administration. (c and d) EMG recordings using a blunt micropipette filled with1 m NaCl are shown during administration of l-CTX to the medium bathing (c) a WT and (d) a 15-week-old R6 ⁄ 2 FDB muscle, showing only a small reductionin amplitude of extracellularly recorded action potentials. (The calibration for d is the same as that shown for c.) Insets directly below the EMG in c and d showexpanded views of extracellular action potentials recorded before and after toxin administration. (e) An intracellularly recorded muscle fiber action potential from anR6 ⁄ 2 muscle fiber is shown after 2 h incubation in l-CTX. The trace in (f) shows an example of a spontaneous mEPP recorded from a 16-week R6 ⁄ 2 FDB musclefiber that was so large it triggered an action potential, even though this recording was made in the presence of l-CTX.



full-blown action potential responses, apparently unaffected by l-CTXtreatment. However, others showed nonovershooting action potentials,with positive-going inflexions on the upstroke of the EPP (Fig. 11e).In some fibers, large mEPPs triggered action potentials that were alsoresistant to l-CTX (Fig. 11f).

To quantify the extent of l-CTX resistance, we measured themaximum rate of rise of the first intracellularly evoked responserecorded in each muscle fiber. FDB muscle fiber action potentialstypically have rates of rise > 60 mV ⁄ms at room temperature, beforetreatment with l-CTX. In WT muscles, 90% of EPPs evoked by nervestimulation after treatment with conotoxin had maximum rates of rise(dV ⁄ dt) < 30 mV ⁄ms (53 ⁄ 58 muscle fibers from eight muscles).

Overall, lCTX reduced the rate of rise of intracellularly recordedmuscle fiber responses in older R6 ⁄ 2 mice: � 42% of muscle fibers in15–18-week-old R6 ⁄ 2 mice (38 ⁄ 90 fibers from eight muscles)showed rates of rise in their nerve-evoked responses that remained> 30 mV ⁄ms; that is, below the lower bound for most EPPs inunblocked WT muscles. In the remainder dV ⁄ dt was < 30 mV ⁄m.Thus, almost half of the muscle fibers in severely affected R6 ⁄ 2mouse FDB muscles showed evidence of significant l-CTXresistance.

Anode-break action potentials

Resistance to tetrodotoxin and l-CTX are properties of rSkM2(NaV1.5) sodium channels that appear in WT muscle fiber membranesafter denervation (White et al., 1991; Chen et al., 1992). Becauseatrophic muscle fibers from R6 ⁄ 2 mouse FDB muscles resembled

denervated or paralysed muscles with respect to the l-CTX resistance,we tested them for anode-break excitation (Marshall & Ward, 1974). Ina sample of 10 fibers recorded from one 16-week R6 ⁄ 2 FDBmuscle, all10 fibers showed clear anode-break excitation (Fig. 12). By compar-ison, > 90% of WT muscle fibers showed no anode-break excitation.

ACh responses

In the light of the other denervation-like physiological and pharma-cological characteristics of R6 ⁄ 2 muscles, we examined them for‘supersensitivity’ to ACh. Figure 12c shows alternating cycles ofresponses to single twitch and 30 Hz tetanic stimulation in an R6 ⁄ 2muscle before, during and after adding ACh (10 lm) to the bathingmedium. This caused a muscle contracture equivalent in magnitude tothe resting twitch response, that is, about one-third of the tetanictension response. Similar contractures were evoked in two other 18-week R6 ⁄ 2 muscles. In two WT muscles, as expected, the AChcontracture was < 10% of the nerve-evoked tetanic tension. Thus,R6 ⁄ 2 muscles were more than three times as sensitive as WT musclesto bath-applied ACh.Finally we tested the response of individual fibers to iontophoretic

application of ACh to extrajunctional muscle fiber membranes(Fig. 12d). Extrajunctional sensitivity to ACh was not seen in WTmuscle fibers (data not show). However, graded depolarizations wereproduced in three out of 10 fibers in one 18-week-old R6 ⁄ 2 muscle.These fibers were innervated, because we also observed spontaneousmEPPs in the recordings. This incidence of supersensitive fibers isconsistent with the sensitivity to bath-applied ACh and the

15 ms

10 µM ACh

dc

ba

2 mV

60 s

4 mV

60 ms

50 mV

5 nA

Fig. 12. Atrophic R6 ⁄ 2 muscles showed anode-break excitation and ACh supersensitivity. (a and b) These characteristics were absent from WT muscles(a) and presymptomatic R6 ⁄ 2 mice (not shown), but strong hyperpolarizing current injections induced action potentials on break of the anodal current in virtually allseverely affected R6 ⁄ 2 mouse FDB muscle fibers (b). (c and d) Muscles from mice at late stages of the disease (15-week-old) were supersensitive to ACh, as shown by(c) contractures in response to bath-applied ACh or (d) graded, iontophoretic application of ACh to muscle during intracellular recording. In (c), FDB muscletwitches and tetani were induced by nerve stimulation, then the bathing medium was substituted with medium containing 10 lm ACh. This produced a musclecontracture that reached � 80% of the twitch contraction response (� 30% of the tetanus evoked by 50 Hz stimulation). Traces in (d) show membrane depolarizationin response to graded 2-ms iontophoretic pulses of ACh applied from a micropipette manipulated over the extrajunctional membrane of the impaled fiber.



other denervation-like properties measured above, such as l-CTXresistance.

Discussion

The R6 ⁄ 2 transgenic mouse model of HD shows motor abnormalitiesthat appear at � 8 weeks of age and progress insidiously until they aremanifest overtly by � 12 weeks of age (for summary, see Fig. 13).From this time onwards, the mice deteriorate steadily until theiruntimely death at � 16–20 weeks of age. The present study shows thatthe R6 ⁄ 2 mouse has a complex phenotype with respect to neuromus-cular structure and function. Both neural and muscular abnormalitieswere detected. These changes were paradoxical. For example, there isno evidence for paralysis or functional denervation until the lateststages of the disease, yet R6 ⁄ 2 muscle fibers undergo profoundatrophy and many show membrane characteristics of denervated orparalysed muscle fibers. Conversely, intact nerve terminals andterminal Schwann cells show no evidence of constitutive sprouting,in spite of the atrophy and other denervation-like state of R6 ⁄ 2 musclefibers. In addition, synaptic transmission fails completely at increasingnumbers of neuromuscular junctions as the disease progresses.Together, these data suggest that neuromuscular abnormalities couldcontribute deleteriously to motor function in R6 ⁄ 2 mice and may alsohelp explain their sudden deaths.

Denervation-like characteristics of R6 ⁄ 2 muscles and their rolein muscle atrophy

Remarkably, in spite of their strong, functional motor innervation,R6 ⁄ 2 muscle fibers resemble denervated muscles in several importantrespects (atrophy, reduced resting membrane potential, resistance toan NaV1.4 sodium-channel blocker, anode-break action potentials).However, other features of R6 ⁄ 2 muscle (most motor nerve terminalsfully occupying ACh receptor pretzels at motor endplates, robust EPPresponses to nerve stimulation, absence of spontaneous fibrillation)were at odds with a denervated state. Thus, while the atrophy waspronounced, the absence of characteristic muscle pathology wouldappear to rule out the possibility that the atrophy was due to adenervation-like pathology. For example, there was no fiber-typegrouping or compensatory hypertrophy of innervated fibers seen(Pachter & Eberstein, 1992). We also ruled out other neuropatholo-gical causes of the atrophy. Neurogenic disorders with skeletal muscleinvolvement show characteristic abnormalities such as angulatedatrophic fibers, fiber-type grouping, group atrophy or target fibers.However, none of these abnormalities were found. There was also nopathology consistent with a myopathic origin of the atrophy becausethere was no myofiber necrosis, myophagocytosis, regeneration,rounded atrophic fibers, fiber hypertrophy and splitting, centralizednuclei, fibrosis, ring fibers, vacuoles or any signs of inflammation.The muscle atrophy is unlikely to be a consequence of malnutritionbecause it is evident weeks before the mice show disrupted feedingpatterns. Further, in malnutrition there is typically a relativelyselective atrophy of type II fibers (Anderson, 1985) that was notseen in the R6 ⁄ 2 muscles. It also seems unlikely that the atrophy isrelated to the late-onset diabetes of the R6 ⁄ 2 mice (Hurlbert et al.,1999). Although electrophysiological changes have been reported indiabetes, they are not the same as those we see here. For example,neuromuscular junctions in diabetic rats show greater resistance totetanic stimulation with fewer failures of action potential propagationthan in healthy rats (Schiller & Rahamimoff, 1989). Moreover, AChrelease is depressed by up to 45% in chronically diabetic rats

(Constantini et al., 1987). In the R6 ⁄ 2 mouse there was no evidenceof reduction in quantal content at functional junctions, even inseverely atrophic muscle. Thus, there is no compelling evidence thateither structural or functional denervation (paralysis) plays anysignificant role in the progression of atrophy. This is in fact veryintriguing given that some of the muscle fiber properties resembledenervation (e.g. l-CTX resistance).The cause of the decline in resting membrane potential is unclear.

One prediction is that intracellular K+ ion concentrations of R6 ⁄ 2muscles should be reduced. Such measurements were not performed inthe present study but would be interesting to examine in future studies,particularly if combined with experiments on sensitivity to changes inextracellular ion concentrations. Although changes in gene expressionin skeletal muscle have been reported (Luthi-Carter et al., 2002), it isnot yet know whether NaV1.5 sodium channels are up-regulated inatrophic R6 ⁄ 2 muscles, nor whether any of the many other proteinsassociated with the orchestrated response of muscles to denervation,such as expression of ACh receptor gamma subunits, are also alteredin these muscles. It would be interesting to apply genetic microarrayanalysis of muscle mRNA expression patterns in R6 ⁄ 2 musclesbecause this would establish the extent of differences in pattern ofgene expression comparing WT innervated or denervated muscle.

Morphological abnormalities in neuromuscular junctions

Abnormalities in nerve terminal morphology and physiology, thoughpresent, were surprisingly infrequent given the substantial andprogressive muscle atrophy. This is in marked contrast to, forexample, the morphological and physiological impairment in nerveterminal structure and function in other neurological mutants whichundergo muscle atrophy, such as wobbler mice (Harris & Ward, 1974)smn mice (Cifuentes-Diaz et al., 2002) pmn mice (Ferri et al., 2003)and wasted mice (R.R. Ribchester, T.H. Gillingwater and C.M.Abbott, unpublished observations). There was also little resemblanceto the widespread retraction of motor nerve terminals that occurs, withaccompanying progressive weakening of synaptic transmission, inmutant or transgenic mice with slow Wallerian nerve degeneration(Ribchester et al., 1995; Gillingwater et al., 2002; Gillingwater et al.,2003).There were a number of morphological abnormalities in R6 ⁄ 2

junctions that might have contributed to abnormal physiology. Forexample, ultrastructural and fluorescence microscopy of motorendplates in mice older than 12 weeks showed several unusualfeatures, including invagination of Schwann cell processes into theterminal, and abnormal neurofilament organization. Further, in somemuscles, only slender neurofilaments penetrated the motor nerveterminals; in others, we saw wispy assemblages of untangledneurofilaments in preterminal axons. There was also evidence atsome junctions for abnormal preterminal axon branching and thesurprising finding of some endplates in older muscles that appeareddevoid of ACh receptors. These changes would be consistent withaberrant function of the endplates. However, none of these adequatelyexplain the electrophysiological deficits.Although nerve function is abnormal, the electrophysiological data

suggest that most of the muscle atrophy in R6 ⁄ 2 muscle up to 12–14 weeks of age is caused by an intrinsic deterioration in muscleproperties, rather than by progressive structural or functional dener-vation. At late stages there was concurrent physiological evidence ofincreasing numbers of junctions showing progressive failure ofneuromuscular transmission, which could reflect independent neuralabnormalities. Thus, one possible explanation for the morbidity and



Fig. 13. Summary diagram showing the relationship between timing of appearance of pathological features in the brains and skeletal muscle in R6 ⁄ 2 mice. Dataused in compiling this figure come from (a) this study and from the studies of (b) Mangiarini et al. (1996), (c) Carter et al. (1999), (d) Lione et al. (1999), (e)Sathasivam et al. (1999), (f) Murphy et al. (2000), (g) Morton et al. (2000), (h) Meade et al., 2002) and (i) Klapstein et al., 2001). Abbreviations used are MWM,Morris water maze; NII, neuronal intranuclear inclusion; ENNI, extranuclear neuronal inclusion.



mortality of R6 ⁄ 2 mice over the age of 15 weeks is a collapse ofskeletal muscle function, caused by both myogenic atrophy andneurogenic failure of neuromuscular transmission.HD is considered to be a disease solely of the brain, despite the

peripheral expression of huntingtin. However, our findings heresuggest that abnormalities of either motor neuron or skeletal musclefunction could contribute directly to motor symptoms in this disease.In addition to the uniform atrophy observed in R6 ⁄ 2 mouse muscles,we found some significant abnormalities in sprouting responses. Ingrade 2–4 HD brain, dendritic changes consistent with both regen-eration and degeneration (Graveland et al., 1985; Ferrante et al., 1991;Sotrel et al., 1993) have been reported. However, in two transgenicmodels where dendritic changes have been studied (R6 ⁄ 2 and HDfull-length cDNA mice), degenerative changes but not sproutingresponses have been reported (Guidetti et al., 2001; Klapstein et al.,2001). It would be interesting to see whether the neurons in R6 ⁄ 2brains show a typical sprouting response to direct injury or whetherthey, like the peripheral neurons, show abnormal sprouting responses.The most parsimonious explanation for our findings is that the

polyQ-huntingtin transgene affects motoneurons, their presynapticmotor nerve terminals and skeletal muscle fibers independently. Infact, the peculiar combination of pathology we see in the neuromus-cular junction suggests that there is a functional disconnection of thehomeostatic feedback mechanisms that normally regulate neuromus-cular function. Thus, skeletal muscle fibers do not appear to respondnormally to the trophic input from their motor nerve terminals, and ⁄ orthe motor neurons do not respond to neurotrophic feedback from themuscle fibers. It is tempting to speculate that a similar dysregulation offeedback mechanisms may also be present in the CNS. This mayconstitute (or produce) breakdown in the mutual trophic interactionsthat normally maintain nerve and muscle. It has been shownpreviously that skeletal muscle is a target of polyglutamine-relatedperturbations in gene expression (Luthi-Carter et al., 2002). Furtheranalysis of patterns of gene expression in R6 ⁄ 2 muscle could helpresolve this issue, or at least show to what extent the Huntington’smutation causes dysregulation of muscle gene expression. Forexample, it would be interesting to determine whether the changesin l-CTX sensitivity were due to changes of sodium channel subunitexpression or subunit composition. Targeted expression of the R6 ⁄ 2transgene selectively in muscle or motoneurons, or utilization ofchimeric mice or mice with xenografts of muscle, might also help toestablish whether muscle atrophy is a direct consequence of themutation. In the light of published abnormalities in synaptic vesicleproteins in the brains of R6 ⁄ 2 mice, notably complexin II (Morton &Edwardson, 2001), it is interesting to speculate that the causes of thediverse array of neuromuscular abnormalities may be a consequenceof abnormal trafficking, integration or function of other synapse-specific proteins. Such abnormalities might, for example, underpin thereduced mEPP frequency and abnormally high coefficient of variationin the amplitude distribution of mEPPs we observed in some R6 ⁄ 2muscle fibers.For the time being, the clinical relevance of our studies remains

speculative. While some studies on human skeletal muscle functionhave been done, most of those have focused on descending control ofmotor function. Interestingly, EMG studies in HD patients havereported abnormalities in motor unit activity (Harper et al., 1991) andlong latency responses (Noth et al., 1985; Leblhuber et al., 1991;Berardelli et al., 1999; Siedenberg et al., 1999). Although thesedeficits have usually been attributed to defects in cortical processing(Noth et al., 1985; Abbruzzese & Berardelli, 2003), not all of them canbe explained by alterations in the primary sensory pathways(Siedenberg et al., 1999). However, systematic studies of skeletal

muscle pathology or function in HD patients have not been carriedout. In the light of the present study, this would clearly be worth doingin the future.

Conclusions

When taken together our data suggest that, despite the inexorable anduniform progression of muscle atrophy in R6 ⁄ 2 mice from � 8 weeksof age, most neuromuscular junctions remained competent in theirfunction. However, the function of increasing numbers of thesejunctions deteriorated with age, evidently through a combination ofpre- and postsynaptic impairment. The progression of abnormalitieswas not linear; indeed, most abnormalities were not seen consistentlyuntil 15 weeks of age, that is, during the last 1–2 weeks of life. Thissuggests that one possible explanation for the morbidity and mortalityof R6 ⁄ 2 mice over the age of 15 weeks is failure of skeletal muscletransmission subsequent to a collapse both in muscle fiber strength andin neuromuscular transmission. Irrespective of the cause, it is clear thatthere is significant and progressive pathology of muscles andneuromuscular junctions in the postsymptomatic R6 ⁄ 2 mouse whichseems likely to contribute to several aspects of motor dysfunction. Ifthe neuromuscular junction deficits we see in the mice are present inhumans then it seems likely that abnormalities in peripheral neuro-muscular function would contribute significantly to the progression ofHD, particularly in the later stages.

Acknowledgements

This work was supported by grants from the Hereditary Disease Foundation(A.J.M.), The Wellcome Trust and the MRC (R.R.R.) We thank Mrs W.Leavens for histology and Mr A. Thomson for technical assistance withconfocal microscopy.

Abbreviations

CTX, l-conotoxin; 4DL, fourth deep lumbrical (muscles); a-BTX,a-bungarotoxin; ACh, acetylcholine; EDL, extensor digitorum longus; EMG,electromyograph(ic); EPP, endplate potential; FDB, flexor digitorum brevis;mEPP, miniature endplate potential; SDH, succinate dehydrogenase; WT, wildtype.

References

Abbruzzese, G. & Berardelli, A. (2003) Sensorimotor integration in movementdisorders. Mov. Disord., 18, 231–240.

Anderson, J.R. (1985) Atlas of Skeletal Muscle Pathology. MTP Press Limited,Lancaster, UK.

Barry, J.A. & Ribchester, R.R. (1995) Persistent polyneuronal innervation inpartially denervated rat muscle after reinnervation and recovery fromprolonged nerve conduction block. J. Neurosci., 15, 6327–6339.

Bates, G., Harper, P.S. & Jones, L. (2002) Huntington’s Disease, 3rd edn.Oxford University Press, Oxford, UK.

Bekoff, A. & Betz, W.J. (1977) Physiological properties of dissociated musclefibers obtained from innervated and denervated adult rat muscle. J. Physiol.(Lond.), 271, 25–40.

Berardelli, A., Noth, J., Thompson, P.D., Bollen, E.L., Curra, A., Deuschl, G.,van Dijk, J.G., Topper, R., Schwarz, M. & Roos, R.A. (1999) Pathophysiol-ogy of chorea and bradykinesia in Huntington’s disease. Mov. Disord., 14,398–403.

Betz, W.J., Caldwell, J.H. & Ribchester, R.R. (1979) The size of motor unitsduring post-natal development of rat lumbrical muscle. J. Physiol. (Lond.),297, 463–478.

Betz, W.J., Caldwell, J.H. & Ribchester, R.R. (1980) Sprouting of active nerveterminals in partially inactive muscles of the rat. J. Physiol. (Lond.), 303,281–297.



Betz, W.J., Mao, F. & Bewick, G.S. (1992) Activity-dependent fluorescentstaining and destaining of living vertebrate motor nerve terminals.J. Neurosci., 12, 363–375.

Blanco, G., Coulton, G.R., Biggin, A., Grainge, C., Moss, J., Barrett, M.,Berquin, A., Marechal, G., Skynner, M., van Mier, P., Nikitopoulou, A.,Kraus, M., Ponting, C.P., Mason, R.M. & Brown, S.D. (2001) Thekyphoscoliosis (ky) mouse is deficient in hypertrophic responses and iscaused by a mutation in a novel muscle-specific protein. Hum. Mol. Genet.,10, 9–16.

Braga, M.F., Anderson, A.J., Harvey, A.L. & Rowan, E.G. (1992) Apparentblock of K+ currents in mouse motor nerve terminals by tetrodotoxin,mu-conotoxin and reduced external sodium. Br. J. Pharmacol., 106,91–94.

Brown, M.C., Holland, R.L. & Hopkins, W.G. (1981) Motor nerve sprouting.Annu. Rev. Neurosci., 4, 17–42.

Carter, R.J., Hunt, M.J. & Morton, A.J. (2000) Environmental stimulationincreases survival in mice transgenic for exon 1 of the Huntington’s diseasegene. Mov. Disord., 15, 925–937.

Carter, R.J., Lione, L.A., Humby, T., Mangiarini, L., Mahal, A., Bates, G.P.,Dunnett, S.B. & Morton, A.J. (1999) Characterization of progressive motordeficits in mice transgenic for the human Huntington’s disease mutation.J. Neurosci., 19, 3248–3257.

Chen, L.Q., Chahine, M., Kallen, R.G., Barchi, R.L. & Horn, R. (1992)Chimeric study of sodium channels from rat skeletal and cardiac muscle.FEBS Lett., 309, 253–257.

Cifuentes-Diaz, C., Nicole, S., Velasco, M.E., Borra-Cebrian, C., Panozzo, C.,Frugier, T., Millet, G., Roblot, N., Joshi, V. & Melki, J. (2002) Neurofilamentaccumulation at the motor endplate and lack of axonal sprouting in a spinalmuscular atrophy mouse model. Hum. Mol. Genet., 11, 1439–1447.

Constantini, S., Schiller, Y., Cohen, A.M. & Rahamimoff, R. (1987)Pathophysiology of the neuromuscular junction in diabetic rats. Isr. J.Med. Sci., 23, 101–106.

Costanzo, E.M., Barry, J.A. & Ribchester, R.R. (1999) Co-regulation ofsynaptic efficacy at stable polyneuronally innervated neuromuscularjunctions in reinnervated rat muscle. J. Physiol. (Lond.), 521, 365–374.

Costanzo, E.M., Barry, J.A. & Ribchester, R.R. (2000) Competition at silentsynapses in reinnervated skeletal muscle. Nat. Neurosci., 3, 694–700.

Culling, C.F.A., Allison, R.T. & Barr, W.T. (1985) Cellular PathologyTechnique, 4th edn. Butterworths, London.

Cummins, T.R., Aglieco, F. & Dib-Hajj, S.D. (2002) Critical moleculardeterminants of voltage-gated sodium channel sensitivity to mu-conotoxinsGIIIA ⁄ B. Mol. Pharmacol., 61, 1192–1201.

Davies, S.W., Turmaine, M., Cozens, B.A., DiFiglia, M., Sharp, A.H.,Ross, C.A., Scherzinger, E., Wanker, E.E., Mangiarini, L. & Bates, G.P.(1997) Formation of neuronal intranuclear inclusions underlies theneurological dysfunction in mice transgenic for the HD mutation. Cell, 90,537–548.

Djousse, L., Knowlton, B., Cupples, L.A., Marder, K., Shoulson, I. & Myers,R.H. (2002) Weight loss in the early stages of Huntington’s disease.Neurology, 59, 1325–1330.

Ferrante, R.J., Kowall, N.W. & Richardson, E.P. Jr (1991) Proliferative anddegenerative changes in striatal spiny neurons in Huntington’s disease: acombined study using the section-Golgi method and calbindin D28kimmunocytochemistry. J. Neurosci., 11, 3877–3887.

Ferri, A., Sanes, J.R., Coleman, M.P., Cunningham, J.M. & Kato, A.C. (2003)Inhibiting axon degeneration and synapse loss attenuates apoptosis anddisease progression in a mouse model of motoneuron disease. Curr. Biol., 13,669–673.

Filipe, M.I. & Lake, B.D. (1983) Histochemistry in Pathology. ChurchillLivingstone, Edinburgh, UK.

Fischer, L.R., Culver, D.G., Tennant, P., Davis, A.A., Wang, M., Castellano-Sanchez, A., Khan, J., Polak, M.A. & Glass, J.D. (2004) Amyotrophic lateralsclerosis is a distal axonopathy: evidence in mice and man. Exp. Neurol.,185, 232–240.

Frey, D., Schneider, C., Xu, L., Borg, J., Spooren, W. & Caroni, P. (2000) Earlyand selective loss of neuromuscular synapse subtypes with low sproutingcompetence in motoneuron diseases. J. Neurosci., 20, 2534–2542.

Gillingwater, T.H., Thomson, D., Mack, T.G., Soffin, E.M., Mattison, R.J.,Coleman, M.P. & Ribchester, R.R. (2002) Age-dependent synapse with-drawal at axotomised neuromuscular junctions in Wld (s) mutant andUbe4b ⁄ Nmnat transgenic mice. J. Physiol. (Lond.), 543, 739–755.

Gillingwater, T.H., Ingham, C.A., Coleman, M.P. & Ribchester, R.R. (2003)Ultrastructural correlates of synapse withdrawal at axotomized neuromus-cular junctions in mutant and transgenic mice expressing the Wld gene.J. Anat., 203, 265–176.

Graveland, G.A., Williams, R.S. & DiFiglia, M. (1985) Evidence fordegenerative and regenerative changes in neostriatal spiny neurons inHuntington’s disease. Science, 227, 770–773.

Guidetti, P., Charles, V., Chen, E.Y., Reddy, P.H., Kordower, J.H., Whetsell,W.O. Jr, Schwarcz, R. & Tagle, D.A. (2001) Early degenerative changes intransgenic mice expressing mutant huntingtin involve dendritic abnormalitiesbut no impairment of mitochondrial energy production. Exp. Neurol., 169,340–350.

Harper, P.S. (1991) Huntington’s disease. 1st Ed. W.B. Saunders Co. Ltd.,London, UK.

Harris, J.B. & Ribchester, R.R. (1979) The relationship between end-plate sizeand transmitter release in normal and dystrophic muscles of the mouse.J. Physiol. (Lond.), 296, 245–265.

Harris, J.B. & Ward, M.R. (1974) A comparative study of ‘denervation’ inmuscles from mice with inherited progressive neuromuscular disorders. Exp.Neurol., 42, 169–180.

Hickey, M.A. & Chesselet, M.F. (2003) The use of transgenic and knock-in miceto study Huntington’s disease. Cytogenet. Genome Res., 100, 276–286.

Hurlbert, M.S., Zhou, W., Wasmeier, C., Kaddis, F.G., Hutton, J.C. & Freed,C.R. (1999) Mice transgenic for an expanded CAG repeat in theHuntington’s disease gene develop diabetes. Diabetes, 48, 649–651.

Jack, J.J.B., Noble, D. & Tsien, R.W. (1974) Electric current flow in excitablecells. Oxford University Press, Oxford.

Katz, B. & Thesleff, S. (1957) On the factors which determine the amplitude ofthe miniature end-plate potential. J. Physiol. (Lond.), 137, 267–278.

Klapstein, G.J., Fisher, R.S., Zanjani, H., Cepeda, C., Jokel, E.S., Chesselet,M.F. & Levine, M.S. (2001) Electrophysiological and morphologicalchanges in striatal spiny neurons in R6 ⁄ 2 Huntington’s disease transgenicmice. J. Neurophysiol., 86, 2667–2677.

Leblhuber, F., Windhager, E., Reisecker, F. & Rittmannsberger, H. (1991) Longlatency EMG responses in early diagnosis of Huntington’s chorea. Eur. Arch.Psychiatry Clin. Neurosci., 241, 113–114.

Li, R.A., Ennis, I.L., Xue, T., Nguyen, H.M., Tomaselli, G.F., Goldin, A.L. &Marban, E. (2003) Molecular basis of isoform-specific micro-conotoxinblock of cardiac, skeletal muscle, and brain Na+ channels. J. Biol. Chem.,278, 8717–8724.

Lin, W., Burgess, R.W., Dominguez, B., Pfaff, S.L., Sanes, J.R. & Lee, K.F.(2001) Distinct roles of nerve and muscle in postsynaptic differentiation ofthe neuromuscular synapse. Nature, 410, 1057–1064.

Lione, L.A., Carter, R.J., Hunt, M.J., Bates, G.P., Morton, A.J. & Dunnett, S.B.(1999) Selective discrimination learning impairments in mice expressing thehuman Huntington’s disease mutation. J. Neurosci., 19, 10428–10437.

Luthi-Carter, R., Hanson, S.A., Strand, A.D., Bergstrom, D.A., Chun, W.,Peters, N.L., Woods, A.M., Chan, E.Y., Kooperberg, C., Krainc, D., Young,A.B., Tapscott, S.J. & Olson, J.M. (2002) Dysregulation of gene expressionin the R6 ⁄ 2 model of polyglutamine disease: parallel changes in muscle andbrain. Hum. Mol. Genet., 11, 1911–1926.

Mack, T.G., Reiner, M., Beirowski, B., Mi, W., Emanuelli, M., Wagner, D.,Thomson, D., Gillingwater, T., Court, F., Conforti, L., Fernando, F.S., Tarlton,A., Andressen, C., Addicks, K., Magni, G., Ribchester, R.R., Perry, V.H. &Coleman,M.P. (2001)Wallerian degeneration of injured axons and synapses isdelayed by a Ube4b ⁄ Nmnat chimeric gene. Nat. Neurosci., 4, 1199–1206.

Mangiarini, L., Sathasivam, K., Seller, M., Cozens, B., Harper, A., Hethering-ton, C., Lawton, M., Trottier, Y., Lehrach, H., Davies, S.W. & Bates, G.P.(1996) Exon 1 of the HD gene with an expanded CAG repeat is sufficient tocause a progressive neurological phenotype in transgenic mice. Cell, 87,493–506.

Marshall, M.W. & Ward, M.R. (1974) Anode break excitation in denervated ratskeletal muscle fibers. J. Physiol. (Lond.), 236, 413–420.

McLachlan, E.M. & Martin, A.R. (1981) Non-linear summation ofend-plate potentials in the frog and mouse. J. Physiol. (Lond.), 311, 307–324.

Meade, C.A., Deng, Y.P., Fusco, F.R., Del Mar., N., Hersch, S., Goldowitz, D.& Reiner, A. (2002) Cellular localization and development of neuronalintranuclear inclusions in striatal and cortical neurons in R6 ⁄ 2 transgenicmice. J. Comp. Neurol., 449, 241–269.

Miura, H., Oda, K., Endo, C., Yamazaki, K., Shibasaki, H. & Kikuchi, T.(1993) Progressive degeneration of motor nerve terminals in GAD mutantmouse with hereditary sensory axonopathy. Neuropathol. Appl. Neurobiol.,19, 41–51.

Morton,A.J.&Edwardson J.M. (2001) Progressive depletion of complexin II in atransgenicmousemodel of Huntington’s disease. J. Neurochem., 76, 166–172.

Morton, A.J., Lagan, M.A., Skepper, J.N. & Dunnett, S.B. (2000)Progressive formation of inclusions in the striatum and hippocampus ofmice transgenic for the human Huntington’s disease mutation. J. Neurocy-tol., 29, 679–702.



Murphy, K.P.S., Carter, R.J., Lione, L.A., Mangiarini, L., Mahal, A., Bates,G.P., Dunnett, S.B. & Morton, A.J. (2000) Abnormal synaptic plasticity andimpaired spatial cognition in mice transgenic for exon 1 of the humanHuntington’s disease mutation. J. Neurosci., 20, 5115–5123.

Nakanishi, T. & Norris, F.H. (1970) Motor fibers in rat sural nerve. Exp.Neurol., 26, 433–435.

Noth, J., Podoll, K. & Friedemann, H.H. (1985) Long-loop reflexes in smallhand muscles studied in normal subjects and in patients with Huntington’sdisease. Brain, 108, 65–80.

Pachter, B.R. & Eberstein, A. (1992) Long-term effects of partial denervation onsprouting and muscle fiber area in rat plantaris. Exp. Neurol., 116, 246–255.

Patel, A.N., Razzak, Z.A. & Dastur, D.K. (1969) Disuse atrophy of humanskeletal muscles. Arch. Neurol., 20, 413–421.

Pun, S., Sigrist, M., Santos, A.F., Ruegg, M.A., Sanes, J.R., Jessell, T.M.,Arber, S. & Caroni, P. (2002) An intrinsic distinction in neuromuscularjunction assembly and maintenance in different skeletal muscles. Neuron, 34,357–370.

Reynolds, M.L. & Woolf, C.J. (1992) Terminal Schwann cells elaborateextensive processes following denervation of the motor endplate.J. Neurocytol., 21, 50–66.

Ribchester, R.R., Mao, F. & Betz, W.J. (1994) Optical measurements ofactivity-dependent membrane recycling in motor nerve terminals ofmammalian skeletal muscle. Proc. R. Soc. Lond. B Biol. Sci., 255, 61–66.

Ribchester, R.R., Tsao, J.W., Barry, J.A., Asgari-Jirhandeh, N., Perry, V.H. &Brown, M.C. (1995) Persistence of neuromuscular junctions after axotomy inmice with slow Wallerian degeneration (C57BL ⁄WldS). Eur. J. Neurosci., 7,1641–1650.

Sanberg, P.R., Fibiger, H.C. & Mark, R.F. (1981) Body weight and dietaryfactors in Huntington’s disease patients compared with matched controls.Med. J. Aust., 1, 407–409.

Sathasivam, K., Hobbs, C., Turmaine, M., Mangiarini, L., Mahal, A., Bertaux,F., Wanker, E.E., Doherty, P., Davies, S.W. & Bates, G.P. (1999) Formationof polyglutamine inclusions in non-CNS tissue. Hum. Mol. Genet., 8,813–822.

Schiller, Y. & Rahamimoff, R. (1989) Neuromuscular transmission indiabetes: response to high-frequency activation. J. Neurosci., 9, 3709–3719.

Siedenberg, R., Goodin, D.S. & Aminoff, M.J. (1999) Changes of forearmEMG and cerebral evoked potentials following sudden muscle stretch inpatients with Huntington’s disease. Muscle Nerve, 22, 1557–1563.

Son, Y.J., Trachtenberg, J.T. & Thompson, W.J. (1996) Schwann cells induceand guide sprouting and reinnervation of neuromuscular junctions. TrendsNeurosci., 19, 280–285.

Sotrel, A., Williams, R.S., Kaufmann, W.E. & Myers, R.H. (1993) Evidence forneuronal degeneration and dendritic plasticity in cortical pyramidal neuronsof Huntington’s disease: a quantitative Golgi study. Neurology, 43, 2088–2096.

Taxt, T. (1983) Cross-innervation of fast and slow-twitch muscles bymotor axons of the sural nerve in the mouse. Acta Physiol. Scand., 117,331–341.

The Huntington’s Disease Collaborative Research Group. (1993) A novel genecontaining a trinucleotide repeat that is expanded and unstable onHuntington’s disease chromosomes. Cell, 72, 971–983.

White, M.M., Chen, L.Q., Kleinfield, R., Kallen, R.G. & Barchi, R.L. (1991)SkM2, a Na+ channel cDNA clone from denervated skeleta1 muscle,encodes a tetrodotoxin-insensitive Na+ channel. Mol. Pharmacol., 39, 604–608.

Wood, S.J. & Slater, C.R. (1997) The contribution of postsynaptic folds to thesafety factor for neuromuscular transmission in rat fast- and slow-twitchmuscles. J. Physiol. (Lond.), 500, 165–176.



Documents

Progressive abnormalities in skeletal muscle and ... · Progressive abnormalities in skeletal muscle and neuromuscular junctions of transgenic mice expressing the Huntington’s disease