European Journal of Neuroscience, Vol. 20, pp. 3092–3114, 2004
ª Federation of European Neuroscience Societies
Progressive abnormalities in skeletal muscle and
neuromuscular junctions of transgenic mice expressing
the 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
1
Division of Neuroscience, University of Edinburgh, George Square, Edinburgh EH8 9JZ, UK
Department of Pharmacology, University of Cambridge, Tennis Court Road, Cambridge CB2 1PD, UK
2
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, we
wondered whether the muscle changes arise from primary pathology. We used R6 ⁄ 2 transgenic mice for our studies. Unlike
denervation atrophy, skeletal muscle atrophy in R6 ⁄ 2 mice occurs uniformly. Paradoxically however, skeletal muscles show agedependent denervation-like abnormalities, including supersensitivity to acetylcholine, decreased sensitivity to l-conotoxin, and
anode-break action potentials. Morphological abnormalities of neuromuscular junctions are also present, particularly in older R6 ⁄ 2
mice. Severely affected R6 ⁄ 2 mice show a progressive increase in the number of motor endplates that fail to respond to nerve
stimulation. Surprisingly, there was no constitutive sprouting of motor neurons in R6 ⁄ 2 muscles, even in severely atrophic muscles
that showed other denervation-like characteristics. In fact, there was an age-dependent loss of regenerative capacity of motor
neurons in R6 ⁄ 2 mice. Because muscle fibers appear to be released from the activity-dependent cues that regulate membrane
properties and muscle size, and motor axons and nerve terminals become impaired in their capacity to release neurotransmitter and
to respond to stimuli that normally evoke sprouting and adaptive reinnervation, we speculate that in these mice there is a progressive
dissociation of trophic signalling between motor neurons and skeletal muscle. However, irrespective of the cause, the abnormalities
at neuromuscular junctions we report here are likely to contribute to the pathological phenotype in R6 ⁄ 2 mice, particularly in late
stages of the disease.
Introduction
Huntington’s disease (HD) is a fatal neurodegenerative disorder
characterized by progressive decline in motor and cognitive function
with insidious onset in the third to fifth decade. In adults, the first motor
symptom of HD is usually chorea, but the motor deficits progress in
advanced disease to rigidity, bradykinesia and dystonia. By the end
stages, the HD patient is usually bedridden with very limited capacity for
voluntary 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 polyglutamine
repeat in the protein huntingtin), the mechanism underlying the
pathology is unknown. The pathological changes in the brains of
HD patients have been well described, and the striking neurodegeneration seen in the caudate and putamen has, for many years,
focused attention on the mechanisms underlying pathology in these
regions. However, with the cloning of the gene came the discovery
that the expression of the HD gene and its protein product
huntingtin is not restricted to the brain (The Huntington’s Disease
Collaborative Research Group, 1993). Indeed, huntingtin expression
Correspondence: Dr Jenny Morton, as above.
E-mail: [email protected]
Received 16 June 2004, revised 10 September 2004, accepted 27 September 2004
doi:10.1111/j.1460-9568.2004.03783.x
is not even neuron-specific, but is found in many tissues including
heart and skeletal muscle. The role of huntingtin in these tissues is
unknown, although many patients exhibit signs of peripheral motor
pathology, including abnormal eye movements, difficulty swallowing, gait abnormality, dysarthria and skeletal muscle wasting (for
references, see Bates et al., 2002). Further, skeletal muscle atrophy
(which undoubtedly contributes to weight loss) is observed in many
HD patients despite an adequate diet and feeding (Sanberg et al.,
1981). Because the general health of HD patients declines as their
disorder progresses, it has generally been assumed that changes in
body mass are secondary to other symptoms. However, significant
body weight changes are measurable in early (Djousse et al., 2002)
as well as late stage HD patients. This suggests that weight loss
due 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, the
possibility that a peripheral pathology contributes directly to these
symptoms 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 for
the human HD mutation (Mangiarini et al., 1996). R6 ⁄ 2 mice exhibit
a progressive neurological phenotype that includes abnormal
Abnormal neuromuscular junction function in HD mice 3093
involuntary movements, tremor and progressive deterioration of motor
and cognitive function (Mangiarini et al., 1996; Carter et al., 1999;
Lione et al., 1999). As well, they exhibit a number of peripheral
symptoms which replicate those seen in HD, in particular, wasting of
skeletal muscle (Sathasivam et al., 1999).
The cause of skeletal muscle atrophy in R6 ⁄ 2 mice and HD patients
is unknown. However, skeletal muscle atrophy is an indicator of nerve
and ⁄ or muscle pathology in a wide variety of diseases, and very subtle
atrophies can be strongly indicative of pathology. In this study
therefore we sought to establish whether the muscle atrophy observed
in the R6 ⁄ 2 mouse is part of the primary pathology associated with the
HD mutation or whether it is secondary to other previously
unrecognized features of the disorder, for example, impairment of
neuromuscular transmission. Our data suggest that both motor neurons
and muscle fibers are independently affected by the HD mutation and
that neuromuscular changes may contribute to the pathological
phenotype in HD, particularly in late stages of the disease.
Materials and methods
the motor endplate, or where the synaptic currents originate; synaptic
potentials of virtually identical magnitude and time course are
recorded at all points along the length of the fiber.) The deep
lumbrical muscles of the hind foot were used because they are
particularly amenable to fluorescence immunocytochemistry (because
they are very thin, and therefore there are minimal problems with
penetration and diffusion of fixative and antibodies for immunostaining). Nerve terminals and endplates are readily observable in wholemount preparations, either in regular fluorescence or confocal
microscopes. Finally, the fourth deep lumbrical muscle has a dual
nerve supply in rodents (Nakanishi & Norris, 1970; Betz et al., 1979;
Taxt, 1983; Costanzo et al., 2000), one via an anastomosis of a branch
of the sural nerve with the lateral plantar nerve, the other supplied by
the same nerve (tibial nerve) as the FDB muscle. Section of the tibial
nerve in anaesthetized animals thus renders FDB and the medial
lumbrical muscles completely denervated, but the fourth deep
lumbrical muscle is subject to a controlled partial denervation, making
the muscle ideal for studies of motor nerve sprouting. We also made
whole mounts of the triangularis sterni muscle, an exceptionally thin
respiratory muscle, well suited to immunocytochemistry.
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 colony
established in the Department of Pharmacology, University of
Cambridge, and maintained by back-crossing transgenic males onto
female CBA · C57Bl ⁄ 6 F1 mice. Some experiments were carried out
using mice that were transferred to Edinburgh. In all cases, dry chow
was supplemented by twice daily moist chow (mash) from weaning
onwards, and lowered waterspouts were also provided. We have
shown previously that this regime improved health and survival of
R6 ⁄ 2 mice (for further details of husbandry see Carter et al., 2000).
Genotyping was carried out using a polymerase chain reaction based
on 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 used
in the study.
Surgery
R6 ⁄ 2 (N ¼ 18) and WT (N ¼ 16) mice were used for experiments to
study the neuromuscular response to denervation or reinnervation.
Reinnervation experiments were done after crushing the entire sciatic
nerve. Partial denervation experiments were done by cutting the tibial
nerve, 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 or
tibial nerve was exposed and a 1–2-mm section was either cut (but not
removed), or crushed using fine forceps. Mice were then kept for
1–40 days before being killed by cervical dislocation.
A number of different muscles from the hind limb and the chest
were used in this study. The hind limb muscles were: flexor digitorum
brevis (FDB), the peroneal, soleus and extensor digitorum longus
(EDL) muscles from the anterior calf, and the deep lumbrical muscles
of the hind foot. Different muscles were used as appropriate. For
example, FDB is a superficial muscle from the foot which is well
suited to location and electrophysiological recording of synaptic
responses. (This is on account of the short length of the muscle fibers;
< 500 lm in length. As a consequence, they are ‘isopotential’ with
respect to membrane potential changes induced locally at any point
along their length. Thus, it does not matter where the intracellular
micropipette tip is located with respect to longitudinal distance from
Histology
Peroneal muscles of the hind limb were dissected and snap-frozen in
liquid isopentane cooled over liquid nitrogen. Serial transverse
sections (20 lm) were cut from the belly of each muscle using a
cryostat. For fiber typing, sections were stained histochemically for
enzyme activity of NADH diaphorase or succinate dehydrogenase
(SDH), both standard markers for the oxidative enzymes that
characterize type I fibers (Filipe & Lake, 1983). Parallel sections
were prepared for standard histological analysis by staining with
Harris’s haematoxylin and eosin (stains cytoplasm and nuclei), Cresyl
Violet (stains Nissl substance), Weigert’s iron haematoxylin (stains
nuclei) and van Geison’s stain (stains connective tissue). Selected
sections were also stained with Oil Red O (stains lipid) and the
periodic acid–Schiff reaction (stains glycogen). For detailed methods,
see Culling et al. (1985). Unless otherwise stated, all reagents for
histological 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 each
section, fibers were classified into type I or type II fibers according to
their NADH diaphorase reaction, and the diameters of at least 75 type I
and 75 type II fibers were measured and recorded using the methodology described by Patel et al. (1969). Proportions of fiber type were
assessed in three sections from each muscle. All the fibers within three
500 · 500-lm square fields were classified according to their NADH
diaphorase reaction, and the number of each type was counted using an
overlaid grid. To visualize intranuclear inclusions, cryosections of freshfrozen tissues were cut onto gelatinized slides, fixed for 30 min in 4%
paraformaldehyde and processed for immunocytochemistry. Sections
were incubated with a rabbit antiubiquitin antibody (DAKO) at 1 : 2000
dilution. A horseradish peroxidase-conjugated second antibody
(1 : 1000 dilution, DAKO) was used and immunoreactive components
were visualized using diaminobenzidine (Sigma, UK).
Neuromuscular junction staining
FDB, lumbrical and triangularis sterni muscles were prepared for
immunocytochemistry by fixing in 0.1 m PBS containing 4% paraformaldehyde for 30–40 min. Acetylcholine (ACh) receptors were labelled
by incubation for up to 30 min in 5 lg ⁄ mL TRITC-conjugated
a-bungarotoxin (a-BTX: Molecular Probes, USA). Muscles were
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
3094 R. R. Ribchester et al.
washed in PBS and blocked in 4% BSA and 0.5% Triton-X in 0.1 m PBS
for 30 min before incubation overnight in primary antibodies directed
against either the 165-kDa neurofilament proteins (mouse monoclonal
2H3), the synaptic vesicle protein SV2 (mouse monoclonal; both
1 : 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 washing
for 30 min in blocking solution (see above), muscles were incubated for
4 h in a 1 : 200 dilution of appropriate (antimouse IgG or antirabbit)
secondary antibodies conjugated to either FITC or TRITC (Diagnostics
Scotland or DAKO), as indicated in the relevant figures. Muscles were
then whole-mounted in Vectashield (Vector Laboratories, Burlingame,
CA, USA). Some preparations were stained for cholinesterase by a
modified Karnovsky–Roots method, described in Harris & Ribchester
(1979). Fiber diameter and endplate area analysis were made using the
public domain NIH Image program (developed at the US National
Institutes of Health and available on the Internet at http://rsb.info.nih.gov/nih-image/). For endplate area analysis, the region covered by the
endplate was outlined and measured. This procedure overestimates the
area of motor nerve terminal contact, but it is sufficiently robust to test
the correlation between endplate size and muscle fiber size (Harris &
Ribchester, 1979). Fiber diameter was measured within 100 lm of the
endplate.
Some preparations were ‘vital’ stained with the dye FM1-43, a
marker of endocytosis (Molecular Probes), a marker of endocytosis in
functioning nerve terminals (Betz et al., 1992; Ribchester et al., 1994;
Barry & Ribchester, 1995). Preparations were incubated in oxygenated
depolarizing saline (K+ increased to 50 mm, Na+ decreased by 45 mm;
concentrations of the other ions in the bathing medium were as given
under Electrophysiology, below) containing 4 lm FM1-43, for
5–15 min, washed and then imaged in situ. Immunofluorescent and
vitally stained preparations were imaged using either a standard
fluorescence microscope (Olympus) or a laser scanning confocal
microscope (Biorad Radiance 2000). Confocal z–series were merged
using Lasersharp (Biorad) software.
Electron microscopy
FDB muscles were fixed in ice-cold 0.1 m phosphate buffer containing 4% paraformaldehyde and 2.5% glutaraldehyde for 4 h. Preparations were then washed in 0.1 m phosphate buffer, postfixed in 1%
osmium tetroxide solution for 45 min and dehydrated through an
ascending series of ethanol solutions before embedding in Durcupan
resin (Sigma, UK) and sectioning at 75–90 nm. Sections were
collected on Formvar-coated grids (Agar Scientific, UK), stained with
uranyl acetate and lead citrate and viewed in a Philips CM12 TEM.
EM negatives taken between 2000· and 60000· were scanned at
600 dpi using a Linoscan 1200 (Heidelberg Instruments, Heidelberg,
Germany) equipped with a transparency adaptor, before importing into
Adobe Photoshop for analysis and presentation.
Electrophysiology
Intracellular recordings were made from isolated FDB nerve–muscle
preparations. In initial experiments, FDB muscles were isolated in
Cambridge early in the morning, bathed in oxygenated HEPESbuffered physiological saline and couriered the same day to
Edinburgh, where they were then transferred to bicarbonate-buffered
saline (below) for electrophysiological recording the same afternoon,
as described previously (Mack et al., 2001). The isolated muscles were
pinned 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 transferred
from Cambridge to Edinburgh and killed, and muscles were directly
dissected into the bicarbonate-buffered medium. Muscle contractions
were 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 medium
throughout subsequent recordings. Normally, up to 30 muscle fibers
per muscle were sampled using microelectrodes filled with 4 m
potassium acetate (impedance 40 MW), according to standard
techniques. Spontaneous and evoked endplate potentials were recorded using either WPI M707 (WP Instruments Inc.) or Axoclamp 2B
amplifiers (Axon Instruments) and stored and analysed on a PC using
either WinWCP v3.0.8 software (developed and distributed by Dr
John Dempster, Strathclyde University), Spike-2 (Cambridge Electronic Designs, Cambridge, UK), or MiniAnalysis (Synaptosoft,
Atlanta, USA). In some experiments, input resistance was measured
by applying current to the recording microelectrode via a Wheatstonebridge circuit built into the recording amplifier. Constant current was
applied to set the resting membrane potential at )80 mV. The
additional steady-state membrane hyperpolarization evoked by superimposed weak hyperpolarizing current pulses (1–5 nA, 30–100 ms
duration) was then measured. In some fibers, stronger current pulses
were injected to test for ‘anode-break’ excitation of the muscle fibers
on termination of the current pulse (Marshall & Ward, 1974). In other
experiments, iontophoretic responses to ACh were tested using
micropipettes filled with 1 m ACh (> 100 MW input impedance). A
holding current of 1–3 nA was applied to the pipettes to prevent
leakage of ACh. Carefully reducing the holding current produced slow
depolarization of the muscle fiber impaled with the recording
microelectrode when the tip of the iontophoretic pipette was in the
vicinity of its endplate. After restoring the holding current, the
electrode was displaced 100 lm to place its tip over the extrajunctional 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 made
using blunt (1–2 MW) pipettes filled with 1 m NaCl. The pipette tip
was inserted into the belly of the FDB muscles and electromyographic
(EMG) responses were evoked by suprathreshold stimulation of the
muscle nerve (nominally 10–30 V, 0.1–0.5 ms duration). In some
experiments, muscle tension recordings were made by pinning the
proximal tendon of FDB muscles to the Sylgard-lined base of the
recording chamber. The distal tendons were gathered and tied with a
short silk suture then attached to a sensitive force transducer (Akers,
Olso, Norway) connected to a custom Wheatstone-bridge circuit and
preamplifier. The output was filtered and further amplified using
Neurolog equipment (Digitimer, UK). Resting muscle length was
adjusted 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 medium
during recording of either evoked EMG or muscle tension responses.
Statistical analysis
Data are presented as mean ± SEM in each group, when n ¼ number
of muscle fibers and N ¼ number of muscles or animals used, or as
median, interquartile range and 5–95% outliers. Where appropriate,
data were analysed either by unpaired Student’s t-test, Mann–Whitney
test or one-way anova. Dunnett’s post hoc test was used to determine
levels of significance following anova.
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
Abnormal neuromuscular junction function in HD mice 3095
Results
Experiments were performed over a period of 44 months, from 2000
to 2004. Initially, muscles for physiological recording were dissected
in Cambridge and couriered to Edinburgh in physiological saline for
analysis and further processing the same day. In later experiments,
surgery, physiological measurements, immunocytochemistry, histochemistry and electron microscopy were made on muscles isolated
from R6 ⁄ 2 mice and their WT littermates following their delivery to
Edinburgh. Over the course of the study, it became evident that the
delay 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 by
the end. Mice did not normally live beyond 16 weeks at the start of the
study, but routinely survived to 18 weeks or older by the end. This
improved longevity occurred in spite of a spontaneous and progressive
increase in the CAG repeat length in the colony over the period of the
study (data not shown) and is attributed primarily to improvements in
husbandry (Carter et al., 2000). None the less, the data are presented
here in relation to the chronological age of the mice, with ‘severely
affected’ 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 ⁄ 2
mice begins to slow and by 12 weeks they begin to lose weight (see
below). At the same time they show an increasing number of abnormal
motor signs, such as excessive hind limb grooming and tremor. Like
many other mutant mice with neurological dysfunction, R6 ⁄ 2 mice
also exhibit abnormal hind-limb reflex responses to tail lifting: hip
flexion and toe clasping rather than extension. By 12 weeks, R6 ⁄ 2
mice are noticeably weaker and less active than their WT littermates
and they perform poorly in behavioural tasks requiring strength. For
example, they have difficulty swimming, performing on the rotorod or
lifting 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 their
home cage they appear generally healthy, and are not overtly different
from their WT littermates.
In the present study, body weights of R6 ⁄ 2 mice at 12 weeks were
21.9 ± 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 g
compared 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 mice
continue to grow. By 15 weeks, the mice show a marked lordokyphosis (hunchedback spine; Fig. 1a and b), poor mobility and
pronounced muscle weakness. However, flexion reflex responses to
tail lifting persisted. Overt motor symptoms such as spontaneous
hindlimb grooming are present by 12 weeks. In R6 ⁄ 2 mice, 1.2 ± 0.6
instances of hindlimb grooming were recorded (in a 10-min observation period in the open field; N ¼ 17) compared with 0.3 ± 0.3
instances recorded in WT mice (N ¼ 16). By 15 weeks, 7.7 ± 2.7
instances of hindlimb grooming were recorded in R6 ⁄ 2 mice (in
10 min) compared with 0.2 ± 0.2 instances in WT mice. The motor
symptoms progress until the premature death of the mice, usually
between 16 and 20 weeks of age.
Skeletal muscle atrophy
By 12 weeks of age, R6 ⁄ 2 mice show visible atrophy of hind limb
skeletal muscles compared with WT littermates and, by 15 weeks, this
muscle atrophy is very severe (Fig. 1c and d). To examine the
pathology of the skeletal muscle in more detail, cryosections of
gluteus maximus and peroni from 15-week-old mice were processed
and 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 showed
normal morphology with haematoxylin and eosin, Cresyl Violet, Van
Geison’s, Oil Red O and periodic acid–Schiff stains. The frequency
and prominence of nuclei was normal, and there were no centrally
located nuclei. Other pathological signs that were sought, but not
found, included central cores, nemaline rods, granular cytoplasm (in
haematoxylin and eosin staining), basophilic rims, target fibers, ring
fibers, cytoplasmic vacuoles, basophilic blebs, concentric lamellae,
fibrosis and thickened endomysium (data not shown). Although
atrophic, the muscle fibers looked healthy and there was no evidence
of cell death, such as inflammatory cell infiltrates, fibro-fatty change,
or regenerating fibers. Oil Red O revealed no evidence of lipid
accumulation. Apart from the diffuse atrophy and fiber type changes
(see below), there was no evidence of any of the other changes
typically associated with muscle pathology.
The extent of muscle atrophy was measured in both transverse
sections (Fig. 1e and f) and teased muscle fiber preparations of peroneal,
SOL and EDL muscles (Fig. 2). Some transverse sections were stained
histochemically for NADH diaphorase to distinguish type I from type II
muscle fibers (Fig. 1c and d). By 8 weeks, muscle fiber diameter was
slightly, though significantly, smaller in R6 ⁄ 2 mice than WT mice
(Fig. 1i and j). However, muscle fiber diameter collapsed between 8 and
12 weeks. By 16 weeks, there was a severe, generalized atrophy of all
muscle fibers in R6 ⁄ 2 mice that we examined and diameters of muscle
fibers from R6 ⁄ 2 mice were half the size of age-matched WT muscles.
For instance, in teased muscle fiber preparations of EDL muscles, fiber
diameter 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 I
and type II muscle fibers were atrophic. In transverse sections of the
peroneal muscles, the diameter of type I and II muscle fibers in WT
muscles were 25.4 ± 0.3 and 41.3 ± 0.3 lm, respectively, compared
with 15.9 ± 0.2 and 17.4 ± 0.2 lm in R6 ⁄ 2 (n ¼ 675 muscle fibers in
each case). Although atrophy occurred in both fiber types (see also
Sathasivam et al., 1999), the proportion of type I and II fibers was
altered. In WT peroneal muscles, 51 ± 2% of fibers were type I compare
with 81.8 ± 2.1% in the R6 ⁄ 2 mouse. However, there were no signs of
degeneration, nor was there any marked change in grouping of muscle
fibers within the muscle. Thus it appears that type II fibers were
converted to type I during the process of muscle atrophy rather than the
change occurring by another process (e.g. fiber splitting).
Fiber diameter and motor endplate area
Muscle atrophy was measured in teased fiber preparations of R6 ⁄ 2
mouse EDL, soleus and FDB muscles, after staining histochemically
for cholinesterase at motor endplates (Harris & Ribchester, 1979). This
method typically stains the rims of junctions, delineating the extent of
secondary folds. There were no discernible differences in the pattern or
density of cholinesterase staining in R6 ⁄ 2 muscles (Figs 2a and b, and
3). Every fiber showed a single motor endplate, and the ‘pretzel’-like
structure of the endplates were similar to those seen in WT muscle.
Further, there was no overt fragmentation or expansion of endplate
areas 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.
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
3096 R. R. Ribchester et al.
By 16 weeks, R6 ⁄ 2 muscle fiber diameters were 20–40% less than WT
muscle diameters. This atrophy was evident in all three muscles but
greatest in EDL (Fig. 2c). Interestingly, endplate areas also shrank in
R6 ⁄ 2 muscles as muscle atrophy progressed (Fig. 2d), but the ratio of
endplate area to fiber diameter was unaltered, as evident from plots of
endplate 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 mouse
models of HD is the presence of abnormal aggregates of ubiquitinated
protein (Davies et al., 1997). Inclusions have been observed in
quadriceps muscle (Sathasivam et al., 1999). To check for the
presence of inclusions in the muscles we immunostained for ubiquitin
in both whole mounts and transverse muscle sections. WT muscles
showed no evidence of ubiquitin accumulation. Some myonuclei in
R6 ⁄ 2 EDL and soleus muscles stained immunopositive for intranuclear inclusions. The morphological appearance of these inclusions
was 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 weeks
they were present in 20% of EDL muscle fibers (Fig. 1g and h). A
similar percentage ( 20%) of myonuclei was immunopositive for
ubiquitin in whole-mount immunofluorescent specimens of the
triangularis sterni muscle. This included nuclei in the motor endplate
region of the muscle fibers (Fig. 3b).
Morphology of neuromuscular junctions
ACh receptors at motor endplates of R6 ⁄ 2 muscles were stained with
TRITC-a-BTX. Co-immunostaining for NF ⁄ SV2 showed that, at all
ages and in all muscles examined, the endplates in R6 ⁄ 2 muscles were
pretzel-shaped and mononeuronally innervated by single axon collaterals in alignment with postsynaptic junctional folds, as in WT muscles
(Fig. 3c–h). This appearance suggests that the adult neuromuscular
innervation pattern develops normally, and this pattern is sustained
throughout the life of R6 ⁄ 2 transgenic mice. Myelinating and perisynaptic terminal Schwann cells were also of normal appearance and there
was no evidence of any constitutive Schwann cell or nerve terminal
sprouting (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 their
incidence was generally very low (estimated at fewer than 5% of
endplates in all muscles). The abnormalities included: long
preterminal axon branches (Fig. 4a); thin, wispy or ‘untwisted’
neurofilaments (Fig. 4b); poor penetration of neurofilaments into
nerve terminals (Fig. 4c); and accumulations of neurofilaments in
preterminal axonal swellings (Fig. 4d). Similar abnormalities have
been reported in mouse models of dying-back neuropathy (Miura
et al., 1993) and motor neuron disease (Frey et al., 2000; Fischer
et al., 2004). Motor nerve terminal abnormalities included partially
occupied motor endplates, with exposed regions of ACh receptor
staining (Fig. 4e). Although this was observed rarely (in three out
of > 600 neuromuscular junctions in three lumbrical muscles
examined; 0.5%), this feature is notable because it has also been
reported in mouse models of motor neuron disease (Cifuentes-Diaz
et 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 a
significant reduction in or absence of endplate staining of ACh
receptors (Fig. 4f). This abnormal feature was not observed in
lumbrical muscles of R6 ⁄ 2 mice at any stage although it is consistent
with physiological failure of synaptic transmission observed in
recordings from FDB muscles (see below). Absence of receptor
staining could not be explained by failure of the TRITC-a-BTX toxin
to penetrate the muscle because endplates in the same confocal plane
were clearly stained (Fig. 4f).
Some lumbrical muscles were vitally stained with FM1-43, a dye
which highlights regions of nerve terminals that contain recycling
synaptic vesicles (Ribchester et al., 1994; Gillingwater et al., 2002).
Almost all neuromuscular junctions in R6 ⁄ 2 mice took up FM1-43 in
response to depolarizing stimuli, indicating they were able to recycle
synaptic vesicles. As in the immunostained material, we saw
occasional instances of partial occupancy of endplates by FM1-43loaded terminals (data not shown).
Electron microscopy
We examined endplates in FDB muscles from two 12-week R6 ⁄ 2
mice, seeking evidence for early changes in nerve terminal ultrastructure. Of 44 endplates examined in detail, 33 appeared normal and 11
showed abnormalities, including membrane inclusions or vacuoles,
pre- or postsynaptic detachment, and insinuation of terminal Schwann
cell processes into synaptic cleft (arrows, Fig. 5a). Two endplates were
found where junctional folds appeared abnormally wide (Fig. 5a and
b). Synaptic vesicles appeared normal but there were vacuolated
inclusions in the presynaptic nerve terminals and the density of
mitochondria appeared abnormally high (Fig. 5a and b).
Myelinated axons in intramuscular nerves appeared normal
(Fig. 5b, inset), confirming the normal appearance of axons in
whole-mounts immunostained and studied using confocal microscopy.
Nerve regeneration and sprouting
Motor nerve endings sprout when muscles atrophy as a result of
muscle paralysis or disuse (Betz et al., 1980; Brown et al., 1981; Pun
et al., 2002). We were therefore surprised by the normal appearance of
most R6 ⁄ 2 neuromuscular junctions in spite of the significant, uniform
and progressive atrophy of the muscle. We hypothesized that atrophic
R6 ⁄ 2 muscles might be incapable of generating nerve regeneration or
sprouting signals, or that motoneurons in these muscles were
incapable of responding to such stimuli. To test this, we first
examined 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 and
lordokyphosis 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 groups
from (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 the
muscles 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 symptomatic
R6 ⁄ 2 limb muscle. (g) There were no ubiquitinated deposits in the myonuclei of WT muscles. (h) About 20% of R6 ⁄ 2 myonuclei contain ubiquitinated
intranuclear 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 significantly
in 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 agematched 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).
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3098 R. R. Ribchester et al.
a
b
WT
c
R6/2
d
750
40
endplate area (µm2)
fiber diameter (µm)
50
**
30
**
*
20
10
0
500
*
250
**
**
0
Wt
WT
R6/2
R6/2
Wt
WT
FDB
R6/2
R6/2
Wt
WT
EDL
R6/2
R6/2
Wt R6/2
R6/2 WT
Wt R6/2
R6/2 WT
Wt R6/2
R6/2
WT
FDB
FDB
SOL
e
EDL
EDL
SOL
SOL
f
1 250
endplate area (µm2)
endplate area (µm2)
1250
1000
750
500
250
0
1 000
750
500
250
0
0
10
20
30
40
fiber diameter (µm)
50
60
0
10
20
30
40
50
60
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 and
endplate 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 significantly
reduced (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 progression
of 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). In
panels 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) or
brain. 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 and
synaptic terminals in R6 ⁄ 2 muscles also showed diffuse immunostaining for ubiquitin. (c and d) Immunostaining for S100 (green) shows no discernible abnormality
in the organization of either myelinating or terminal Schwann cells. (e–h) Neurofilament–SV2 staining (green) shows near-normal appearance of axons, motor nerve
terminals 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).
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3100 R. R. Ribchester et al.
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 triangularis
sterni muscle whose motor nerve terminal is supplied by a long preterminal axon branch (arrow); (b) neurofilaments that are ‘untwisted’ in this triangularis sterni
terminal (arrow), a feature of 5% of terminals; (c) thinning of neurofilaments with poor penetration into a lumbrical nerve terminal; (d) a lumbrical terminal
supplied by an axon showing swollen accumulations of neurofilaments (arrows), with an ultra-fine preterminal branch supplying the motor endplate; (e) a partially
occupied endplate (arrow) (the inset shows a partially occupied lumbrical neuromuscular junction contacted by only two synaptic boutons, leaving the remainder of
the 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).
by crushing the tibial nerve in 6-week-old animals, and studied the
innervation pattern of muscles physiologically using FM1-43
staining and microelectrode recording. In six R6 ⁄ 2 mice studied
5–6 weeks after the nerve crush was done, flexion reflexes had
returned and, in isolated preparations, FM1-43 stained regenerated
motor nerve terminals (Fig. 5c and d). Intracellular recordings from
two of the muscles showed spontaneous miniature endplate
potentials (mEPPs), evoked EPPs and ⁄ or action potentials in
15 ⁄ 20 and 19 ⁄ 20 fibers, respectively. Thus, motor neurons in
young R6 ⁄ 2 mice are capable of axon regeneration and neuromuscular synapse repair.
Reinnervation studies were not possible in the oldest, most severely
affected R6 ⁄ 2 mice because they did not live long enough for axon
regeneration to occur after complete nerve section. However, we
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
Abnormal neuromuscular junction function in HD mice 3101
Fig. 5. Ultrustructural abnormalities were evident in some R6 ⁄ 2 presynaptic terminals but synaptic vesicle recycling appeared largely normal. Electron
micrographs 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 and
infiltration of Schwann cell processes (arrows) into the synaptic cleft, and the presynaptic terminals also contain vacuolated structures. The intramuscular nerves
appear 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-aBTX staining show that motor axons are competent to regenerate functional synapses. This was confirmed by electrophysiological recording and recycled synaptic
vesicles (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).
examined whether motor axons in R6 ⁄ 2 muscles older than 12 weeks
were able to generate compensatory nerve sprouts in response to
partial denervation. Sprouting of motor axons is normally preceded by
sprouting of terminal Schwann cells (Reynolds & Woolf, 1992; Son
et al., 1996) and also, in mice, by down-regulation of the intensity of
S100 immunostaining (W.J. Thompson, personal communication).
The fourth deep lumbrical (4DL) muscles in rodents receive motor
innervation by axons running in both the tibial–lateral plantar and
sural nerves (Nakanishi & Norris, 1970; Betz et al., 1979; Taxt, 1983).
Section of the tibial nerve in anaesthetized animals thus renders FDB
and the medial lumbrical muscles completely denervated but the 4DL
muscle is subject to a controlled partial denervation, making the
muscle ideal for studies of motor nerve sprouting of the sural nerve.
Tibial nerves were cut under anaesthesia in nine R6 ⁄ 2 mice (four at
9–14 weeks and five at 15–18 weeks) and eight age-matched WT
mice. 4DL muscles were immunostained 7 days later and examined
for 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 to
sprout as their muscle became atrophic. Figure 6a–d shows images
taken from 4DL muscles 7 days after partially denervation, stained
for S100 protein. Taking account of the settings of laser power and
the gain and offset on the photomultiplier tubes in the confocal
microscope, the attenuation of the S100 immunostaining signal was
similar in WT and R6 ⁄ 2 mice. All muscles showed strong
indications of Schwann cell and axonal sprouting (Fig. 6e–h). To
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3102 R. R. Ribchester et al.
quantify the sprouting of axons, we scored the number of sprouts
growing either into or out of a-BTX-stained ACh receptor pretzels.
Collateral sprouting from nodes of Ranvier of intact intramuscular
nerve axons was observed in all 9–14 week muscles and in four out
of five muscles from severely affected mice older than 15 weeks
(Fig. 6e–h). We pseudorandomly selected confocal microscope fields
straddled by an intramuscular nerve branch. In the four muscles from
the 9–14-week-old group of R6 ⁄ 2 mice, 17 endplates out of 71
sampled (24%) received collateral axonal sprouts from intact sural
nerve motor axons, compared with 12 out of 53 (23%) of endplates
in WT littermates. In the muscles from R6 ⁄ 2 mice older than
15 weeks, we observed that 13 endplates out of 50 analysed (26%)
received collateral axonal sprouts. However, the sprouts were
discernibly finer than in WT muscles, suggesting a qualitative
impairment. Interestingly, in the WT muscles we also saw several
instances of terminal sprouting (outgrowths from nerve terminals
extending to nearby denervated endplates; arrows in Fig. 6f).
However, we did not observe any of this form of sprouting in any of
four partially denervated muscles from two severely affected R6 ⁄ 2
mice older than 16 weeks.
Together, the morphological data suggest that the innervation
pattern and morphological appearance of neuromuscular junctions in a
variety of R6 ⁄ 2 muscles is quite normal between 8 and 12 weeks of
age, despite the decline in muscle mass during this period. However,
significant and increasing instances of abnormal nerve terminals were
detected in coincidence with increasing severity of the disease
and there was a weakened sprouting response following partial
denervation in muscles from mice older than 12 weeks. Most notably,
whereas a compensatory sprouting reaction in motor nerve terminals
might have been expected in response to the pronounced atrophy seen
in 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 have
focused on central nervous system defects in synaptic transmission, as
measured behaviourally and electrophysiologically. Because R6 ⁄ 2
mice show deteriorating cognitive and behavioural function yet remain
mobile despite their severe muscle wasting and weight loss, it is
unclear whether muscle atrophy is a consequence of central or
peripheral neural defects or whether it is caused by intrinsic changes in
skeletal muscle properties. In preliminary tests we measured forelimb
grip strength. These showed that both WT and severely affected
(15–17-week-old) R6 ⁄ 2 mice were motivated to exert consistent
voluntary force, but R6 ⁄ 2 mice showed some fatigue on repeated
testing (data not shown). However, such tests do not distinguish
neuropathic changes from myopathic ones. We therefore studied
passive membrane properties of R6 ⁄ 2 muscle fibers (resting membrane potential, input resistance and time constant) as well as synaptic
properties (mEPPs and EPPs). We also examined the sensitivity of
muscle fibers to bath-applied and iontophoretically applied ACh, and
the sensitivity of action potentials to the selective muscle NaV1.4
sodium channel blocker l-CTX.
Resting membrane potentials
The mean resting membrane potential of FDB muscle fibers from
WT mice was )72.2 ± 3.2 mV (SEM, N ¼ 6 muscles). This was
significantly different from muscle fibers of presymptomatic R6 ⁄ 2
mice aged < 14 weeks ()62.8 ± 1.52; P < 0.001; N ¼ 10 muscles).
There was a further deterioration of resting membrane potential in
R6 ⁄ 2 muscle fibers as the animals became overtly symptomatic and
the muscle fibers more atrophic (Fig. 7a). By 15–18 weeks, muscle
fiber resting potentials were )55.2 ± 4.3 mV (N ¼ 10 muscles).
This decline was not an artefact of impalement of the atrophic
muscle fibers because we made the recordings using high impedance microelectrodes filled with potassium acetate (> 40 MW). Low
resting membrane potentials were also evident in R6 ⁄ 2 muscles
when using ultra-sharp, very high impedance ( 100 MW) microelectrodes, which also registered normal resting potentials in WT
muscles.
Input resistance and time constant
The small diameter of atrophic muscle fibers would be expected to
confer a high input resistance and prolonged membrane time constant,
as measured by the ratio and time course of the steady-state change in
membrane potential in response to injection of rectangular constantcurrent pulses. Thus, input resistance is proportional to d)3 ⁄ 2, where d
is the muscle fiber diameter (Katz & Thesleff, 1957). FDB muscle
fibers in 15–18-week R6 ⁄ 2 mice are 40% narrower than in WT
mice (see above). As a consequence, input resistance should be about
twice as great. Input resistance and membrane time constant were
estimated using a single microelectrode and Wheatsone bridge method
(Fig. 7b–e). Though imperfect, the single-electrode and bridge method
minimizes damage to muscle fibers and allows effective comparisons
between experimental and control groups of muscles (Barry &
Ribchester, 1995). There was no signficant difference between input
resistances of presymptomatic, nonatrophic R6 ⁄ 2 muscles and those
of WT (data not shown). However, by 15–18 weeks, as expected from
the 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 three
mice) compared with WT (2.34 ± 0.27 MW; n ¼ 27 fibers, N ¼ 3
muscles 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). FDB
muscle fibers are equivalent to short cables with open-circuited ends,
and are isopotential along their length in response to focal constant
current injection (e.g. by synaptic currents, or via intracellular
micropipettes; Bekoff & Betz, 1977). Biophysically, they are similar
in their overall passive membrane behaviour to spherical cells. The
membrane time constant was therefore estimated from the time taken
to reach 63% of the steady-state membrane potential displacement
during the same constant current injections (Jack et al. 1974). In
presymptomatic R6 ⁄ 2 muscles the membrane time constant was not
significantly different from WT (data not shown). However, in
atrophic muscle fibers from 15–18-week R6 ⁄ 2 mice, with higher input
resistances 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 becomes
advanced. Defined settings of the confocal laser power, gain and offset demonstrated that Schwann cells down-regulate S100 protein in response to section of the
tibial 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). Neurofilament
staining (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 with
age-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).
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Abnormal neuromuscular junction function in HD mice 3103
WT
S100
a
R6/2
b
Power = 6.5%
Gain = 4.1
Offset = 1.9
c
Power = 4.7%
Gain = 4.9
Offset = -3.9
d
Power = 8.1%
Gain = 4.9
Offset = -3.9
NF/SV2
Power = 6.5%
Gain = 4.1
Offset = 1.9
e
f
g
h
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3104 R. R. Ribchester et al.
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 severely
affected. 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 measure
membrane 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).
threefold greater: 6.53 ± 0.72 ms (SEM, n ¼ 10 fibers) compared
with 2.22 ± 0.19 ms (n ¼ 17) in WT muscles.
Spontaneous transmitter release
Measurements of spontaneous mEPPs were made in 137 muscle fibers
from 12 muscles of R6 ⁄ 2 mice aged 8–18 weeks. In the younger
mice, mEPP frequencies were in the normal range (mean ± SEM
0.23 ± 0.2 ⁄ s at 18 C, n ¼ 40 fibers from two muscles; compared
with 0.16 ± 0.04 ⁄ s, 25 fibers from two age-matched WT muscles).
mEPP amplitude, rise time and decay time constants were also within
the 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. Betweenevent interval histograms of mEPPs in 18-week-old mice showed
an exponential time course, indistinguishable from the Poisson
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Abnormal neuromuscular junction function in HD mice 3105
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) WT
and (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-whisker
plots summarizing overall median, interquartile range and 95% outliers of mEPP amplitude and frequency in (h) severely affected R6 ⁄ 2 mice and (g) their WT
littermates. Overall, mEPP amplitude was significantly increased about two-fold, as expected from the difference in muscle fiber input resistance in atrophic FDB
muscle 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).
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: median
frequency was 0.27 ⁄ s (interquartile range 0.17–0.5 ⁄ s) compared with
0.60 ⁄ 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 ⁄ 2
mouse neuromuscular junctions. However, further analysis of the
characteristics of spontaneous transmitter release frequency may be
warranted, especially in the light of the finding that at least one synapsespecific protein thought to modulate exocytosis (complexin II) is downregulated in the R6 ⁄ 2 mouse brain (Morton & Edwardson, 2001).
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
3106 R. R. Ribchester et al.
As expected from the measurements of input resistance, mEPP
amplitudes were markedly increased as the phenotype of the R6 ⁄ 2
mice deteriorated. In R6 ⁄ 2 mice older than 15 weeks, mEPPs were
typically 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 mV
n ¼ 11 fibers) from two age-matched WT muscles (P < 0.001,
Mann–Whitney). The coefficient of variation of mEPP amplitude in
some R6 ⁄ 2 muscle fibers was numerically higher (mean ± SEM
0.47 ± 0.27) but not statistically significantly different from WT
(0.34 ± 0.06; P > 0.1, t-test). The instances where the variability
seemed higher may thus have been merely a consequence of the better
signal-to-noise ratio for detection of mEPPs in R6 ⁄ 2 muscle fibers on
account of their high input resistance. However, we cannot rule out at
this 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 the
repolarization phase of the normalized average mEPP were consistently 2–3 times longer in R6 ⁄ 2 than WT.
Evoked EPP characteristics
Muscles were pretreated with l-CTX GIIIB (2.5 lm), with the
intention of blocking muscle action potentials and thereby exposing
the full amplitude and time course of nerve-evoked EPPs. The l-CTX
was very effective in WT muscles, but we experienced considerable
difficulties with older R6 ⁄ 2 muscles because they were resistant to the
blocking effects of the toxin (see below). Movement artefacts were
evident in many of the recordings, and action potentials rather than
EPPs were frequently registered. However, continuous incubation in
l-CTX eventually produced sufficient block to enable recording of
EPPs (Fig. 9). In most fibers, robust EPPs with a low coefficient of
variation were recorded, indicating a high quantal content (Fig. 9b).
We found no instances of random ‘failures’ of synaptic transmission in
any of the trains of EPPs recorded from single muscle fibers, at any
stage in the deterioration of R6 ⁄ 2 mice. For instance, EPP trains were
sustained during repetitive high-frequency (> 30Hz) stimulation
(Fig. 9c) as in WT muscles and there was no evidence for any
systematic change in mean quantal content. Highly variable responses
(including random occurrences of ‘failures’ in response to nerve
stimulation) would have indicated a low quantal content and weak
synaptic transmission. This was never observed in any of the
recordings from > 500 muscle fibers recorded during the course of
this study. Analysis of EPP amplitudes using the coefficient of
variation method indicated that the overall mean quantal content
of EPPs in atrophic muscle fibers was not statistically significantly
different from WT (data not shown). Voltage-clamp analysis of
synaptic currents would perhaps clarify whether there are any subtle
differences in quantal content of EPPs in R6 ⁄ 2 muscle fibers but,
based on the voltage recordings, any such differences are unlikely to
be physiologically significant.
Despite the absence of significant progressive deterioration in EPP
quantal content, there was an increase in the number of muscle fibers
that did not respond at all to nerve stimulation in severely affected
R6 ⁄ 2 mice (older than 14 weeks). Only two out of 206 fibers in 16
WT muscles did not show evoked EPPs or action potentials when the
muscle nerve was stimulated supramaximally. However, in 269
recordings from 18 muscles in R6 ⁄ 2 mice aged 15–18 weeks, 52 of
the fibers (i.e. 20% overall) were consistently unresponsive to
a
b
10 mV
10 mV
10 ms
c
10 ms
d
10 mV
10 mV
10 ms
200 ms
Fig. 9. Evoked EPPs in R6 ⁄ 2 muscles are robust. Typical superimposed serial
traces 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 low
coefficient of variation in amplitude (high quantal content), but note the longer
decay time course in the R6 ⁄ 2 muscle fiber. (c) R6 ⁄ 2 fibers showed robust
responses to high frequency (50 Hz) repetitive stimulation. (d) The prolonged
decay time constant of R6 ⁄ 2 EPPs could not be explained by inactive
cholinesterase because, after a 10-min incubation in the anticholinesterase
neostigmine, all EPPs in R6 ⁄ 2 muscles were significantly prolonged. This
recording was made from a different fiber but the same muscle as in (b).
repeated stimulation and failed to show evoked EPP responses. This
was not a consistent feature between mice or muscles because in eight
out of 18 of the muscles all fibers impaled gave evoked responses.
Nevertheless, complete failure of evoked synaptic transmission was
evident in muscles from half of the severely affected R6 ⁄ 2 mice. It
remains to be established whether this was due to abnormal nerve
conduction or complete pre- or postsynaptic failure of neurotransmitter
release or action.
In spite of the high input resistance of atrophic R6 ⁄ 2 muscle
fibers and the concomitant increase in mEPP amplitudes, the
amplitudes of evoked EPPs were not significantly different from
those in WT muscles (Fig. 10a). However, resting potentials of the
R6 ⁄ 2 muscles were reduced (see above, Fig. 8), and differences in
EPP amplitude could have been obscured by the nonlinear
summation of synaptic potentials as the membrane potential
approached the transmitter null (i.e. reversal) potential (McLachlan
& Martin, 1981). We therefore measured the membrane potential at
the peak of the EPP response. A nonlinear polynomial curve fitted to
the data showed a trend towards increased synaptic efficacy in the
ageing R6 ⁄ 2 mice that did not reach statistical significance (EPPs
depolarized muscle fibers to )40 ± 5 mV (N ¼ 10 muscles) in older
R6 ⁄ 2 mice compared with )48 ± 4 mV in WT mice (N ¼ 4
muscles; P > 0.05; t-test). There were similar trends in EPP latency
and time-to-peak (Fig. 10c and d), although only the time-to-halfdecay of EPPs in atrophic R6 ⁄ 2 muscles was statistically significantly different: about three times longer in R6 ⁄ 2 mice older than
15 weeks compared with WT.
Effect of anticholinesterase
The prolongation of EPPs as R6 ⁄ 2 mice age is not a consequence of
any functional reduction in cholinesterase activity. Histochemically,
cholinesterase distribution at endplates was unaffected (see Fig. 2).
Further, the addition of the cholinesterase inhibitor neostigmine
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
Abnormal neuromuscular junction function in HD mice 3107
b
Age (weeks)
R6/2
age (weeks)
40
0
30
Em at peatk (ms)
EPP amplitude (mV)
a
20
10
0
6
9
12
15
6
9
WT
12
15
18
12
15
18
-10
-20
-30
-40
-50
WT
d
c
time to peatk (ms)
4
latency (ms)
9
-60
18
R6/2
age (weeks)
Age (weeks)
3
2
1
0
2.0
1.5
1.0
0.5
0.0
6
9
12
15
18
WT
WT
R6/2
(weeks)
Age age
(weeks)
R6/2 age (weeks)
WT
WT
f
e
1.0
7.5
5.0
frequency
EPP half decay (ms)
6
2.5
WT
R6/2
0.5
0.0
6
9
12
15
R6/2 age (weeks)
18
WT
WT
0.0
0.0
2.5
5.0
7.5
10.0
Half-Decay
time(ms)
(ms)
half-decayTime
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 the
EPP after allowing for the depolarized resting membrane potential of R6 ⁄ 2 muscle fibers (see Fig. 7), showing that evoked synaptic strength did not change
substantively 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 significant
change 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 in
a–e are best-fit second-order polynomials. (f) Cumulative distributions of relative frequencies of EPP half-decay times from all muscle fibers recorded in muscles
from R6 ⁄ 2 mice (solid line) aged 15–18 weeks were significantly different from WT (dotted line; P < 0.01, Kolmagorov–Smirnov test).
(10 lm) to the bathing medium prolonged the decay time of EPPs in
R6 ⁄ 2 mice about seven-fold, showing that the cholinesterase in
unblocked muscles was functional. The mean ± SD half-decay time
was 22.95 ± 7.06 ms (n ¼ 4 fibers) in one 16-week R6 ⁄ 2 muscle
after neostigmine compared with 3.34 ± 1.54 ms (n ¼ 7) in the same
muscle 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 at
concentrations of 1–3 lm (Cummins et al., 2002; Li et al., 2003).
At this concentration, the toxin is ineffective on axons and motor
nerve terminals, which therefore release physiologically normal
amounts of neurotransmitter in response to nerve stimulation
(Wood & Slater, 1997; Costanzo et al., 1999; but see Braga
et al., 1992).
Incubation of WT muscles in l-CTX routinely and visibly blocked
nerve-evoked muscle contractions within 5–30 min and this block
normally persisted for 1–4 h after returning muscles to normal
physiological saline. As described above, robust endplate potentials
were recorded from virtually all WT muscle fibers during the period of
muscle paralysis induced by the toxin. Muscles from R6 ⁄ 2 mice aged
< 11 weeks were also sensitive to l-CTX; muscle fiber action
potentials were blocked, and robust nerve-evoked EPPs remained.
However, isolated FDB muscles from R6 ⁄ 2 mice older than 12 weeks
were consistently resistant to the blocking effect of l-CTX, showing
visible, moderate-to-strong contractions in response to nerve stimulation, 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 muscle
tension and EMG currents from FDB muscles. Twitches and tetani
declined following l-CTX administration. In two WT muscles,
tension declined to 10% of the initial response (Fig. 11a). In one
16-week R6 ⁄ 2 muscle, tension declined to only 35% of the initial
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
3108 R. R. Ribchester et al.
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 fibers
leaving 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-CTX
to 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 before
adding the toxin while the lower traces show recordings made 15 min after toxin administration. (c and d) EMG recordings using a blunt micropipette filled with
1 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 reduction
in 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 show
expanded views of extracellular action potentials recorded before and after toxin administration. (e) An intracellularly recorded muscle fiber action potential from an
R6 ⁄ 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 muscle
fiber that was so large it triggered an action potential, even though this recording was made in the presence of l-CTX.
response (Fig. 11b), still producing visible contractions that appeared
quite strong under the dissecting microscope.
Next, we recorded extracellular action potentials using 1 m NaClfilled blunt-tipped ( 2–20 lm) glass micropipettes inserted into the
belly of the FDB muscles. In three R6 ⁄ 2 preparations recorded in this
fashion, l-CTX depressed nerve-evoked muscle fiber inward currents
by < 5%, 28% (Fig. 12a) and 48%, whereas extracellular responses
were reduced by 54% and 76% of initial current amplitude in two WT
muscles (Fig. 11c and d). No contractions were visible in the case of
these two WT muscles. The residual responses were therefore caused
by endplate currents. As expected, adding 10 lm a-BTX to the
bathing medium completely blocked junctional ACh receptors and
abolished all evoked responses and contractions in both R6 ⁄ 2 and WT
muscles (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
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
Abnormal neuromuscular junction function in HD mice 3109
a
b
50 mV
5 nA
15 ms
c
d
4 mV
10 µM ACh
60 ms
2 mV
60 s
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 all
severely 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 muscle
twitches and tetani were induced by nerve stimulation, then the bathing medium was substituted with medium containing 10 lm ACh. This produced a muscle
contracture that reached 80% of the twitch contraction response ( 30% of the tetanus evoked by 50 Hz stimulation). Traces in (d) show membrane depolarization
in response to graded 2-ms iontophoretic pulses of ACh applied from a micropipette manipulated over the extrajunctional membrane of the impaled fiber.
full-blown action potential responses, apparently unaffected by l-CTX
treatment. 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 also
resistant to l-CTX (Fig. 11f).
To quantify the extent of l-CTX resistance, we measured the
maximum rate of rise of the first intracellularly evoked response
recorded in each muscle fiber. FDB muscle fiber action potentials
typically have rates of rise > 60 mV ⁄ ms at room temperature, before
treatment with l-CTX. In WT muscles, 90% of EPPs evoked by nerve
stimulation 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 recorded
muscle fiber responses in older R6 ⁄ 2 mice: 42% of muscle fibers in
15–18-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 in
unblocked WT muscles. In the remainder dV ⁄ dt was < 30 mV ⁄ m.
Thus, almost half of the muscle fibers in severely affected R6 ⁄ 2
mouse FDB muscles showed evidence of significant l-CTX
resistance.
Anode-break action potentials
Resistance to tetrodotoxin and l-CTX are properties of rSkM2
(NaV1.5) sodium channels that appear in WT muscle fiber membranes
after denervation (White et al., 1991; Chen et al., 1992). Because
atrophic 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). In
a sample of 10 fibers recorded from one 16-week R6 ⁄ 2 FDB muscle, all
10 fibers showed clear anode-break excitation (Fig. 12). By comparison, > 90% of WT muscle fibers showed no anode-break excitation.
ACh responses
In the light of the other denervation-like physiological and pharmacological characteristics of R6 ⁄ 2 muscles, we examined them for
‘supersensitivity’ to ACh. Figure 12c shows alternating cycles of
responses to single twitch and 30 Hz tetanic stimulation in an R6 ⁄ 2
muscle before, during and after adding ACh (10 lm) to the bathing
medium. This caused a muscle contracture equivalent in magnitude to
the resting twitch response, that is, about one-third of the tetanic
tension response. Similar contractures were evoked in two other 18week R6 ⁄ 2 muscles. In two WT muscles, as expected, the ACh
contracture was < 10% of the nerve-evoked tetanic tension. Thus,
R6 ⁄ 2 muscles were more than three times as sensitive as WT muscles
to 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 WT
muscle fibers (data not show). However, graded depolarizations were
produced in three out of 10 fibers in one 18-week-old R6 ⁄ 2 muscle.
These fibers were innervated, because we also observed spontaneous
mEPPs in the recordings. This incidence of supersensitive fibers is
consistent with the sensitivity to bath-applied ACh and the
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
3110 R. R. Ribchester et al.
other denervation-like properties measured above, such as l-CTX
resistance.
Discussion
The R6 ⁄ 2 transgenic mouse model of HD shows motor abnormalities
that appear at 8 weeks of age and progress insidiously until they are
manifest overtly by 12 weeks of age (for summary, see Fig. 13).
From this time onwards, the mice deteriorate steadily until their
untimely death at 16–20 weeks of age. The present study shows that
the R6 ⁄ 2 mouse has a complex phenotype with respect to neuromuscular structure and function. Both neural and muscular abnormalities
were detected. These changes were paradoxical. For example, there is
no evidence for paralysis or functional denervation until the latest
stages of the disease, yet R6 ⁄ 2 muscle fibers undergo profound
atrophy and many show membrane characteristics of denervated or
paralysed muscle fibers. Conversely, intact nerve terminals and
terminal Schwann cells show no evidence of constitutive sprouting,
in spite of the atrophy and other denervation-like state of R6 ⁄ 2 muscle
fibers. In addition, synaptic transmission fails completely at increasing
numbers of neuromuscular junctions as the disease progresses.
Together, these data suggest that neuromuscular abnormalities could
contribute deleteriously to motor function in R6 ⁄ 2 mice and may also
help explain their sudden deaths.
(Constantini et al., 1987). In the R6 ⁄ 2 mouse there was no evidence
of reduction in quantal content at functional junctions, even in
severely atrophic muscle. Thus, there is no compelling evidence that
either structural or functional denervation (paralysis) plays any
significant role in the progression of atrophy. This is in fact very
intriguing given that some of the muscle fiber properties resemble
denervation (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 ⁄ 2
muscles should be reduced. Such measurements were not performed in
the present study but would be interesting to examine in future studies,
particularly if combined with experiments on sensitivity to changes in
extracellular ion concentrations. Although changes in gene expression
in skeletal muscle have been reported (Luthi-Carter et al., 2002), it is
not yet know whether NaV1.5 sodium channels are up-regulated in
atrophic R6 ⁄ 2 muscles, nor whether any of the many other proteins
associated with the orchestrated response of muscles to denervation,
such as expression of ACh receptor gamma subunits, are also altered
in these muscles. It would be interesting to apply genetic microarray
analysis of muscle mRNA expression patterns in R6 ⁄ 2 muscles
because this would establish the extent of differences in pattern of
gene expression comparing WT innervated or denervated muscle.
Morphological abnormalities in neuromuscular junctions
Denervation-like characteristics of R6 ⁄ 2 muscles and their role
in muscle atrophy
Remarkably, in spite of their strong, functional motor innervation,
R6 ⁄ 2 muscle fibers resemble denervated muscles in several important
respects (atrophy, reduced resting membrane potential, resistance to
an NaV1.4 sodium-channel blocker, anode-break action potentials).
However, other features of R6 ⁄ 2 muscle (most motor nerve terminals
fully occupying ACh receptor pretzels at motor endplates, robust EPP
responses to nerve stimulation, absence of spontaneous fibrillation)
were at odds with a denervated state. Thus, while the atrophy was
pronounced, the absence of characteristic muscle pathology would
appear to rule out the possibility that the atrophy was due to a
denervation-like pathology. For example, there was no fiber-type
grouping or compensatory hypertrophy of innervated fibers seen
(Pachter & Eberstein, 1992). We also ruled out other neuropathological causes of the atrophy. Neurogenic disorders with skeletal muscle
involvement show characteristic abnormalities such as angulated
atrophic fibers, fiber-type grouping, group atrophy or target fibers.
However, none of these abnormalities were found. There was also no
pathology consistent with a myopathic origin of the atrophy because
there was no myofiber necrosis, myophagocytosis, regeneration,
rounded atrophic fibers, fiber hypertrophy and splitting, centralized
nuclei, fibrosis, ring fibers, vacuoles or any signs of inflammation.
The muscle atrophy is unlikely to be a consequence of malnutrition
because it is evident weeks before the mice show disrupted feeding
patterns. Further, in malnutrition there is typically a relatively
selective atrophy of type II fibers (Anderson, 1985) that was not
seen in the R6 ⁄ 2 muscles. It also seems unlikely that the atrophy is
related to the late-onset diabetes of the R6 ⁄ 2 mice (Hurlbert et al.,
1999). Although electrophysiological changes have been reported in
diabetes, they are not the same as those we see here. For example,
neuromuscular junctions in diabetic rats show greater resistance to
tetanic stimulation with fewer failures of action potential propagation
than in healthy rats (Schiller & Rahamimoff, 1989). Moreover, ACh
release is depressed by up to 45% in chronically diabetic rats
Abnormalities in nerve terminal morphology and physiology, though
present, were surprisingly infrequent given the substantial and
progressive muscle atrophy. This is in marked contrast to, for
example, the morphological and physiological impairment in nerve
terminal structure and function in other neurological mutants which
undergo 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 resemblance
to the widespread retraction of motor nerve terminals that occurs, with
accompanying progressive weakening of synaptic transmission, in
mutant 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. For
example, ultrastructural and fluorescence microscopy of motor
endplates in mice older than 12 weeks showed several unusual
features, including invagination of Schwann cell processes into the
terminal, and abnormal neurofilament organization. Further, in some
muscles, only slender neurofilaments penetrated the motor nerve
terminals; in others, we saw wispy assemblages of untangled
neurofilaments in preterminal axons. There was also evidence at
some junctions for abnormal preterminal axon branching and the
surprising finding of some endplates in older muscles that appeared
devoid of ACh receptors. These changes would be consistent with
aberrant function of the endplates. However, none of these adequately
explain 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 muscle
properties, rather than by progressive structural or functional denervation. At late stages there was concurrent physiological evidence of
increasing numbers of junctions showing progressive failure of
neuromuscular transmission, which could reflect independent neural
abnormalities. Thus, one possible explanation for the morbidity and
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
Abnormal neuromuscular junction function in HD mice 3111
Fig. 13. Summary diagram showing the relationship between timing of appearance of pathological features in the brains and skeletal muscle in R6 ⁄ 2 mice. Data
used 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.
ª 2004 Federation of European Neuroscience Societies, European Journal of Neuroscience, 20, 3092–3114
3112 R. R. Ribchester et al.
mortality of R6 ⁄ 2 mice over the age of 15 weeks is a collapse of
skeletal muscle function, caused by both myogenic atrophy and
neurogenic failure of neuromuscular transmission.
HD is considered to be a disease solely of the brain, despite the
peripheral expression of huntingtin. However, our findings here
suggest that abnormalities of either motor neuron or skeletal muscle
function 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. In
grade 2–4 HD brain, dendritic changes consistent with both regeneration and degeneration (Graveland et al., 1985; Ferrante et al., 1991;
Sotrel et al., 1993) have been reported. However, in two transgenic
models where dendritic changes have been studied (R6 ⁄ 2 and HD
full-length cDNA mice), degenerative changes but not sprouting
responses have been reported (Guidetti et al., 2001; Klapstein et al.,
2001). It would be interesting to see whether the neurons in R6 ⁄ 2
brains show a typical sprouting response to direct injury or whether
they, like the peripheral neurons, show abnormal sprouting responses.
The most parsimonious explanation for our findings is that the
polyQ-huntingtin transgene affects motoneurons, their presynaptic
motor nerve terminals and skeletal muscle fibers independently. In
fact, the peculiar combination of pathology we see in the neuromuscular junction suggests that there is a functional disconnection of the
homeostatic feedback mechanisms that normally regulate neuromuscular function. Thus, skeletal muscle fibers do not appear to respond
normally to the trophic input from their motor nerve terminals, and ⁄ or
the motor neurons do not respond to neurotrophic feedback from the
muscle fibers. It is tempting to speculate that a similar dysregulation of
feedback mechanisms may also be present in the CNS. This may
constitute (or produce) breakdown in the mutual trophic interactions
that normally maintain nerve and muscle. It has been shown
previously that skeletal muscle is a target of polyglutamine-related
perturbations in gene expression (Luthi-Carter et al., 2002). Further
analysis of patterns of gene expression in R6 ⁄ 2 muscle could help
resolve this issue, or at least show to what extent the Huntington’s
mutation causes dysregulation of muscle gene expression. For
example, it would be interesting to determine whether the changes
in l-CTX sensitivity were due to changes of sodium channel subunit
expression or subunit composition. Targeted expression of the R6 ⁄ 2
transgene selectively in muscle or motoneurons, or utilization of
chimeric mice or mice with xenografts of muscle, might also help to
establish whether muscle atrophy is a direct consequence of the
mutation. In the light of published abnormalities in synaptic vesicle
proteins in the brains of R6 ⁄ 2 mice, notably complexin II (Morton &
Edwardson, 2001), it is interesting to speculate that the causes of the
diverse array of neuromuscular abnormalities may be a consequence
of abnormal trafficking, integration or function of other synapsespecific proteins. Such abnormalities might, for example, underpin the
reduced mEPP frequency and abnormally high coefficient of variation
in the amplitude distribution of mEPPs we observed in some R6 ⁄ 2
muscle fibers.
For the time being, the clinical relevance of our studies remains
speculative. While some studies on human skeletal muscle function
have been done, most of those have focused on descending control of
motor function. Interestingly, EMG studies in HD patients have
reported abnormalities in motor unit activity (Harper et al., 1991) and
long latency responses (Noth et al., 1985; Leblhuber et al., 1991;
Berardelli et al., 1999; Siedenberg et al., 1999). Although these
deficits have usually been attributed to defects in cortical processing
(Noth et al., 1985; Abbruzzese & Berardelli, 2003), not all of them can
be 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 carried
out. In the light of the present study, this would clearly be worth doing
in the future.
Conclusions
When taken together our data suggest that, despite the inexorable and
uniform progression of muscle atrophy in R6 ⁄ 2 mice from 8 weeks
of age, most neuromuscular junctions remained competent in their
function. However, the function of increasing numbers of these
junctions deteriorated with age, evidently through a combination of
pre- and postsynaptic impairment. The progression of abnormalities
was not linear; indeed, most abnormalities were not seen consistently
until 15 weeks of age, that is, during the last 1–2 weeks of life. This
suggests that one possible explanation for the morbidity and mortality
of R6 ⁄ 2 mice over the age of 15 weeks is failure of skeletal muscle
transmission subsequent to a collapse both in muscle fiber strength and
in neuromuscular transmission. Irrespective of the cause, it is clear that
there is significant and progressive pathology of muscles and
neuromuscular junctions in the postsymptomatic R6 ⁄ 2 mouse which
seems likely to contribute to several aspects of motor dysfunction. If
the neuromuscular junction deficits we see in the mice are present in
humans then it seems likely that abnormalities in peripheral neuromuscular function would contribute significantly to the progression of
HD, 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 with
confocal 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, wild
type.
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