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The Journal of Neuroscience, December 1, 2001, 21(23):9112-9123
Changes in Cortical and Striatal Neurons Predict Behavioral and
Electrophysiological Abnormalities in a Transgenic Murine Model of
Huntington's Disease
Genevieve A.
Laforet1,
Ellen
Sapp3,
Kathryn
Chase2,
Charmian
McIntyre3,
Frederick M.
Boyce3,
Mary
Campbell2,
Beth A.
Cadigan2,
Lori
Warzecki2,
Danilo A.
Tagle4,
P. Hemachandra
Reddy4,
Carlos
Cepeda5,
Christopher R.
Calvert5,
Eve S.
Jokel5,
Gloria J.
Klapstein5,
Marjorie A.
Ariano6,
Michael S.
Levine5,
Marian
DiFiglia3, and
Neil
Aronin2
Departments of 1 Psychiatry and 2 Medicine,
University of Massachusetts Medical School, Worcester, Massachusetts
01655, 3 Department of Neurology, Massachusetts General
Hospital, Boston, Massachusetts 02114, 4 Genetics and
Molecular Biology Branch, National Genome Research Institute, National
Institutes of Health, Bethesda, Maryland 20892, 5 Mental
Retardation Research Center, University of California at Los Angeles,
Los Angeles, California 90095, and 6 Department of
Neuroscience, Chicago Medical School, North Chicago, Illinois 60064
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ABSTRACT |
Neurons in Huntington's disease exhibit selective morphological
and subcellular alterations in the striatum and cortex. The link
between these neuronal changes and behavioral abnormalities is unclear.
We investigated relationships between essential neuronal changes that
predict motor impairment and possible involvement of the
corticostriatal pathway in developing behavioral phenotypes. We
therefore generated heterozygote mice expressing the N-terminal one-third of huntingtin with normal (CT18) or expanded (HD46, HD100)
glutamine repeats. The HD mice exhibited motor deficits between 3 and
10 months. The age of onset depended on an expanded polyglutamine
length; phenotype severity correlated with increasing age. Neuronal
changes in the striatum (nuclear inclusions) preceded the onset of
phenotype, whereas cortical changes, especially the accumulation of
huntingtin in the nucleus and cytoplasm and the appearance of
dysmorphic dendrites, predicted the onset and severity of behavioral
deficits. Striatal neurons in the HD mice displayed altered responses
to cortical stimulation and to activation by the excitotoxic agent
NMDA. Application of NMDA increased intracellular Ca2+ levels in HD100 neurons compared with wild-type
neurons. Results suggest that motor deficits in Huntington's disease
arise from cumulative morphological and physiological changes in
neurons that impair corticostriatal circuitry.
Key words:
cortex; Huntington's disease; NMDA; neuronal morphology; striatum; transgenic mice
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INTRODUCTION |
Huntington's disease is a fatal
autosomal dominant neurodegenerative disease characterized by movement
abnormalities, psychiatric disturbances, and dementia (Huntington,
1872 ). The gene mutation in Huntington's disease comprises a
lengthening of a series of CAG repeats beyond 35 in a gene encoding a
protein called huntingtin (Huntington's Disease Collaborative Research
Group, 1993). Although mutant huntingtin has a widespread distribution
in neuronal and non-neuronal tissues, neurons in the striatum and
cortex layers 3, 5, and 6 demonstrate characteristic cellular and
subcellular pathological changes, including formation of nuclear
inclusions and dystrophic neurites containing N-terminal mutant
huntingtin (Vonsattel et al., 1985; DiFiglia et al., 1997; Vonsattel
and DiFiglia, 1998).
In murine models of Huntington's disease the cellular features differ
among transgenic models and from morphological changes observed in the
Huntington's disease patient brain. Mice expressing very truncated
transgenes develop widespread nuclear inclusions (Davies et al., 1997;
Schilling et al., 1999), far more than exist in the HD patient brain
(DiFiglia et al., 1997). Mice with full-length mutant huntingtin have
fewer inclusions than in the human condition (Reddy et al., 1998;
Hodgson et al., 1999). Striatal cell loss, a hallmark of change in the
patient brain, is not striking in most Huntington's disease mouse
models, except in some older mice with full-length mutant huntingtin
that are hypokinetic or akinetic (Reddy et al., 1998, 1999). Still,
current murine models of Huntington's disease manifest behavioral
changes. Thus, neuronal dysfunction as implicated by abnormal neuronal
morphology, and not cell death, might underlie movement abnormalities
in Huntington's disease.
To address these issues, we generated transgenic mice with 18, 46, and
100 CAG repeats in a human huntingtin cDNA that encoded the first
one-third of the huntingtin protein. The CAG repeat lengths are
consistent with the normal human (18), adult form of Huntington's
disease (46), and the juvenile form (100). This size of huntingtin
protein causes the same cellular pathology as full-length mutant
huntingtin in transfected clonal medium spiny neurons (Kim et al.,
1999; Kegel et al., 2000). The morphologic and subcellular
abnormalities in vitro mimic those in striatum and cortex in
Huntington's disease. Thus, we have evidence that this portion of
huntingtin with an expanded polyglutamine repeat is sufficient to
reproduce morphological changes of Huntington's disease.
On the basis of morphological, behavioral, and electrophysiological
evidence from our transgenic murine model of Huntington's disease,
changes in cortical neurons, including nuclear and cytoplasmic accumulation of huntingtin and the appearance of dysmorphic dendrites, were correlated the most closely with the onset of motor
deficits. Striatal neurons in transgenic mice had marked alterations in electrophysiological responses to corpus callosum stimulation (corticostriatal pathway) and in the activation of NMDA receptors. Thus, we propose that the effects of mutant huntingtin on cortical neurons play a critical role in striatal neuronal dysfunction in
Huntington's disease and in its neurologic impairment.
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MATERIALS AND METHODS |
Transgene design
The initial 3 kb of the human huntingtin-encoding cDNA sequence
IT15 (Huntington's Disease Collaborative Research Group, 1993) were
cloned into transfer plasmid AcMNPV downstream from the rat neuron-specific enolase promoter and its first intron (Fig.
1A). An
eight-amino-acid FLAG epitope tag was included just 5' to the IT15
sequence to permit immunologic detection of the transgene. Three
different CAG repeat lengths, 18, 46, and 100, were inserted into the
sequence. The construct included an SV40 polyadenylation signal. The
full 3 kb length of the IT15 cDNA used in the transgene was
sequenced.
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Figure 1.
Design and expression analysis of transgenes CT18,
HD46, and HD100. A, Map of construct used for transgene
vector. Bases 316-3210 of the human huntingtin cDNA sequence
(IT15), modified to contain 18, 46, or 100 CAG repeats,
were cloned into AcMNPV transfer plasmid NSE4-BV. A rat neuron-specific
enolase promoter regulates expression. An N-terminal epitope tag, FLAG,
was included to permit differential identification of the
transgene-encoded huntingtin product. Northern blot and RT-PCR analyses
confirm RNA expression from CT18, HD46, and HD100 transgenes.
B, The Northern blot demonstrates transgene expression
in different HD46 and HD100 lineages (denoted above the
lanes). B, Left and
Top right, Total RNA (10 µg) per lane.
B, Bottom right, Total RNA (30 µg) per lane. An arrowhead ( ) marks
the bands corresponding to transgene-encoded RNA. Wild-type control
lanes show no transgene-derived message.
C, RT-PCR results from CT18, HD46, and HD100 lineages. Lane M, A
100 base pair ladder; lane C, no RNA control. An
arrowhead () marks the position of the expected
RT-PCR product. Expression is seen in all transgenic lineages but is
absent from the transgene-negative (WT) control.
D, Western blot analysis of huntingtin
(top) and FLAG (bottom) immunoreactivity
in WT, CT18, HD46, and HD100 mice. D,
Top, A product of the expected size (~120 kDa,
black arrowhead) is present in the transgenic mice, but
not in WT. Native huntingtin is denoted with a white
arrow. The signal intensity for the ~120 kDa product is
stronger in the HD mice compared with CT. Huntingtin is detected with
antiserum Ab1. D, Bottom, A product of
the expected size (~120 kDa, black arrow) is present
in the transgenic mice, but not in WT.
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Generation and genotyping of transgenic mice
Transgene DNA for microinjection was treated with
PacI, purified, and used to microinject oocyte pronuclei
from C57BL/6×SJL F1 hybrid mice. Transgenic mice
were generated and maintained in the Transgenic Core Facility at the
University of Massachusetts Medical School. Genomic DNA from
F0 mice was extracted from tail biopsies and
characterized initially by Southern blot analysis, using a cDNA IT15
probe. After transgene-positive founders were identified, lineages were
screened by Southern blot or PCR analysis. Estimation of transgene
insertion site and copy number followed the procedure of Reddy et al.
(1998).
Detecting expression of the huntingtin transgene
Evidence for transcription of the transgene was provided by
analysis of total RNA in Northern blots and RT-PCR. Western blot analysis was performed as reported previously (Aronin et al., 1995).
Behavioral analysis
All transgenic animals were maintained in microisolator cages
with free access to food and water under pathogen-free conditions in a
12 hr light/dark cycle in the Animal Medicine Facility at the
University of Massachusetts Medical School. After weaning, the males
were housed singly, and the female littermates were housed grouped.
Care and handling of experimental animals complied fully with
institutional, Society for Neuroscience, and federal guidelines
for humane treatment. Behavioral analyses were performed in the
mornings, circadian time CT 1-6 hr.
Animals were evaluated neurologically with four tests: rotarod
performance, clasping, activity level, and gait. Evaluation of
clasping, activity level, and gait were subjective. Two observers tested 474 mice so that the testing procedures would be performed consistently. We cannot state with certainty that the observers did not
know or suspect the HD genotype, because WT mice rarely displayed
behavioral changes under 12 months of age. We used only robust changes
in clasping, activity, and gait to minimize observer bias. Observations
were made at the same examination to minimize age as a variable.
Rotarod performance. This analysis used a rotarod apparatus
(Economex Rotarod, Columbus Instruments, Columbus, OH) to assess the
ability of the animals to stay on a rotating rod through successive 2 min trials at increasing speeds. The mice were pretrained on the
rotarod for five trials at low speed (214 cm/min) for up to 120 sec.
Animals then were tested for one trial each at medium (277 cm/min) and
high speed (340 cm/min). The rotarod score (0-120 sec) equaled the
mean number of seconds the animal stayed on the rotarod at low speed
averaged with the mean of the two higher speed trials. For inclusion in
the hazard analysis, rotarod scores of <60 sec were considered
abnormal. Rotarod scores had a bimodal distribution, with clusters of
HD mice below 60 sec and with control and wild-type mice above 60 sec.
To validate our rotarod protocol, we tested 181 different mice (wild
type, CT18, HD46, and HD100) in the rotarod protocol of Clark et al.
(1997) and found no differences in the rotarod results when
compared with our method.
Clasping. Mice were held suspended by the tail for up to 1 min at a height of 15 cm. Clasping was defined as grabbing forepaws alone or forepaws and hindlimbs simultaneously. A presence of clasp
required paws to be pressed against the chest with the spine in a
lordotic position. Clasp was scored as its presence or absence.
Activity level. Hyperactivity was defined as circling or
rapid to-and-fro movement, also known as stereotypic behavior.
Hypoactivity was defined as paucity of spontaneous movement. The mice
were not perturbed to elicit activity. If an animal did not meander voluntarily in the cage, the observer picked up the cage, which was
sufficient to elicit meandering in most mice. Activity was scored as
the presence or absence of hypoactivity or hyperactivity.
Gait. Abnormal gaits included wide-based gait, walking with
an arched posture, or slow gait with tremulousness. Gait was scored as
the presence or absence of an abnormal gait. For emphasis, only robust
changes in clasping, activity, or gait would be detected with the
scoring of these behaviors.
Morphological analysis
Immunohistochemistry antibodies. Anti-huntingtin
rabbit antisera were made with the N terminus of huntingtin (Ab1) and
an internal epitope of huntingtin (Ab585; amino acids 585-725). Both antisera were characterized as reported previously (DiFiglia et al.,
1995).
Immunohistochemical procedure. After perfusion with 4%
paraformaldehyde in PBS, the brains of the mice were removed,
post-fixed in paraformaldehyde, and sectioned at 40 µm. A series of
treatments with primary antiserum, biotinylated anti-rabbit antiserum,
and avidin-biotin horseradish peroxidase complex followed. The
chromogen was diaminobenzidine. Details were described by DiFiglia et
al. (1995). Controls included omission of primary antisera and
competitive incubation with 10 µg of huntingtin peptide.
Light microscopy. Examination of brain sections was
performed without previous knowledge of the genotype or phenotype of mice.
Cell counts and cross-sectional area measurements. The cell
counts were obtained in seven wild-type controls (mean age, 9.4 months,
range 3-17 months), three CT18 mice (all 18 months), and seven HD100
mice (mean age, 8.7 months, range 3-17 months), using a stereological
method described by Hyman et al. (1998). Briefly, 40 µm sections were
cut through the entire striatum and stained with cresyl violet; every
sixth section was assessed, totaling 10-12 sections per striatum. At
1000× magnification, all neurons in 100 × 100 µm fields were
counted in every sixth field throughout the section. In all, 50-60
fields were counted for each brain from which an average and SD were
determined. For measurement of cross-sectional area, digital computer
images of 25 striatal neurons per mouse were acquired through the
microscope with a 100× objective lens and Sigma Scan software (Jandel
Scientific, San Rafael, CA). The mean values from five controls (three
wild type and two CT18) and six HD mice (five HD100 and one HD46) were analyzed with a two-tailed Student's t test. The ages of
the controls (11 months ± 2.4) and the HD mice (8.1 months ± 4.8) were not significantly different.
Assessment of dendrite morphology in biocytin-filled neurons.
To provide a set of unbiased estimates, three observers, who were
blinded to the genotype from which the neurons were obtained, counted
J-dendrites and wavy dendrites in all recovered cells on a set of
qualitative scales (for J-dendrites: 0 = dendrites extend
normally, 1 = one or two dendrites begin to turn, 2 = one or
two dendrites make close to a complete turn, and 3 = three or more
dendrites make close to a complete turn; for wavy dendrites: 0 = dendrites extend normally, 1 = one or two have moderately wavy
dendrites, 2 = two or three have obviously wavy dendrites, 3 = three or more dendrites have obvious waviness, 4 = there are many wavy dendrites, some severe, and 5 = almost all dendrites are
severely wavy). There was a high counting reliability for both measures.
Electrophysiology
Slice preparation. Preparation of tissue was similar
to procedures described previously (Cepeda et al., 1998; Levine et al., 1999). Twenty-six mice (17 HD100 and 9 WT) were used for
electrophysiology. The mice were decapitated. After dissection
the brains were placed in ice-cold oxygenated artificial CSF (aCSF)
[containing (in mM) 130 NaCl, 26 NaHCO3, 3 KCl, 5 MgCl2,
1.25 NaH2PO4, 1 CaCl2, and 10 glucose, pH 7.2-7.4]. Coronal
brain sections containing the striatum were cut (350 µm) and placed
in oxygenated aCSF, but with 2 mM each
CaCl2 and MgCl2. After 1 hr
the slice was transferred to a recording chamber.
Current clamp. In the recording chamber the slices were
perfused constantly with oxygenated aCSF [containing (in
mM) 124 NaCl, 2 MgSO4, 2 CaCl2, 5 KCl, 1.25 NaH2PO4, 26 NaHCO3, and 10 glucose at 31-32°C, with 50 µM picrotoxin to block the responses mediated by
activation of GABAA receptors] in an atmosphere
of warm, moist 95% O2/5%
CO2. Responses of individual cells were recorded
by sharp microelectrodes filled with 3 M
K+-acetate, 5 mM KCl, and 2%
w/v biocytin (Sigma, St. Louis, MO) to label dendritic and axonal
processes. After the experiment the slice was fixed in 4%
paraformaldehyde overnight and processed according to published
protocols (Kita and Armstrong, 1991). Resting membrane potentials were
recorded only after the cell had recovered from penetration and had
stabilized (at least 15 min after impalement). The current-voltage
relationship was determined from the responses to incrementally
increasing square wave hyperpolarizing and depolarizing current pulses
delivered via the recording electrode. To examine synaptic responses,
we placed a bipolar stimulating electrode in the corpus callosum
to activate the corticostriatal fibers preferentially. Stimuli (100 µsec duration, variable intensity from 100 µA to 5 mA) were
delivered every 5 sec, and input-output relationships were determined.
Responses were averages of five traces.
Whole-cell voltage clamp. Neurons were visualized in the
slice with a fixed-stage upright microscope (Olympus, Model BX50WI) and
submerged in continuously flowing oxygenated aCSF (25°C, 4 ml/min).
Cells were visualized with a 40× water immersion lens that used
differential interference contrast optics and were illuminated with
near-infrared light (790 nm; Ealing Optics, Holliston, MA). The image
was detected with an infrared-sensitive CCD camera. Patch electrodes
(3-6 M ) were filled with the following internal solution (in
mM): 130 Cs-methanesulfonate, 10 CsCl, 4 NaCl, 1 MgCl2, 5 Mg-ATP, 5 EGTA, 10 HEPES, 0.5 GTP, 10 phosphocreatine, and 0.1 leupeptin, pH 7.25-7.3, with an osmolality of 280-290 mOsm. An
Axopatch 200A (Axon Instruments, Foster City, CA) was used for
recording. A 3 M KCl agar bridge was inserted between the
extracellular solution and the Ag-AgCl indifferent electrode. Tight
seals (2-5 G) from visualized medium-sized cells were obtained by
applying negative pressure. Recordings were not corrected for junction
potentials, which ranged from 2-3 mV. Leak subtraction was performed
on-line with the Axopatch 200B amplifier. The membrane was disrupted
with additional suction to obtain the whole-cell configuration. Access
resistances ranged from 10 to 25 M and were compensated 60-85%.
Cells were held at  70 mV to minimize contributions of voltage-gated
currents. Voltage-gated currents also were blocked with the following
solution: 1 µM tetrodotoxin (TTX), 100-200
µM Cd2+, 3 mM
Cs, 20 mM tetraethylammonium (TEA), and 2 mM
4-aminopyridine (4-AP). NMDA was bath applied to the striatal slice.
Non-NMDA receptor activation was blocked with
6-cyano-7-nitroquinoxaline-2, 3-dione (CNQX; 5 µM). Cell
capacitances and input resistances were measured by applying a 10 mV
depolarizing voltage command and by using the membrane test function
integrated in the pClamp8 software (Axon Instruments).
Statistical analysis
For the behavioral analysis, logistical regression (Draper and
Smith, 1998) was used when behaviors were ranked as the presence or
absence of an abnormality (clasping, activity, gait). Linear regression
(Draper and Smith, 1998) was used for rotarod tests. In analyses in
which the number of abnormalities in each mouse was examined, we
converted the rotarod to the presence or absence of abnormality.
Rotarod tests were distributed in a bimodal pattern. The range spanned
48-78 sec between the two modes. We selected 60 sec as the cutoff for
a normal rotarod test. A hazard score was used to compare the rates at
which subsets of mice developed behavioral abnormalities. Hazard score
is a rate that assesses the short-term conditional probability of
developing an abnormality. The rate over a particular time interval is
calculated by using the number of mice in subsets (WT and CT18 vs HD46
and HD100) becoming abnormal over that interval, conditional on the
number of animals that remained healthy at the beginning of the
interval (Collett, 1994). To determine possible correlations between
behavioral deficits and neuropathological findings, we used the
CORR procedure (SAS statistical analysis software; SAS
Institute, Cary, NC) to generate Pearson correlation coefficients. For
analysis of electrophysiological changes, Welch's approximation to the
t test for unequal variances (Welch, 1947) and ANOVA with
Bonferroni correction were used.
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RESULTS |
Verification of mice with huntingtin transgenes
Southern blot analysis confirmed insertion of mutant (HD) and
wild-type (CT) huntingtin transgenes into the mouse genome (transgene map shown in Fig. 1A). Multiple lineages with 18, 46, and 100 CAG repeat lengths were identified. The lineages had a range of copy numbers, estimated from 1-52 (CT18: 1, 5; HD46: 11, 15, 23, 52;
HD100: 3, 5, 6, 10, 11, 12), and different insertion sites (data not
shown). As detected by Northern blot analysis (Fig. 1B) and RT-PCR (Fig. 1C), RNA expression
demonstrated transgene transcription. Western blot analysis that used
anti-huntingtin and anti-FLAG antisera confirmed the presence of a
protein of the expected size for the transgene product (~120 kDa)
(Fig. 1D). Protein expression in Western blot was
less than that of native huntingtin and variable in the HD mice.
Immunohistochemistry with anti-FLAG and anti-huntingtin antibodies
detected nuclear inclusions and other changes in the HD mice,
consistent with the expression of the mutant transgene (see below).
Mutant huntingtin transgenes promote neurological impairments
We studied 474 mice, distributed as wild type (n = 110) and transgenics with 18 CAG (CT18, n = 30), 46 CAG
(HD46, n = 146), and 100 CAG (HD100, n = 188). Animals were tested from all lineages and from generations up
to F6. Neither lineage nor copy number influenced
the neurological outcomes. Age had a separate but weak effect on
neurological outcomes (p < 0.05).
Neurological outcomes were scored as normal versus abnormal for
clasping, gait, and activity; significance was tested by logistic regression. The performance in staying on the rotarod was scored in
seconds, and linear regression was used for statistical analysis. There
was no significant difference between wild-type mice and CT18 mice for
any of the neurological tests. Compared with wild type or CT18, HD46
showed differences in clasping (p < 0.001), gait (p < 0.05), activity
(p < 0.001), and rotarod
(p < 0.001). HD100 mice also demonstrated a
difference in each neurological test (p < 0.001 for each).
The frequency of the number of neurological abnormalities also was
evaluated in a  2 distribution,
regardless of age (Table 1).
Approximately 80% of the wild-type and CT18 mice had no neurological
impairment, whereas 60% of the HD46 mice and 72% of the HD100 mice
had at least one abnormality. Furthermore, the HD46 and HD100 were more likely to have more than one abnormal neurological test.
To determine the rate at which neurological impairments presented, we
generated a hazard score. A hazard score is the measurement of change
in behavior as a function of time. In this formulation the test outcome
was either normal or abnormal. Rotarod scores of <60 sec were
considered abnormal (derivation in Materials and Methods). Figure
2 shows that for both wild-type and CT18
mice there is little likelihood of developing a behavioral abnormality for the first 12 months of life, but the rate of developing
neurological deficits increases after 12 months. However, the HD46 and
the HD100 transgenic mice demonstrate an increase in the rate of
developing an abnormality by 3 months. The rate increases through the
lifespan of the population of mice.
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Figure 2.
Hazard score for WT/CT18 versus HD46/HD100. Hazard
scores calculate the rate of development of neurological deficits over
time. Mice with normal polyglutamine length (WT, CT18  ) were
compared with transgenics with expanded polyglutamines (HD46, HD100
 ). WT and CT18 have a near-zero hazard rate for the first 12 months;
the rate rises thereafter, suggesting late development of severe
symptoms. Conversely, the hazard rate for HD46 and HD100 begins to
increase in the first month and continues to climb, indicating that
phenotypic expression of HD transgenes with expanded polyglutamines
begins early and increases with age.
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Neuropathological changes in the HD mice: accumulation of
huntingtin, dysmorphic dendrites, vacuoles, membrane blebs, and
atrophy
We evaluated the localization of huntingtin in HD, CT18, and WT
mice. Huntingtin staining detected with Ab1 and Ab585 antisera was
increased markedly in the somatodendritic cytoplasm of cortical and
striatal neurons of HD46 (n = 9) and HD100
(n = 26) compared with CT18 (n = 6) and
WT (n = 4) mice (Fig.
3a). Cortical pyramidal neurons in the HD mice had dysmorphic dendrites, which were
characterized by marked retraction and disorientation of the apical
dendrite (Fig. 3b-d). Cytoplasmic huntingtin accumulation
and dysmorphic changes were seen in the cortex of all HD lineages, were
especially prevalent in frontal and cingulate cortices, and occurred
variably and infrequently in piriform and hippocampal cortices.
Intracytoplasmic vacuoles and plasma membrane blebs appeared in cell
bodies and dendrites of some of the HD neurons that had a cytoplasmic
accumulation of huntingtin (Fig. 3e). Dysmorphic dendrites
were not present in hippocampal pyramidal neurons of five of eight mice
examined, were sparse in two mice, and extensive in a severely affected mouse. This severely affected mouse also showed dysmorphic dendrites in
some cerebellar Purkinje cells and shrunken neurons in hippocampus and
cerebellum. Except for the most severely affected HD mouse, other brain
regions known to be relatively uninvolved at early stages of
Huntington's disease had no or mild neuropathological changes when
compared with cortex and striatum. Thus, dysmorphic dendrites were not
evident in other brain regions of most HD mice, despite increased
somatodendritic cytoplasmic labeling for huntingtin in the neurons
there.
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Figure 3.
Increased huntingtin immunoreactivity and
dysmorphic dendrites in HD mice. a, WT mouse (8.5 months, rotarod, normal behavior) exhibits low to moderate levels of
huntingtin labeling in the cytoplasm and proximal dendrites of
pyramidal neurons. b-e, HD mice have intense labeling
for huntingtin in the cytoplasm and proximal dendrites
(b, HD100, L63, 4.5 months, RR = 10, +clasp;
c, HD46, L17, 12 months, RR = 66, +clasp;
d, HD100, L63, 7 months, RR = 51; e,
HD100, L46, 12 months, RR = 26). Some neurons also show diffuse
nuclear staining (d). Dysmorphic dendrites show
marked retraction (arrows in c, d) and
disorientation (arrowhead in d). Membrane
blebbing occurs in cell bodies and dendrites (arrow in
e). Staining was performed with anti-huntingtin antibody
Ab585. Scale bars: a, b, 25 µm;
c-e, 10 µm.
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Huntingtin labeling in neuronal nuclei was widespread in layers 2/3, 5, and 6 of the cortex and throughout the striatum of HD46 and HD100 mice,
but not of CT18 and WT mice (Figs.
4a, 5a,b). The
proportion of neurons with diffuse nuclear labeling was significantly greater (Student's t test, p < 0.0001) in
the striatum (mean, 80% of neurons) than in the cortex (mean, 43% of
neurons; see Fig. 6A, top). Intranuclear
inclusions were detected in most HD46 and HD100 mice (Figs.
4b, 5d-f) with anti-huntingtin antisera Ab1 and Ab585 as well as with anti-ubiquitin and anti-FLAG antisera in
some of the older, more severely affected mice (Fig.
5e,f). Ab1 robustly
detects inclusions in cortical and striatal neurons of Huntington's
disease brain and in other transgenic mouse models (Davies et al.,
1997; Levine et al., 1999). Ab585 detects inclusions in the cortex of
some Huntington's disease brains (Sapp et al., 1999) and in clonal
striatal cells after overexpression of truncated or full-length mutant
huntingtin (Kim et al., 1999). In parallel with Huntington's disease
juvenile and adult-onset patients (Sapp et al., 1997; Gutekunst et al.,
1999), neurons with inclusions were more frequent in the cortex and
striatum of HD100 mice (mean, 23%) than in the cortex and striatum of
HD46 mice (mean, 7%). Inclusions were significantly more frequent in
the cortex (mean, 19% of neurons) than in the striatum of HD mice
(mean, 6% of neurons; Student's t test, p < 0.01; Fig. 6A,
bottom), a characteristic also noted in adult-onset
Huntington's disease brain (Sapp et al., 1999). Neurons with
inclusions also had prominent, diffuse staining for huntingtin in the
nucleus (20-30% of neurons with inclusions) and cytoplasm (98% of
neurons with inclusions).
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Figure 4.
Cortical neurons with huntingtin immunoreactivity
in HD mice. a, HD46 mouse (L14, 14 months, RR = 14, +clasp, hypoactive, broad-based gait) has diffuse nuclear huntingtin
labeling in layer 5 pyramidal neurons. Some neurons have marked
blebbing of the plasma membrane (arrows).
b, Nuclear inclusions in pyramidal cells of HD100 mouse
(L59, 7 months, RR = 15, +clasp). One of the neurons
(top) also has diffuse nuclear labeling. c,
d, Layer 6 from a transgenic control mouse with 18 glutamines
in huntingtin (c, L3, 7 months, normal behavior) and an
HD100 mouse (d, L59, 7 months, RR = 15, +clasp).
Compared with the neurons in c, the neurons in
d are reduced in size and show increased huntingtin
immunoreactivity in nuclei. cc, Corpus callosum.
Staining for huntingtin was performed with antibody Ab585. Scale bars:
a-d, 25 µm.
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Figure 5.
Striatum of HD mice. a,
Presymptomatic HD100 mouse (L46, 5 months, normal behavior) shows
diffuse nuclear labeling in medium-sized striatal neurons, which was
not seen in wild-type or HD18 mice. b, c, Severely
affected HD46 mouse (L14, 14 months, RR = 14, +clasp, hypoactive,
broad-based gait) shows marked shrinkage of striatal neurons in
b (compare with a). Atrophy of the
striatum shown in c is comparable with grade 2 brain in
Huntington's disease. d, Nuclear inclusion
(arrow) and marked perinuclear accumulation of
huntingtin appear in large striatal neuron from HD100 mouse (L46, 5 months, RR = 96, +clasp). e, f, Nuclear inclusions
(arrows) detected with anti-ubiquitin antibody in
e and with anti-FLAG antibody in f in
medium-sized striatal neurons of affected HD100 mouse (L63, 11 months,
+clasp, hypoactive, broad-based gait). Immunostaining for huntingtin in
a, b, and d was performed
with Ab585. Scale bars: a, b, 25 µm; c,
1 mm; d, 10 µm; e, f, 25 µm.
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Figure 6.
Quantitative assessment of dysmorphic
dendrites, diffuse nuclear labeling, nuclear inclusions, and cell
density in relation to phenotype. A, Diffuse nuclear
labeling and neuronal intranuclear inclusions in cortex and striatum.
Mice are identified by HD type (46 or 100) and age (months) on the
abscissa. A, Top, Neurons with diffuse
nuclear labeling for huntingtin in the cortex and striatum of HD46 and
HD100. RR score is shown at the top of the bars, from
highest to lowest. There are significantly more neurons with diffuse
nuclear labeling in the striatum than in the cortex
(p < 0.01). Impaired rotarod performance is
correlated positively with the extent of diffuse nuclear labeling in
the cortex (p < 0.03), but not in the
striatum. A, Bottom, Intranuclear inclusions in the
cortex and striatum in the same set of HD mice. Inclusions are
significantly more frequent in the cortex than in the striatum
(p < 0.01) and are correlated with better
rotarod scores (p < 0.004).
B, Extent of dysmorphic morphology in huntingtin-labeled
cortical neurons of control and HD46 and HD100 mice
(J-dendrites, wavy dendrites) versus rotarod performance. A rating
scale (0, +1, +2) for dysmorphic dendrites was used (see Materials and
Methods), with +2 assigned to brains with the most severely dysmorphic
neurons. The asterisk indicates an animal with severe
dendritic pathology (+2) that was unable to perform on the rotarod
because it developed seizures when it was tested. There is a
significant correlation between higher degrees of dysmorphic changes
and lower RR scores (p < 0.0001).
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Both medium-sized and large striatal neurons had inclusions. Large
neurons, which account for only 1-2% of all striatal neurons and
generally are spared in Huntington's disease, had a much higher frequency of inclusions than medium-sized cells. In an analysis of five
HD100 mice (ages 3-11 months) with normal and abnormal behaviors,
27-70% of the large striatal neurons that labeled for huntingtin had
intranuclear inclusions (mean, 47%). Electron microscopic analysis
revealed that the nuclear inclusions formed in the cortex and striatum
of HD mice were composed of granules and filaments (results not shown)
and appeared to be similar to those seen in Huntington's disease brain
(DiFiglia et al., 1997).
Huntingtin-immunoreactive neurons appeared smaller or reduced in number
in the striatum (Fig. 5b) and in layer 6 cortex (Fig. 4c,d) in the majority of the HD mice. Measurements of
cross-sectional area showed that huntingtin-immunoreactive striatal
neurons in the HD mice were significantly smaller than in wild-type and
HD18 mice (control, 151 ± 14.9, SE; HD, 71.3 ± 4.10;
p < 0.0003). One notable HD46 mouse with a 31%
reduction in neuronal density showed marked atrophy of the striatum
characteristic of a grade 2 HD brain (Fig. 5c). Nonetheless,
stereological analysis in striatum revealed no consistent difference in
HD100 compared with WT or CT18 (HD100, n = 7; CT18,
n = 3; WT, n = 7; HD100 = 94% of
WT and 91% of CT18 neurons per striatum; NS).
In HD mice in which changes in the cortex and striatum had been
observed, we examined the presence of nuclear changes in huntingtin labeling in cerebellum, hippocampus, substantia nigra, and brainstem. None of these other regions is known to be affected in the early stages
of Huntington's disease but can be involved at advanced stages of the
disease (Vonsattel and DiFiglia, 1998). In 11 of 14 HD mice that were
studied, nuclear inclusions appeared in a few cells in one or more of
the regions. Diffuse nuclear labeling was not evident in neurons in any
of these regions, except for one mouse, which had the most severe motor
impairment. In this mouse (shown in Fig. 5b,c) diffuse
nuclear labeling occurred in ~25% of cerebellar Purkinje cells and
hippocampal pyramidal neurons. Thus, except for the most severely
affected mice, nuclear changes in huntingtin immunoreactivity were not
prominent in other brain regions compared with the cortex and striatum.
We compared the occurrence of different neuropathological changes in
the cortex and striatum with behavioral phenotype. The higher the
dysmorphic dendrite score (Fig. 6B; see Materials and Methods), the worse the rotarod score (p < 0.0001) and the greater the number of neurological deficits
(p < 0.0001). In cortex the prevalence of
diffuse nuclear labeling (Fig. 6A, top)
correlated with worse rotarod scores (Pearson correlation coefficient,
r = 0.74; p < 0.03) and a higher
number of neurological deficits (r = 0.79;
p < 0.002). In contrast, diffuse nuclear labeling in striatal neurons did not predict neurological impairments
(r = 0.44 for rotarod and r = 0.41 for
number of deficits; both NS). The frequency of nuclear inclusions (Fig.
6A, bottom) was not associated with worse
rotarod scores in either brain region, but in cortex it was correlated
with better rotarod performance (r = 0.74;
p < 0.004). Shrinkage and loss of neurons in cortical layer 6 also did not appear to correlate with rotarod performance, although these were not quantified.
Thus we found that the mutant huntingtin transgene led to multiple
neuropathological changes in neurons in the striatum and cortex. In
addition to the striatum, the cortex was a site of early and
progressive neuropathology in the HD mice. Two findings, the extent of
dendritic abnormalities revealed by extensive cytoplasmic labeling and
diffuse nuclear localization of huntingtin in cortical neurons, were
the most predictive of the degree of neurological impairment. In
contrast, the formation of huntingtin-positive nuclear inclusions had
no predictive value.
Impaired responsiveness to corticostriatal stimulation correlates
with alterations in dendrite morphology
Striatal neurons from WT (n = 13) and HD100 mice
(n = 12) showed similar intracellular responses to
current injections (Fig. 7A),
with no significant differences in average resting membrane potential
(RMP) or action potential amplitudes. Mean input resistance in the HD
neurons was slightly lower than in WT neurons (28.3 ± 4.5 vs
40.9 ± 4.5 M in HD100 and WT, respectively). Stimulation of
subcortical white matter corpus callosum, which was in the slice
preparation, evoked slightly smaller synaptic responses (EPSPs) in
striatal neurons of HD mice compared with neurons of WT mice [mean
amplitude and half-amplitude duration: for HD mice, 7.3 ± 1.3 mV
and 7.9 ± 0.4 msec; for WT mice, 7.8 ± 1.6 mV and 8.8 ± 0.9 msec (Fig. 7B)]. Of more importance, there was a
statistically significant rightward shift in the input-output curve,
indicating that more current was required to produce responses in
neurons from HD100 mice (p < 0.05; Fig.
7C).
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Figure 7.
Electrophysiological characteristics of striatal
neurons in HD transgenic mice. A, Responses to
intracellular square wave hyperpolarizing and depolarizing current
pulses in WT and HD100 striatal neurons. Amplitudes and durations of
pulses are shown in the bottom traces (for clarity, the
voltage responses induced by the highest intensity pulses are not shown
for WT). B, EPSPs evoked by increasing intensities of
stimulation of WT and HD100 striatal neurons. C,
Input-output relationships for synaptic responses evoked in WT and
HD100 striatal neurons. Each point represents the mean
intensity (± SE) to evoke synaptic responses that were 20, 50, and
80% of the maximum amplitude. D, Responses to NMDA.
Peak currents (left), current densities
(middle), and the percentage of changes in intracellular
Ca2+ (right) in WT
(n = 6 neurons, 3 WT mice) and HD100 mice
(n = 14 neurons, 4 HD mice) after stimulation of
striatal neurons with 50 µM NMDA. Values are means
and ± SE. Asterisk denotes p < 0.05 for peak current and current density and p < 0.025 for Ca2+, using Welch's approximation to
the t test (Welch, 1947). E, Current
density-voltage relationships in response to a ramp voltage command
before and after the bath application of 50 µM NMDA in
HD100 (n = 5) and WT (n = 5)
striatal neurons. Current density-voltage relationships were
calculated as the response to the agonist minus a baseline response in
the absence of the agonist. NMDA current densities were significantly
larger in transgenic than in control animals from 50 to +10 mV
(p < 0.05). F, Time course
of changes in mean NMDA-induced current densities (± SE). HD100 group
was divided into least- and most-affected neurons (see Results for
details and justifications). Asterisks in
F and G indicate that the most-affected
HD100 group was significantly different from its respective WT and
least-affected HD100 group (p < 0.05). The
horizontal line indicates the application of NMDA in
F and G. G, Time course of
mean percentage of changes (± SE) in Ca2+
concentration in most- and least-affected HD100 and WT neurons.
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Striatal neurons were labeled by intracellular injection of
biocytin (Fig. 8A,B).
The labeled cells were medium sized and displayed relatively similar
spine densities in HD and WT mice. However, neurons from HD100 mice had
significantly more dendrites with endings that curved back toward the
soma (J-dendrites) and/or had sharp bends (wavy dendrites) compared
with WT mice (Fig. 8B,C). Mean J-dendrites and wavy
scores in HD mice were 1.39 ± 0.27 and 3.32 ± 0.23, respectively, and in WT were 0.62 ± 0.13 (p < 0.05) and 2.23 ± 0.17 (p < 0.001), respectively. In the HD mice,
scores of J-dendrites and wavy dendrites were correlated significantly with each other (HD: r = 0.589, p < 0.05, n = 12; WT: r = 0.03, n = 13). These scores also were correlated with the
amount of current that was necessary to induce 20% of the maximum
synaptic responses (J-dendrite score vs current: r = 0.87, p < 0.05, n = 5; wavy dendrite
score vs current: r = 0.93, p < 0.05, n = 5). These results indicated that altered morphology
and abnormal synaptic physiology covaried in the HD neurons. The number
of J-dendrites, but not wavy dendrites, also correlated with the
current that was necessary to produce synaptic responses in WT mice,
suggesting that the former property might covary with synaptic input in
general.
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Figure 8.
Examples of biocytin-filled medium-sized spiny
striatal neurons (A-C) and cortical neurons
(D-G) in WT (A, D, F) and
HD100 mice (B, C, E, G). B, HD neuron has
J-dendrites with curved endings (filled
arrowheads) and wavy dendrites with sharp bends (open
arrows). Scale bar in B also applies to
A. C, High magnifications of wavy
dendrites with sharp bends (arrowheads at
top and bottom left) and J-, or recurved,
ending (arrowhead in bottom right).
D, Cortical WT neuron has smooth dendrites and extension
of the apical dendrite to the pial surface (arrow).
E, A cortical pyramidal neuron from an HD mouse with a
disoriented apical dendrite (arrow). F,
Higher magnification of a cortical pyramidal neuron in the WT mouse
shown in D. Note the smooth apical dendrite
(arrow). G, Higher magnification of HD
neuron shown in E (rotated to fit better in the
rectangular frame). Note the small sharp bends (arrow
points to apical dendrite). Scale bar in E also applies
to D.
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Cortical neurons in HD and WT mice were filled with
biocytin to examine dendrite morphology. Labeled WT neurons
(n = 3) had the typical appearance of pyramidal cells,
with an extensive basal plexus and a smooth apical dendrite that
extended toward the pia (Fig. 8D,F). In
contrast, cortical neurons in HD100 mice displayed dendritic
abnormalities (four of eight recovered cells; Fig.
8E,G), including beading, small sharp bends
(n = 4; Fig. 8G), and misaligned and
markedly bifurcated apical dendrites (Fig. 8E,
arrow). These results were consistent with the dysmorphic
changes seen in the cortex of HD mice with huntingtin labeling.
Response to NMDA receptor activation is altered in striatal neurons
of HD100 mice
Excitotoxicity may contribute to neuronal death in HD and other
neurodegenerative disorders and may involve alterations in NMDA-type
glutamate receptors and a rise in intracellular
Ca2+ (Coyle and Schwarcz, 1976; McGeer and
McGeer, 1976; Choi, 1988; DiFiglia, 1990; Beal et al., 1991; Albin and
Greenamyre, 1992; Brown et al., 1997). We examined electrophysiological
properties of NMDA receptors in neurons from striatal slices of WT and
HD100 mice. Peak membrane currents, regardless of time, induced by the activation of NMDA receptors were evaluated with whole-cell
voltage-clamp recording (see Materials and Methods). Peak membrane
currents induced by NMDA (50 µM, 3 min duration) were
significantly larger in HD100 mice (n = 14 neurons)
compared with wild-type controls (n = 6 neurons;
p < 0.05; Fig. 7D). To normalize for cell
size, we determined NMDA current density (peak current divided by cell capacitance; Alzheimer et al., 1993). HD mice displayed significantly increased NMDA peak current density compared with WT
(p < 0.05; Fig. 7D).
In a subgroup of neurons from HD100 (n = 5) and
wild-type controls (n = 5) we evaluated possible
changes in voltage dependence. NMDA currents were examined with a ramp
protocol [voltage command from 70 to +30 mV (5 sec) and then back to
90 mV (1 sec); all data were collected from the downward ramp]
before and after bath application of NMDA. Current density-voltage
relationships were calculated as the response to the agonist minus a
baseline response in the absence of the agonist. We found that NMDA
current densities were significantly larger in transgenic than in
control animals from 50 to +10 mV (p < 0.05;
Fig. 7E). Both groups displayed peak current densities
between 30 and 20 mV; no obvious changes in voltage dependence were apparent.
Finally, we evaluated intracellular Ca2+
flux induced by exposure to NMDA, using fura-2 as a
Ca2+ indicator. Basal
Ca2+ levels were not statistically
different between HD100 and WT. However, with treatment with NMDA,
Ca2+ influx was significantly greater in
HD100 mice than in WT mice (p < 0.025; Fig.
7D). In HD100 mice two populations of cells emerged: one
population with peak NMDA currents and densities very similar to those
obtained from WT animals and a second population of neurons with much
higher peak currents and current densities. Seven of 14 neurons in the
HD100 mice accounted for the marked increase in NMDA peak currents and
current density by displaying changes that were three- to sixfold
greater than the other neurons, which had values similar to WT. Because
there appeared to be two populations of neurons in HD100 mice, we
divided these neurons into two groups (most-affected and least-affected
neurons) to analyze the time course of the effects of the NMDA
application. Current density was increased significantly only in the
most-affected neurons from 3 to 7 min after NMDA application
(p < 0.05; Fig. 7F). Although there was considerable cell-to-cell variation, the most-affected neurons displayed significantly larger increases in intracellular Ca2+ from 4 to 5 min after NMDA
application (p < 0.05; Fig. 7G).
Thus the presence of electrophysiological abnormalities in response to
NMDA was marked but variable in the HD100 mice, thereby defining a
subpopulation of neurons more affected by the transgene. Moreover, neurons with altered response to NMDA occurred in an HD100 mouse with
no deficits and in two HD100 mice with motor impairments (one with
hyperactivity and one with hyperactivity and clasp). This finding
suggests that altered function of NMDA receptors in striatal neurons
may precede the onset of motor deficits. Unfortunately, the morphology
of striatal neurons with altered NMDA receptor function could not be
evaluated with biocytin because of the presence of the
Ca2+ indicator fura-2. Because NMDA was
applied in the bath and the changes were examined over an 8 min period
during and after application of the agonist, it is conceivable that the
observed effects were attributable to a combination of NMDA receptor
activation as well as changes in receptor desensitization,
inactivation, or rundown.
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DISCUSSION |
We sought to identify critical neuropathological and functional
neuronal changes that explain the onset and progression of the
Huntington's disease phenotype. We used heterozygote transgenic mice
that express a large N-terminal fragment of the human huntingtin gene
with normal or expanded CAG repeats. Transfection of this size of
mutant human huntingtin in clonal striatal cells recapitulates in
vitro many of the subcellular neuronal changes in Huntington's disease brain (Kim et al., 1999; Sapp et al., 1999; Kegel et al., 2000). In transgenic HD mice we found that nuclear inclusion
formation alone is insufficient to cause the aberrant phenotype;
instead, abnormal dendritic morphology and diffuse nuclear labeling in cortex are aligned with behavioral changes. Furthermore, the size of
huntingtin expressed in our transgenic HD model (1000 amino acids) with
expanded polyglutamines suffices to cause the neuropathological changes
found in Huntington's disease. We propose that cortical involvement in
our HD transgenic mice, as corroborated by neuropathological and
electrophysiological findings, contributes to the development of
phenotype in the HD mice. We speculate that cortical changes in the
human are fundamental to the onset and progression of phenotype in
Huntington's disease.
Abnormal neuronal morphology and diffuse nuclear huntingtin in the
cortex predict the presence of clinical disease in our transgenic HD
mice. In contrast, nuclear accumulation of huntingtin in the striatum
is comparable in HD mice regardless of the severity of motor deficits.
This characteristic of striatal neuropathology can occur early and
might contribute to the clinical course, but it is insufficient to
predict the onset of behavioral deficits. Striatal neurons from HD mice
displayed abnormal electrophysiological responses to cortical
stimulation and to NMDA receptor activation. These abnormal responses
were separate from the extent of neurological impairment. Thus, even
before the onset of outward phenotype, mutant huntingtin could affect
cortical inputs to the striatum and thereby alter function in
corticostriatal pathways.
Although Huntington's disease often has been considered primarily a
process of striatal neurodegeneration, there have been indications that
the cortex contributes to the disease diathesis and that mutant
huntingtin affects corticostriatal circuitry early in the disease.
Marked nuclear and cytoplasmic accumulation of mutant huntingtin in
cortical pyramidal neurons coincides with dendritic abnormalities in
postmortem brain of patients with low-grade striatal pathology
(DiFiglia et al., 1997; Sapp et al., 1997). Furthermore, abundant
(mutant) huntingtin is evident in numerous cortical dystrophic axons
that project to the striatum (Sapp et al., 1999). In Golgi
impregnations the cortical neurons as well as the striatal neurons show
abundant dendritic changes in Huntington's disease postmortem brain
(Graveland et al., 1985; Sotrel et al., 1993). Clinical findings also
implicate cortical involvement. Metabolic dysfunction in cortex can
occur early in the course of the disease (Sax et al., 1996), and
cognitive disturbances can appear before motor deficits (Lawrence et
al., 1996). In a "knock-in" model of Huntington's disease in which
an expanded N-terminal region of mutant huntingtin was inserted into
murine huntingtin, behavioral changes were not observed despite the
presence of nuclear inclusions in the striatum (Wheeler et al., 2000). The mice had no neuronal changes in the cortex. In contrast, Lin et al.
(2001) generated a knock-in mouse with 150 CAG repeats that exhibited
predominantly striatal huntingtin nuclear inclusions with a modicum of
cortical involvement. These animals developed late-onset behavioral
changes. In all, these findings support the centrality of the cortex in
the genesis of the Huntington's disease phenotype.
Use of the neuron-specific enolase promoter directed transgene
expression to neurons. Endogenous huntingtin was detected in neurons
and eluded detection by immunohistochemistry in glia. Similarly,
huntingtin mRNA was found in high abundance by in situ hybridization (Landwehrmeyer et al., 1995) in neurons but in very low
amounts in glia. A primary dysfunction of glia in Huntington's disease
has been proposed on the basis of neuropathologic grade-dependent changes in the astrocyte-specific GLT1 glutamate transporter (Arzberger et al., 1997). Reactive astrocytosis is a feature of striatal pathology
in Huntington's disease and a criterion for the grading system of
striatal pathology (Vonsattel et al., 1985). How the expression of
mutant huntingtin in glia might contribute to astroglial changes in
Huntington's disease brain is unclear.
Our results in transgenic HD mice support in vitro data
(Saudou et al., 1998; Kim et al., 1999) that nuclear inclusions do not
play an early or decisive role in cell death. In a conditional mouse
model of HD involving highly expanded CAG repeats in exon 1 of
huntingtin, Yamamoto et al. (2000) showed that the formation of nuclear
inclusions in the striatum was reversible with the transcriptional
repression of the transgene. Therefore, the initial accumulation of
mutant huntingtin in inclusions did not trigger inexorable cell death.
Our findings indicate that cellular dysfunction in Huntington's
disease might require an accumulation of mutant huntingtin over a long
duration, thus also extending therapeutic opportunities.
Electrophysiological data obtained in slice preparations from HD100
mice demonstrated that striatal neurons had an altered response to
cortical stimulation and NMDA receptor activation but a normal average
resting membrane potential and action potential amplitudes. These
results indicate a fundamental defect in NMDA receptor function. We
found that striatal neurons in R6/2 mice with huntingtin exon 1 transgene (Mangiarini et al., 1996) also displayed altered responses to
NMDA receptor activation (Cepeda et al., 2000) and to cortical
stimulation (Levine et al., 1999; Klapstein et al., 2000). However, the
presence of depolarized resting membrane potentials in these cells
makes it difficult to interpret the significance of changes in NMDA
receptor responsiveness.
The marked changes in dendritic architecture (J- and wavy dendrites)
observed in cortical and striatal neurons of the HD100 mice could
contribute to the reduced synaptic responsiveness of striatal neurons
to cortical stimuli. In cortical neurons the alterations in dendritic
geometry and the distribution of ion channels significantly influence
neuronal firing patterns, especially those dependent on glutamatergic
neurotransmission (Henze et al., 1996; Mainen and Sejnowski, 1996).
Excitotoxicity as a mechanism for the pathogenesis of Huntington's
disease has been posited previously (Coyle and Schwarcz, 1976;
DiFiglia, 1990; Albin and Greenamyre, 1992). In vivo
treatment of rat striatum with quinolinic acid, an NMDA receptor
agonist, created a pattern of cell loss reminiscent of Huntington's
disease (Beal et al., 1991). We found that NMDA receptor activation
significantly elevated peak current densities and levels of
intracellular Ca2+ in striatal neurons of
HD100 mice compared with wild-type mice. The latter effect could lower
the threshold for excitotoxic injury. In addition, cell swelling, an
early event in the excitotoxic cascade (Choi, 1988), is seen in some
neurons from HD100 mice after the application of NMDA in
vitro (M. Levine, unpublished observations). Similar findings have
been observed in striatal cells from transgenic mice with huntingtin
exon 1 (R6/2) and a knock-in HD mouse with no behavioral phenotype
(Levine et al., 1999).
In other experimental systems the expression of mutant huntingtin has
been shown to alter electrophysiological properties mediated by NMDA
receptor activation and to render cells more susceptible to
excitotoxicity-related cellular changes. Coexpression of NMDA receptor
subtypes NR1 and NR2B with mutant huntingtin into human embryonic
kidney cells produced larger currents after NMDA receptor
activation than did coexpression of these receptors with wild-type
huntingtin (Chen et al., 1999) and increased excitotoxic cell death
(Zeron et al., 2001). Current studies indicate that NMDA receptor
activation is excitotoxic in our HD100 and HD46 mice (A. Petersen, K. Chase, Z. Puschban, M. DiFiglia, P. Brundin, and N. Aronin,
unpublished data). Unexpectedly and inexplicably, striatal neurons in
the R6/2 mice are resistant to the neurotoxic effects of quinolinic
acid (Hansson et al., 1999).
Caveats understandably attend the use of transgenic models. Different
promoters, transgene size, CAG repeat length, and genetic background
might influence characteristics of particular models. Our murine model
used a large portion of mutant N-terminal huntingtin; exon 1 and
full-length huntingtin have provided inadequate neuropathology to
pursue our goals. Most huntingtin transgenics use promoters that are
foreign to mice [portions of human (Mangiarini et al., 1996);
CMV-based (Reddy et al., 1998); human YAC (Hodgson et al., 1999);
prion-based (Schilling et al., 1999)]. We used a neuron-specific promoter that by mRNA estimations regulated mutant huntingtin gene
expression differently from endogenous huntingtin gene expression but
provided morphologic changes in neurons that are characteristic of
Huntington's disease. Finally, several mouse strains have been used in
generating HD models. Ours has a mixed background of two strains. With
nearly 500 mice tested in multiple lineages, we suggest that the mutant
huntingtin, rather than clustered strain differences, accounts for our
observed motor impairments. Furthermore, preliminary studies in our
laboratory indicate that transgenic mice with full-length huntingtin
HD100 in C57BL/6 develop behavioral abnormalities at the same age as
HD100 transgenic mice with N-terminal one-third of the protein in
C57BL/6×SJL.
Our findings point to a multi-tiered process of dysfunction in cortical
and striatal neurons; this process produces the HD mouse phenotype.
Accumulation of mutant huntingtin in nuclear and cytoplasmic
compartments occurs contemporaneously with the onset of behavioral
abnormalities as detected in our transgenic HD mice. Dendritic
pathology in the cortex, electrophysiological abnormalities in the
striatum, and disruption of corticostriatal circuitry were detected in
mice at early and later times. Accumulated mutant huntingtin might
maintain the neuron in a state of chronic vulnerability. Cortical and
striatal neurons respond to the persistent presence of mutant
huntingtin in a highly reproducible pattern by forming J- and wavy
dendrites. How mutant huntingtin causes J- and wavy dendrites is
unknown, but effects of the mutant protein on the cytoskeleton and
vesicle trafficking could be implicated in altering dendritic
architecture (Aronin et al., 1999). Striatal neurons in our HD mice
concurrently become especially sensitive to NMDA receptor activation
and Ca2+ flux into the neuron increases.
Collectively, these quantifiable characteristics of the HD mouse
phenotype should be valuable in identifying therapeutic approaches that
slow or reverse neuronal dysfunction.
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FOOTNOTES |
Received March 30, 2001; revised Sept. 12, 2001; accepted Sept. 12, 2001.
This work was supported by National Institutes of Health (NIH) Grant NS
38194 (N.A.) and Grants NS 16367 and NS 35711 (M.D.), by Department of
Defense Grant USAMRMC 98292059 (N.A.), by a University of Massachusetts
Medical School (UMMS) institutional award (N.A.), by the Huntington's
Disease Society of America (M.D., G.J.K.), and by the Hereditary
Disease Foundation (N.A., M.D., D.A.T., M.A.A., M.S.L). G.A.L. is a
fellow of the Howard Hughes Medical Institute. We thank Dr. Steven
Jones, Dr. Janet Stein, and the staff in the UMMS transgenic core for
their able advice and assistance. We thank Steven Baker in the UMMS
statistics core and Robert Lew for their advice on statistical analysis
and Yun Kim for help with cell counts. We also thank the NIH Diabetes
and Endocrine Research Center at UMMS (Grant DK32520) for its support
of the Peptide Core and the Transgenic Core.
Correspondence should be addressed to Dr. Neil Aronin, Department of
Medicine, University of Massachusetts Medical School, 55 Lake Avenue
North, Worcester, MA 01655. E-mail: Neil.Aronin{at}umassmed.edu.
| |
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ABSTRACT
INTRODUCTION
Gene
