1Department of Physiology, New York Medical College, Valhalla, New York 10595; 2Howard Hughes Medical Institute, Baylor College of Medicine, Houston, Texas 77030; and 3Department of Pathology, University of Minnesota, Minneapolis, Minnesota 55455
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ABSTRACT |
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Inoue, Takafumi, Xi Lin, Kristi A. Kohlmeier, Harry T. Orr, Huda Y. Zoghbi, and William N. Ross. Calcium Dynamics and Electrophysiological Properties of Cerebellar Purkinje Cells in SCA1 Transgenic Mice. J. Neurophysiol. 85: 1750-1760, 2001. Cerebellar Purkinje cells (PCs) from spinocerebellar ataxia type 1 (SCA1) transgenic mice develop dendritic and somatic atrophy with age. Inositol 1,4,5-trisphosphate receptor type 1 and the sarco/endoplasmic reticulum Ca2+ ATPase pump, which regulate [Ca2+]i, are expressed at lower levels in these cells compared with the levels in cells from wild-type (WT) mice. To examine PCs in SCA1 mice, we used whole-cell patch clamp recording combined with fluorometric [Ca2+]i and [Na+]i measurements in cerebellar slices. PCs in SCA1 mice had Na+ spikes, Ca2+ spikes, climbing fiber (CF) electrical responses, parallel fiber (PF) electrical responses, and metabotropic glutamate receptor (mGluR)-mediated, PF-evoked Ca2+ release from intracellular stores that were qualitatively similar to those recorded from WT mice. Under our experimental conditions, it was easier to evoke the mGluR-mediated secondary [Ca2+]i increase in SCA1 PCs. The membrane resistance of SCA1 PCs was 3.3 times higher than that of WT cells, which correlated with the 1.7 times smaller cell body size. Most SCA1 PCs (but not WT) had a delayed onset (about 50-200 ms) to Na+ spike firing induced by current injection. This delay was increased by hyperpolarizing prepulses and was eliminated by 4-aminopyridine, which suggests that this delay was due to enhancement of the A-like K+ conductance in the SCA1 PCs. In response to CF stimulation, most PCs in mutant and WT mice had rapid, widespread [Ca2+]i changes that recovered in <200 ms. Some SCA1 PCs showed a slow, localized, secondary Ca2+ transient following the initial CF Ca2+ transient, which may reflect release of Ca2+ from intracellular stores. Thus, with these exceptions, the basic physiological properties of mutant PCs are similar to those of WT neurons, even with dramatic alteration of their morphology and downregulation of Ca2+ handling molecules.
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INTRODUCTION |
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Spinocerebellar ataxia type 1 (SCA1) is an autosomal dominant neurological disorder characterized by
ataxia and brain stem dysfunction. Like several other neurodegenerative
diseases, SCA1 is caused by the expansion of a CAG
trinucleotide repeat, which results in an expanded polyglutamine tract
in its gene product, ataxin-1 (Orr et al. 1993).
It is well established that polyglutamine expansion confers
neurotoxicity to the native proteins, which leads to the degeneration
of selective groups of neurons. In SCA1, the consistent
neuropathological finding is loss of Purkinje cells (PCs) in the
cerebellar cortex and loss of neurons in pontine nuclei (Gilman
et al. 1996
). As part of an effort to understand the role of
the expanded CAG polyglutamine tracts in the pathogenesis of SCA1,
lines of transgenic mice that express either a normal human
SCA1 allele with 30 glutamine (30Q; e.g., the A02 line) or
an expanded allele with 82 glutamine (82Q; e.g., the B05 line) have
been established (Burright et al. 1995
; Clark et
al. 1997
). Lines of transgenic animals that expressed a wild
type (WT) human SCA1 allele did not develop the typical SCA1
phenotypes, but lines of transgenic mice that expressed the mutant form
of the SCA1 gene developed ataxia and PC pathology.
Importantly, the eventual development of ataxia is not attributable to
cell death per se but to cellular dysfunction and morphological
alterations that occur long before neuronal death (Clark et al.
1997
). To gain insight into the mechanism underlying cerebellar
dysfunction, we previously adopted a subtractive cloning strategy to
evaluate the alteration of gene expression in the cerebellums of SCA1
mice and found that several key neuronal genes are specifically altered in the PCs of mutant cerebellums prior to any clinicopathologic manifestations (Lin et al. 2000
). It is noteworthy that
some of these genes are important for regulating intracellular
Ca2+ dynamics. At postnatal day 14 (P14), the
inositol 1,4,5-trisphosphate receptor type 1 (IP3R1) and sarco/endoplasmic reticulum
Ca2+ ATPase (SERCA2) are greatly reduced. By
P21-P28, type 1 inositol phosphate 5-phosphatase, excitatory amino
acid transporter type 4 (EAAT4), and transient receptor potential type
3 (TRP3) are dramatically downregulated (Lin et al.
2000
). Therefore it was of interest to study the
electrophysiological and synaptic properties of SCA1 PCs, particularly
those that affect stimulus-evoked
[Ca2+]i changes, since
they might be sensitive to alterations in the expression levels of
these molecules.
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Methods |
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Transgenic mice
Adult (3-7 mo old) heterozygous SCA1 transgenic mice (B05 line)
and WT littermates were used. Transgene configuration and establishment
of the B05 line carrying a mutant SCA1 allele with 82 CAG
repeats have been described in Burright et al. (1995).
Slice preparation and electrophysiology
Sagittal cerebellar slices 300 µm thick were prepared
according to standard procedures (Inoue et al. 1998;
Tsubokawa and Ross 1997
). PCs were visually identified
under differential interference contrast (DIC) optics using a 40×
water immersion objective (numeric aperture 0.80) attached to
an upright microscope (BX50WI, Olympus). A model P-97 puller (Sutter
Instrument, Novato, CA) was used to pull patch electrodes from
fiber-filled capillary tubing with an outside diameter of 1.5 mm (no.
1511-M, Friedrich and Dimmock, Millville, NJ). The resistance of the
patch electrodes was 3-6 M
when they were filled with an
intracellular solution composed of (in mM) 130 K-gluconate, 10 Na-gluconate, 4 NaCl, 2 Mg-ATP, 0.3 Na-GTP, 0.2 bis-fura-2 (Molecular
Probes, Eugene, OR), and 10 HEPES (pH 7.2). Bis-fura-2 was replaced
with 0.3 mM of the lower affinity indicator fura-6F (Molecular Probes)
in experiments measuring parallel fiber (PF) responses. The composition
of the artificial cerebrospinal fluid (ACSF) bathing solution
was (in mM) 124 NaCl, 2.5 KCl, 2 CaCl2, 2 MgCl2, 1.25 NaH2PO4, 26 NaHCO3, and 20 glucose. The bathing solution was
kept at about 32°C and bubbled continuously with a mixture of 95%
O2 and 5% CO2. Recordings were made with an AxoClamp 2A amplifier (Axon Instruments, Foster, CA)
in the current-clamp mode, digitized at 20 KHz, and stored in a
computer. For stimulation of climbing fiber (CF) and PF responses, a
glass pipette (tip diameter of 5-10 µm) filled with standard saline
was used. Square pulses (0.4 ms duration) were applied for focal stimulation.
Soma size measurements
DIC images of PC somata were taken with a video camera (VE1000, DAGE MTI, Michigan, IN) and stored in a computer with a frame grabber (Axon Image Lightning 2000, Axon Instruments). The projected areas of the PC somata were measured as numbers of pixels and converted to area.
Calcium concentration measurements
High-speed fluorescence measurements using a cooled charge
coupled device (CCD) camera were made as described in
Lasser-Ross et al. (1991). Fluorescence was excited and
detected using the 40× objective. The use of an Opti-Quip Model 1600 (Highland Mills, NY) power supply (ripple <0.02%) and a Hamamatsu
L2481-01 75-W Mercury-Xenon arc lamp improved the excitation intensity
stability. Typical intensity fluctuations were <0.3%. Excitation of
bis-fura-2 or fura-6F was at 380 nm selected with a 15-nm-wide
interference filter (Omega Optical, Brattleboro, VT). Fluorescence
changes due to bleaching and baseline drift were corrected by
subtracting an identical measurement without stimulation. Optical
changes are expressed as
F/F (change in
fluorescence from resting levels divided by the resting fluorescence
corrected for autofluorescence). The contribution of background
(autofluorescence) was corrected by making comparable measurements at
locations in the slice away from the filled cell and then subtracting
this measurement from the experimental ones.
Sodium concentration measurements
Dynamic changes in
[Na+]i were measured
using the Na+-sensitive indicator sodium binding
benzofuran isophthalate (SBFI) (Callaway and Ross
1997; Lasser-Ross and Ross 1992
; Minta
and Tsien 1989
). These experiments were made using the same
techniques and apparatus used for measuring
[Ca2+]i changes, with the
exception that the Ca2+ indicator was replaced by
2 mM SBFI (Molecular Probes) and 0.5 mM EGTA in the pipette
solution. Excitation and emission wavelengths for detecting SBFI
fluorescence were the same as for bis-fura-2 and fura-6F.
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RESULTS |
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Shape of PCs, size of somata, and membrane resistance
Most mutant PCs had reduced dendritic arbors that were
clearly revealed in fluorescence images of cells filled with bis-fura-2 (Fig. 1). Their somata differed from
the somata of WT cells in shape, size, and location, as has been
reported (Clark et al. 1997). In WT mice, the cell
bodies of PCs are found in a clear layer between the granule cells and
the molecular layer (PC layer). In SCA1 mice, the somata were more
widely distributed. Many were found in the middle of the molecular
layer (Fig. 2A) while others were located in the PC layer. The cell bodies of the mutant PCs appeared to be smaller than those of control PCs when observed with DIC
optics (Fig. 2A). The projected areas of the somata of mutant and WT PCs were 170 ± 46 µm2
(n = 50) and 286 ± 58 µm2
(n = 33), respectively (mean ± SD,
P < 0.01, t-test). The smaller size of the
cell bodies, together with the reduced dendritic arborization, suggested that the input impedance
(Rin) of SCA1 PCs would be higher than
the impedance of WT cells, even if the specific membrane resistance
(Rm) of the two cell types was the
same. To test this hypothesis, we measured
Rin with small hyperpolarizing current pulses in the soma. Rin was estimated
as the peak voltage response divided by the current. A correction for
the sag current (Ih) (Crepel
and Penit-Soria 1986
) was not applied since this response was
the same in SCA1 and WT PCs (Fig. 2B).
Rin of the mutant PCs was much higher
than that of control PCs (147 ± 75 M
, n = 57, and 46 ± 13 M
, n = 43; P < 0.01). The soma sizes and membrane conductances
(1/Rin) of both types of PCs were
positively correlated (Fig. 2C), suggesting that the high
Rin of the mutant PCs is primarily due
to their smaller somata.
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Active membrane potentials
We next observed the pattern of action potentials evoked by
depolarizing current pulses applied with a patch pipette on the soma.
In normal PCs (n = 36), fast action potentials
(Na+ spikes) were generated when the
depolarization reached a threshold (Fig.
3A, top left). The frequency
of the Na+ spikes increased as depolarizing
current increased. With larger currents, spikes stopped firing and a
plateau potential appeared (Fig. 3A, bottom left). This
pattern of Na+ spikes and plateau potentials
accords well with that described for guinea-pig PCs
(Llinàs and Sugimori 1980). We also observed slower Ca2+ spikes (Llinàs and
Sugimori 1980
) in some SCA1 Purkinje neurons (n = 5/41 cells) and WT Purkinje neurons (n = 9/35 cells)
(data not shown).
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Mutant PCs (n = 53) also showed Na+ spikes and plateau potentials (Fig. 3A, right), but there were clear differences in the active potentials. First, less current was needed to reach Na+ spike threshold in the SCA1 neurons. This is primarily due to the larger Rin of the mutant PCs. Second, the kinetics of the Na+ spikes in SCA1 cells were slower than those in WT cells (Table 1 and Fig. 3B). All the kinetic parameters (width at half height, 10-90% rise time, maximum slope of rising phase, 90-10% fall time, and maximum slope of falling phase) were notably different (Table 1). These differences were significant both for the first Na+ spikes evoked just above threshold (Fig. 3B, top traces) and for Na+ spikes in the middle of the train (Fig. 3B, bottom traces).
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Delay in Na+ spike firing
By using moderate amplitude depolarizing pulses, we observed a delayed onset before Na+ spike initiation in many mutant PCs (43/53) (Fig. 3A, right, arrows). This delay ranged from 20 to 300 ms and was characterized as a plateau-like potential with small humps. Although, when threshold depolarization was used, there was a delay before the start of a train of Na+ spikes in WT PCs (Fig. 3A, left, top two traces), in most cases it was a slow, gradual depolarization without a plateau. We occasionally observed that the delay had the plateau and humps characteristic of SCA1 PCs in WT PCs, but the incidence was very low (2/36 cells).
The delay and plateau before spike firing in the SCA1 PCs resembled
that found in other neurons and in PCs of other species (Hounsgaard and Midtgaard 1988; Klee et al.
1995
). In these cells, this response has been attributed to the
fast activation and inactivation of an A-like K+
conductance. We performed several standard tests to see if a similar
mechanism was responsible for the delay in SCA1 PCs. The delayed onset
was blocked by 4-aminopyridine (4-AP) (3 cells by 50 µM and 4 cells by 2 mM; Fig. 4A). In
contrast, 2 mM 4-AP did not affect the latency of spike firing in WT
PCs (Fig. 4A) but slowed spike kinetics (data not shown), as
has been reported for hippocampal neurons (Storm 1990
).
To test whether this alteration of
IA-like current in mutant PCs was due
to a different level of inactivation of the A-like
K+ channels or to a higher expression level of
the channels, we added a hyperpolarizing prepulse before the
depolarization (Fig. 4B). This protocol should completely
activate all A-like channels before stimulation. In mutant PCs, the
delay to spike firing was enhanced by the prepulse. In a mutant cell
(Fig. 4B, right), the firing delay, which
appeared with moderate depolarization (0.1 nA), diminished with
higher-amplitude stimulation (0.3 nA). The prepulse restored the delay
to firing (n = 3/5). In WT PCs, only 2 of 13 cells showed a delay in spike firing with this prepulse protocol. Thus
the delay in Na+ spike firing in mutant PCs was
not due to a difference in the inactivation state of the A-like
conductance. Most likely, the delay is due to a higher level of the
conductance in mutant PCs.
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[Na+]i increases evoked by Na+ spikes
The difference in Na+ spike properties
in mutant PCs could be a consequence of the different electrotonic
properties of these smaller cells and/or a different spatial
distribution of Na+ channels in the cells. One
way to test these possibilities is to examine the spatial pattern of
Na+ entry caused by Na+
spike firing in the PCs. This pattern can be determined by measuring the spike-evoked [Na+]i
changes in the cell at a time before Na+
significantly diffuses away from the sites of entry into the neuron.
Using the Na+-sensitive dye SBFI, we found that
spikes induced [Na+]i increases in
the axon and in the soma that peaked at the time of the last action
potential (n = 16). In contrast, this stimulation protocol caused little increase in the dendrites of all WT cells tested
(8/8) in and half of the mutant PCs tested (8/16) (Fig. 5, A and B). This
spatial distribution matches that found in guinea pig
(Lasser-Ross and Ross 1992) and rat (Callaway and
Ross 1997
) PCs. In the rest of the mutant cells
(n = 8), an additional spike-associated [Na+]i increase,
peaking at the end of the spike train, was clearly observed at the base
of the primary dendrites (Fig. 5C). No increase was observed
in the more distal arborization. Since the falling phase of the
[Na+]i transient in the
primary dendrites was faster than that in the soma, the increase in
[Na+]i in the primary
dendrites was not due to diffusion of Na+ from
the soma but resulted from direct Na+ influx into
the primary dendrites.
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Parallel fiber synaptic responses
Normal PCs receive more than 100,000 parallel fiber connections on
their elaborate dendritic arbors (Palay and Chan-Palay 1974). The shrunken dendrites of SCA1 PCs could alter the
parallel fiber synaptic connections. To test this possibility, we
placed a stimulating electrode over the dye-filled dendrites in the
molecular layer. Stimulation in this region evoked a smoothly graded
excitatory postsynaptic potential (EPSP) as a function of stimulus
intensity (Fig. 6A). The
response showed paired-pulse facilitation in both mutant
(n = 4) and WT (n = 2) PCs (Fig.
6B). These properties are characteristic of this synapse
(Konnerth et al. 1990
; Perkel et al.
1990
). No significant qualitative difference was noted between
the properties of mutant and WT PF responses. However, we did not
undertake a detailed quantitative analysis of these electrical
parameters.
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Parallel fiber-evoked Ca2+ release in dendrites
Calcium release from intracellular stores mediated by
the activation of IP3 receptors is well known in
many cell types (Berridge 1993). This process has been
observed in PC dendrites following bath application of metabotropic
glutamate receptor agonists (Netzeband et al. 1997
;
Vranesic et al. 1991
), flash photolysis of
caged-IP3 (Khodakhah and Ogden
1993
), and by repetitive parallel fiber stimulation (Finch and Augustine 1998
; Takechi et al.
1998
). Since molecules important for regulating
Ca2+ release, such as IP3R1
and SERCA2, were drastically reduced in SCA1 PCs
(Lin et al. 2000
), we tested whether
Ca2+ release by parallel fiber stimulation could
be observed in SCA1 PCs. Seven of fifteen SCA1 PCs showed slow
[Ca2+]i increases in
addition to a rapid Ca2+ transient following a
short train of parallel fiber stimuli (3-8 pulses at 50 Hz). The
latency of the slow
[Ca2+]i increase was
100-300 ms and the increase was confined to the stimulated dendritic
area (Fig. 7). The time course and
location of this slow
[Ca2+]i increase
coincides well with the characteristics of
IP3-mediated Ca2+ release
reported in normal rats (Finch and Augustine 1998
) and mice (Takechi et al. 1998
). Consistent with this
interpretation, we found that the secondary
[Ca2+]i increase was
blocked by 1 mM (R, S)-
-methyl-4-carboxyphenylglycine (MCPG) (Fig.
7, n = 3). MCPG blocks group I metabotropic glutamate receptors and associated IP3 mobilization
(Conn and Pin 1997
). The secondary increase is unlikely
to be due to Ca2+ entry through voltage-sensitive
Ca2+ channels in the dendrites since there was no
observed voltage change at the recording electrode at the time of the
secondary [Ca2+]i
increase. In addition, the secondary increase persisted when the cell
was hyperpolarized with current from the patch electrode. However, it
is possible that a small part of the increase was due to a localized
potential change in the dendrites that was invisible at the soma. No
Ca2+ release was observed in control WT PCs from
older animals (0 of 7 cells). However, in later experiments, where we
preloaded the Ca2+ stores with long depolarizing
pulses, we observed Ca2+ release in a younger WT
PC (1 of 1 cell) and also in PCs from another WT strain of mice (C57B6,
4 mo old; 3 of 4 cells, data not shown).
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CF synaptic responses
Stimulation of the CF PC synapse evoked a large and complex spike
in an all-or-none fashion, as has been shown previously (Llinàs and Sugimori 1980). This CF-evoked EPSP
elicits a [Na+]i increase
in the dendritic shafts mainly due to Na+ influx
through
-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid
(AMPA)-type glutamate receptors located at the CF-PC synapse (Callaway and Ross 1997
; Lasser-Ross and Ross
1992
). Thus the topological distribution of the
[Na+]i transient by CF
stimulation is a good indication of the distribution of CF-PC synapses
in the PC dendrite. We measured the
[Na+]i changes by CF
stimulation (Fig. 8). In WT PCs,
[Na+]i increased over
thick dendrites (n = 5) as has been observed in other
species (Callaway and Ross 1997
; Lasser-Ross and
Ross 1992
). In SCA1 PCs, the same pattern of
[Na+]i changes was
observed by CF stimulation (n = 7). Thus we conclude that CF innervates the thick portion of dendrites in SCA1 PCs just as
it does in WT PCs, although the dendritic arbor is deformed.
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The CF-evoked EPSP elicited a
[Ca2+]i transient mainly
in the dendrites of both WT and mutant PCs (Fig.
9, A and B), as has been reported (Callaway et al. 1995; Miyakawa et
al. 1992
). Since IP3R1 and the SERCA2
pump are severely underexpressed in the SCA1 PCs (Lin et al.
2000
), we carefully measured the characteristics of these
CF-induced Ca2+ transients. The average time
constant of decay of the
[Ca2+]i transient in SCA1
PCs was 112 ± 56 ms (mean ± SD, n = 11), which was not significantly different from that in WT PCs (140 ± 71 ms, n = 14). The amplitude of the transient depends
on many factors. These include the density of
Ca2+ channels, the surface-to-volume ratio in the
dendrites, and the buffering power of the cytoplasm. Therefore it is
difficult to draw conclusions from a comparison of this parameter in
different cells. Nevertheless, we found no qualitative difference in
the Ca2+ transient amplitude in SCA1 and WT PCs.
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In 4 of 16 SCA1 PCs, a secondary slow Ca2+ transient was observed after the typical initial fast [Ca2+]i change following CF stimulation. This slow transient was found in only part of the dendritic tree, usually on a thick branch (Fig. 9B). The latency of the initial Ca2+ transient was <0.50 ms; the latency of the secondary Ca2+ transient ranged between 180 and 300 ms. The amplitude of the secondary [Ca2+]i increase was larger than that of the initial [Ca2+]i increase at the locations in dendrites where it occurred. There was no detectable change in membrane potential during this secondary Ca2+ transient. This slow transient was never detected in WT PCs (n = 17) or in young SCA1 PCs (5-6 wk old, n = 8). However, we did not follow the protocol of preloading the stores in these experiments. Therefore we cannot rule out the possibility that CF activation in WT PCs can release Ca2+.
Unfortunately, the low incidence of the CF-induced slow Ca2+ transient prevented us from undertaking a detailed examination of the underlying mechanism. However, the close similarity of this [Ca2+]i increase to that observed following PF stimulation suggests that it was due to IP3-mediated release following activation of metabotropic glutamate (mGlu) receptors.
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DISCUSSION |
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In this study we compared the basic electrophysiological and
Ca2+ handling properties of PCs in SCA1 and WT
mice that were 3-7 mo old. Perhaps the most remarkable finding was how
similar the properties were in the normal and mutant PCs in spite of
their morphologic and molecular differences (Burright et
al. 1995; Clark et al. 1997
; Lin
et al. 2000
). After three months of age, SCA1 transgenic mice
develop severe ataxia and their PCs have reduced soma size, loss of
dendritic arborization, and mislocalization in the molecular layer
(heterotopia). Moreover, the expression of a number of PC-specific
genes is dramatically reduced, including two critical
Ca2+-regulating molecules
(IP3R1 and SERCA2). The finding that these two
genes, which have opposing functions in regulating intracellular [Ca2+]i, are both
downregulated was perplexing and raised questions about the
physiological consequences of these molecular changes. Na+ and Ca2+ spike firing,
CF and PF synaptic inputs, paired-pulse facilitation, and the time
course and amplitude of CF-induced
[Ca2+]i changes were
similar in the two cell types. Ca2+ release
following repetitive PF stimulation was also preserved in SCA1 PCs and
occasionally was observed following CF stimulation. These findings
might provide insight about the molecular changes. It is possible that
only one of the two genes (either SERCA2 or IP3R1) is downregulated because of the mutation
and that the downregulation of the second is compensatory. The
similarity of the PF responses is particularly interesting because the
greatly reduced dendritic arbor of the SCA1 PCs probably means that
these cells receive many fewer inputs than the 100,000+ inputs that
have been reported for normal PCs (Palay and Chan-Palay
1974
). It is within the context of this overall similarity that
we discuss the differences between the SCA1 and WT neurons.
Small soma size and high Rin
The input resistance (Rin) of
SCA1 PCs was significantly higher than the
Rin of WT PCs. The most likely reason
for this increase is the smaller somata and reduced dendritic
arborization of the SCA1 cells. If the specific membrane resistance
(Rm) of both cell types were the same,
this correlation would result. We tried to quantitate this relationship
by measuring the cross-sectional area of the soma. This area should be
proportional to the surface area of the cell body if the cell bodies
have similar shapeswhich they appear to have. We found that the
cross-sectional area and the input conductance
(1/Rin) were strongly correlated (Fig.
2). However, Rin depends on current
flowing through the dendritic membrane as well as the somatic membrane,
although the somatic component should dominate (Rapp et al.
1994
). Since we could not measure the dendritic contribution to
either the surface area or Rin, we
could not be more precise about the constancy of
Rm. A similar value for
Rm in SCA1 and WT PCs suggests that
none of the molecular consequences of the polyglutamine expansion have an important impact on the molecules that determine
Rm.
One consequence of the higher Rin in SCA1 PCs is that the synaptic currents activated by single PFs will cause a greater potential change in the PC. Therefore activation of fewer PFs will be required to reach the threshold for spike firing. Since the dendrites are much smaller in these cells, they probably receive fewer inputs than do normal PCs. The need for fewer active inputs to reach threshold appears to be a homeostatic mechanism intended to preserve some aspects of PC function in SCA1 mice.
Delay in Na+ spike firing and other parameters affecting spike kinetics
The delay in Na+ spike firing in SCA1 PCs
was a surprise since none of the known molecular or morphological
changes pointed toward this alteration. The most likely explanation for
this delay is an enhancement of an A-like K+
conductance. The rapid activation and inactivation of this conductance (Connor and Stevens 1971) are known to cause similar
delays in other cells (Storm 1990
). Support for this
conclusion comes from experiments in which the delay was eliminated by
4-AP, a known blocker of this conductance (see Hoffman et al.
1997
). In SCA1 PCs, the delay in spike firing was restored or
enhanced by a hyperpolarizing prepulse (Fig. 4B), which
suggests that the A-like conductance is partially inactivated at rest
in mutant PCs. In contrast, most WT PCs did not show a delay in spike
firing even with a hyperpolarizing prepulse, indicating that a fully
active A-like conductance is not enough to generate a delay in spike
firing in normal PCs. This shows that the enhanced A-like conductance
in SCA1 PCs is not merely due to a difference in the inactivation state
of the conductance but that it may result from increased expression of IA-like channels or from altered expression
of accessory modulatory proteins for voltage-gated
K+ channels (An et al. 2000
).
In addition to the delay in spike firing, other aspects of
Na+ spike kinetics were altered in SCA1 PCs.
These include spike width and rate of rise (Table 1). We have no clear
explanation for these significant differences. Some of the variation
may be due to the differences in cell morphology since the need to
charge the cell membrane will affect the rise time and fall time of the spike independent of the properties of the channels. The spatial distribution of the channels also can affect spike kinetics. In several
experiments, we measured the spatial distribution of the spike-evoked
[Na+]i changes. This
distribution should reflect the distribution of active
Na+ channels. We found that the
[Na+]i increase in half
of the tested SCA1 PCs and in all of the WT PCs was confined to the
soma and axon. This result is consistent with previous measurements in
rats and guinea pigs (Callaway and Ross 1997;
Lasser-Ross and Ross 1992
) and is also consistent with experiments that found that Na+ propagation into
PC dendrites is completely passive (Stuart and Hausser
1994
). The results from the other half of the tested SCA1 PCs
are interesting. In these PCs we found clear spike-evoked [Na+]i increases in the
proximal dendrites. We did not find dendritic [Na+]i increases in any
WT cell. This result suggests that the spatial distribution of
Na+ channels in SCA1 PCs may not be as tightly
regulated as in WT cells.
It is not clear whether the differences in the kinetics of the Na+ spikes in SCA1 and WT PCs have any significant physiological consequences. Propagation in the axon should be insensitive to small variations in spike parameters. Differences in spike width could affect the release of neurotransmitter. However, we have no information about spike parameters in the terminal arborization.
Biphasic [Ca2+]i transients evoked by parallel fiber stimulation
We observed an initial rapid Ca2+ transient
and a localized slow
[Ca2+]i increase
following a brief train of PF stimulation in SCA1 PCs. The slow
[Ca2+]i increase was
blocked by MCPG, a blocker of Group I metabotropic glutamate receptors.
A similar biphasic pattern of
[Ca2+]i increase by PF
stimulation has been reported in rat PCs (Finch and Augustine
1998) and murine PCs (Takechi et al. 1998
). In
those reports the secondary Ca2+ transient was
attributed to Ca2+ release from
IP3 receptor-operated intracellular stores
because the [Ca2+]i
increase was blocked by intracellular heparin. The unreliability in
generating the secondary response prevented us from doing similar experiments. Nevertheless, the similarity in the stimulation pattern (50-60 Hz, 5-15 pulses), the delay following stimulation (0.1-0.3 s), the localized pattern of
[Ca2+]i increase, and the
sensitivity to MCPG make it likely that the second slow phase in our
experiments also was due to Ca2+ release from
intracellular Ca2+ stores. We observed this
Ca2+ release in half of the SCA1 PCs (7/15
cells). This incidence is lower than the 100% reported by
Takechi et al. (1998)
. The discrepancy may arise from
differences in experimental procedure. In particular, we did not
preload the stores with voltage pulses before PF stimulation, a
procedure followed by Finch and Augustine (1998)
. In
support of this possibility we did not observe
Ca2+ release by PF stimuli in seven WT PCs in
early experiments. But we did observe release in later experiments on
WT neurons when we preloaded the PCs (n = 3). However,
the main conclusion is that PF-mediated Ca2+
release was observed in SCA1 mice, indicating that mGlu receptors, IP3 receptors, and other molecules responsible
for controlling release are present and functional in these mutant animals.
[Ca2+]i transients evoked by CF stimulation
We observed two kinds of Ca2+ transients in
response to CF stimuli in SCA1 PCs (Fig. 9). One was a fast transient
that peaked at about the time of the CF electrical response and decayed
rapidly. This transient was widely distributed in the PC dendrites and closely resembled the CF-evoked Ca2+ transients
in other cells (Callaway et al. 1995; Miyakawa et al. 1992
). In other cell types, the removal rate of
Ca2+ following transient elevation has been
attributed to a combination of cytoplasmic buffering and pumps (plasma
membrane and endoplasmic reticulum) (Airaksinen et al.
1997
; Markram et al. 1995
; Neher and
Augustine 1992
). The similarity in removal rates in SCA1 and WT
PCs implies that these mechanisms are equally effective in both types
of cells. Since SERCA2 is downregulated in SCA1 PCs, our results imply
that uptake by SERCA2 is not the rate-limiting step for restoring the
resting [Ca2+]i level
after CF excitation. It is noteworthy that the expression of another
calcium pump, plasmalemmal Ca2+-ATPase (PMCA2),
which is abundantly expressed in cerebellar PCs, is not reduced in SCA1
PCs (Lin et al. 2000
).
The second kind of Ca2+ transient was a slower
and larger signal that was restricted to parts of the dendritic tree,
suggesting Ca2+ release. The localization to
regions near the thick dendrites is consistent with the location of CF
synapses on the cell (Larramendi and Victor 1967).
Because of the low incidence of this response, we were not able to
undertake a detailed pharmacological analysis of the underlying
mechanism of this secondary
[Ca2+]i increase. This
phenomenon was not seen in WT PCs or in young (5-6 wk old) mutant PCs
in this study and it has never been reported in other species following
CF stimulation (Callaway et al. 1995
).
One argument in favor of a release mechanism is the localized spatial
distribution of the secondary response. The primary response, which has
been shown to be caused by the generation of a
Ca2+ spike in the dendrites (Callaway et
al. 1995; Llinàs and Sugimori 1980
), is
widely distributed in the dendrites. This distribution is consistent
with the generation and propagation of this spike over the dendrites.
In contrast, a spatially restricted response is consistent with the
localized generation of IP3 following mGluR activation, as also observed following repetitive PF stimulation (see
Parallel fiber-evoked Ca2+ release in
dendrites). In addition, the secondary
[Ca2+]i increase had a
slow time course and there was no apparent membrane potential change
underlying the Ca2+ transient, as was also
observed in the PF response. Because of this similarity, it is likely
that the secondary CF-evoked Ca2+ transients
observed in this study were also caused by release from calcium stores.
However, further experiments are needed to definitively establish this conclusion.
In summary, the electrophysiological responses and synaptically activated [Ca2+]i changes in SCA1 PCs are similar to those in WT PCs even though the expression of some important Ca2+-regulating molecules is different in the two cell types. The most significant exceptions are the secondary [Ca2+]i increase observed following some CF responses, the higher A-like K+ conductance in SCA1 PCs, and the higher Rin of SCA1 PCs. The differences we observed may be adapted to compensate for the atrophied dendrites of the SCA1 PCs. Alternatively, the heterotopic PCs and the atrophic molecular layer suggest that the organization of the PF inputs to the SCA1 PCs are altered. These observations suggest that the mechanisms underlying ataxia in SCA1 transgenic mice may reside in changes to cerebellar neuronal circuitry rather than in alterations of the intrinsic electrophysiological properties of the PCs.
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ACKNOWLEDGMENTS |
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We thank Dr. Akinori Kuruma for help with some of the calcium release experiments.
This research was supported by National Science Foundation Grant IBN-9514266, National Institute of Neurological Disorders and Stroke Grants NS-27699 and NS-16295, and by the Howard Hughes Medical Institute. T. Inoue was supported by a Postdoctoral Fellowship for Research Abroad from the Japan Society for the Promotion of Science (JSPS). K. A. Kohlmeier was supported by National Institute of Mental Health Grant MH-18825.
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FOOTNOTES |
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Address for reprint requests: W. N. Ross (E-mail: ross{at}nymc.edu).
Received 14 July 2000; accepted in final form 21 December 2000.
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REFERENCES |
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