Department of Anatomy and Neurobiology, Medical College of Ohio, Toledo, Ohio 43614-5804
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ABSTRACT |
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McNerney, Mary Ellen,
Desiree Pardi,
Phyllis C. Pugh,
Qiang Nai, and
Joseph F. Margiotta.
Expression and Channel Properties of -Bungarotoxin-Sensitive
Acetylcholine Receptors on Chick Ciliary and Choroid Neurons.
J. Neurophysiol. 84: 1314-1329, 2000.
Cell-specific expression of nicotinic acetylcholine receptors (AChRs)
was examined using ciliary and choroid neurons isolated from chick
ciliary ganglia. At embryonic days 13 and 14 (E13,14) the neurons can
be distinguished by size, with ciliary neuron soma diameters exceeding
those of choroid neurons by about twofold. Both neuronal populations
are known to express two major AChR types:
3*-AChRs recognized by
mAb35, that contain
3,
5,
4, and occasionally
2 subunits,
and
-bungarotoxin (
Bgt)-AChRs recognized and blocked by
Bgt,
that contain
7 subunits. We found that maximal whole cell current
densities (I/Cm) mediated
by
Bgt-AChRs were threefold larger for choroid compared with ciliary
neurons, while
3*-AChR current densities were similar in the two
populations. Different densities of total cell-surface
Bgt-AChRs
could not explain the distinct
Bgt-AChR response densities
associated with ciliary and choroid neurons. Ciliary ganglion neurons
display abundant [125I]-
Bgt binding
(
106 sites/neuron), but digital fluorescence
measurements revealed equivalent site densities on both populations.
AChR channel classes having single-channel conductances of
30, 40, 60, and 80 pS were present in patches excised from both ciliary and
choroid neurons. Treating the neurons with
Bgt selectively abolished
the 60- and 80-pS events, identifying them as arising from
Bgt-AChRs. Kinetic measurements revealed brief open and long closed
durations for
Bgt-AChR channel currents, predicting a very low
probability of being open (po) when
compared with 30- or 40-pS
3*-AChR channels. None of the channel
parameters associated with the 60- and 80-pS
Bgt-AChRs differed
detectably, however, between choroid and ciliary neurons. Instead
calculations based on the combined whole cell and single-channel
results indicate that choroid neurons express approximately threefold
larger numbers of functional
Bgt-AChRs (NF) per unit area than do ciliary
neurons. Comparison with total surface
[125I]-
Bgt-AChR sites
(NT), reveals that
NF/NT
1 for both neuron populations, suggesting that "silent"
Bgt-AChRs predominate. Choroid neurons may therefore express a
higher density of functional
Bgt-AChRs by recruiting a larger
fraction of receptors from the silent pool than do ciliary neurons.
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INTRODUCTION |
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Neurons are typically
segregated into populations that perform specific functions. In a
well-studied motor system, two neuronal populations in the chick
ciliary ganglion serve different effector roles, each reflected in a
distinct neuronal phenotype (Dryer 1994; Marwitt
et al. 1971
). Ciliary neurons have large cell bodies and
innervate striated muscle fibers in the iris and ciliary body to
control the light reflex and visual accommodation. By contrast, choroid
neurons are smaller and innervate smooth muscle fibers controlling
blood flow in vessels of the choroid coat. The two populations also
differ in expression of individual genes. For example, somatostatin
immunoreactivity is detected only in choroid neurons (De Stefano
et al. 1993
; Epstein et al. 1988
), while ciliary neurons express more transcript for active agrin isoforms than do
choroid neurons (Smith and O'Dowd 1994
).
The divergence between ciliary and choroid neuron populations extends
to the structure and function of synapses formed on them by
preganglionic terminals arising from midbrain neurons (Dryer and
Chiappinelli 1987; Martin and Pilar 1963
;
Ullian et al. 1997
). Ciliary neurons receive cholinergic
synaptic input from single, large calyciform terminals that produce
large, nonsummating excitatory synaptic currents (EPSCs) that, in most
cases, decay biexponentially. Choroid neurons receive multiple, small
cholinergic boutons, producing smaller EPSCs that summate and decay
monoexponentially. For both neuron populations, however, only two
nicotinic acetylcholine receptor (AChR) classes contribute to the EPSCs
(Ullian et al. 1997
; Zhang et al. 1996
).
-Bungarotoxin (
Bgt)-AChRs are responsible for the rapidly
decaying EPSC phase and are likely to be important in maintaining
reliable ganglionic transmission (Chang and Berg 1999
).
Bgt-AChRs contain predominantly
7-subunits, are recognized and
blocked by
Bgt, and are abundant in ciliary ganglion extracts (Blumenthal et al. 1999
; Chiappinelli and
Giacobini 1978
; Conroy and Berg 1995
;
Pugh et al. 1995
). Except for a preliminary report (McNerney and Margiotta 1993
), the single-channel
properties of
Bgt-AChRs on neurons isolated from the ciliary
ganglion at any developmental age have not been studied, and the
numbers of surface receptors per neuron is unknown. The second major
receptor class, termed
3*-AChRs, mediate the slowly decaying EPSC
phase and have been estimated to be present at levels ~10-fold lower
than
Bgt-AChRs (Blumenthal et al. 1999
). These
receptors contain
3,
5, and
4 subunits and occasionally
2
subunits (Conroy and Berg 1995
; Vernallis et al.
1993
) and are neither recognized nor blocked by
Bgt but can
be detected with a number of
-subunit specific antibodies, including
mAb35 (Conroy and Berg 1995
; Vernallis et al.
1993
). Previous single-channel experiments revealed two
functional subtypes of
3*-AChRs having conductances of 25-30 and 40 pS, both of which were blocked by N-Bgt (Margiotta and Gurantz
1989
), a snake toxin that recognizes the same receptors on the
neurons as mAb35.
Given the distinct phenotypes of choroid and ciliary neurons, we
wondered whether the levels or functional properties of Bgt- and/or
3*-AChRs would also be different in the two cell populations. Thus
ciliary and choroid neurons were identified by established differences
in cell soma size, and
Bgt- and
3*-AChRs on each population were
compared. The findings demonstrate that ciliary ganglion neurons
express a restricted array of receptor classes and suggest they can
regulate the relative densities of functional
Bgt-AChRs in a
cell-specific manner.
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METHODS |
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Cell and substrate preparation
Ciliary and choroid neurons were dissociated from stage 40 (Hamburger and Hamilton 1951) embryonic day 13 or 14 (E13,14) chick ciliary ganglia using collagenase A treatment and
mechanical trituration procedures as previously described
(Margiotta and Gurantz 1989
; Margiotta and Pardi
1995
). For electrophysiological and fluorescence measurements,
dissociated neurons were plated on coated, 12- or 15-mm-diam glass
coverslips (Fisher Scientific, Houston, TX) and for binding studies in
16-mm-diam plastic tissue culture wells, both at densities of 1-4
ganglion equivalents (3,700 neurons/E14 ganglion) (Pilar et al.
1980
) per coverslip or well. Before use in experiments, the
neurons were allowed to equilibrate at 37°C in recording solution
(RS) containing (in mM) 145.0 NaCl, 5.3 KCl, 5.4 CaCl2, 0.8 MgSO4, 5.6 glucose, and 5.0 HEPES (pH 7.4) supplemented with 10% heat-inactivated
horse serum (RS+hs) for 2-4 h.
To coat glass coverslips, they were first acid-washed, then treated
with poly-D-lysine in 0.13 M borate buffer (pH 8.5), washed four times with distilled water, and air-dried. For coverslips used in
fast perfusion experiments, 70-150 kDa poly-D-lysine was applied at 1 µg/ml for 1 min (21-23°C). Using this minimal coating protocol, the neuron somata became loosely attached, allowing them to
be lifted above the substrate in subsequent fast-perfusion experiments
(see following text). For all other studies, stronger neuron attachment
was achieved by coating glass coverslips or tissue culture wells with
300 kDa poly-D-lysine at 0.5-1.0 mg/ml for 12-16 h
(4°C).
Neuron size measurements
Ciliary and choroid neurons were identified based on previously
established differences in cell body size (Landmesser and Pilar
1974; Pilar and Tuttle 1982
; Smith and
O'Dowd 1994
). To selectively label ciliary neuron cell bodies,
ciliary nerve axons were stained with the fluorescent dye
1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate
(DiI; Molecular Probes, Eugene, OR). Briefly, ciliary nerves from
dissected ganglia were draped over a 2-3 mm petroleum jelly (Vaseline)
barrier, and the cut ends were placed in distilled water at room
temperature for 10 min and then in RS containing 3% DiI (vol/vol;
dissolved in 95% ethanol at 1 mg/ml). Cell bodies usually became
labeled with DiI within 12-16 h at 37°C. Neurons from DiI-labeled
ganglia were dissociated and plated as described above, and examined
with fluorescence and DIC optics using a Zeiss Axiophot microscope.
Labeled neurons were identified with fluorescence microscopy, and major
and minor axis lengths of labeled neuronal cell bodies measured with an
eyepiece micrometer using DIC optics. Similar measurements were made on
neurons from unstained ganglia. In both cases, average axis lengths
were used as an estimate of neuronal diameter, the values compiled into
histograms and analyzed using commercial software.
Electrophysiology
WHOLE CELL RECORDING.
AChR currents were collected from visually identified ciliary or
choroid neurons at room temperature (21-24°C) using fast agonist
microperfusion (Jonas 1995) coupled with whole cell
recording, as we recently described (Pardi and Margiotta
1999
). Briefly, nicotine (20 µM) or ACh (500 µM) was
dissolved in RS, and agonist and RS streams were delivered by gravity
flow from the paired channels of theta (
) glass tubing (1.6 mm OD,
BT-15-10, Sutter Instruments, Novato, CA) pulled to an OD of
100-120 µm. The
tubing was mounted to a piezoelectric device
(Burleigh Instruments, Model LSS-3100), allowing an individual neuron
suspended in the RS stream to be rapidly exposed to the adjacent
agonist stream by a translational step triggered from the recording
software. Junction current experiments with open tip patch pipettes
revealed that movement of the laminar interface separating streams of
150 and 75 mM NaCl flowing from the
tubing occurs in <1 ms.
Agonists were also applied by pressure ejection (5-10 psi) from
large-bore (2-5 µm diam) patch pipettes (Choi and Fischbach
1981
; Margiotta and Gurantz 1989
;
Margiotta et al. 1987a
,b
). The latter method was used
for single-channel studies (see following text) and, occasionally, for
whole cell experiments to induce slow,
3*-AChR currents. Whole cell
3*-AChR currents and decay kinetics induced in this way were
indistinguishable from those obtained using fast ACh perfusion, and
results obtained using both approaches have been combined.
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(1) |
SINGLE-CHANNEL RECORDING.
Single AChR channels were studied in outside-out patches excised from
ciliary or choroid neurons. Patches were exposed to RS containing ACh
(2 or 5 µM) or nicotine (0.5 µM) by gentle pressure microperfusion
(2-5 psi) for 10-30 s, at holding potentials ranging from 60 to
140 mV. Patch currents were filtered at a cutoff frequency
(fc) of 5.8-8.0 kHz using an 8-pole
Bessel filter (902 LPF; Frequency Devices, Haverhill, MA) and sampled
onto computer disk at
5 × fc
using a TL-1 interface and pClamp 6.0 software (Axon Instruments) or an
ITC-16 interface and Pulsefit software (Instrutech Systems, Long
Island, NY). The vast majority of single-channel currents occurred as
unitary events; those corrupted by the simultaneous opening of other
channels were not included for analysis. Many of the AChR openings
associated with
Bgt-AChRs appeared to have durations <100 µs. To
accurately resolve the events, we used the criterion that only events
above a preset amplitude having durations more than two times the rise
time (Tr = 0.3321/fc) of the Bessel filter would
be accepted for analysis (Colquhoun and Sigworth 1995
).
SINGLE-CHANNEL CURRENT ANALYSIS.
Single-channel currents, obtained from choroid and ciliary neuron
patches, were compiled into histograms and fit with Gaussian functions
having means and standard deviations that defined the amplitude limits
for four channel classes. Channel conductances () were determined
from the slopes of mean channel current versus voltage (I-V)
plots. The I-V plots also provided a measure of the reversal
potential (Er) for each channel class,
determined by linear extrapolation. In patches used for kinetic
analysis, single AChR channel currents were collected at a single
holding potential (typically
100 mV), and channel conductances
calculated using mean Er values
determined from patches where full I-V plots were obtained
(Table 2). Open-duration histograms were constructed for the accepted
events in each AChR class, and fit with exponential functions to obtain
a measure of the mean channel open time (
o). Steady-state open probabilities for each AChR channel conductance class (Popen,x) were calculated using
Popen,x = (
t,x/T)/Nx,
where T represents the total record length,
t,x is the summed open durations of channel
class x, and Nx is the number of
channels of that conductance class in the patch.
Nx was estimated from the number of
current levels in each patch recording that corresponded to conductance
class x, and by visual inspection of the records was usually 1.
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(2) |
Fluorescence detection of AChRs
Bgt- and
3*-AChRs were visualized using fluorescence
methods modified from Blumenthal et al. (1999)
. To
detect
Bgt-AChRs, freshly dissociated E13,14 ciliary ganglion
neurons were treated with biotinylated
Bgt (20 nM; Molecular Probes)
in recording solution (RS) containing 10% horse serum
(RS+hs) and incubated at 37°C for 2 h.
After washing twice with RS+hs and twice with RS,
cells were fixed with 1-2% paraformaldehyde for 30 min, rinsed three
times with RS+hs, and then incubated with
Cy3-conjugated streptavidin (2 µg/ml; Jackson Laboratories, Bar
Harbor, ME) in RS+hs for 45 min at room
temperature. Cells were then rinsed twice with
RS+hs and twice with RS, and the coverslips were
dipped in distilled water and mounted on glass slides with Vectashield
(Vector Laboratories, Burlingame, CA). To detect
3*-AChRs, neurons
were treated mAb35 (100 nM; A generous gift from Dr. D. K. Berg)
in RS+hs containing 17% rabbit serum at room
temperature for 1.5 h. After three rinses in
RS+hs, cells were incubated with
biotinylated-SP-conjugated rabbit anti-rat IgG (5 µg/ml, Jackson
Laboratories) in RS+hs for 45 min at room
temperature, washed three times in RS+hs,
incubated with Cy3-conjugated streptavidin (2 µg/ml) in
RS+hs, and then rinsed three times with
RS+hs and twice with RS. Cells were then fixed in
1-2% paraformaldehyde for 10 min and washed and mounted as for
Bgt
staining. Previous studies have demonstrated the specificities of these
probes for fluorescence detection of AChRs on chick ciliary ganglion
neurons (Jacob et al. 1984
; Wilson Horch and
Sargent 1995
).
Neurons were examined with DIC and reflected light fluorescence using
an Olympus BX50 microscope equipped with a UplanFL ×40 objective (0.75 N.A.) and optics appropriate for Cy-3 detection (510-550 nm band-pass
filter, 570-nm dichroic mirror, 590-nm barrier filter). After focusing
on the cell surface, 16-bit DIC and fluorescence images were acquired
from identified ciliary and choroid neurons using a cooled digital CCD
camera (SenSys, Model KAF-1400; Photometrics, Tucson AZ) under the
control of IP Lab software (Version 3.0 Scanalytics; Reading, PA). For
subsequent quantification of the fluorescence signal from each neuron
studied, an elliptical region of interest (ROI) was placed around the
neuron perimeter (defined by the DIC image), and the fluorescence
intensity values for each pixel within the ROI were summed. The summed
intensity value was then divided by the ROI area yielding fluorescence
intensity per unit area (total fluorescence density). The background
fluorescence density was obtained for an identical ROI area acquired
from an adjacent unlabeled region of the same image, and the value
subtracted to yield net fluorescence density (total minus background)
for each neuron. A similar immunofluorescence detection and CCD
quantification approach was used previously to determine relative
levels of surface Bgt- and
3*-AChRs on freshly dissociated
ciliary ganglion neurons isolated at various stages of development
(Blumenthal et al. 1999
). In this earlier study, tests
of the SenSys CCD camera demonstrated that the probe concentrations
used here are in the linear portion of the camera's fluorescence
detection range. Digitized images were prepared for presentation using
Photoshop 4.0 (Adobe Systems, San Jose, CA) after conversion to TIFF format.
Bgt binding assay
NEURON ISOLATION.
Immediately after trituration, dissociated ciliary ganglion neurons
were plated in triplicate, 16-mm-diam plastic culture wells (at 1-4
ganglion equivalents per well in a final volume of 400 µl). This
plating procedure gives a uniform distribution of neurons, but because
some are invariably lost during the dissociation, the actual number of
neurons per well was determined by counting 5-10 randomly chosen
microscope fields for each condition. Calculated yields were typically
50-75%, based on a total of 3,700 neurons (2,300 choroid +1,400
ciliary) expected per E14 ciliary ganglion (Pilar et al.
1980).
NEURON MORPHOMETRY.
To determine the relative numbers and surface areas of choroid and
ciliary neurons, the lengths of major and minor axes of 200 randomly
selected neurons were measured. The axes were used to calculate soma
surface areas (S) based on the area equation for an oblate
spheroid {S = 2
a2 +
(b2/
) log [(1 +
)/ (1
)]} where a and b are the lengths of the major and minor semiaxes, and
is the ellipticity given by
= (a2
b2)1/2/a
(Beyer 1987
), and the calculated S values
were compiled into histograms. The surface area histograms were well
described by the sum of two Gaussian functions describing choroid
(60-70%) and ciliary neuron populations, with ciliary neurons having
mean surface areas three- to fourfold larger than choroid neurons.
BINDING.
After equilibration for 2 h at 37°C, neurons were incubated in
RS+hs containing 0-130 nM
[125I]-Bgt at 37°C for 2 h and rinsed
three times with RS+hs.
[125I]-
Bgt was obtained from DuPont NEN
(Wilmington, DE) at an initial specific activity of 209 cpm/fmol.
Nonspecific binding was determined in parallel triplicate wells by
including 1 µM unlabeled toxin with the
[125I]-
Bgt For quantification of
125I radioactivity, the wells were scraped in 0.6 N NaOH, and the radioactivity counted in an LKB Wallac gamma counter
(Model 1275 Minigamma, Gaithersburg, MD).
Materials
Fertilized white leghorn chicken eggs were obtained from Hertzfeld Poultry Farm (Waterville, OH) and maintained at 37°C in a forced-air draft incubator at 100% humidity. Any chemicals or reagents not already specified were obtained from Sigma (St. Louis, MO).
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RESULTS |
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Identification of ciliary and choroid neurons
Neurons dissociated from E13,14 ciliary ganglia fall into ciliary
and choroid populations that can be distinguished on the basis of
differences in cell body size (Fig. 1,
A and B). Measurements from 1,300 selected
neurons revealed a cell body diameter distribution best fit by a
bimodal Gaussian function having significantly different peaks at
13 ± 2 and 23 ± 3 µm (mean ± SD), respectively
(P < 0.001). These diameter values are similar to
those previously reported for choroid and ciliary neurons, respectively
(Landmesser and Pilar 1974
). To confirm the correlation
of cell size with each respective neuron population, ciliary neuron
cell bodies were labeled by staining their axons with DiI (Fig. 1,
C and D). DiI-labeled ciliary cell bodies were
uniformly large, having diameters well described by a single Gaussian
function with a peak at 21 ± 4 µm. The size of DiI-stained
ciliary neurons was significantly different from that of the small
cells (P < 0.001) but not of the large cells
(P > 0.1) measured from unstained ganglia. Large cells
were therefore assigned to the ciliary neuron population. Small cells
were assigned to the choroid neuron population because they did not
label with DiI and because the choroid and ciliary neuron populations
in the ganglion have previously been shown to correlate with small and
large soma diameters, respectively (Landmesser and Pilar
1974
; Pilar and Tuttle 1982
). Thus the 95% confidence interval (CI = 1.96 × SD) for the large cell
distribution was used to establish cells with diameter
16 µm as a
criterion size for choroid neurons. Similarly, using the 95% CI for
the small cell distribution would yield a criterion size of
17 µm for cells to be assigned to the ciliary population. In practice, however, only the largest cells (with diameters
23 µm) were
considered ciliary neurons. This was done because previous studies
indicated that the two populations in adult chickens overlap in size
somewhat, with some neurons of intermediate size scoring as choroid
cells by ultrastructural criteria (De Stefano et al.
1993
; Pilar et al. 1980
).
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Whole cell AChR currents from choroid and ciliary neurons
To maximally activate Bgt- or
3*-AChRs, choroid and ciliary
neurons were transiently exposed to saturating doses of nicotine (20 µM) or ACh (500 µM). Nicotine was used to assay whole cell responses mediated by
Bgt-AChRs (Fig.
2) because it provided good temporal
separation of
Bgt- and
3*-AChR currents (Blumenthal et al.
1999
; Pardi and Margiotta 1999
) due to the
higher affinity of
Bgt-AChRs for nicotine (Zhang et al.
1994
). To determine if specific
Bgt-AChR responses differed
in ciliary and choroid populations, whole cell current values were
normalized to membrane capacitance (Cm), which was presumed to be
proportional to membrane surface area and therefore to cell size. In
both neuron populations, fast application of 20 µM nicotine induced a
biphasic response featuring a large, rapidly decaying response
component, and a smaller, slowly decaying response component (Fig. 2,
A and B). We confirmed that the rapidly decaying
component represents activation of
Bgt-AChRs since it was completely
abolished after treating neurons with
Bgt (e.g., Fig. 2C;
see also Fig. 3C) as previously described (Blumenthal
et al. 1999
; Pardi and Margiotta 1999
;
Zhang et al. 1994
). The peak values of
Bgt-AChR
currents (If) were similar in choroid
and ciliary neurons (Fig. 2, A and B, insets),
and overall If did not differ
significantly between the two populations (P > 0.1, Table 1). After normalizing for
population differences in Cm, however,
the similar values for If translated
to substantially larger peak values of
Bgt-AChR current density
(If/Cm)
associated with choroid neurons when compared with ciliary neurons
(Fig. 2, A and B, filled arrows). Overall, this
difference was threefold (P < 0.001, Table 1). While
3*-AChR currents were studied in isolation using ACh applied in the
presence of
Bgt (see following text), they could also be resolved as
the slowly decaying current component
(Is±i) induced by nicotine. In
contrast with the cell-specific difference observed in
Bgt-AChR
current density, Is±i values induced
by nicotine were scaled proportionally in choroid and ciliary neurons
(
484 ± 57 and
1,959 ± 150 pA, respectively,
P < 0.001), and the resulting measures of
Is±i/Cm (e.g., Fig. 2, A and B, open arrows) did not
differ detectably between the two neuron populations (
53 ± 6 and
67 ± 7 pA/pF, respectively, P > 0.1).
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|
We were initially concerned that the difference in Bgt-AChR current
density might reflect nonsynchronous exposure to agonist across the
cell surface, a potential limitation of the fast perfusion method that
might be expected to be more severe for larger cells. This concern now
seems unfounded for two reasons. First, the
tubing pore diameter
(50 µm) used to deliver agonist exceeds the mean diameter of ciliary
neurons by twofold. Second, if ciliary neurons were more susceptible to
size-related limitations of the perfusion system, they would be
expected to display slower kinetics of AChR current activation and
desensitization. We measured the time required for
Bgt-AChR currents
to attain peak value (Tp = 4-10 ms)
and the time constant describing the
Bgt-AChR desensitization (
f = 5-11 ms) following nicotine exposure but
detected no significant cell-specific differences in these parameters
(Fig. 2; P > 0.1, Table 1). Thus the larger
Bgt-AChR current density observed for choroid neurons cannot be
readily explained by technical limitations of the perfusion system and
is likely to represent a true difference between
Bgt-AChRs on the
two neuron populations.
Fast perfusion experiments with nicotine indicated a difference between
choroid and ciliary neurons in Bgt- but not
3*-AChR current
densities. To further explore this finding, we examined
3*-AChRs in
isolation after treating neurons with 60 nM
Bgt to block
Bgt-AChRs. Identified neurons were tested as in Fig. 2, except that
500 µM ACh was used as the agonist, and
Bgt was present (Fig.
3). Under these
conditions,
3*-AChRs are activated in isolation from
Bgt-AChRs
(Blumenthal et al. 1999
; Pardi and Margiotta
1999
) producing a peak whole cell current represented by either
a single slow component or the sum of intermediate and slow components
(Is±i) having time constants
i and
s (Fig. 3,
A and B). In the presence of
Bgt, the
isolated, peak
3*-AChR currents induced by ACh were scaled
proportionally to neuron cell size (Fig. 3, A and B,
insets) resulting in current density values
(Is±i/Cm)
that were not detectably different between choroid and ciliary neurons
(Fig. 3, A and B, open arrows; P > 0.1, Table 1). To confirm the specificity of
Bgt, values of
Is±i/Cm
obtained for choroid and ciliary neurons (using nicotine in the
presence of toxin or ACh in its absence) were compared. The observation
that
Is±i/Cm
values were indistinguishable between choroid and ciliary neurons for
both ACh and nicotine, while
If/Cm
was dramatically reduced by >90% in both cases (Fig. 3C)
indicates
Bgt is specific in blocking
If without affecting Is±i. The time required for
3*-AChR currents to attain peak value and the associated decay time
constants were also measured and found to be generally similar for the
two neuron populations. For both choroid and ciliary neurons,
Tp was
10 ms and
s was
2 s, while
i
for choroid neurons (90 ms) reflected a somewhat faster decay than the
value obtained for ciliary neurons (P < 0.05, Table
1). The small difference detected in
i is
nevertheless likely to be specific since the methods employed permit
adequate resolution of much faster desensitization processes (see Fig. 2 and METHODS). Taken together, the findings indicate that
the peak whole cell response density attributable to
Bgt-AChRs is threefold larger in choroid compared with ciliary neurons, while that
attributable to
3*-AChRs did not differ detectably between the two
populations.
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Comparison of Bgt- and mAb35-site densities on choroid and
ciliary neurons
We next used a fluorescence method (Blumenthal et al.
1999) to determine if the larger
Bgt-AChR responses seen for
choroid neurons could be explained by a higher overall density of
Bgt binding sites expressed on the neuronal cell surface (Fig.
4). Neurons were incubated with
biotinylated
Bgt and then with Cy3-conjugated streptavidin to
visualize
Bgt-AChRs. Neurons in ciliary and choroid populations were
identified based on the criteria described in Fig. 1, and both
displayed specific labeling (Fig. 4A) compared with control
preparations where the cells were treated with 100 µM
D-tubocurarine chloride (dTC; Fig. 4B) or where
the toxin was omitted (data not shown). There was, however, no obvious
difference in the intensity or the pattern of
Bgt labeling on the
two neuronal types. Both ciliary and choroid neurons displayed a
nonhomogeneous distribution of
Bgt-AChRs at about the same overall
surface density. Specific
3*-AChR density was also similar for the
two cell types, as seen by using mAb35 as the label (Fig. 4,
C and D). The higher fluorescence intensity
associated with mAb35 (relative to
Bgt) probably reflects signal
amplification due to the utilization of a biotinylated secondary
antibody (See METHODS). To quantify the level of AChR
labeling on the two neuron populations, we compared the average
fluorescence intensity of the labeling for
Bgt- and
3*-AChRs on
the two neuron populations using digital microscopy. In accord with the
images in Fig. 4, A-D, there was no detectable difference
in
Bgt- or mAb35 net fluorescence density between ciliary and
choroid neurons (Fig. 4E). The lack of a difference in
Bgt- net fluorescence density between ciliary and choroid neurons
did not represent a sensitivity limitation of digital microscopy.
Reducing the Cy3-conjugated streptavidin concentration threefold by
molar replacement with FITC-conjugated streptavidin resulted in a
2.1 ± 0.2-fold reduction in average fluorescence intensity
compared with control neurons treated with normal levels of
Cy3-conjugated streptavidin (P < 0.001;
n = 16 neurons for each condition). These findings
indicate that the higher
7-AChR response density for choroid over
ciliary neurons (Fig. 2; Table 1) cannot be explained by a gross
cell-specific difference in total
7-AChR surface density.
|
Surface labeling with [125I]-Bgt reveals an
abundance of
Bgt sites
We next measured the number of surface Bgt-AChRs present on
ciliary and choroid neurons. Freshly isolated E14 ciliary ganglion neurons displayed saturable surface binding of
[125I]-
Bgt having a
KD of 1.1 ± 0.2 nM
(n = 2) and a Bmax of
7.67 ± 0.03 fmol per ganglion equivalent (n = 2, Fig. 5A). To determine the
relationship between number of neurons and binding sites, the number of
ganglion equivalents was varied, and these studies yielded a linear
relationship (Fig. 5B) predicting a similar surface site
density (7.47 fmol per ganglion equivalent). These results are
consistent with previous findings from whole ganglion extracts, where
detection of surface, intracellular and presynaptic
Bgt sites
revealed 15-20 fmol of
Bgt bound per E14 ciliary ganglion (Chiappinelli and Giacobini 1978
; Conroy and Berg
1995
; Pugh et al. 1995
). Based on digital
imaging experiments, we determined that 94 ± 1% of the net
Bgt fluorescence was associated with neuron cell bodies; only a tiny
fraction was associated with bits of membrane debris or nonneuronal
cells (data not shown; n = 10 fields from 2 experiments). Normalizing the number of sites for the yield of cells in
each experiment and correcting for the small amount of binding to
debris and nonneuronal cells permitted calculation of 1.14 ± 0.01 × 106
Bgt-sites per neuron
(n = 3 experiments). Because the density of
Bgt
sites was equivalent on the two neuron populations (Fig. 4E), we were able to calculate the number of
Bgt sites
per average choroid and ciliary neuron. Major and minor axis
measurements from individual neurons in the same wells revealed the
relative distribution of choroid and ciliary neurons in each well and
allowed calculation of cell areas. The area and distribution
measurements were then used to calculate the numbers of
Bgt binding
sites on choroid and ciliary neurons, assuming the label had access to
the entire cell surface in both cases. Such calculations revealed, on
average, 6.88 × 105
Bgt sites per
choroid neuron and 2.41 × 106
Bgt sites
per ciliary neuron.
|
Single AChR currents from ciliary and choroid neurons reveals four distinct channel classes
A higher Bgt-AChR affinity for nicotine cannot explain the
larger response density of choroid over ciliary neurons since nicotine
applied at 20 µM is known to generate near-maximal
Bgt-AChR responses from ciliary neurons (Zhang et al. 1994
). We
therefore conducted excised patch experiments to determine if the
higher
Bgt-AChR responses of choroid neurons resulted from
cell-specific differences in the permeation or kinetic properties of
individual receptors. Four distinct AChR channel amplitude classes were
detected in outside-out patches excised from choroid or ciliary neurons challenged with recording solution containing 2-5 µM ACh (Fig. 6, A and B, Table
2) or 0.5 µM nicotine. The events were
mediated by nicotinic AChRs since they were absent in patches
pretreated with 100 µM dTC and not seen in patches challenged with
recording solution lacking agonist (data not shown). The AChR channel
events were classified according to their conductance (
), determined from the slope of single channel current amplitude versus voltage plots
(e.g., Fig. 6C). Two previously described event classes (Margiotta and Gurantz 1989
) displaying slope
conductances of approximately 30 and 40 pS (Table 2) were routinely
observed. Two additional AChR channel classes that we noted previously
but did not characterize because of their brief open durations
(Margiotta and Gurantz 1989
) were also present in most
patches. In the present studies, we were able to record these latter
events at multiple voltages to determine that they display slope
conductances of ~60 and 80 pS (Fig. 6C; Table 2). Each of
the four channel classes displayed an extrapolated current reversal
potentials (Er) indicative of similar
cationic selectivities. Mean Er values
were about
10 mV for the 30- and 40-pS channels, as previously
described, and nominally closer to 0 mV for the 60- and 80-pS channels
(Table 2). As expected for AChR permeation (Gardner et al.
1984
),
and Er values
obtained for the four channel classes using 0.5 µM nicotine as
agonist were similar to those obtained using ACh (n = 4 patches; data not shown). In addition, there was no detectable difference between choroid and ciliary neurons in mean values of
or
Er (P > 0.1 for both)
for each channel class, suggesting that ion permeation through AChRs is
similar in the two neuronal populations.
|
|
To further explore potential cell-specific differences in channel
properties and more fully characterize Bgt-AChRs, the open time,
open probability, and opening frequency of each AChR conductance class
were determined using ACh as the agonist (Fig. 6D and Table 2). As with permeation properties, however, these kinetic parameters did not differ detectably between ciliary and choroid neurons (Table
2), and results from both populations are combined in the following
description. For each channel class, open time constants (
o) were derived by fitting exponential
functions to open-duration histograms, while open probabilities
(Popen), and opening frequencies (Fopen) were calculated directly from
the records (see METHODS). As previously shown, the open
time distributions for the 30- and 40-pS AChR events were best fit by
the sum of two exponential functions (Margiotta and Gurantz
1989
). The open time constants (
o,1
and
o,2) were ~100 µs and 2 ms,
respectively, and as before, the brief-duration openings represented a
minority contribution for both event classes (Table 2). The 30- and
40-pS events occurred with an overall
Fopen of ~5 and 18 s
1 and with
Popen values of 0.011 and 0.067, respectively. In contrast, the 60- and 80-pS openings were uniformly
brief, with
o for the 80-pS events (
200
µs) scoring about twofold larger (P < 0.001) than
those of the 60-pS events (Fig. 6, A and D; Table
2). The brief open durations of 60- and 80-pS events, combined with
opening frequencies of 7 and 5 s
1, respectively,
resulted in their making a much smaller net contribution to the total
record open time (Popen
0.002)
than either 30- or 40-pS events.
Bgt blocks 60- and 80-pS AChR channels
To correlate the channel conductance classes with AChRs of
established subunit composition and toxin sensitivity, neurons were
treated with Bgt (60 nM). Toxin block was assessed by comparing Fopen and
Popen for each conductance class in
patches from treated and untreated neurons. Results from choroid and
ciliary neurons were again indistinguishable and are combined in the
following description. In each of six patches from neurons treated with 60 nM
Bgt, both Popen (Fig.
7, A and B) and
Fopen (not shown) associated with the
60- and 80-pS events dropped to zero. The blockade of 60- and 80-pS
AChR channels was specific since values for
Fopen and
Popen for the 30- or the 40-pS events
were not detectably changed in
Bgt-treated patches when compared
with the values obtained in 13 control patches (P > 0.1 for both). This finding indicates that the 60- and 80-pS events
arise from
Bgt-AChRs, and extends the results from whole cell
experiments where 60 nM
Bgt blocked the initial, rapidly decaying
current density component attributable to
Bgt-AChRs but not the slow
component attributable to
3*-AChRs. The experiments further suggest
that the 30- and 40-pS events arise from
3*-AChRs, which do not
contain
7-subunits and are neither recognized nor blocked by
Bgt.
|
Bgt-AChR opening kinetics
We analyzed the closed durations between 60-pS channel openings
induced by 5 µM ACh in outside-out patches excised from choroid and
ciliary neurons (Fig. 8) to gain more
information about the rate constants governing Bgt-AChR channel
activation (k
n,
, and
=
o
1) based on Scheme 2. The analytical approach described by Colquhoun and Hawkes
(Colquhoun and Hawkes 1981
; Colquhoun and Sakmann
1981
; Colquhoun and Sigworth 1995
; Sine
and Steinbach 1986
) was employed because it is more amenable to
records with few events (
200-300) such as those obtained here, than
is single-channel ensemble analysis (Dionne and Leibowitz
1982
; Leibowitz and Dionne 1984
), which we used
previously to study
3*-AChR kinetics (Margiotta and Gurantz 1989
; Margiotta et al. 1987a
,b
). The basic
assumption inherent in both approaches is that a single channel having
just closed to AnR, and still fully occupied by agonist, is more likely
reenter the open state than channels are to enter the open state from any of the other n closed states. Scheme 2 provides a generally valid description of the processes governing
ligand-gated channel activation, but receptor desensitization would
require that additional considerations be made. While
Bgt-AChRs do
desensitize rapidly after exposure to high concentrations of agonist
(e.g., Fig. 2),
Bgt-AChR desensitization does not appear significant
under the low concentration conditions of our single-channel
experiments. First, although the opening rate of 60- and 80-pS events
(Fopen) was low using 5 µM ACh,
especially when compared with 40 pS AChRs (Table 2), the 60- and 80-pS
events occurred at a reasonably steady rate that did not vary by more
than a factor of 2 throughout most recordings. Second, when nicotine is
applied to neurons at 0.5-2.0 µM, whole cell currents attributable
to
Bgt-AChRs are sustained (Zhang et al. 1994
; Nai
and Margiotta, unpublished data). Third, if 5 µM ACh did cause
Bgt-AChR channels to desensitize significantly, the closed-duration
distributions would be expected to display several components
reflecting the slow process of recovery from desensitization
(Sine and Steinbach 1987
), but these were not observed
(see following text).
|
According to Bgt-AChR activation described by Scheme 2, the distribution of closed durations at low agonist concentration should be represented by the sum of a few exponential functions with
one or two slow component(s) representing time spent by all channels in
the n + 1 closed states and a fast component representing the more rapid transitions of a single channel between
AnR and AnR* governed by opening and closing
transition rate constants
and
. Reconstructed records portraying
only events corresponding to 60-pS channels in a choroid patch (Fig. 8,
A and B) clearly reveal "bursts" of openings,
with each opening separated by short "gaps" far more brief than the
average closed duration. Histograms of all closed durations separating
60-pS
Bgt-AChR channel openings from the same patch (Fig.
8C) indicate these two kinetic components have fast
[
1 = 1/(
+ k
n)] and slow
(
2) time constants of 6 and 54 ms,
respectively. In this example, a calculation of the critical gap length
that would minimize misclassifying short and long closed durations
(Colquhoun and Sigworth 1995
) was 7 ms, and the mean
number of gaps per burst (m =
/k
n) determined as 1.2. The
findings predict values of 87 and 74 s
1 for
and
k
n, respectively, for 60-pS
channels in this choroid neuron patch. From the distribution of 60-pS
channel open times (Fig. 6D), a value of 7462 s
1 was determined for
, and po
[po =
/(
+
); see below]
calculated as 0.012. Similar values were obtained in five other patches
from four ciliary and one other choroid neuron predicting a mean
po value of 0.013 for 60-pS
Bgt-AChRs on both neuron populations (Table
3). In three of the same six patches,
single-channel ensemble analysis (Dionne and Leibowitz
1982
; Leibowitz and Dionne 1984
) predicted
similar values for
and po (73 s
1 and 0.010, respectively). Unfortunately, it was not possible to reliably study
80-pS channel closed durations due to the lower numbers of these events
usually obtained. The observation that Popen values were indistinguishable
for 80-pS relative to 60-pS channels (Table 2; P > 0.1), however, suggests similar rate constants describe the activation
process of both channel classes.
|
Choroid neurons would exhibit a higher Bgt-AChR response density
than ciliary neurons if they expressed a higher density of functional
Bgt-AChRs. This would occur if the number of functional
Bgt-AChRs
(NF) on both populations was far less
than the total number (NT) determined
from [125I]-
Bgt binding studies (Fig. 5; see
following text) and the fraction of functional receptors were regulated
in a cell-specific manner. In principle,
NF can be calculated (Margiotta
and Gurantz 1989
; Margiotta et al. 1987a
,b
)
using NF = If/poi
(Hille 1992
) where If
is the maximal
Bgt-sensitive whole cell current (
3,500 and
4,100
pA for choroid and ciliary neurons, respectively; Table 1),
i is the single
Bgt-AChR channel current (
4.9 pA at
70 mV), and po is the probability
that the channels will be in the open state (0.013). The calculation
predicts NF = 55,000 for choroid neurons, and 64,600 for ciliary neurons, much smaller than the total
number of
Bgt sites per choroid and ciliary neuron determined in
Fig. 5 (6.88 × 105 and 2.41 × 106, respectively). Three assumptions are
inherent in using this calculation to determine
NF: The first is that, under the
conditions of the whole cell experiments,
Bgt-AChRs rapidly achieve
full occupancy (AnR). This seems
reasonable since agonists are applied by fast perfusion having an
estimated delay time <1 ms, well below the 10-ms
Td time required to produce full
current activation (Table 1). Also, if
Bgt-AChRs display similar
k
n values for different
agonists as do muscle-type AChRs (Sine and Steinbach
1986
), the
10 µM EC50 for
Bgt-AChR activation by nicotine (Zhang et al. 1994
)
predicts a KD based on
n = 1 or 3 sites per receptor (see following text) of
10 or 2.6 µM, and since KD = k
n/k+n,
a rapid association rate constant
(k+n) of 0.8 or 3.1 × 107
M
1
s
1. The derived
k+n rate constant is similar to
that for muscle AChRs obtained using similar physiological approaches (Dionne and Leibowitz 1982
) and about one-half that
measured directly for Torpedo AChRs using rapid-mixing
ultrafiltration methods (Boyd and Cohen 1980
). The
second assumption is that the po
values, derived for low agonist concentrations, apply at the high
agonist concentration used in the whole cell studies (e.g., Figs. 2 and 3). Given that the saturating agonist doses result in most
receptors rapidly achieving the fully liganded state, Scheme
2 reduces to AnR
AnR*
with the probability of being in the open state,
AnR*, given by
po =
/(
+
) (Sine and
Steinbach 1987
). Unlike Popen (Table 2), po is independent of
agonist concentration since it depends solely on the rate constants
governing the transitions between
AnR and
AnR*. Thus
po determined using Scheme
2 at low doses of agonist should also apply for high agonist
concentrations, as was previously shown for AChRs on clonal BC3H-1
cells (Sine and Steinbach 1986
, 1987
). The third
assumption that a mean value of
4.9 pA can be used for the
contribution of 60- and 80-pS channels to the whole cell current at
70 mV also seems reasonable. Similar values for
NF
(NF = NF60 + NF80) were obtained for choroid
(57,000) and ciliary (67,000) neurons when they were calculated
independently for 60- and 80-pS channels
(If = NF60poi60 + NF80poi80;
i60 =
4.2 pA and
i80 =
5.6 pA) using the ratio of 60- to 80-pS events per patch (1.75) as a measure of the relative abundance
NF60/NF80 of the channel classes. Considering that ciliary neurons have approximately equal to threefold larger surface area than choroid neurons (Table 1, Fig. 1) the 64,600 and 55,000 functional
Bgt-AChRs present on ciliary and choroid neurons, respectively, result in choroid
neurons expressing a 2.6-fold higher density of functional
Bgt-AChRs. Thus the 3.0-fold larger
Bgt-AChR response density seen for choroid neurons (Fig. 2, Table 1) can be accounted for by a
higher density of functional
Bgt-AChRs on this neuron population.
Functional Bgt-AChRs are a small fraction of total
Bgt-AChRs on both neuron populations
The total number of Bgt-AChRs per choroid or ciliary neuron
(NT) was evaluated from the number of
Bgt sites obtained in surface binding studies (e.g., Fig. 5). Since
the ratio of sites bound by toxin for each
Bgt-AChR on ciliary
ganglion neurons is unknown, it was estimated from previous studies
performed on native receptors from other neuronal preparations.
Biochemical experiments using subunit-specific antibodies to recognize
purified receptor obtained from PC-12 cells and rat brain neurons
(Chen and Patrick 1997
; Drisdel and Green
2000
) indicate that
Bgt-AChRs are homopentamers consisting
solely of
7 subunits. While each is presumed to have a site
recognized by
Bgt,
7 subunit monomers do not bind
Bgt
(Pugh et al. 1995
) and for assembled receptors in rat
brain neurons, the molar ratio of
Bgt binding sites to
7 subunit
was 0.2, suggesting that each
7 homopentamer contains only one
Bgt-accessible site (Chen and Patrick 1997
). As for ciliary ganglion neurons, which express the same subset of AChR subunits as do PC-12 cells (Blumenthal et al. 1997
), the
Hill coefficient for
Bgt competition binding to
Bgt-AChRs on
intact neurons is
1.0 (Vijayaraghavan et al. 1992
).
This finding also suggests
Bgt binds to a single site on its
receptor, although the results do not exclude the possibility of
multiple sites displaying little or no cooperativity (Limbird
1996
). In contrast, functional studies with nicotine reveal a
dose-response relation for
Bgt-AChRs on ciliary ganglion neurons
having a Hill slope of 2-3 (Zhang et al. 1994
).
Similarly, Hill slopes for nicotine or ACh binding to native
Bgt-AChRs on PC-12 cells were both 2.4 (Rangwala et al.
1997
), suggesting, as do the functional studies on ciliary ganglion neurons, that the
Bgt-AChRs in these preparations may have
three agonist binding sites per receptor. Given these considerations, we estimated a range of one to three available
Bgt sites for each
Bgt-AChR on ciliary ganglion neurons. Based on this range, and the
numbers of surface [125I]-
Bgt sites per
neuron soma (Fig. 5), NT can be
predicted as 2.29-6.88 × 105 per average
choroid neuron and 0.80-2.41 × 106 per
average ciliary neuron. The derived value of
NF for choroid neurons (55,000) would
therefore represent 8-24% of NT,
while for ciliary neurons the same argument would predict that
NF (64,600) represented only 3-8% of
NT. By such considerations, choroid
and ciliary neurons would appear to express
Bgt-AChRs in substantial excess of the numbers required to elicit a maximal current, suggesting that the majority of
Bgt-AChRs on both populations is "silent". The higher
NF/NT
ratio associated with choroid neurons can then be interpreted as a form
of cell-specific regulation to control the expression of functional
Bgt-AChRs.
![]() |
DISCUSSION |
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---|
The present findings extend the distinction between ciliary
ganglion neuron populations to include differential expression of
specific functional AChR classes and provide new information about the
identity, abundance, and single-channel properties of Bgt-AChRs. We
determined that nicotinic response densities attributable to
Bgt-AChRs are threefold larger on choroid than on ciliary neurons,
while those attributable to
3*-AChRs are similar for the two
populations. The cell-specific difference in
Bgt-AChR response
density did not result from physical limitations of the fast agonist
perfusion method. Such limitations would have produced longer response
latencies and slower rates of AChR desensitization for ciliary compared
with choroid neurons, neither of which were observed. In spite of their
larger
Bgt-AChR response, choroid neurons did not display a
detectably higher density of surface
Bgt sites determined either by
[125I]-
Bgt binding or by digital
fluorescence microscopy when compared with ciliary neurons, although a
threefold difference would have been readily detected. Moreover,
analysis of single-channel records obtained from choroid and ciliary
patches failed to detect any cell-specific difference in either the
permeation or the kinetic properties of individual
Bgt-AChR channels
expressed on the two neuronal populations. Instead the findings lead us
to conclude that a large pool of surface
Bgt-AChRs exists on both
neuronal populations, with most being "silent," and only a minor
fraction being functional in the sense that they are immediately
available as pathways for transmembrane ionic flux. Differences in
Bgt-AChR response density can then be explained by cell-specific
regulation of the fraction of functional
Bgt-AChRs from the much
larger total pool of surface receptors. This cell-specific regulation may be related to the fact that choroid and ciliary neurons innervate different peripheral targets and receive distinct forms of synaptic input. Normal synaptic inputs may be particularly critical for maintaining the ratio of total to functional surface AChRs because denervation reduces total surface AChRs on ciliary ganglion neurons (Arenella et al. 1993
) without grossly affecting
functional AChRs, assessed by whole cell ACh-induced currents
(Schwartz-Levey et al. 1995
). Thus cell-specific,
input-derived signals, perhaps working in conjunction with
target-derived signals, are reasonable candidates to explain the
differential regulation of functional
Bgt-AChRs seen here for
choroid and ciliary neuron populations.
Analysis of single 3*- and
Bgt-AChR channels provided new
insights about the properties and respective subunit composition of the
four channel species present on ciliary ganglion neurons and was
instrumental to the conclusions summarized in the preceding text.
Previous co-precipitation studies using subunit specific antibodies
indicated two broad AChR classes on ciliary ganglion neurons, each
represented by two identifiable subtypes. The majority of
3*-AChRs
contain
3,
4, and
5 subunits, but ~20% also contain
2
subunits (Vernallis et al. 1993
). In contrast, most
Bgt-AChRs contain
7 subunits (but not
3,
5,
4, or
2
subunits), while ~5% of
Bgt-AChRs (
T/35-AChRs) contain neither
7 nor any of the known neuronal receptor subunits (Pugh et
al. 1995
). Our single-channel experiments reveal four
functional AChR channel classes characterized by conductances of 30, 40, 60, and 80 pS that can be correlated with the four molecular
species on the basis of their sensitivity to blockade by
Bgt. Since
Bgt does not recognize
3*-AChRs (Vernallis et al.
1993
), the 60- and 80-pS AChR channels blocked by the toxin can
be considered, by definition, to be members of the
Bgt-AChR class.
Based on the co-precipitation findings cited in the preceding text,
either the 60- or 80-pS conductance channels could represent
7
subunit homopentamers and the other
T/35-AChRs. Since the functional
status of
Bgt-AChRs lacking
7 has not been established, however,
and other uncharacterized receptor subunits may be present in the
ganglion, it is also possible that the 60- and/or 80-pS conductance
channels represent
7-AChR subtypes that assemble in two different
stoichiometric combinations with other subunits (Vernallis et
al. 1993
). A third possibility, suggested by the requirements
for expression of functional
Bgt-AChRs in oocytes and cell lines
(Chen et al. 1998
; Rakhilin et al. 1999
)
is that both channel classes represent
7 homopentamers that are
differentially altered by posttranslational subunit modification,
resulting in channels having different biophysical properties. The
present findings do not permit us to distinguish between these
interesting possibilities. Since
Bgt-AChRs and
3*-AChRs are the
only known AChRs present on ciliary ganglion neurons, the 30- and 40-pS
channel events are likely to reflect activation of
3*-AChRs. While
this correlation of 30- and 40-pS channels with
3*-AChRs and 60- and 80-pS channels with
Bgt-AChRs on the basis of
Bgt-sensitivity seems reasonable, further studies are necessary before each of the four
channel classes can be identified with a unique molecular subtype.
The 60- and 80-pS conductance values we obtained for Bgt-AChR
channels agree well with the 73-pS AChR channel class attributed to the
Bgt-sensitive nicotine response in rat hippocampal neurons (Castro and Albuquerque 1993
). Open-duration analysis
for the 73-pS channel revealed a 0.12-ms mean open time (Castro
and Albuquerque 1993
) similar to the 0.12- and 0.19-ms
o values for 60- and 80-pS
Bgt-AChR
channels reported in Table 2. The
Bgt-AChR channel conductances we
observed also agree well with the 45-pS value obtained for chick
7
homopentamers expressed in Xenopus oocytes (Revah et
al. 1991
), which, after correcting for the lower cation concentration of the amphibian recording solution (Hille
1992
), would be ~60 pS. The 60- and 80-pS conductances
reported here differ, however, from the 19- and 32-pS values reported
for chick
7 subunits expressed in human BOSC 23 cells
(Ragozzino et al. 1997
). In this case, the multiple
conductances are puzzling since only
7 subunits were expressed,
suggesting that endogenous AChR-like subunits coassembled with the
7
subunits or that individual
7 subunits were altered by different
posttranslational modifications. Moreover, the filter settings and
sampling rates employed (2 and 10 kHz, respectively) would have
precluded detection of brief openings such as those seen here for the
60- and 80-pS
Bgt-AChR channels. Somewhat surprisingly, the
conductance values we find for
Bgt-AChRs in acutely dissociated
chick parasympathetic neurons are also different from the 18-pS value
reported for
Bgt-sensitive,
7-containing AChRs detected on chick
sympathetic neurons in culture (Yu and Role 1998
). This
disparity may reflect cell-specific differences in the
7-subunit
content of
7-AChRs on parasympathetic and central neurons versus
sympathetic neurons (Yu and Role 1998
) or be related to
different levels or processing of
7-subunit transcript expression seen for neurons from the ganglion compared with those maintained in
culture (Corriveau and Berg 1994
). Relevant to the
latter interpretation, we recently found that similar large,
rapidly-desensitizing,
Bgt-sensitive whole cell currents can be
induced by 20 µM nicotine in freshly dissociated embryonic ciliary or
sympathetic ganglion neurons (S. Thomasey and J. F. Margiotta,
unpublished data), whereas in culture such rapidly desensitizing
Bgt-sensitive currents are absent in sympathetic neurons (Yu
and Role 1998
) and greatly attenuated in ciliary ganglion
neurons (M. Chen and J. F. Margiotta, unpublished results).
An explanation for the higher Bgt-AChR response density of choroid
over ciliary neurons emerged after deriving rate constants governing
60-pS AChR unbinding (k
n = 80 s
1), opening (
= 95 s
1), and closing
(
= 7470 s
1), and
the predicted probability of being in the open state at high agonist
concentration [po =
/(
+
) = 0.013] from Scheme 2 using ACh as the agonist.
The derived rate constants appear to reflect the process of
Bgt-AChR
activation uncorrupted by desensitization and are quite distinct from
those obtained for other AChRs using the same model-dependent
analytical approaches. In E13,14 ciliary ganglion neurons
(Margiotta and Gurantz 1989
; Margiotta, unpublished
observations), for example, values of
k
2,
, and
were 1,150, 1,275, and 5,230 s
1,
respectively, for 30-pS AChRs (n = 3;
po = 0.20) and 1,380, 1,430, and 1,155 s
1 for 40-pS AChRs
(n = 6; po = 0.54).
For clonal BC3H-1 muscle cells (Sine and Steinbach 1986
)
values of k
2,
, and
were 900, 150-1,200, and 20-50
s
1, respectively
(po = 0.93), while for snake muscle
(Dionne and Leibowitz 1982
; Leibowitz and Dionne
1984
), they were 3,400-6,000, 750-825, and 520-740
s
1
(po = 0.5) and for frog muscle
(Colquhoun and Sakmann 1981
) were 2,000, 14,000 and 40 (po
1.0). Thus when compared with
muscle AChRs or even other neuronal AChRs, the 60-pS
Bgt-AChRs
are unique, displaying the slowest rate constant for agonist
dissociation and opening and the fastest rate constant for channel
closing. These combined factors will result in prolonged dwell times
for 60-pS AChRs in the fully occupied state
(AnR, Scheme 2), when compared with other AChR channels, leading to a very low
po. In spite of this low
po, whole cell currents attributable
to
Bgt-AChRs are substantial on ciliary and choroid neurons (Table
1), suggesting the presence of an adequately large functional receptor
pool on both neuron populations. After normalizing
NF for the threefold larger surface
area of ciliary neurons, the density of functional
Bgt-AChRs is seen
to be ~2.6-fold higher on choroid compared with ciliary neurons. This
value is in excellent agreement with the threefold higher response
density attributable to
Bgt-AChRs on choroid neurons.
Our conclusion that most Bgt-AChRs are silent on both ciliary and
choroid populations is based partly on single-channel kinetic measurements, using 5 µM ACh as the agonist, which predict that NF = 3-24% of
NT. Recent single-channel experiments
using 0.5 µM nicotine predict a qualitatively similar outcome, but in
this case functional receptors represented an even smaller fraction of
NT
(po = 0.08;
NF = 0.4-3.9% of
NT) (Nai and Margiotta, unpublished data). Thus for both agonists, differences in
Bgt-AChR density can
be explained by cell-specific regulation of the fraction of functional
Bgt-AChRs relative to the much larger pool of total (silent + functional) receptors. Since a similar conclusion was previously made
concerning
3*-AChRs (Blumenthal et al. 1999
; Margiotta and Gurantz 1989
), the physiological
implications of silent neuronal AChRs warrant consideration. Recent
studies have shown that lynx1, a small (11 kDa) prototoxin, is
associated with neurons in the rodent cortex, cerebellum and
hippocampus (sites where
7 is also expressed) and displays
structural motifs resembling
Bgt (Miwa et al. 1999
).
Lynx1 was shown to modulate the function of chick
4
2 AChRs
expressed in oocytes with similar effects also reported for
7-AChRs,
suggesting it may recognize endogenous
Bgt-AChRs. Silent AChRs might
serve as targets for lynx1 or similar prototoxins and thereby direct
neurite outgrowth or other cell surface functions important in neuronal
differentiation and/or synapse formation. Another possibility is that
silent receptors represent a reserve that can become functional in
response to cues generated by developmental interactions or by
activation of intracellular signaling pathways. We previously showed,
for example, that silent
3*-AChRs on ciliary ganglion neurons can be
rapidly converted to a functional state by increasing intracellular cAMP (Margiotta et al. 1987b
), and that this ability is
developmentally regulated (Margiotta and Gurantz 1989
).
A similar conversion mechanism may apply for
Bgt-AChRs on the
neurons and for muscle AChRs expressed in Xenopus oocytes,
which can be rescued from entry into a deep desensitized state by
cAMP-dependent protein kinase (PKA) phosphorylation (Paradiso
and Brehm 1998
). Cyclic AMP- and PKA-dependent mechanisms also
enhance Xenopus muscle AChR currents (Fu
1993
; Lu et al. 1993
) and neuronal currents
mediated by glutamate (Greengard et al. 1991
), glycine,
and GABA (Smart 1997
) receptors. In most of these cases,
the enhanced responses have been shown to involve an increase in
receptor po, but the possible
contribution of receptor conversion from silent to functional status
has not been assessed. In the case of ciliary ganglion neurons, AChR
regulation mediated by second-messenger cascades may be relevant in
vivo since receptor function is enhanced after applying
neuropeptides that stimulate cAMP production via adenylate cyclase (AC)
(Gurantz et al. 1994
; Margiotta and Pardi
1995
). One such neuropeptide is pituitary AC activating
polypeptide (PACAP), which is abundant in the ciliary ganglion, and
binds to a receptor that couples through both AC and phospholipase-C
signal cascades (Margiotta and Pardi 1995
; Pardi
and Margiotta 1999
). Because these pathways exert rapid, opposing effects on
Bgt-AChRs (Pardi and Margiotta
1999
), their selective activation by endogenous PACAP could
transiently regulate the degree of mismatch between functional and
nonfunctional
Bgt-AChRs present on choroid and ciliary neurons.
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ACKNOWLEDGMENTS |
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We thank A. Burns, M. Chen, and J. Dittus for expert technical assistance and Dr. Marthe Howard for valuable discussions.
This work was supported by National Institute of Neurological Disorders and Stroke Grant NS-24417 to J. F. Margiotta.
Present addresses: M. E. McNerney, Pharmacia-Upjohn Laboratories, Kalamazoo, MI 49001; D. Pardi, Dept. of Medicine, Mount Sinai School of Medicine, 1 Gustave Levy Place, New York, NY 10029.
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FOOTNOTES |
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Address for reprint requests: J. F. Margiotta, Dept. of Anatomy and Neurobiology, Medical College of Ohio, Block HSB, 3035 Arlington Ave., Toledo, OH 43614-5804 (E-mail: jmargiotta{at}mco.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 14 February 2000; accepted in final form 17 May 2000.
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REFERENCES |
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