Expression and Channel Properties of alpha -Bungarotoxin-Sensitive Acetylcholine Receptors on Chick Ciliary and Choroid Neurons

Mary Ellen McNerney, Desiree Pardi, Phyllis C. Pugh, Qiang Nai, and Joseph F. Margiotta

Department of Anatomy and Neurobiology, Medical College of Ohio, Toledo, Ohio 43614-5804


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

McNerney, Mary Ellen, Desiree Pardi, Phyllis C. Pugh, Qiang Nai, and Joseph F. Margiotta. Expression and Channel Properties of alpha -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: alpha 3*-AChRs recognized by mAb35, that contain alpha 3, alpha 5, beta 4, and occasionally beta 2 subunits, and alpha -bungarotoxin (alpha Bgt)-AChRs recognized and blocked by alpha Bgt, that contain alpha 7 subunits. We found that maximal whole cell current densities (I/Cm) mediated by alpha Bgt-AChRs were threefold larger for choroid compared with ciliary neurons, while alpha 3*-AChR current densities were similar in the two populations. Different densities of total cell-surface alpha Bgt-AChRs could not explain the distinct alpha Bgt-AChR response densities associated with ciliary and choroid neurons. Ciliary ganglion neurons display abundant [125I]-alpha Bgt binding (approx 106 sites/neuron), but digital fluorescence measurements revealed equivalent site densities on both populations. AChR channel classes having single-channel conductances of approx 30, 40, 60, and 80 pS were present in patches excised from both ciliary and choroid neurons. Treating the neurons with alpha Bgt selectively abolished the 60- and 80-pS events, identifying them as arising from alpha Bgt-AChRs. Kinetic measurements revealed brief open and long closed durations for alpha Bgt-AChR channel currents, predicting a very low probability of being open (po) when compared with 30- or 40-pS alpha 3*-AChR channels. None of the channel parameters associated with the 60- and 80-pS alpha 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 alpha Bgt-AChRs (NF) per unit area than do ciliary neurons. Comparison with total surface [125I]-alpha Bgt-AChR sites (NT), reveals that NF/NT 1 for both neuron populations, suggesting that "silent" alpha Bgt-AChRs predominate. Choroid neurons may therefore express a higher density of functional alpha Bgt-AChRs by recruiting a larger fraction of receptors from the silent pool than do ciliary neurons.


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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). alpha -Bungarotoxin (alpha 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). alpha Bgt-AChRs contain predominantly alpha 7-subunits, are recognized and blocked by alpha 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 alpha 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 alpha 3*-AChRs, mediate the slowly decaying EPSC phase and have been estimated to be present at levels ~10-fold lower than alpha Bgt-AChRs (Blumenthal et al. 1999). These receptors contain alpha 3, alpha 5, and beta 4 subunits and occasionally beta 2 subunits (Conroy and Berg 1995; Vernallis et al. 1993) and are neither recognized nor blocked by alpha Bgt but can be detected with a number of alpha -subunit specific antibodies, including mAb35 (Conroy and Berg 1995; Vernallis et al. 1993). Previous single-channel experiments revealed two functional subtypes of alpha 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 alpha Bgt- and/or alpha 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 alpha Bgt- and alpha 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 alpha Bgt-AChRs in a cell-specific manner.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 (theta ) glass tubing (1.6 mm OD, BT-15-10, Sutter Instruments, Novato, CA) pulled to an OD of approx 100-120 µm. The theta  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 theta  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, alpha 3*-AChR currents. Whole cell alpha 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.

Whole cell currents induced by agonist microperfusion were collected at a holding potential of -70 mV using Axopatch amplifiers (1B or 1D; Axon Instruments, Burlingame, CA), filtered at 1-2 kHz, and digitized at 2-4 kHz. Patch pipettes were pulled from Corning 8161 glass capillary tubing and had tip impedances in recording solution of 1-3 MOmega . The patch pipette solution contained (in mM) 145.6 CsCl, 1.2 CaCl2, 2.0 EGTA, 15.4 glucose, 5 Na-HEPES, and 1.0 ATP (pH 7.3). Computer platforms running either BASIC-23 (Digital Equipment, Maynard, MA) or pClamp 6.0 (Axon Instruments) were used for whole cell data acquisition and analysis. Membrane capacitance compensation was achieved by eliminating the capacitive current transient in response to a -10-mV pulse using the amplifier series resistance (Rs) and capacitance (Cm) controls, thereby obtaining a measure of Rs (<= 4 MOmega ). Records were compensated off-line for Rs errors as described previously (Margiotta and Gurantz 1989; Pardi and Margiotta 1999). Rs compensation allowed more accurate measurement of agonist-induced whole-cell currents; however none of the conclusions presented here were changed when results were reanalyzed without its use.

The decay of the agonist-induced current from its peak value is complex and has previously been described as containing a very fast component and one to two slower components (Pardi and Margiotta 1999; Zhang et al. 1994). The decay kinetics were well characterized by fitting the sum of 2 or 3 exponential functions to the digitized current at time t (It) decaying from Ip (t = 0), as previously described (Pardi and Margiotta 1999) and given by
<IT>I</IT><SUB><IT>t</IT></SUB><IT>=</IT><IT>A</IT><SUB><IT>f</IT></SUB><IT> exp</IT>(−<IT>t</IT><IT>/&tgr;<SUB>f</SUB></IT>)<IT>+</IT><IT>A</IT><SUB><IT>i</IT></SUB><IT> exp</IT>(−<IT>t</IT><IT>/&tgr;<SUB>i</SUB></IT>)<IT>+</IT><IT>A</IT><SUB><IT>s</IT></SUB><IT> exp</IT>(−<IT>t</IT><IT>/&tgr;<SUB>s</SUB></IT>) (1)
In Eq. 1, Af, Ai, and As indicate the t = 0 amplitudes of the fast, intermediate, and slow current components, and tau f, tau i, and tau s represent their respective decay time constants. An intermediate component was not obvious in all cells, and three components were assigned only if the standard deviation of the predicted fit (usually 30-60 pA) was lower than that obtained using two components. In experiments involving nicotine, we were most interested in the fast component, and its amplitude (If) was estimated both from the fit value (Af) and by subtracting the summed slower component amplitude(s) (Is±i = As, or Is±i = As + Ai) from the total peak current (Ip). If values presented here were obtained using the latter method and were within 10% of those obtained using the former approach. For ACh applications, alpha Bgt was present to block If, and the whole cell current fit to Eq. 1 with intermediate and/or slow component(s) and associated decay time constant(s). In all experiments, component current values were normalized for neuron size by dividing each value by the membrane capacitance (Cm) determined from the patch clamp controls. The statistical significance (P < 0.05) of differences in any agonist response parameter for ciliary and choroid neurons was determined using Student's two-tailed, unpaired t-test.

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 alpha 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 (gamma ) 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 (tau o). Steady-state open probabilities for each AChR channel conductance class (Popen,x) were calculated using Popen,x = (Sigma t,x/T)/Nx, where T represents the total record length, Sigma 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.

To gain more information about the opening kinetics of alpha Bgt-AChRs, closed-duration distributions were analyzed in more detail for the alpha Bgt-sensitive 60-pS channel class. We assumed alpha Bgt-AChRs (and alpha 3*-AChRs) obey a modified Castillo and Katz activation scheme (Castillo and Katz 1957) such that channels can exist in n +1 closed states (R, A1R, A2R ... AnR), but only 1 open state (AnR*) given by
(2)
In Scheme 2, n is the number of agonist molecules required to bind before the channel can open (n = 2 for alpha 3*-AChRs, and unknown for alpha Bgt-AChRs); k-n and k+n are the agonist binding and unbinding rate constants (with KD k-n/k+n) and beta  and alpha  (alpha  = 1/tau o) represent the transition rate constants governing channel opening and closing. Closed-duration distributions were fit with two exponential functions having brief and long-duration dwell times (tau 1 and tau 2) and analyzed according to the method of Colquhoun and Hawkes (Colquhoun and Hawkes 1981; Colquhoun and Sakmann 1981; Sine and Steinbach 1986), which is based on the assumption that the briefest channel closures (tau 1) represent rapid transitions between AnR and AnR* and that such transitions occur in bursts separated by much longer duration closures having mean dwell times tau 2. Under this scheme, the mean brief closure time (tau 1) will be given by tau 1 = 1/(k-n beta ) and the mean number of brief channel closures per burst (m) by m = beta /k-n.

Fluorescence detection of AChRs

alpha Bgt- and alpha 3*-AChRs were visualized using fluorescence methods modified from Blumenthal et al. (1999). To detect alpha Bgt-AChRs, freshly dissociated E13,14 ciliary ganglion neurons were treated with biotinylated alpha 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 alpha 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 alpha 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 alpha Bgt- and alpha 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.

alpha 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 approx 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 = 2pi a2 + pi (b2/epsilon ) log [(1 + epsilon )/ (1 - epsilon )]} where a and b are the lengths of the major and minor semiaxes, and epsilon  is the ellipticity given by epsilon  = (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]-alpha Bgt at 37°C for 2 h and rinsed three times with RS+hs. [125I]-alpha 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]-alpha 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).


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 approx 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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Fig. 1. Ciliary and choroid neuron can be identified by soma diameter. A: differential interference contrast (DIC) view of 2 neuron cell bodies representative of choroid (left) and ciliary (right) neurons after dissociation from E14 ciliary ganglia. B: size distribution of ciliary ganglion neuron cell body diameters (n = 1280; binwidth 2.2 µm). The histogram was well described by the sum of 2 Gaussian functions (---, r2 = 0.87) predicting mean (± SD) neuron diameters of 13 ± 2 µm and 23 ± 3 µm for the 2 populations. The individual functions are depicted as - - -. C: mixed fluorescence (top) and DIC (bottom) views of a field of dissociated neurons from ganglia where ciliary axons were stained with 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI) to selectively label the ciliary neuron population. Due to the intense DiI fluorescence, the diameter of the labeled neuron appears larger in the fluorescence view than in the accompanying DIC view, from which size measurements were made. The labeled (ciliary) neuron shown (bottom right) has a diameter of 24 µm while that for the unlabeled neuron (top left) is ~14 µm. D: distribution of DiI-labeled (ciliary) neuron cell body diameters (n = 123; binwidth 2.2 µm). The histogram was well described by a single Gaussian function (---, r2 = 0.93) predicting a mean ciliary neuron diameter of 21 ± 3 µm. Note the different magnifications in A and C; scale bars in both represent 25 µm.

Whole cell AChR currents from choroid and ciliary neurons

To maximally activate alpha Bgt- or alpha 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 alpha Bgt-AChRs (Fig. 2) because it provided good temporal separation of alpha Bgt- and alpha 3*-AChR currents (Blumenthal et al. 1999; Pardi and Margiotta 1999) due to the higher affinity of alpha Bgt-AChRs for nicotine (Zhang et al. 1994). To determine if specific alpha 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 alpha Bgt-AChRs since it was completely abolished after treating neurons with alpha 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 alpha 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 alpha 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 alpha 3*-AChR currents were studied in isolation using ACh applied in the presence of alpha 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 alpha 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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Fig. 2. alpha -Bungarotoxin (alpha Bgt)-acetylcholine receptor (AChR) response densities are threefold larger in choroid compared with ciliary neurons. Whole cell records are displayed from representative E14 choroid (A) and ciliary (B and C) neurons identified by the size criteria described in the text and Fig. 1. Neurons were held at -70 mV and exposed to 20 µM nicotine by fast perfusion, as indicated by the bar above each record. Except for the insets in A and B, nicotine-induced responses were normalized for the different sizes of ciliary and choroid neurons by dividing each digitized current value by Cm (pA/pF), and thereby reflect current densities. Time scale below C applies to A-C. The open and filled arrows indicate peak values of the alpha 3*- (Is±i/Cm) and alpha Bgt- (If/Cm) AChR current density components, respectively. Insets: the records at expanded time resolution and scaled to 100% of the total peak current, with calibrations shown at bottom right. Also shown are the respective times to achieve peak response (Tp), fits to Eq. 1 (filled dots), and the associated values of tau f for each neuron. The record in C shows the nicotine response of a ciliary neuron after 1 h treatment with 60 nM alpha Bgt. Note that alpha Bgt abolishes the initial, fast component of the response, but leaves the slow, alpha 3*-AChR-mediated component unaffected. Similar results were obtained for choroid neurons (data not shown). Cell diameters and Cm were, respectively, 7 µm and 9.4 pF in A, 30 µm and 31.7 pF in B, and 27 µm and 28.4 pF in C.


                              
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Table 1. AChR response parameters of choroid and ciliary neurons

We were initially concerned that the difference in alpha 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 theta  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 alpha Bgt-AChR currents to attain peak value (Tp = 4-10 ms) and the time constant describing the alpha Bgt-AChR desensitization (tau 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 alpha 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 alpha Bgt-AChRs on the two neuron populations.

Fast perfusion experiments with nicotine indicated a difference between choroid and ciliary neurons in alpha Bgt- but not alpha 3*-AChR current densities. To further explore this finding, we examined alpha 3*-AChRs in isolation after treating neurons with 60 nM alpha Bgt to block alpha Bgt-AChRs. Identified neurons were tested as in Fig. 2, except that 500 µM ACh was used as the agonist, and alpha Bgt was present (Fig. 3). Under these conditions, alpha 3*-AChRs are activated in isolation from alpha 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 tau i and tau s (Fig. 3, A and B). In the presence of alpha Bgt, the isolated, peak alpha 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 alpha 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 alpha Bgt is specific in blocking If without affecting Is±i. The time required for alpha 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 approx 10 ms and tau s was approx 2 s, while tau 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 tau 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 alpha Bgt-AChRs is threefold larger in choroid compared with ciliary neurons, while that attributable to alpha 3*-AChRs did not differ detectably between the two populations.



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Fig. 3. alpha 3*-AChR response densities of choroid and ciliary neurons are indistinguishable. Whole cell alpha 3*-AChR responses are displayed from representative E13 choroid (A and C) and ciliary (B) neurons. Records in A and B were obtained using fast perfusion as in Fig. 2 except that alpha 3*-AChR responses were isolated by pretreating neurons with 60 nM alpha Bgt and using 500 µM ACh as agonist. The open arrows indicate peak values of the alpha 3*-AChR current normalized to membrane capacitance (Is±i/Cm). Insets: the records at an expanded time base and scaled to 100% of the total peak current with calibrations shown at bottom right. Also shown are the respective times to achieve peak response (Tp), fits to Eq. 1 (filled circles), and the associated values of tau i and tau s for each neuron. Cell diameters and Cm were, respectively, 8 µm and 9.5 pF in A, and 28 µm and 30.0 pF in B. The bar graph in C shows the specificity of alpha Bgt for If over Is±i using either 20 µM nicotine or 500 µM ACh as agonist. If/Cm (black bar) and Is±i/Cm (gray bar) agonist response values obtained from neurons treated with alpha Bgt (50 nM, 1 h) are plotted as percents of the values from untreated controls (100%, solid line ± SE) in the same experiments. For both nicotine and ACh tests, alpha Bgt reduced If/Cm by >90% (P < 0.01) without a detectable effect on Is±i/Cm measured from the same neurons (n = 9-12 alpha Bgt-treated, and 6-14 control neurons for each).

Comparison of alpha 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 alpha Bgt-AChR responses seen for choroid neurons could be explained by a higher overall density of alpha Bgt binding sites expressed on the neuronal cell surface (Fig. 4). Neurons were incubated with biotinylated alpha Bgt and then with Cy3-conjugated streptavidin to visualize alpha 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 alpha Bgt labeling on the two neuronal types. Both ciliary and choroid neurons displayed a nonhomogeneous distribution of alpha Bgt-AChRs at about the same overall surface density. Specific alpha 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 alpha 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 alpha Bgt- and alpha 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 alpha Bgt- or mAb35 net fluorescence density between ciliary and choroid neurons (Fig. 4E). The lack of a difference in alpha 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 alpha 7-AChR response density for choroid over ciliary neurons (Fig. 2; Table 1) cannot be explained by a gross cell-specific difference in total alpha 7-AChR surface density.



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Fig. 4. Distributions of alpha Bgt- and alpha 3*-AChRs on ciliary and choroid neurons. Neurons were dissociated from E13,14 ganglia and labeled with biotinylated alpha Bgt (20 nM) to detect alpha Bgt-AChRs (A) or with mAb35 (100 nM) to detect alpha 3*-AChRs (C), as described in METHODS. Fields containing neurons from nonspecific controls (B and D) were processed as in A and C except that in B neurons received 100 µM D-tubocurarine chloride (dTC) in conjunction with the biotinylated alpha Bgt, while in D mAb35 was omitted from the initial labeling step. Choroid and ciliary neurons are present at the top and bottom, respectively, of each panel. Cells were viewed by fluorescence microscopy and images acquired with the level of focus set at the cell surface. Exposure and filter settings for the images in each pair (A and B and C and D) were identical. Scale bar: 15 µm. E: alpha Bgt-AChR and alpha 3*-AChR densities do not differ on choroid and ciliary neurons. Images were acquired from neurons such as those depicted in A-D and quantified as described in METHODS. For each probe, values represent the mean (± SE) net fluorescence density and thereby provide a basis for comparing the average density of alpha Bgt- or alpha 3*-AChRs on ciliary vs. choroid neurons. Note that while the mAb35 net fluorescence density signal was higher than that for alpha Bgt, there was no significant difference (P > 0.1) in the net fluorescence density of either probe on ciliary neurons () compared with choroid neurons (). Results depicted represent measurements from 15 to 27 ciliary and choroid neurons labeled with alpha Bgt or mAb35 from 2 to 3 separate experiments each.

Surface labeling with [125I]-alpha Bgt reveals an abundance of alpha Bgt sites

We next measured the number of surface alpha Bgt-AChRs present on ciliary and choroid neurons. Freshly isolated E14 ciliary ganglion neurons displayed saturable surface binding of [125I]-alpha 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 alpha Bgt sites revealed 15-20 fmol of alpha 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 alpha 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 alpha Bgt-sites per neuron (n = 3 experiments). Because the density of alpha Bgt sites was equivalent on the two neuron populations (Fig. 4E), we were able to calculate the number of alpha 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 alpha 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 alpha Bgt sites per choroid neuron and 2.41 × 106 alpha Bgt sites per ciliary neuron.



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Fig. 5. Quantifying surface alpha Bgt-AChRs on ciliary ganglion neurons. To determine the total number of surface alpha Bgt binding sites (Bmax) and their affinity (KD) for the toxin, E14 ciliary ganglion neurons were plated and incubated with the indicated concentrations of [125I]-alpha Bgt (A). Nonspecific binding was determined from wells including 1 µM unlabeled alpha Bgt. Aliquots of the incubation mixtures were counted to assure the actual concentrations of free label. The total number and size distribution of the neurons were determined by cell counts and axes measurements in randomly selected fields from parallel wells (see METHODS). Scatchard analysis (A, inset) of the binding isotherm (A) yielded a KD of 0.96 nM and a Bmax of 7.64 fmol/ganglion equivalent (1 ganglion equivalent = 2,300 ch +1,400 ci = 3,700 neurons). Similar results were seen in an additional experiment. To determine the linearity of the assay, differing numbers of ciliary ganglion neurons were plated and subjected to binding with 30 nM 125[I]-alpha Bgt (± 1 µM unlabeled toxin). Specific binding was linear relative to the number of cells plated (B) with a slope of 7.47 fmol/ganglion equivalent (r2 = 0.99). Again, the total number and size distribution of the neurons were determined by cell counts in randomly selected fields in parallel wells.

Single AChR currents from ciliary and choroid neurons reveals four distinct channel classes

A higher alpha 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 alpha Bgt-AChR responses from ciliary neurons (Zhang et al. 1994). We therefore conducted excised patch experiments to determine if the higher alpha 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 (gamma ), 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), gamma  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 gamma  or Er (P > 0.1 for both) for each channel class, suggesting that ion permeation through AChRs is similar in the two neuronal populations.



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Fig. 6. Identification of 4 AChR channel classes on ciliary ganglion neurons. Single-channel currents were collected from excised, outside-out neuron patches exposed to 5 µM ACh. A: 4 AChR channel amplitude classes (1-4) from a representative choroid neuron patch held at -100 mV are displayed. Some events are labeled with arrows in the top panel. These events are displayed at expanded time resolution in the bottom panel traces, except for the smallest event (1), which was obtained from a different record segment. In both panels, note that the largest events (3 and 4) display very brief open durations. Calibration: 4 pA and 40 ms (top), 4 pA and 1 ms (bottom). Currents were sampled at 25-µs intervals and filtered at 6.8 kHz such that events with durations <100 µs were excluded from analysis. B: amplitude histogram for events in the patch depicted in A (n = 787 events; 0.25 pA binwidth). Peaks labeled with arrows correspond to the 4 event amplitude classes. C: current-voltage plot for the same patch with the slope conductances for each amplitude class indicated. Extrapolated current reversal potentials (in mV) for each of the event classes (1-4) were 1.1 (1), -15.2 (2), -9.1 (3), and -13.5 (4). Similar conductance and reversal potential values were obtained from 5 ciliary and 4 choroid patches, and mean values (± SE) are compiled in Table 2. D: open-duration distributions for channel openings corresponding to the 60 (3)-and 80 (4)-pS AChR events are displayed from another choroid neuron patch held at -100 mV. The distributions were well fit (r2 > 0.9 for both) with single-exponential functions predicting open time constants (tau o) of 135 and 217 µs for the 60 (3)- and 80 (4)-pS AChR channel event classes (n > 200 events for each; binwidth 68 µs). Currents were sampled at 34-µs intervals and filtered at 5.8 kHz so that events <115 µs were excluded. Legend applies to both C and D. Similar distributions were obtained from 7 ciliary and 6 choroid patches, and mean tau o values (± SE) for each of the 4 event classes are compiled in Table 2.


                              
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Table 2. Basic AChR channel properties for ciliary and choroid neurons

To further explore potential cell-specific differences in channel properties and more fully characterize alpha 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 (tau 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 (tau o,1 and tau 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 tau o for the 80-pS events (approx 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 approx  0.002) than either 30- or 40-pS events.

alpha 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 alpha 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 alpha 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 alpha 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 alpha Bgt-AChRs, and extends the results from whole cell experiments where 60 nM alpha Bgt blocked the initial, rapidly decaying current density component attributable to alpha Bgt-AChRs but not the slow component attributable to alpha 3*-AChRs. The experiments further suggest that the 30- and 40-pS events arise from alpha 3*-AChRs, which do not contain alpha 7-subunits and are neither recognized nor blocked by alpha Bgt.



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Fig. 7. Channels (60 and 80 pS) are blocked by alpha Bgt. Single AChR channel currents were obtained as in Fig. 6. The AChR channel current amplitude histogram in A is from an excised patch from a neuron treated with alpha Bgt (60 nM; 1 h). Labeled arrows indicate the channel current amplitudes expected for the indicated channel class. Note that the current amplitudes expected for 60- and 80-pS channel classes (3 and 4) were abolished by alpha Bgt, while the relative abundance of events having amplitudes corresponding to the 30- and 40-pS channel classes (1 and 2) were similar to untreated controls (e.g., Fig. 6B). B summarizes the toxin effects. The bar plots show mean Popen values in toxin for the indicated channel classes expressed as a mean (± SE) percent of the value obtained in patches from control neurons not treated with toxin (n = 13). Open triangles and bars indicate controls; filled triangles and bars indicate toxin treatment (n = 6). Asterisks indicate a significant reduction (P < 0.05) in Popen for the indicated channel class in the presence of the toxin compared with untreated controls (100%). Note that alpha Bgt reduced Popen for the 60- and 80-pS channels to 0% of control levels. The toxin did not detectably change Popen for the 30- or 40-pS events (P > 0.1 for both).

alpha 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 alpha Bgt-AChR channel activation (k-n, beta , and alpha  = tau 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 (approx 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 alpha 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 alpha Bgt-AChRs do desensitize rapidly after exposure to high concentrations of agonist (e.g., Fig. 2), alpha 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 alpha Bgt-AChRs are sustained (Zhang et al. 1994; Nai and Margiotta, unpublished data). Third, if 5 µM ACh did cause alpha 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).



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Fig. 8. alpha Bgt-AChR opening kinetics. Results presented are for 60-pS alpha Bgt-AChR events, isolated from all other events, from the patch depicted in Fig. 6. The reconstituted record segment in A shows the timing and mean amplitude for openings (downward deflections) that correspond to the 60-pS channel class in the event amplitude histogram in Fig. 6B (mean ± SD = -5.1 ± 0.8 pA) with the dashed lines delimiting the range of accepted amplitudes. Solid bars above the baseline indicate 3 "bursts" as defined in the text, 1 of which is shown at higher temporal resolution in B. The vertical calibration bar represents 2 pA and applies to both A and B. C: closed-duration distribution with the inset showing all closed durations for the entire 45-s record (n = 359 60-pS events), and the main panel showing only durations <200 ms. The distribution is well described by the sum (solid line) of 2 exponential components (r2 = 0.86), each depicted by dashed lines. The fast component (tau 1 = 6 ms; A1 = 36) represented 26% of the total histogram area (tau 2 = 55 ms; A2 = 15).

According to alpha 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 beta  and alpha . 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 alpha Bgt-AChR channel openings from the same patch (Fig. 8C) indicate these two kinetic components have fast [tau 1 = 1/(beta  + k-n)] and slow (tau 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 = beta /k-n) determined as 1.2. The findings predict values of 87 and 74 s-1 for beta  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 alpha , and po [po = beta /(beta  + alpha ); 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 alpha 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 beta  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.


                              
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Table 3. Kinetic analysis of 60-pS alpha Bgt-AChR channel openings

Choroid neurons would exhibit a higher alpha Bgt-AChR response density than ciliary neurons if they expressed a higher density of functional alpha Bgt-AChRs. This would occur if the number of functional alpha Bgt-AChRs (NF) on both populations was far less than the total number (NT) determined from [125I]-alpha 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 alpha Bgt-sensitive whole cell current (-3,500 and -4,100 pA for choroid and ciliary neurons, respectively; Table 1), i is the single alpha 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 alpha 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, alpha 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 alpha Bgt-AChRs display similar k-n values for different agonists as do muscle-type AChRs (Sine and Steinbach 1986), the approx 10 µM EC50 for alpha 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 left-arrow alpha  right-arrowbeta   AnR* with the probability of being in the open state, AnR*, given by po = beta /(beta  + alpha ) (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 alpha Bgt-AChRs present on ciliary and choroid neurons, respectively, result in choroid neurons expressing a 2.6-fold higher density of functional alpha Bgt-AChRs. Thus the 3.0-fold larger alpha Bgt-AChR response density seen for choroid neurons (Fig. 2, Table 1) can be accounted for by a higher density of functional alpha Bgt-AChRs on this neuron population.

Functional alpha Bgt-AChRs are a small fraction of total alpha Bgt-AChRs on both neuron populations

The total number of alpha Bgt-AChRs per choroid or ciliary neuron (NT) was evaluated from the number of alpha Bgt sites obtained in surface binding studies (e.g., Fig. 5). Since the ratio of sites bound by toxin for each alpha 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 alpha Bgt-AChRs are homopentamers consisting solely of alpha 7 subunits. While each is presumed to have a site recognized by alpha Bgt, alpha 7 subunit monomers do not bind alpha Bgt (Pugh et al. 1995) and for assembled receptors in rat brain neurons, the molar ratio of alpha Bgt binding sites to alpha 7 subunit was 0.2, suggesting that each alpha 7 homopentamer contains only one alpha 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 alpha Bgt competition binding to alpha Bgt-AChRs on intact neurons is approx 1.0 (Vijayaraghavan et al. 1992). This finding also suggests alpha 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 alpha 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 alpha 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 alpha 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 alpha Bgt sites for each alpha Bgt-AChR on ciliary ganglion neurons. Based on this range, and the numbers of surface [125I]-alpha 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 alpha Bgt-AChRs in substantial excess of the numbers required to elicit a maximal current, suggesting that the majority of alpha 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 alpha Bgt-AChRs.


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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 alpha Bgt-AChRs. We determined that nicotinic response densities attributable to alpha Bgt-AChRs are threefold larger on choroid than on ciliary neurons, while those attributable to alpha 3*-AChRs are similar for the two populations. The cell-specific difference in alpha 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 alpha Bgt-AChR response, choroid neurons did not display a detectably higher density of surface alpha Bgt sites determined either by [125I]-alpha 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 alpha Bgt-AChR channels expressed on the two neuronal populations. Instead the findings lead us to conclude that a large pool of surface alpha 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 alpha Bgt-AChR response density can then be explained by cell-specific regulation of the fraction of functional alpha 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 alpha Bgt-AChRs seen here for choroid and ciliary neuron populations.

Analysis of single alpha 3*- and alpha 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 alpha 3*-AChRs contain alpha 3, beta 4, and alpha 5 subunits, but ~20% also contain beta 2 subunits (Vernallis et al. 1993). In contrast, most alpha Bgt-AChRs contain alpha 7 subunits (but not alpha 3, alpha 5, beta 4, or beta 2 subunits), while ~5% of alpha Bgt-AChRs (alpha T/35-AChRs) contain neither alpha 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 alpha Bgt. Since alpha Bgt does not recognize alpha 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 alpha Bgt-AChR class. Based on the co-precipitation findings cited in the preceding text, either the 60- or 80-pS conductance channels could represent alpha 7 subunit homopentamers and the other alpha T/35-AChRs. Since the functional status of alpha Bgt-AChRs lacking alpha 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 alpha 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 alpha Bgt-AChRs in oocytes and cell lines (Chen et al. 1998; Rakhilin et al. 1999) is that both channel classes represent alpha 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 alpha Bgt-AChRs and alpha 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 alpha 3*-AChRs. While this correlation of 30- and 40-pS channels with alpha 3*-AChRs and 60- and 80-pS channels with alpha Bgt-AChRs on the basis of alpha 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 alpha Bgt-AChR channels agree well with the 73-pS AChR channel class attributed to the alpha 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 tau o values for 60- and 80-pS alpha Bgt-AChR channels reported in Table 2. The alpha Bgt-AChR channel conductances we observed also agree well with the 45-pS value obtained for chick alpha 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 alpha 7 subunits expressed in human BOSC 23 cells (Ragozzino et al. 1997). In this case, the multiple conductances are puzzling since only alpha 7 subunits were expressed, suggesting that endogenous AChR-like subunits coassembled with the alpha 7 subunits or that individual alpha 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 alpha Bgt-AChR channels. Somewhat surprisingly, the conductance values we find for alpha Bgt-AChRs in acutely dissociated chick parasympathetic neurons are also different from the 18-pS value reported for alpha Bgt-sensitive, alpha 7-containing AChRs detected on chick sympathetic neurons in culture (Yu and Role 1998). This disparity may reflect cell-specific differences in the alpha 7-subunit content of alpha 7-AChRs on parasympathetic and central neurons versus sympathetic neurons (Yu and Role 1998) or be related to different levels or processing of alpha 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, alpha 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 alpha 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 alpha 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 (beta  = 95 s-1), and closing (alpha  = 7470 s-1), and the predicted probability of being in the open state at high agonist concentration [po = beta /(beta  + alpha ) = 0.013] from Scheme 2 using ACh as the agonist. The derived rate constants appear to reflect the process of alpha 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, beta , and alpha  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, beta , and alpha  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 approx  1.0). Thus when compared with muscle AChRs or even other neuronal AChRs, the 60-pS alpha 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 alpha 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 alpha 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 alpha Bgt-AChRs on choroid neurons.

Our conclusion that most alpha 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 alpha Bgt-AChR density can be explained by cell-specific regulation of the fraction of functional alpha Bgt-AChRs relative to the much larger pool of total (silent + functional) receptors. Since a similar conclusion was previously made concerning alpha 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 alpha 7 is also expressed) and displays structural motifs resembling alpha Bgt (Miwa et al. 1999). Lynx1 was shown to modulate the function of chick alpha 4beta 2 AChRs expressed in oocytes with similar effects also reported for alpha 7-AChRs, suggesting it may recognize endogenous alpha 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 alpha 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 alpha 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 alpha Bgt-AChRs (Pardi and Margiotta 1999), their selective activation by endogenous PACAP could transiently regulate the degree of mismatch between functional and nonfunctional alpha Bgt-AChRs present on choroid and ciliary neurons.


    ACKNOWLEDGMENTS

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.


    FOOTNOTES

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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