Nicotinic Synapses Formed between Chick CiliaryGanglion Neurons in Culture Resemble ThosePresent on the Neurons in Vivo

Min Chen, Phyllis C. Pugh, Joseph F. Margiotta

Department of Anatomy and Neurobiology, Medical College of Ohio, 3035 Arlington Avenue, Toledo,Ohio 43614

Received 30 July 2000; accepted 7 February 2001

ABSTRACT: We studied nicotinic synapses be-tween chick ciliary ganglion neurons in culture to learnmore about factors influencing their formation and re-ceptor subtype dependence. After 4–8 days in culture,nearly all neurons displayed spontaneous excitatorypostsynaptic currents (sEPSCs), which occurred atabout 1 Hz. Neurons treated with tetrodotoxin displayedminiature EPSCs (mEPSCs), but these occurred at lowfrequency (0.1 Hz), indicating that most sEPSCs areactually impulse driven. The sEPSCs could be classifiedby decay kinetics as fast, slow, or biexponential and,reminiscent of the situation in vivo, were mediated bytwo major nicotinic acetylcholine receptor (AChR) sub-types. Fast sEPSCs were blocked bya-bungarotoxin(aBgt), indicating dependence onaBgt-AChRs, most ofwhich are a7 subunit homopentamers. Slow sEPSCswere unaffected byaBgt, and were blocked instead bythe a3/b2-selectivea-conotoxin-MII ( aCTx-MII), indi-cating dependence ona3*-AChRs, which lack a7 and

contain a3 subunits. Biexponential sEPSCs were medi-ated by both aBgt- and a3*-AChRs because they hadfast and slow components qualitatively similar to thosecomprising simple events, and these were reduced byaBgt and blocked by aCTx-MII, respectively. Fluores-cence labeling experiments revealed bothaBgt- anda3*-AChR clusters on neuron somata and neurites. Co-labeling with antisynaptic vesicle protein antibody sug-gested that somea3*-AChR clusters, and a fewaBgt-AChR clusters are associated with synaptic sites, as isthe casein vivo. These findings demonstrate the utility ofciliary ganglion neuron cultures for studying the regu-lation of nicotinic synapses, and suggest that mixedAChR subtype synapses characteristic of the neuronsinvivo can form in the absence of normal inputs ortargets. © 2001 John Wiley & Sons, Inc. J Neurobiol 47: 265–279,

2001

Keywords:acetylcholine receptor; subtype;a-conotoxin-MII; a-bungarotoxin; synapse; cell culture

INTRODUCTION

Nicotinic acetylcholine receptors (AChRs) are widelyexpressed in the vertebrate nervous system, with agrowing list of nine a and threeb subunit genescurrently identified that produce several known homo-and heteromeric receptor subtypes (Sargent, 1993;

McGehee and Role, 1995). In spite of the wide ex-pression and diversity of AChRs, fast excitatory trans-mission mediated by nicotinic receptors was rarelyobserved in the CNS, and thought to be confined tomotoneuron-Renshaw cell synapses in the spinal cord(Curtis and Ryall, 1966; Duggan et al., 1976). Theresulting search for novel functional roles for neuro-nal AChRs has revealed they “modulate rather thanmediate” nicotinic transmission (reviewed by Roleand Berg, 1996; MacDermott et al., 1999), regulateneurite outgrowth and neuron death (Pugh and Berg,1994; Pugh and Margiotta, 2000), and may even in-

Correspondence to:J.F. Margiotta (jmargiotta@mco.edu).Contract grant sponsor: NIH; contract grant number: NS24417

(J.F.M.).© 2001 John Wiley & Sons, Inc.

265

fluence synapse formation (Vetter et al., 1999; re-viewed by Broide and Leslie, 1999; Jones et al.,1999). Recent studies have uncovered a wider CNSdistribution of postsynaptic, AChR-mediated syn-apses than previously thought, however, with nico-tinic transmission now evident in hippocampus(Alkondon et al., 1998; Frazier et al., 1998), nucleusambiguus (Zhang et al., 1993), visual cortex (Roeriget al., 1997), lateral spiriform nucleus (Nong et al.,1999), and retina (Feller et al., 1996).

Because nicotinic AChRs are implicated in severalneurological diseases (reviewed by Lindstrom, 1997;Newhouse and Kelton, 2000), in nicotine-induced ad-dictive/reinforcement behaviors (Dani and Heine-mann, 1996), and in the developmental processesoutlined above, understanding mechanisms relevantto the formation and regulation of interneuronal syn-apses where transmission is mediated by such recep-tors represents a potentially important, yet open ques-tion. Because of their accessibility and simplicity,autonomic ganglion preparations have served as mod-els for nicotinic synapses in the nervous system (e.g.,O’Lague et al., 1978; Willard and Nishi, 1985). Inchick ciliary ganglia, for example, fast excitatorychemical transmission is mediated exclusively by nic-otinic AChRs (Martin and Pilar, 1963), and synapticarchitecture, components, and resultant function allundergo patterned changes influenced by cell-cell in-teractions throughout development (reviewed byLandmesser and Pilar, 1978; Pilar and Tuttle, 1982;Dryer, 1994). By embryonic day 8 (E8) cholinergicinputs from the accessory occulomotor nucleus(Cowan and Wenger, 1968) have functionally inner-vated all neurons in the ganglion. Between E8 andE21 (hatching) synaptic morphology becomes pro-gressively more complex, and transmission efficiencyincreases (Landmesser and Pilar, 1972, 1978; Dryer,1994). During the same developmental period, AChRsubunit transcript and protein levels markedly in-crease (Corriveau and Berg, 1993), and AChR bio-physical properties, functional status, and capacity forregulation by second messengers all undergo dramaticchanges (Margiotta and Gurantz, 1989). Recent stud-ies indicate that two major AChR classes are presentat single functional synapses in mature ganglia(Zhang et al., 1996; Ullian et al., 1997; Chang andBerg, 1999). One AChR class is recognized andblocked bya-bungarotoxin (aBgt-AChRs), which ishighly abundant ('106 surface sites/neuron; McNer-ney et al., 2000), with 95% containinga7 subunits,and lacking any other known subunits (Vernallis etal., 1993; Pugh et al., 1995).aBgt-AChRs (alsotermed a7-AChRs) are present on perisynaptic so-

matic spines (Jacob and Berg, 1983; Shoop et al.,1999), mediate the rapid phase of biexponential exci-tatory postsynaptic currents (EPSCs) in both ciliaryand choroid neurons, and, while not necessary for 1:1transmission (Zhang et al., 1996), are important forsustaining transmission at frequencies as low as 1 Hz(Chang and Berg, 1999). AChRs in the other class(a3*-AChRs) are less abundant (63 104 surfacesites/neuron; Margiotta and Gurantz, 1989), not rec-ognized or blocked byaBgt, lacka7 subunits, and areinstead composed ofa3, a4, a5, and occasionallyb2AChR subunits (Vernallis et al., 1993; Pugh et al.,1995).a3*-AChRs are present at postsynaptic densi-ties and on perisynaptic somatic spines, and mediatethe slowly decaying phase of the EPSC in both ciliaryand choroid neurons, and are essential for unitaryganglionic transmission. Denervation experiments in-dicate that presynaptic inputs and peripheral targetsboth powerfully influence AChR expression in ciliaryganglion neurons (Jacob and Berg, 1988; Arenella etal., 1993). Moreover, properties of both interneuronalnicotinic synapses and AChRs are influenced by thepresence and type of target present in sympatheticneuron cultures (Devay et al., 1999). It is presentlyunknown, however, whether normal inputs and targetsare required for the expression of mixed receptorsynapses such as those mediated byaBgt- anda3*-AChRs on ciliary ganglion neurons.

Studying the formation and regulation of nicotinicsynapses on ciliary ganglion neuronsin situ is prob-lematic because the neurons are innervated early indevelopment, and not easily accessible to direct, long-term manipulation. Such studies would be simplifiedby a reliable cell culture system, and ciliary ganglionneurons can, in fact, be maintained for weeks on asubstratum of lysed fibroblasts in dissociated cell cul-ture without target cells when grown in medium con-taining 3% (v/v) embryonic eye extract (Nishi andBerg, 1981). By both morphological and electrophys-iological criteria, nicotinic synapses form between theneurons in such cultures (Crean et al., 1982; Margiottaand Berg, 1982), but the fibroblast substratum is fre-quently unstable on the glass coverslips preferred inelectrophysiological experiments. To study nicotinicsynapses present on the neurons in more detail, weimproved the culture system by coating glass cover-slips with polyornithine and laminin (Bruses et al.,1995; see Methods). The neurons adhere and extendneurites on this substrate for up to 2 weeks, and wenow report that interneuronal synapses formed after'4–8 days in culture closely resemble those presentat an equivalent developmental agein vivo. In partic-ular, we report that EPSCs mediated byaBgt- and

266 Chen et al.

a3*-AChRs are retained, suggesting that the forma-tion and/or maintenance of such mixed-receptor syn-apses can occur in the absence of normal muscletargets or preganglionic inputs.

METHODS

Cell Culture

Ciliary ganglion neuron cultures were prepared and main-tained under sterile conditions as previously described(Margiotta et al., 1987a,b), except that a polyornithine/laminin substratum was used (Bruses et al., 1995), allowingbetter neuron adhesion to glass coverslips. First, 12 mmdiameter, #1 glass coverslips (Fisher Scientific, Pittsburgh,PA) were acid-washed, incubated for 16 h at 4°C with 0.13M borate buffer (pH 8.0) containing 0.2 mg/mL poly-DL-ornithine (#P0671, Sigma, St. Louis, MO). They were thenwashed with distilled water. The coated coverslips werethen each treated for 4 h at 37°C with 150mL minimumessential medium (MEM; # 11090-08, GIBCO BRL, Rock-ville, MD; see below) containing 0.03mg/mL mouse lami-nin (Collaboarative Biomedical, Bedford, MA). Lamininstocks were prepared at 1.33 mg/mL in 0.05M Tris/0.15MNaCl (pH 7.4), frozen in 50mL aliquots, and diluted inMEM immediately before use. After coating, the polyorni-thine/laminin-coated coverslips were washed 43 withMEM, placed in 16 mm culture wells, and covered with 400mL complete culture medium. The complete culture me-dium consisted of MEM that was supplemented with 100U/mL penicillin, 100 mg/mL streptomycin, 2 mM glu-tamine, 10% (vol/vol) heat inactivated horse serum (allcomponents from GIBCO-BRL), and 3% embryonic chickeye extract (Nishi and Berg, 1981). Ciliary ganglia weredissected from stage 34–35 (embryonic day 8) chick em-bryos (Hamburger and Hamilton, 1951), hemisected, di-gested with 0.05% trypsin, and the neurons dissociated bymechanical trituration. The dispersed neurons were plated ata density of one to two ganglion equivalents per coatedcoverslip. Coverslip cultures were maintained in the 16 mmwells at 37°C in 95% air, 5% CO2, and received freshculture medium every 2–3 days.

Electrophysiology

After 1 to 8 days, coverslip cultures were moved to the stageof an inverted microscope (Axiovert 135, Carl Zeiss, Thorn-wood, NY) where neurons were visualized with differentialinterference contrast optics, and whole-cell recordings wereobtained at 21–24°C using patch clamp methods previouslydescribed (Margiotta et al., 1987a,b). The recording solution(RS; derived from Dichter and Fischbach, 1977) contained(in mM): 145.0 NaCl, 5.3 KCl, 5.4 CaCl2, 0.8 Mg(SO4), 5.6glucose, and 5.0 mM HEPES (pH 7.4), and was sometimesfurther supplemented with 10% horse serum (RS1hs). Theinclusion of horse serum did not detectably affect any of the

results presented here. Patch pipettes were fabricated fromglass capillary tubing (Corning 8161, WP Instruments,Tampa, FL) and had tip impedances of 2–3 MV when filledwith (in mM): 145.6 CsCl, 1.2 CaCl2, 2.0 EGTA, 15.4glucose, and 5 Na-HEPES (pH 7.3). Membrane currentswere collected from neuron somata using Axopatch 200B or1D patch clamps (Axon Instruments, Burlingame, CA), andwere digitized, usually at 100ms intervals, using a CheshireData interface (Indec Systems, Inc., Sunnyvale, CA) and alaboratory computer equipped with an LSI-11/73 processor(Digital Equipment Corp., Maynard, MA). After establish-ing the whole-cell recording configuration, Na1 and Ca12

currents were evoked by 17.5 ms step depolarizations to240 through130 mV in 15 mV increments from theholding potential (typically270 mV). Neurons were thenheld at 270 mV without stimulation for 2–5 min whilespontaneous EPSCs were collected. During data acquisition,only those portions of the current record containing synapticcurrents exceeding a preset amplitude threshold (usually210 pA) were collected.

The sEPSCs were analyzed off-line using modifiedBASIC-23 programs originally written for single AChRchannel currents (provided by Dr. Vince Dionne, MarineBiological Laboratory, Woods Hole, MA). For initial anal-ysis, EPSC peaks were selected and the time of occurrencefor each stored, and used to determine sEPSC incidence andfrequency. Records with fewer than two sEPSCs were givenan incidence value (I1) of zero, and those with two or moreevents an incidence value of one. The cumulative sEPSCfrequency (F1) was determined for sEPSC positive neurons(I1 5 1) from the total number of events divided by theelapsed time between first and last events. In some cases,the occurrence of events was examined for stationarity bycounting the number of events in successive time bins of15–30 s. Because many sEPSCs occurred on the fallingphase of the previous event, only the occurrence times ofeach detectable event peak were measured in every record.In records where the sEPSCs were well separated, selectedevents were then reanalyzed to quantify their peak ampli-tudes and decay kinetics. Some sEPSCs appeared to decaymonoexponentially from peak, and obey simple fast or slowkinetics with the current at timet after peak (I t) predictedby

I t 5 Af exp~ 2 t/tf! or by It 5 As exp~ 2 t/ts!. (1)

The decay phases of other sEPSCs were clearly more com-plex, and appeared to require fast and slow (biexponential)kinetics, withI t predicted by

I t 5 Af exp~ 2 t/tf! 1 As exp~ 2 t/ts!. (2)

Thus, sEPSC decay phases were quantified by repetitivelydisplaying Equation 1 or 2 over the fast and/or slow decayphase(s) of each event, while manually adjusting the (t5 0) amplitude (Af and/orAs) and time constant (tf and/or

Neuronal Nicotinic Synapses in Culture 267

ts) parameters to achieve the best fit to the data. The list ofkinetic parameters for each selected event in a record wasthen transferred to a Power Macintosh 850 computer usingVersaterm Pro (v. 2.1, Synergy Software) and commercialsoftware used for statistical analysis (Excel 98, Microsoft)and construction and fitting of histograms (Kaleidagraph v.3.5, Synergy). Differences in parameter values betweencontrol and test neuron populations were evaluated forstatistical significance (p , .05) using Student’s unpaired,two-tailedt test. All replicate values are reported in the textas the mean6 S.E.M.

As mentioned above, the decision to treat the decay ofindividual sEPSCs as mono- or biexponential, and the sub-sequent evaluation of the “goodness-of-fit” relied on visualcriteria. To evaluate the suitability of this approach, 26events, visually identified as fast, slow, or biexponential,were refit with mono- and biexponential functions (data notshown) using nonlinear regression and the Levenburg-Mar-quardt optimization algorithm (Prism v. 2.0, GraphPad).TheF-test was then employed to determine which functionbest described the current decay. For sEPSCs where thedecay was visually classified as monoexponential (n 5 7fast and 9 slow events), only one of the fits from each wassignificantly improved (F . 10; p , .05) using thebiexponential function. For sEPSCs visually classified asdecaying biexponentially (n 5 10), all but onewere sig-nificantly improved using the biexponential function. Theone case where the fit of a visually classified biexponentialsEPSC was not significantly improved by fitting with thebiexponential function had a clear fast component whichwas presumably too small ('10% of peak) for the algorithmto detect. Thus, the visual- and algorithm-based classifica-tions of sEPSCs as mono- or biexponential were in gener-ally good agreement. The two approaches were also similarin assessing goodness-of-fit because they generated kineticparameter values (tf, Af, ts, and As) that differed, onaverage, by only 4–19% in the 26 events studied. Thesetests validate our approaches for discriminating events asmono- or biexponential and for estimating the parametervalues that describe sEPSC decay kinetics.

Visualizing Components of NicotinicSynapses

AChRs and presynaptic terminals were visualized in ciliaryganglion neuron cultures grown for 5–6 days using conven-tional fluorescence methods (e.g., McNerney et al., 2000).To detectaBgt-AChRs, cultures were first incubated for24 h in medium containing unconjugated streptavidin (SA,6 mg/mL), then washed (33 with RS1hs), treated withbiotinylatedaBgt (20 nM; Molecular Probes, Eugene, OR)in RS1hs at 37°C for 2 h, and washed again. The cultureswere then incubated with Cy3-conjugated streptavidin (2mg/mL; Jackson Laboratories, Bar Harbor, ME) in RS1hs

for 45 min at room temperature (RT). Cells were thenwashed, fixed (Histochoice, Amresco, Solon, OH) for 20min, and rinsed with PBS. To detecta3*-AChRs, neurons

were treated with mAb35 (100 nM; a generous gift from Dr.D.K. Berg, University of California, San Diego) in RS1hs

containing 17% rabbit serum at RT for 1.5 h, and thenwashed (33 in RS1hs). Cells were then incubated withbiotinylated-SP-conjugated rabbit antirat IgG (5mg/mL,Jackson Laboratories) in RS1hs for 45 min at RT, washed,incubated with Cy3-conjugated streptavidin (2mg/mL) inRS1hs, washed, and then rinsed once with PBS. Cells werethen fixed for 20 min and rinsed with PBS as above foraBgt. After staining with either AChR probe, presynapticterminals were labeled by first blocking cultures with PBScontaining 10% normal goat serum (NGS) and then incu-bating with the anti-SV2 antibody (mAb 10h, Developmen-tal Studies Hybridoma Bank, University of Iowa) in PBScontaining 4% NGS (PBS-NGS) overnight at 4°C. Cellswere then rinsed with PBS containing 0.3% TX-100 (PBS-TX) and incubated with Alexa Fluor 488 conjugated goatantimouse IgG (Molecular Probes) in PBS-NGS for 1 h atRT. After washing with PBS-TX, the coverslips weremounted on glass slides using Vectashield (Vector Labora-tories, Burlingame, CA).

Neurons were examined with DIC and reflected lightfluorescence using an Olympus BX50 microscope equippedwith a UplanFL 403 objective (0.75 N.A.) and opticsappropriate for detection of Cy3 or Alexa Fluor 488 fluoro-phores. After focusing on the neuron (usually at or near thetop surface), 16 bit DIC and fluorescence images wereacquired using a cooled digital CCD camera (SenSys,Model KAF-1400, Photometrics, Tucson, AZ) under thecontrol of IP Lab software (Version 3.0, Scanalytics, Read-ing, PA). Digitized images were prepared for presentationusing Photoshop 5.0 (Adobe Systems, San Jose, CA) afterconversion to TIFF format, and output to a Kodak DS8650PS color printer (Rochester, NY).

Materials

Fertilized white Leghorn chicken eggs were obtained fromHertzfeld Poultry Farms (Waterville, OH) and maintained at37°C in a forced air draft incubator at 100% humidity. Mostreagents were purchased from Sigma. Sera, biotinylatedIgG, etc., were obtained from Jackson Biolabs.

RESULTS

Spontaneously Active NicotinicSynapses

Functional nicotinic synapses are readily detected onciliary ganglion neurons grown in dissociated cellculture. Whole-cell recordings obtained at270 mVroutinely revealed spontaneous inward currents thattended to occur in bursts [Fig. 1(A,B)]. For E8 neu-rons maintained in culture for 4–8 days, the incidenceof neurons displaying such currents (I1) was 89

268 Chen et al.

6 2% (n 5 257), and thecurrents occurred inpositive cells at an overall average frequency [F1;e.g., Fig. 1(B)] of 0.756 0.06 Hz (n 5 228).

Reversal potential estimates and pharmacologicaltests both indicated that the currents are mediated bynicotinic AChRs. Consistent with the'210 mV re-versal potential for single AChR channels on ciliaryganglion neurons in culture (Margiotta et al., 1987b),individual spontaneous inward current amplitudes be-came progressively smaller as the holding potentialwas changed from2100 to 270 mV, and were un-detectable between220 and 0 mV [e.g., Fig. 1(C)].Tubocurarine (dTC, 25mM) a competitive antagonistfor nicotinic AChRs, blocked the currents to nearcompletion, reducingI1 to nearly zero, andF1 in theone positive cell by about fivefold, relative to un-treated controls tested in parallel [Fig. 2(A,D)]. Incontrast, bath perfusion with atropine (ATR, 1mM), amuscarinic AChR antagonist, had no detectable effecton eitherI1 or F1 compared to the levels recorded inthe same neurons before treatment [Fig. 2(B,D)].These findings mirror results obtained previously us-ing sharp-electrode voltage recordings (Margiotta andBerg, 1982) and indicate that the inward currentsrepresent spontaneous excitatory postsynaptic cur-rents (sEPSCs) that arise from nicotinic AChR-medi-ated synapses formed between the neurons in culture.

In spite of their spontaneous appearance, the bulkof sEPSCs required impulse-driven release of ACh[Fig. 2(C,D)]. In the presence of tetrodotoxin (TTX, 1mM) to block presynaptic impulse activity, the inci-dence of neurons displaying spontaneous miniatureEPSCs (mEPSCs) was 806 9% (n 5 20), avalueindistinguishable from that for sEPSCs in controlneurons tested in parallel (n 5 23; p . .1). ThemEPSC frequency in positive, TTX-treated neurons(0.116 0.05 Hz;n 5 16) was reduced, however, to'41% of that for sEPSCs obtained from controlstested in parallel. The low mEPSC frequency is sim-ilar to that observed for newly hatched chicks ininvivo where transmission is quantal, and is character-ized by a low level of spontaneous release (Martin andPilar, 1964). This similarity indicates that, withoutimpulse activity, terminals of ciliary ganglion neuronsin culture, like those of the normal preganglionicinputs, release ACh quanta at a low basal rate. TTXtreatment also altered the distribution of sEPSC am-plitudes [Fig. 2(E)]. For control neurons, the sEPSCamplitudes ranged from about210 to 280 pA, hav-ing an overall mean of227 6 2 pA (n 5 15). Thedistribution of sEPSC amplitudes appeared multimo-dal, with an initial peak usually apparent at211 to219 pA (mean5 215 6 1 pA; n 5 11). Incontrast,and consistent with quantal release, EPSC amplitudesrecorded from TTX-treated neurons were distributedaround a unimodal value of210 to 216 pA (mean

Figure 1 Ciliary ganglion neurons in culture display spon-taneous synaptic currents. (A) Synaptic currents are shownfor a neuron grown for 5 days in culture. The three tracesrepresent noncontiguous segments of the same data recordacquired without stimulation, at a holding potential of270mV. Note that the currents are heterogeneous in peak am-plitude and decay kinetics. (B) Frequency histogram con-structed using peak current occurrence time data from therecord in (A). Note that the numbers of events in each 15 sbin are unequal, indicating the currents occurred in bursts.The dashed line indicates the overall average spontaneoussynaptic current frequency (F1) for this neuron (0.9 Hz).(C) Reversal potential estimation. Spontaneous currentswere recorded from another neuron held at three differentvoltages. Note that the synaptic currents tend to be large at2100 mV (top), smaller at270 mV (middle), and unde-tectable at 0 mV (lower), indicating they reverse near 0 mV.Calibration bars: 25 pA and 20 ms in (A); 25 pA and 100 msin (C).

Neuronal Nicotinic Synapses in Culture 269

5 213 6 1 pA; n 5 4). This mean EPSC amplitudein TTX was indistinguishable from that of the firstapparent peak in the multimodal amplitude distribu-tion typical of control neurons (215 pA; p . .1), butabout half that of the overall mean sEPSC amplitudefor control neurons (227 pA; p , .05). Taken to-gether, these results indicate that the sEPSCs ob-served in culture are mediated by nicotinic AChRs,with most depending on presynaptic impulses, and aminor TTX-resistant fraction likely to representmEPSCs that arise from spontaneous impulse-inde-pendent, ACh release.

Time Course of sEPSC Appearance inCulture

sEPSCs, indicative of functional nicotinic synapseformation, were first detected in about 40% of neu-rons tested after 1 day in culture. Between days 1 and2, however,I1 had increased to 70% and remained at80–100% for the duration of the period examined[through day 8; Fig. 3(A)]. The frequency of sEPSCsin positive neurons (F1) was very low at day 1 (0.03Hz), but increased steadily to reach a plateau betweendays 4 and 6 ('0.6 Hz), and again between days 7 and8 ['1 Hz; Fig. 3(B)]. The observation that mostneurons failed to score as EPSC-positive after 1 day inculture but did so after 2 days coincided with theinitial appearance and elaboration of neurite contactsarising from adjacent neurons in the culture [compareFig. 3(A,C–E)]. During the next few days, both thedensity of the neuritic arbor andF1 increased steadily[compare Fig. 3(B,F–H)]. Thus, the developmentalincrease inI1 andF1 after the first 2 days in culturemay reflect increased levels of innervation of individ-ual neurons by multiple inputs, a greater frequency ofrelease from single inputs, or a combination of theseeffects.

Individual sEPSCS DisplayHeterogeneous Decay Kinetics

As noted above, sEPSCs recorded from ciliary gan-glion neurons after 4–8 days in culture usually oc-

Figure 2 EPSC pharmacology. (A–C) sEPSCs recordedfrom ciliary ganglion neurons in the presence of: (A) 100mM d-tubocurarine (1dTC), (B) 1 mM atropine (1Atro-pine), or (C) 1mM tetrodotoxin (1TTX) (lower traces) aredepicted. Control records (top traces) were obtained fromthe same neuron before perfusion with the drug (B) or fromneurons in the same culture not exposed to the drug (A, C).Calibration bars apply to (A–C) [horizontal5 50 ms; ver-tical 5 20 pA for (A) and (C), 40 pA for (B)]. (D) Summaryof drug effects. Open bars indicate sEPSC incidence (I1);shaded bars represent mean sEPSC frequency (F1), bothexpressed as percentages (6 S.E.M.) of mean values ob-tained from controls within the same experiments. Asterisksindicate a significant difference (p , .05, by Student’sttest) from control values. Results in (A–D) were compiledfrom 8–23 neurons in two to four separate experiments. (E)TTX alters the distribution of sEPSC amplitudes. The am-plitude distribution was typically broad and multimodal incontrol neurons (solid line), but narrower and unimodal inneurons treated with TTX (dashed line). Fitting the controlamplitude distribution depicted (n 5 258 events) requiredmultiple Gaussian functions predicting modal values of211, 217, and224 pA (r2 5 0.99). By contrast, theamplitude distribution in TTX (n 5 102 events) was welldescribed by one Gaussian function predicting a single

mode at 212 pA (r2 5 0.92). For both control andTTX-treated neurons, sEPSCs having amplitudes smallerthan the acquisition threshold (210 pA) were present. Thesesmall events were captured because they occurred in datasegments with suprathreshold events. Similar results wereobtained from a total of 11 control and four TTX-treatedneurons.

270 Chen et al.

curred in bursts of individual and overlapping events.In some records, a substantial number of sEPSCsoccurred in isolation, and selected sEPSCs from thesewere studied for amplitude and kinetic information byfitting exponential functions to the decay of the cur-rent from its peak value (see Methods). IndividualsEPSCs analyzed in this way from seven controlneurons were identified as monoexponential (fast orslow) or biexponential (fast and slow), according toobvious differences in the kinetics of current decay[Fig. 4(A)]. Monoexponential sEPSCs displayed sim-ple kinetics, with the current decay from peak valueadequately described by a single fast or slow expo-nential function. Monoexponential fast and slow sEP-SCs comprised 106 3% and 526 6% of analyzedevents, respectively, and were clearly distinguishable,

because their associated time constants differed by'10-fold (tf 5 0.50 ms andts 5 5.25 ms,Table 1).Biexponential sEPSCs represented 386 4% of eventsanalyzed, and featured an initial rapidly decayingcomponent followed by a slowly decaying compo-nent, each described by distinct time constants (tf

5 0.30 ms andts 5 6.88 ms). The fast and slowdecay time constants obtained here are similar tothose previously reported for evoked EPSCs (Ullian etal., 1997) recorded from ciliary and choroid neuronsin vivo (tf ' 1 ms andts ' 10 ms). In the cultures,the fast and slow time constants describing biexpo-nential sEPSC decay were also qualitatively similar tothose associated with simple fast and slow sEPSCs,respectively, yet the small differences observed weresignificant (p , .05 for each). For biexponential

Figure 3 Time course of sEPSC appearance and elaboration of neuronal processes. (A) Timecourse of sEPSC incidence (I1). Note that nearly all neurons displayed sEPSCs after 2 days inculture. (B) Time course of sEPSC frequency (F1). Note that the sEPSC frequency increased fromnear 0 Hz at 1 day to'0.5 at 4 days, and then increased to a second plateau of'1.0 Hz between6 and 8 days.I1 andF1 values plotted represent cumulative measurements obtained from 9–99neurons in 2–24 cultures for each time point. (C–H) Appearance of neurons at plating (C), and after1 (D), 2 (E), 4 (F), 6 (G), and 8 (H) days in culture. Note the extensive elaboration of neuronalprocesses between days 2 and 4 overlaps with the initial rise inF1 occurring in the same period.Scale bar5 50 mm.

Neuronal Nicotinic Synapses in Culture 271

slow sEPSC components and simple slow sEPSCs,however, the difference in decay kinetics was lessthan 25%, and respective peak amplitudes were notdetectably different. In contrast, the fast componentsof biexponential sEPSCs were twofold smaller anddecayed 40% faster than simple fast sEPSCs (Table

1). The smaller amplitude and faster decay of fastcomponent biexponential sEPSCs compared to simplefast sEPSCs are likely to result from partial maskingof the fast decay phase of biexponential events causedby near-simultaneous slow component activation. Themasking effect results in few points contributing to

272 Chen et al.

the initial fast component falling phase, and wouldtherefore be expected to reduce the efficiency of curvefitting in predicting the fast component’s time courseand amplitude. Apart from this limitation, the simi-larities between simple fast and slow sEPSC ampli-tudes and kinetics and those of the associated com-ponents of biexponential sEPSCs suggest that thelatter represent temporally overlapping fast and slowmonoexponential events. Because biexponential andsimple slow events together comprise 90% of allsEPSCs, such an arrangement predicts that sEPSCshaving a slowly decaying current component predom-inate in the cultures.

Fast and Slow sEPSC DecayComponents Are Mediated by aBgt- anda3*-AChRs

It is unlikely that sEPSCs having fast, slow, andbiexponential decay kinetics are mediated by a singleAChR population. One way such an arrangementmight explain the present findings, however, would beif some sEPSCs recorded in the soma were evoked atdistant, unclamped regions where they activated Na1

channels resulting in the spread of voltage-dependentcurrents into the soma. We tested this possibility byadding QX-314 (lidocaineN-ethyl bromide, 20 mM)

Figure 4 sEPSC heterogeneity and dependence onaBgt- anda3*-AChRs. (A) Heterogeneity insEPSC decay kinetics. The distributions of decay time constants associated with monoexponential(fast or slow) sEPSCs (black or open bars, respectively) and with biexponential sEPSCs having bothfast and slow components (shaded bars) are depicted. The histogram was constructed using a binwidth of 100ms, from a recording where sEPSCs were sufficiently separated to determine decaytime constants from 214 nonoverlapping events. Similar histograms were constructed from fiveother recordings (n 5 147 6 24 events). Theinsetdepicts examples of monoexponential fast andslow sEPSCs and a biexponential sEPSC, with the dashed lines indicating the fitted exponentialfunctions, with associated time constant(s) included. Calibration: 10 pA, 5 ms. (B) Distribution ofdecay time constants and exemplar remaining events (inset) after treating cultures with 60 nM aBgtfor 1 h to block aBgt-AChRs. (C) Distribution of decay time constants and exemplar remainingevents (inset) after treatment with 25 nM aCTx-MII for 10 min to blocka3*-AChRs. Calibrationbars: 10 pA (vertical) and 50 ms (horizontal) in (B) and (C). (D) Summary of toxin effects on sEPSCincidence and frequency. Open bars indicate sEPSC incidence (I1); shaded bars represent meansEPSC frequency (F1), both expressed as percents (6 S.E.M.) of mean values obtained fromcontrols within the same experiments. Asterisks indicate a significant difference (p , .05) fromcontrol values. Results were compiled from 14–17 toxin-treated neurons and 14–23 control neuronsin four separate experiments.

Table 1 Effects ofaBgt and aCTx-MII on Fast, Slow, and Biexponential EPSCs

Condition

Fast EPSCs Slow EPSCs Biexponential EPSCs

Ff

(Hz)tf

(ms)Af

(pA)Fs

(Hz)ts

(ms)As

(pA)F(f1s)

(Hz)tf

(ms)Af

(pA)ts

(ms)As

(pA)

Control 0.08 0.50 217.7 0.44 5.25 220.5 0.33 0.30 29.1 6.88 216.4(0.03) (0.05) (1.0) (0.09) (0.43) (1.8) (0.06) (0.02) (0.6) (0.58) (1.3)

aBgt 0.00* — — 0.48 5.38 218.4 0.08* 0.34 27.6 8.24 213.8(0.00) (0.21) (0.38) (1.1) (0.04) (0.03) (0.6) (0.80) (1.1)

aCTx-MII 0.02 0.39 215.0 0.02* 2.54* 219.2 0.02* 0.27 214.6* 6.62 28.1*(0.01) (0.12) (3.7) (0.00) (1.21) (1.0) (0.01) (0.07) (1.3) (2.82) (1.3)

Subscriptedt andA values indicate sEPSC decay time constants and amplitudes, respectively. Peak amplitudes are depicted for fast andslow monoexponential sEPSCs, while fast and slow component amplitudes obtained from curve fits are depicted for biexponential sEPSCs.Ff, Fs, andF( f1s) indicate the frequencies of unitary fast, slow, or biexponential sEPSCs that were analyzed fort and A in each record(duration5 117–172 s). Note that for control neurons, the sum ofFf, Fs, andF( f1s) (0.85 Hz) differs slightly from the overall event frequency(F1 5 0.75 Hz)because the former represents only the well-separated, unitary events in a record, while the latter includes all observed events(see Methods). Parameter values from seven control and sixaBgt-treated neurons are expressed as the mean with S.E.M. in parentheses.Results from twoaCTx-MII treated neurons are expressed as the mean with S.D. in parentheses. The asterisks indicate a significant difference( p , .05) in theparameter value in toxin-treated compared to control neurons.

Neuronal Nicotinic Synapses in Culture 273

to the patch pipette solution, and allowing the drug todiffuse for 10 min to intracellularly block Na1 chan-nels, at sites near to and distant from the soma (Col-ling and Wheal, 1994). In previous studies LuciferYellow, injected into the soma, filled thin neurites.200 mm distant within 10 min (Margiotta, unpub-lished results). Somatic Na1 currents were blockedfollowing the QX-314 treatment, but sEPSCs re-corded there were unaffected. sEPSCs in six treatedneurons occurred at frequencies that were indistin-guishable from those of controls, and neither the pro-portions of events classified as fast, slow, or biexpo-nential, nor their individual kinetic parameters (tf, Af,ts, andAs) were changed (data not shown). Thus, theheterogeneity of sEPSC decay kinetics is unlikely toarise due to limitations in space clamp, indicatinginstead that synapses in the cultures arise from differ-ential contributions of at least two distinct AChRclasses. In fact, botha3*- and aBgt-AChRs areknown to participate in transmission at somatic syn-apses on ciliary ganglion neuronsin vivo, and therelative contribution of each has been estimated bycomparing evoked mono- and biexponential EPSCdecay times and amplitudes in the presence of toxinsthat block one (Zhang et al., 1996) or both (Ullian etal., 1997) AChR classes. Such studies have demon-strated thataBgt blocks the fastaBgt-AChR-medi-ated component of biexponential EPSCs. In contrast,a-conotoxin-MII (aCTx-MII), a snail toxin recogniz-ing mammaliana3/b2 AChRs expressed in oocytes(Cartier et al., 1996), blocks the slow component ofthe biexponential evoked EPSCs and the monoexpo-nential slow evoked EPSC (both mediated bya3*-AChRs). Our recent findings demonstrate four func-tional AChR channel classes on ciliary ganglionneuronsin vivo having unitary conductances of'30,40, 60, and 80 pS, with the relatively rare 60 and 80pS channel classes selectively blocked byaBgt (Mc-Nerney et al., 2000), and the predominant 40 pS classselectively blocked byaCTx-MII (Nai et al., 2000).Using 1 mM ACh, the 60 and 80 pS channels dis-played uniformly fast kinetics characterized by briefduration unitary openings of 100–200ms, while boththe 30 and 40 pS channels displayed mixed kineticscharacterized by brief and long duration openings ofabout 100ms and 2 ms (McNerney et al., 2000). Inorder to determine the relative contribution of differ-ent AChR subtypes to synaptic transmission in cul-ture, the effects of 60 nM aBgt or 25 nM aCTx-MIIon individual fast, slow, and biexponential sEPSCfrequencies, amplitudes, and decay rates were exam-ined [Fig. 4(B–D), Table 1]. We also assessed toxineffects on the more general parameters,I1 and F1.

aBgt treatment abolished simple fast sEPSCs (Ff

5 0) and reduced the observed frequency of biexpo-nential sEPSCs (Ff1s) by about fourfold, withoutdetectably changing the frequency of slow sEPSCs[Fs; Fig. 4(B), Table 1]. These effects are consistentwith aBgt-AChRs contributing to the fast currentdecay of both simple fast and biexponential sEPSCs.The toxin treatments also resulted in nominal reduc-tions in the amplitude of slow sEPSCs and in the slowcomponent amplitude of the few remaining biexpo-nential sEPSCs (Table 1). Because sEPSCs having aslow component comprise 90% of all events, andthese were not significantly altered byaBgt, the tox-in’s effects on overall sEPSC incidence and frequencywere minor. Thus, inaBgt-treated cultures,I1 wasnot detectably changed, andF1 was reduced by'40% when compared to untreated neurons tested inparallel [Fig. 4(D)]. Contrasting with the relativelyminor effects ofaBgt, aCTx-MII abolished nearly allslow and biexponential sEPSCs, but had no detectableeffect on fast sEPSCs [Fig. 4(C)]. Specifically,aCTx-MII reduced the frequency of slow and biexponentialsEPSCs, both by about 20-fold compared to untreatedcontrols, without significantly changing the frequencyof fast sEPSCs (Table 1). Because slow and slow-component biexponential events comprise nearly allsEPSCs, the effects ofaCTx-MII on overall sEPSCincidence and frequency were dramatic, resulting in athreefold reduction inI1 and a 20-fold reduction inF1 [Fig. 4(D)]. Consistent with its ability to nearlyabolish slow and slow-component biexponentialevents,aCTx-MII reduced the amplitude of the re-maining slow component biexponential sEPSCs,nominally reduced the amplitude of the remainingsimple slow sEPSCs, and decreased their decay timeconstant (Table 1). Taken together, the toxin studiesindicate that the slow EPSCs are mediated bya3*-AChRs, fast sEPSCs are mediated byaBgt-AChRs,and biexponential sEPSCs depend on both AChRclasses. Synapses on the neurons in culture thus ap-pear qualitatively similar to those present on the neu-ronsin vivo in the context of the two nicotinic AChRclasses that mediate nicotinic transmission (Zhang etal., 1996; Ullian et al., 1997).

a3*- and aBgt-AChR Clusters ArePresent on the Neurons in Culture

If a3*- andaBgt-AChRs participate in synaptic trans-mission between ciliary ganglion neurons in culture,some should be clustered in the vicinity of presynapticterminals. We examined the distribution of both

274 Chen et al.

AChR classes with conventional fluorescence micros-copy using mAb35 andaBgt to detecta3*-AChRsand aBgt-AChRs, respectively, as previously de-scribed (McNerney et al., 2000), and anti-SV2 mAb10h to detect presynaptic terminals (Wilson Horchand Sargent, 1995) (Fig. 5). In cultures grown for 5–6days, clusters ofa3*-AChRs were apparent as smallbright puncta or larger blobs of mAb35 immunolabel-ing on the surface of neuron somata [Fig. 5(A,D)].The general pattern of mAb 10h immunolabeling inthe same focal plane was frequently quite similar [Fig.5(B)], and in some cases it appeared that some so-matic a3*-AChR clusters and presynaptic terminalsoverlapped or were at least in close proximity [Fig.5(C)]. a3*-AChR clusters were also prevalent onneurites, but their distribution here appeared less ob-viously correlated with that of presynaptic terminals(not shown). In parallel cultures labeled withaBgt,aBgt-AChR clusters were evident, and while appear-ing smaller than a3*-AChR clusters, these alsoseemed most prevalent on neuron somata [Fig.5(E,H)]. Although less distinct than mAb35 labeling,

the general pattern ofaBgt labeling was not obviouslysimilar to that of mAb 10h labeling [Fig. 5(E,F)], andin only rare instances didaBgt-AChRs and presyn-aptic terminals appear to be in proximity [e.g., Fig.5(G)]. While further experiments using higher reso-lution techniques, such as confocal or electron mi-croscopy, are necessary to better determine the pre-cise relationship between presynaptic terminals anda3*- and/or aBgt-AChRs, two general conclusionscan be drawn from the present results. The first is that,as expected from their participation in fast and slowsEPSC components, botha3*- and aBgt-AChRs areclustered on ciliary ganglion neuron somata in cellculture. The second is that somea3*-, and fewaBgt-AChR somatic clusters appear to be in the vicinity ofpresynaptic terminals. Both conclusions are consistentwith the present study, indicating a major contributionof a3*-AChRs to EPSCs in culture, and an arrange-ment of a3*- and aBgt-AChR clusters at post- andperisynaptic sites, respectively, on ciliary ganglionneuronsin vivo (Jacob and Berg, 1983; Jacob et al.,1984; Wilson Horch and Sargent, 1995).

Figure 5 Arrangement ofa3*- and aBgt-AChR clusters on ciliary ganglion neurons in culture.After 5–6 days, cultures were labeled (see Methods) either with mAb 35 [(A) a3*-AChRs] oraBgt[(E) aBgt-AChRs], both visualized using Cy3 conjugates (red), and then with mAb 10h [(B) and(F)], visualized using an Alexa Fluor 488 conjugate (green). The respective merged images [(C) and(G)] depict regions of apparent AChR1 terminal proximity (red1 green5 yellow). Differentialinterference contrast views of the same neurons are shown in (D) and (H). Within both series, theplane of focus was maintained for each exposure, and grazes the top surface and perimeter of theneuron soma. Note thata3*- andaBgt-AChRs were evident as clusters, but thata3*-AChRs weremore obviously associated with presynaptic sites [(C), bold arrowheads] than wereaBgt-AChRs[(G), outlined arrowhead]. Control fields from the same cultures where mAb35,aBgt, or mAb 10hwas omitted yielded only weak background fluorescence (not shown). Scale bar5 10 mm.

Neuronal Nicotinic Synapses in Culture 275

DISCUSSION

We have shown that nicotinic synapses between chickciliary ganglion neurons in culture resemble thosepresent on the neuronsin vivo. It was previouslydemonstrated that chemical synapses form betweenthe neurons in culture, and that such synapses areexcitatory and mediated exclusively by nicotinicAChRs (Margiotta and Berg, 1982), as is the case forchemical ganglionic transmissionin vivo (Martin andPilar, 1963; Zhang et al., 1996; Ullian et al., 1997).We now report similar parallels between thein vivoand in vitro settings with regard to quantal release oftransmitter from presynaptic terminals, EPSC decaykinetics, receptor subtype dependence for transmis-sion, and arrangement of AChRs at synaptic sites. Inparticular, we find thata3*- andaBgt-AChRs under-lie EPSCs in culture, as they doin vivo, suggestingthat the formation of single and mixed-receptor syn-apses does not require the presence of either thenormal inputs or targets, because both are absent inthe cultures. Thus, while synapses do not occur be-tween ciliary ganglion neuronsin vivo, the cultures,by virtue of their simplicity and ease of manipulation,represent a useful model system for studying forma-tion and regulation of nicotinic synapses in the ner-vous system.

Sharp electrode experiments performed using cili-ary ganglia from newly hatched chicks previouslydemonstrated spontaneous, quantal ACh release re-flected in small (1–5 mV) mEPSPs that occurred atlow frequency (0.1 to 0.2 Hz) in a subset of neurons(Martin and Pilar, 1964). Consistent with low levelsof quantal releasein vivo, miniature synaptic eventswere previously undetectable in ciliary ganglion neu-ron cultures treated with TTX (Margiotta and Berg,1982) and occurred at low frequency ('0.1 Hz) in thepresent experiments. Recent whole-cell studiesin vivoindicate that mEPSCs are either absent or occur atvery low rates in embryonic ciliary ganglion neurons(Sargent, unpublished results). Another study usingidentical approaches also reported mEPSCs in a sub-set of recordings, but these occurred at a much higherfrequency ('6 Hz) (Zhang et al., 1996). The reasonfor the partial discrepancy amongin vivo results isunknown, however, the high mEPSC frequency in theone study might have resulted from injury-inducedACh release. If so, the generally low frequencies ofneuronal mEPSCs seenin vivo and in culture wouldbe consistent with the low rates of quantal transmitterreleaseatthedevelopingneuromuscularjunction(Fisch-bach, 1972).

Pharmacological studies using subunit-specificneurotoxins indicate that slow mono- and biexponen-tial EPSCs in culture are mediated by the samea3*-and aBgt-AChR subtypes that underlie ganglionictransmissionin vivo.aBgt had effects on sEPSCs thatcould be attributed to both post- and presynaptic sitesof action. The ability of the toxin to abolish or dras-tically reduce the frequency of fast mono- or biexpo-nential sEPSCs, respectively, is consistent with apostsynaptic effect of blocking brief open duration 60and 80 pSaBgt-AChR channels on the neurons (Mc-Nerney et al., 2000).aBgt also reduced overallsEPSC frequency (F1) in the cultures by 40%, andnominally reduced the slow component amplitudes ofmono- and biexponential sEPSCs. The reduction inslow component sEPSC amplitudes could not be ex-plained by a block of the long-duration 30 and 40 pSa3*-AChR events presumed to mediate them becausePopen values for such events are unaffected byaBgt(McNerney et al., 2000). One explanation is thataBgtblocks preterminalaBgt-AChRs, present on the neu-rons bothin vivo (Coggan et al., 1997) and in culture(Pugh and Berg, 1994). Such presynapticaBgt-AChRs are known to enhance transmitter release byvirtue of their high Ca21 permeability (McGehee etal., 1995; Gray et al., 1996). If presynapticaBgt-AChRs normally enhanced transmission between theneurons in culture, their blockade would be expectedto reduce overallF1 by increasing failures, and thussomewhat reduce the amplitude of evoked slowEPSCs, effects that were observed in the present ex-periments. A presynaptic blocking mechanism wouldalso explain the absence of an otherwise expectedhigher frequency of slow EPSCs inaBgt-treated neu-rons compared to controls. The inhibitory effects ofaBgt were relatively minor, however, compared tothose ofaCTx-MII, which drastically reducedF1,abolishing nearly all slowly decaying sEPSCs, leav-ing only rapidly decaying monoexponential sEPSCs.For the few remaining slow sEPSCs,aCTx-MII re-duced the slow component amplitude and increasedthe fast component amplitude of biexponentialsEPSCs, and enhanced the decay rate of monoexpo-nential slow sEPSCs. In the case of the remainingbiexponential sEPSCs, we presume the toxin block ofthe slow component sufficiently unmasked the fastcomponent such that its amplitude became fully re-solvable. As for the remaining slow sEPSCs, theenhanced decay rate suggests that the toxin blockedthe underlyinga3*-AChRs by reducing channel opentime. Given the specificity ofaCTx-MII for a3/b2AChRs expressed in oocytes (Cartier et al., 1996) andrecent studies indicating that the toxin blocks the 40

276 Chen et al.

pS a3*-AChR channels on the neurons, without af-fecting 60 and 80 pSaBgt-AChRs (Nai et al., 2000),the findings indicate that 40 pSa3*-AChR channelsmediate 90% of sEPSCs in the cultures. If the fastcomponent of biexponential events was mediatedsolely byaBgt-AChRs, however, a higher frequencyof monoexponential fast events would have been ob-served in the presence ofaCTx-MII than was seen.Because 40 pSa3*-AChRs display both brief- andlong-duration openings (McNerney et al., 2000), apossible explanation for the low frequency of fastevents in the presence ofaCTx-MII is that part of thebiexponential sEPSC fast component is mediated bya3*-AChRs. Thus, while botha3*- andaBgt-AChRsparticipate in synaptic transmission,a3*-AChRs me-diate the bulk of transmission in culture. This situa-tion is similar to that foundin vivo,whereN-Bgt (alsoknown ask-Bgt), a snake toxin that recognizes thesamea3*-AChRs on the neurons as mAb35, blocksganglionic transmission, whileaBgt fails to do so(Chiappinelli, 1983).

The arrangement ofa3*- or aBgt-AChR clusterswith presynaptic terminals on the neurons in culture isalso reminiscent of that found previouslyin vivousingconfocal fluorescence (Wilson Horch and Sargent,1995) and electron microscopic (Jacob and Berg,1983; Jacob et al., 1984) approaches. In these earlierstudies,a3*-AChRs were consistently (Jacob et al.,1984) or occasionally (Wilson Horch and Sargent,1995) aligned adjacent to presynaptic sites, whereasaBgt-AChRs were localized on somatic spines(“pseudodendrites”) outside of the immediatepostsynaptic membrane. The conventional fluores-cence approach we employed does not provide theresolution of either confocal or eletron microscopy.The general correspondence of our results with theinvivo setting is consistent, however, witha3*-AChRmediated synaptic transmission predominantly in thecultures as it doesin vivo.

While similar in function and morphology, nico-tinic synapses between ciliary ganglion neurons inculture can be distinguished from those present on theneuronsin vivo by several criteria. The first and mostobvious is that the210 to280 pA sEPSCs observedin culture are much smaller than the peak-evokedEPSCsin vivo (20.5 to 28 nA; see Zhang et al.,1996; Ullian et al., 1997). The disparity in EPSCamplitudes is consistent with the fourfold larger max-imal ACh responses obtainedin vivo (Margiotta andGurantz, 1989) relative to thosein vitro (Margiotta etal., 1987b), and therefore may arise from lower levelsof AChR subunits and transcripts expressed in culture(Corriveau and Berg, 1994). As previously demon-

strated, the reduced AChR subunit transcript levelsand sEPSC amplitudes in culture may reflect the ab-sence of influences normally provided by targets(Devay et al., 1999) and/or preganglionic inputs (Ja-cob and Berg, 1988; Arenella et al., 1993). Otherexplanations for the smaller EPSC amplitudes mayinvolve mechanisms controlling the extent of AChRclustering, the activation of presynaptic release sites,or the formation of appropriately aligned synapticcontacts. A second difference is that the cultures ap-pear somewhat more permissive, because'10% ofevents were fast monoexponential EPSCs, while suchevents are apparently absentin vivo. This disparitymay also result from the absence of normal targetsand/or inputs, but the possibility that other influencesare involved cannot be excluded. Apart from thesedifferences, the similarities betweenin vitro and invivosettings indicate the culture system will be usefulfor studying processes such as the molecular require-ments for nicotinic synapse formation, and the mech-anisms underlying regulation of nicotinic synapticfunction. In the former case, the cultures can be usedto determine if neuronal synapses can form at new orpre-existing sites, and to identify molecules responsi-ble for organizing AChRs at developing synapses.The availability of pre- and postsynaptic probes andcandidate molecules should make such studies feasi-ble. With regard to regulation of function, recentstudies indicate that sEPSC frequency is greatly en-hanced following activation of calcium induced cal-cium release (CICR; Chen et al., 1998). Ciliary gan-glion cultures will be useful in unraveling the signalpathways that lead to these and other functionalchanges at nicotinic synapses, and will thereby help todefine new mechanisms underlying synaptic plasticityin the nervous system.

We thank Dr. J. Michael McIntosh (University of Utah)for generously supplyinga-conotoxin-MII, and Dr. Peter B.Sargent (University of California, San Francisco) for in-sightful comments and discussions.

REFERENCES

Alkondon M, Pereira EF, Albuquerque EX. 1998.a-bun-garotoxin and methyllycaconitine-sensitive nicotinic re-ceptors mediate fast synaptic transmission in interneuronsof rat hippocampal slices. Brain Res 810:257–263.

Arenella LS, Oliva JM, Jacob MH. 1993. Reduced levels ofacetylcholine receptor expression in chick ciliary gan-glion neurons developing in the absence of innervation.J Neurosci 13:4525–4537.

Neuronal Nicotinic Synapses in Culture 277

Broide RS, Leslie FM. 1999. Thea7 nicotinic receptor inneuronal plasticity. Mol Neurobiol 20:1–16.

Bruses J, Oka S, Rutishauser U. 1995. NCAM-associatedpolysialic acid on ciliary ganglion neurons is regulated bypolysialyltransferase levels and interaction with muscle.J Neurosci 15:8310–8319.

Cartier GE, Yoshikami D, Gray WG, Luo S, Olivera BM,McIntosh JM. 1996. A newa-conotoxin which targetsa3b2 nicotinic acetylcholine receptors. J Biochem 271:7522–7528.

Chang KT, Berg DK. 1999. Nicotinic acetylcholine recep-tors containinga7 subunits are required for reliable syn-aptic transmissionin situ. J Neurosci 19:3701–3710.

Chen M, Yu XH, Margiotta JF. 1998. Regulation of cho-linergic interneuronal synapses. Soc Neurosci Abs 24:335.

Chiappinelli VA. 1983. Kappa-bungarotoxin: a probe forthe neuronal nicotinic receptor in the avian ciliary gan-glion. Brain Res 277:9–22.

Coggan JS, Paysan J, Conroy WG, Berg DK. 1997. Directrecording of nicotinic responses in presynaptic nerveterminals. J Neurosci 17:5798–5806.

Colling SB, Wheal HV. 1994. Fast sodium action potentialsare generated in the distal apical dendrites of rat hip-pocampal CA1 pyramidal cells. Neurosci Lett 172:73–76.

Corriveau RA, Berg DK. 1993. Coexpression of multipleacetylcholine receptor genes in neurons: quantification oftranscripts during development. J Neurosci 13:2662–2671.

Corriveau RA, Berg DK. 1994. Neurons in culture maintainacetylcholine receptor levels with far fewer transcriptsthan in vivo. J Neurobiol 25:1579–1592.

Cowan WM, Wenger E. 1968. Degeneration in the nucleusof origin of the preganglionic fibers to the chick ciliaryganglion following early removal of the optic vesicle. JExp Zool 168:105–123.

Crean G, Pilar G, Tuttle JB, Vaca K. 1982. Enhancedchemosensitivity of chick parasympathetic neurones inco-culture with myotubes. J Physiol 331:87–104.

Curtis DR, Ryall RW. 1966. The synaptic excitation ofRenshaw cells. Exp Brain Res 107:166–170.

Dani JA, Heinemann S. 1996. Molecular and cellular as-pects of nicotine abuse. Neuron 16:905–908.

Devay P, McGehee DS, Yu CR, Role LW. 1999. Target-specific control of nicotinic receptor expression at devel-oping interneuronal synapses in chick. Nature Neurosci2:528–534.

Dichter MA, Fischbach GD. 1977. The action potential ofchick dorsal root ganglion neurones maintained in cellculture. J Physiol 267:281–298.

Dryer SE. 1994. Functional development of the parasym-pathetic neurons of the avian ciliary ganglion: a classicmodel system for the study of neuronal differentiationand development. Prog Neurobiol 43:281–322.

Duggan AW, Hall JG, Lee CY. 1976. Alpha-bungarotoxin,cobra neurotoxin and excitation of Renshaw cells byacetylcholine. Brain Res 107:166–170.

Feller MB, Wellis DP, Stellwagen D, Werblin FS, Shatz CJ.

1996. Requirement for cholinergic synaptic transmissionin the propogation of spontaneous retinal waves. Science272:1182–1186.

Fischbach GD. 1972. Synapse formation between dissoci-ated nerve and muscle cells in low density cell cultures.Dev Biol 28:407–429.

Frazier CJ, Buhler AV, Weiner JL, Dunwiddie TV. 1998.Synaptic potentials mediated viaa-bungarotoxin-sensi-tive nicotinic acetylcholine receptors in rat hippocampalinterneurons. J Neurosci 18:8228–8235.

Gray R, Rajan A, Radcliffe K, Yakehiro M, Dani J. 1996.Hippocampal synaptic transmission enhanced by lowconcentrations of nicotine. Nature 383:713–716.

Hamburger V, Hamilton HL. 1951. A series of normalstages in the development of the chick embryo. J Morphol88:49–92.

Jacob MH, Berg DK. 1983. The ultrastructural localizationof a-bungarotoxin binding sites in relation to synapses onchick ciliary ganglion neurons. J Neurosci 3:260–271.

Jacob MH, Berg DK. 1988. The distribution of acetylcho-line receptors in chick ciliary ganglion neurons followingdisruption of ganglionic connections. J Neurosci 8:3838–3849.

Jacob MH, Berg DK, Lindstrom JM. 1984. Shared antigenicdeterminant between theElectrophorusacetylcholine re-ceptor and a synaptic component on chicken ciliary gan-glion neurons. Proc Natl Acad Sci USA 81:3223–3227.

Jones S, Sudweeks S, Yakel JL. 1999. Nicotinic receptors inthe brain: correlating physiology with function. TrendsNeurosci 22:555–561.

Landmesser L, Pilar G. 1972. The onset and development oftransmission in the chick ciliary ganglion. J Physiol 222:691–713.

Landmesser L, Pilar G. 1978. Interactions between neuronsand their targets duringin vivo synaptogenesis. Fed Proc37:2016–2022.

Lindstrom J. 1997. Nicotinic acetylcholine receptors inhealth and disease. Mol Neurobiol 15:193–222.

MacDermott AB, Role LW, Siegelbaum SA. 1999. Presyn-aptic ionotropic receptors and the control of transmitterrelease. Annu Rev Neurosci 22:443–485.

Margiotta JF, Berg DK. 1982. Functional synapses areestablished between ciliary ganglion neurons in dissoci-ated cell culture. Nature 296:152–154.

Margiotta JF, Berg DK, Dionne VE. 1987a. Cyclic AMPregulates the proportion of functional acetylcholine re-ceptors on chick ciliary ganglion neurons. Proc Natl AcadSci USA 84:8155–8159.

Margiotta JF, Berg DK, Dionne VE. 1987b. The propertiesand regulation of functional acetylcholine receptors onchick ciliary ganglion neurons. J Neurosci 7:3612–3622.

Margiotta JF, Gurantz D. 1989. Changes in the number,function and regulation of nicotinic acetylcholine recep-tors during neuronal development. Dev Biol 135:326–399.

Martin AR, Pilar G. 1964. Quantal components of the

278 Chen et al.

synaptic potential in the ciliary ganglion of the chick.J Physiol 175:1–16.

Martin RA, Pilar G. 1963. Dual mode of synaptic transmis-sion in the avian ciliary ganglion. J Physiol 168:443–463.

McGehee D, Heath M, Gelber S, Devay P, Role LW. 1995.Nicotinic enhancement of fast excitatory synaptic trans-mission in CNS by presynaptic receptors. Science 269:1692–1697.

McGehee DS, Role LW. 1995. Physiological diversity ofnicotinic acetylcholine receptors expressed by vertebrateneurons. Ann Rev Physiol 57:521–546.

McNerney ME, Pardi D, Pugh PC, Nai Q, Margiotta JF.2000. Expression and channel properties ofa-bungaro-toxin-sensitive acetylcholine receptors on chick ciliaryand choroid neurons. J Neurophysiol 84:1314–1329.

Nai Q, Pardi D, Margiotta JF. 2000.a-bungarotoxin anda-conotoxin-MII recognize different nicotinic acetylcho-line receptor channels on chick ciliary ganglion neurons.Soc Neurosci Abs 26:372.

Newhouse PA, Kelton M. 2000. Nicotinic systems in centralnervous systems disease: degenerative disorders and be-yond. Pharm Acata Helv 74:91–101.

Nishi R, Berg DK. 1981. Two components from eye tissuethat differentially stimulate the growth and developmentof ciliary ganglion neurons in cell culture. J Neurosci1:505–513.

Nong Y, Sorenson EM, Chiappinelli VA. 1999. Fast exci-tatory nicotinic transmission in the chick lateral spiriformnucleus. J Neurosci 19:7804–7811.

O’Lague PH, Potter DD, Furshpan EJ. 1978. Studies on ratsympathetic neurons developing in cell culture. III. Cho-linergic transmission. Dev Biol 67:424–443.

Pilar G, Tuttle JB. 1982. A simple neuronal system with arange of uses: the avian ciliary ganglion. In: Hanin I,Goldberg AM, editors. Progress in Cholinergic Biology:Model Cholinergic Synapses. New York: Raven Press.

Pugh PC, Berg DK. 1994. Neuronal acetylcholine receptorsthat binda-bungarotoxin mediate neurite retraction in acalcium-dependent manner. J Neurosci 14:889–896.

Pugh PC, Corriveau RA, Conroy WG, Berg DK. 1995.Novel subpopulation of neuronal acetylcoline receptorsamong those bindinga-bungarotoxin. Mol Pharmacol47:717–725.

Pugh PC, Margiotta JF. 2000. Nicotinic acetylcholine re-ceptor agonists promote survival and reduce apoptosis of

chick ciliary ganglion neurons. Mol Cell Neurosci 15:113–122.

Roerig B, Nelson DA, Katz LC. 1997. Fast synaptic signal-ing by nicotinic acetylcholine and serotonin 5-HT3 re-ceptors in developing visual cortex. J Neurosci 17:8353–8362.

Role LW, Berg DK. 1996. Nicotinic receptors in the devel-opment and modulation of CNS synapses. Neuron 16:1077–1085.

Sargent PB. 1993. The diversity of neuronal nicotinic ace-tylcholine receptors. Annu Rev Neurosci 16:403–443.

Shoop RD, Martone ME, Yamada N, Ellisman MH, BergDK. 1999. Neuronal acetylcholine receptors witha7 sub-units are concentrated on somatic spines for synapticsignaling in embryonic chick ciliary ganglia. J Neurosci19:692–704.

Ullian EM, McIntosh JM, Sargent PB. 1997. Rapid synaptictransmission in the avian ciliary ganglion is mediated bytwo distinct classes of nicotinic receptors. J Neurosci17:7210–7219.

Vernallis AB, Conroy WG, Berg DK. 1993. Neurons as-semble AChRs with as many as 3 kinds of subunits whilemaintaining subunit segregation among subtypes. Neuron10:451–464.

Vetter DE, Liberman MC, Mann J, Barhanin J, Boulter J,Brown MC, Saffiote-Kolman J, Heinemann SF, ElgoyhenAB. 1999. Role ofa9 nicotinic ACh receptor subunits inthe development and function of cochlear efferent inner-vation. Neuron 23:93–103.

Willard AL, Nishi R. 1985. Neurons dissociated from ratmyenteric plexus retain differentiated properties whengrown in cell culture. III. Synaptic interactions and mod-ulatory effects of neurotransmitter candidates. Neuro-science 16:213–221.

Wilson Horch HL, Sargent PB. 1995. Perisynaptic surfacedistribution of multiple classes of nicotinic acetylcholinereceptors on neurons in the chicken ciliary ganglion.J Neurosci 15:7778–7795.

Zhang M, Wang YT, Vyas DM, Neuman RS, Bieger D.1993. Nicotinic cholinoceptor-mediated excitatorypostsynaptic potentials in the rat nucleus ambiguous. ExpBrain Res 96:83–88.

Zhang Z-W, Coggan JS, Berg DK. 1996. Synaptic currentsgenerated by neuronal acetylcholine receptors sensitive toa-bungarotoxin. Neuron 17:1231–1240.

Neuronal Nicotinic Synapses in Culture 279