Neurotrophic molecules are key retrograde influences of cell survival in the developing nervous system, but other influences such as activity are also emerging as important factors. In the avian ciliary ganglion, half the neurons are eliminated between embryonic day 8 (E8) and E14, but it is not known how cell death is initiated. Because systemic application of α7-nicotinic acetylcholine receptor (nAChR) antagonists prevents this cell loss, we examined differences in receptor densities and responses of intracellular calcium to nicotine using the calcium-sensitive dye fura-2. In addition, we determined whether cell-autonomous inhibition of α7 activation in neurons prevented cell death. E8 neurons are heterogeneous with respect to α7-nAChR density, which leads to large increases in [Ca2+]i in some neurons; E8 neurons also exhibit a slower rate of Ca2+ decay after nicotinic stimulation than E13 neurons. Expressing α-bungarotoxin that is tethered to the membrane by a glycosylphosphatidylinositol linkage (GPIαbtx) in ciliary ganglion neurons with the retroviral vector RCASBP(A) blocks increases in intracellular calcium induced by nicotine through α7-nAChRs and prevents neurons from dying. Expression of GPIαbtx in surrounding non-neural tissues, but not in neurons, does not prevent cell loss. Furthermore, the GPIαbtx is not efficiently expressed in the accessory oculomotor neurons, eliminating preganglionic inputs as another site for action of the antagonist. These results support the hypothesis that cholinergic inputs facilitate cell death in the developing autonomic nervous system by activating α7-nAChRs, possibly by leading to increases in intracellular calcium that exceed the threshold for cell survival.
The phenomenon of cell death of postmitotic neurons in the developing vertebrate nervous system has been well described, but the means by which cell death is induced are not well understood (Oppenheim, 1991). Up to 70% of neurons are lost at the time of synapse formation with their targets, during which neurons are dependent on target-derived factors for survival (Davies, 1996). The neurotrophic hypothesis proposes that neurons die because they fail to compete effectively for a limited amount of trophic factor in the target tissue. Both gain- and loss-of-function studies show that many target-derived factors are essential for supporting neuronal survival (Snider, 1994); however, neurons integrate trophic support from more than one source (target and glia), and to do this, they must enter the competition with some advantage involving cell-to-cell differences in intrinsic factors, such as surface receptor expression (for review, see Pettmann and Henderson, 1998). This suggests that the trophic requirements of neurons are more complex and other signals may also be involved in controlling cell death (Eisen and Melancon, 2001).
The ciliary ganglion, a parasympathetic ganglion that innervates the iris, ciliary muscle, and choroidal vasculature in the avian eye (Marwitt et al., 1971), undergoes a period of target-dependent cell loss between embryonic day 8 (E8) and E14 (Landmesser and Pilar, 1974b). However, virtually all ciliary ganglion neurons are rescued from dying by daily application of α7 subunit-containing nicotinic acetylcholine receptor (nAChR)-specific antagonists to the chorioallantoic membrane between E8 and E14 without changes in trophic factor availability or access at target tissues (Meriney et al., 1987; Bunker and Nishi, 2002). This suggests that activation of α7-nAChRs can facilitate cell death independently of the competition for target-derived factors. However, the site at which these antagonists act is not clear because they were delivered throughout the entire chicken embryo.
α7-nAChRs are cation-selective channels that are highly expressed together with heteromeric nAChRs containing α3, α5, β4, and sometimes β2 subunits on ciliary ganglion neurons, where they mediate excitatory transmission (Zhang et al., 1996; Chang and Berg, 1999). In addition, α7-nAChRs have high relative permeability to Ca2+ that rivals that of NMDA receptors, but unlike NMDA receptors, α7-nAChRs can mediate Ca2+ influx at resting or hyperpolarized potentials (for review, see Role and Berg, 1996). This feature enables them to influence a variety of Ca2+-dependent events, including gene expression, neurite retraction, axonal pathfinding, and cell death (Pugh and Berg, 1994; Zheng et al., 1994; Berger et al., 1998; Chang and Berg, 2001; Liu et al., 2006).
This study was undertaken to determine whether activation of α7-nAChRs on ciliary ganglion neurons causes cell death. To do this, we used α-bungarotoxin (αbtx) tethered to the membrane by glycosylphosphatidylinositol (GPI) linkage (Ibanez-Tallon et al., 2004), to determine whether cell-autonomous inhibition of α7-nAChRs prevents both ciliary and choroid neurons from dying. We also examine differences in Ca2+ signaling between neurons before cell death and propose a possible mechanism of α7-nAChR-induced cell death through Ca2+ overload. Our findings highlight the importance of nontraditional roles of nAChRs during neural development.
Materials and Methods
Isolation of neurons.
Ciliary ganglia were isolated and dissociated for imaging, staining, and flow cytometry from E6 to E10 embryos as described previously (Nishi, 1996). E13 ciliary ganglia were dissociated by incubating in Earle's balanced salt solution (EBSS) containing 1 mg/ml collagenase type 2 (catalog #4176; Worthington, Lakewood, NJ), 1 mg/ml hyaluronidase (catalog #2592; Worthington), and 10 mg/ml bovine serum albumin (Sigma, St. Louis, MO) for 15 min at 37°C. The activity of the proteases was terminated by dilution, then the supernatant was removed and ganglia were incubated in EBSS containing 3 mg/ml trypsin (catalog #3703; Worthington) for 3 min at 37°C. Ciliary ganglia were triturated until completely dissociated. After dissociation, cells were preplated on the plastic surface for 30 min at 37°C to remove adherent cells (fibroblast and glia). Neurons were plated on poly-d-lysine/laminin-coated coverslips in complete medium consisting of DMEM (Invitrogen, Carlsbad, CA) containing 10% (v/v) heat-inactivated horse serum (Invitrogen), 2% (v/v) fetal calf serum (Atlanta Biologicals, Lawrenceville, GA), 2% (v/v) chicken eye extract with 50 U/ml penicillin, 50 mg/ml streptomycin, and 2 mm glutamine (Invitrogen). Neurons were used for staining or calcium imaging after allowing them to attach to the coverslip for 2 h after plating. Alternatively, acutely isolated neurons were incubated with αbtx–Alexa 488 or Q211 and immediately processed for flow cytometry as described below.
Immunostaining and live cell labeling.
Dissociated E8 or E13 neurons were incubated while alive with Alexa 488-conjugated αbtx (Invitrogen), diluted 1:500 in complete medium for 30 min at 37°C, or with rabbit anti-αbtx, diluted 1:1000 in complete medium for 30 min on ice, to label only cell-surface molecules. After the incubation, neurons were washed three times in EBSS and fixed in Zamboni's fixative (4% paraformaldehyde and 15% picric acid in 0.1 m sodium phosphate, pH 7.4) for 2 h at 4°C. Coverslips were washed with PBS (137 mm NaCl and 20 mm sodium phosphate, pH 7.4), blocked overnight in blocking buffer [PBS plus 0.5% (v/v) Triton X-100 containing 5% (v/v) horse serum] at 4°C, and incubated overnight in primary antibodies. Secondary antibodies were incubated for 1–2 h at room temperature.
Tissues were prepared for immunohistochemistry as follows. Ciliary ganglia and brainstems from E14 embryos were harvested, fixed in Zamboni's fixative for 48 h at 4°C, washed, and equilibrated in 30% sucrose at 4°C. Tissue was embedded in Microm cryo-embedding compound (Richard Allen Scientific, Kalamazoo, MI), sectioned on a Microm HM 560 cryostat (Richard Allen Scientific) at 30 μm, and collected on Superfrost Plus slides (Fisher Scientific, Pittsburgh, PA). Sections were postfixed in Zamboni's fixative vapors for 15 min at 37°C, submerged in Zamboni's fixative for an additional 15 min at 25°C, washed in PBS, and blocked. Primary antibodies were incubated overnight at 4°C, and secondary antibodies were incubated 2 h at room temperature.
Primary antibodies and the dilutions in blocking buffer at which they were used were as follows: anti-mouse AMV-3C2 (Developmental Studies Hybridoma Bank, Iowa City, IA), which recognizes a viral p19 gag expressed by avian sarcoma and leukemia viruses (Potts et al., 1987), at 1:10; mouse anti-Islet-1, which recognizes a transcription factor expressed in ciliary ganglion neurons (Lee et al., 2001), at 1:100 dilution of the culture supernatant (prepared in the Nishi laboratory from clone 39.4D5; Developmental Studies Hybridoma Bank); mouse anti-Hu C/D (Invitrogen), which recognizes a neuron-specific RNA-binding protein (Marusich and Weston, 1992; Lee et al., 2001), at 1:250 dilution of the culture supernatant; rabbit anti-p27gag (SPAFAS, Norwich, CT), which recognizes avian sarcoma gag p27 (Wang et al., 1976), at 1:1000; rat anti-somatostatin (product number YMC1020; Accurate Chemical and Scientific Corporation, Westbury, NY) diluted 1:100; and rabbit anti-αbtx [a generous gift from Dr. J. Sanes, Washington University, St. Louis, MO (now at Harvard University, Cambridge, MA)] at 1:1000, the specificity of which is shown in supplemental Figure 1 (available at www.jneurosci.org as supplemental material).
Secondary antibodies were as follows: biotinylated anti-mouse (Vector Laboratories, Burlingame, CA) at 1:250; biotinylated anti-rabbit (Vector Laboratories) at 1:250; goat anti-mouse Cy3 (Jackson ImmunoResearch, West Grove, PA) at 1:750; goat anti-rabbit Alexa 488 (Invitrogen) at 1:750; and goat anti-rat Cy3 (Jackson ImmunoResearch) at 1:750. Images of dissociated neurons were acquired with a 60× objective on a DeltaVision (Olympus IX70) deconvolution microscope (Applied Precision, Issaquah, WA) using a CoolSNAP HQ CCD camera (Photometrics, Tucson, AZ). Image stacks were deconvolved using softWoRx image analysis software version 3.5.1 (Applied Precision, Issaquah, WA). Images of ciliary ganglia and midbrain were acquired with a 20× objective using a Nikon (Melville, NY) C1 confocal scanner attached to a Nikon Eclipse E800 microscope (Micro Video Instruments, Avon, MA).
Dissociated neurons were labeled live with 1:500 αbtx–Alexa 488 and 1:3000 mouse anti-Q211 (kindly provided by Dr. H. Rohrer, Max Planck Institute for Brain Research, Frankfurt, Germany), which recognizes a neuron-specific ganglioside (Rohrer et al., 1985). Q211 immunoreactivity was visualized with goat F(ab′)2 anti-mouse IgG conjugated to PE-Cy5.5 (Caltag Laboratories, Burlingame, CA) diluted 1:200. At each developmental stage examined, 40,000 cells were analyzed for their surface α7-nAChR levels by flow cytometry using a Coulter (Miami, FL) Epics XL Analyzer. Generated scatter plots indicated the relative levels of αbtx–Alexa 488 binding of all Q211 positive cells.
Expression of GPIαbtx with a viral vector.
An αbtx that is tethered to the membrane via a GPI linkage (GPIαbtx) was a gift from Drs. I. Ibanez-Tallon and N. Heintz (Rockefeller University, New York, NY) (Ibanez-Tallon et al., 2004). The GPIαbtx sequence was cloned into the Slax13NCO1 shuttle vector using 5′ NcoI and 3′ EcoRI sites (Morgan and Fekete, 1996); the insert was removed from Slax13NCO1 by cutting with ClaI and was cloned into the avian retroviral vector RCASBP(A) (Federspiel and Hughes, 1997). Infective RCASBP(A)–GPI-αbtx viral particles were generated by transfecting DF-1 chicken fibroblast cells with 800 ng of RCASBP(A)–GPIαbtx plasmid using TransIT-LT transfection reagent (Mirus Bio Corporation, Madison, WI). Conditioned media containing viral stocks collected from DF-1 cells were concentrated ∼20-fold by ultracentrifugation at 90,000 × g at 4°C for 3 h (Morgan and Fekete, 1996). Concentrated stocks were titered by limiting dilution and infectivity of cells as measured by staining with p27gag antibody. Stocks with a concentration of >108 infectious particles per milliliter were used for in vivo injection. Viral particles (60–120 nl) were injected into the mesencephalic enlargement of the neural tube of Hamburger/Hamilton stage 8–9 or stage 10–13 embryos using a Nanoject microinjector (Drummond Scientific, Broomall, PA). The shells were sealed with a glass coverslip and sterile vacuum grease and incubated at 37°C to the desired stage.
Acutely isolated ciliary ganglion neurons were loaded with fura-2 AM (Invitrogen) dissolved in DMSO at a final concentration of 5 μm with 2% Pluronic F-127 (Invitrogen). Neurons were loaded at room temperature for 30 min in the dark. Calcium signals were recorded by exposure to alternating wavelength (340 and 380 nm, 50 ms) generated by a xenon light source and a Lambda DG-4 ultra high-speed wavelengths switcher (Sutter Instruments, Novato, CA). Fluorescent responses were recorded using an Orca-ER digital camera (Hammatsu, Bridgewater, NJ). Paired 340/380 ratio images were acquired at 4 s intervals with Metaflour 5.0r5 software (Molecular Devices, Downingtown, PA). Drugs were dissolved in chicken physiological buffer (in mm: 145 NaCl, 5.4 KCl, 0.8 MgCl2, 5.4 CaCl2, 5 glucose, and 13 HEPES, pH 7.4). Voltage-gated sodium and calcium channels were blocked by supplementing the perfusion medium with 600 nm tetrodotoxin (TTX; Tocris, Ellisville, MO) and 200 μm cobalt chloride (Sigma), respectively. A concentration of 10 μm nicotine (Sigma) was applied for 20 s to activate nAChRs. α7-nAChRs were inhibited by perfusing the neurons with 50 nm α-methyllycaconitine citrate hydrate (MLA) (Sigma) for 1 min or preincubating with 50 nm αbtx (Sigma) for 30 min at 25°C. Heteromeric α3/α5/β4/β2 (α3*)-nAChRs were blocked by perfusing neurons for 1 min with 5 μm dihydro-β-erythroidine hydrobromide (DHβE; Sigma). On completion of these experiments, the extent of dye loading was determined by activating voltage-gated calcium channels with a high-potassium perfusion solution (25 mm KCl, with no TTX or cobalt chloride).
Analysis of Ca2+ imaging data.
After the initial recordings were performed, background was subtracted from every image acquired and new ratios were calculated using the Metafluor 5.0r5 software. The ratios were then exported into the Microsoft (Redmond, WA) Excel spreadsheet, and all the calculations were performed using Microsoft Excel. For the calculation of Ca2+ decay rates, traces were normalized so that the baseline was always 0 and the peak calcium signal was always 1, using the equation (F − F 0)/F peak. Baseline fluorescence (F 0) was subtracted from the fluorescence at every time point (F). This value was then divided by peak amplitude fluorescence (F peak). This allowed us to observe changes in the rate of Ca2+ decay between individual neurons.
Serially sectioned ciliary ganglia (cut at 30 μm) were prepared for designed-based stereology as described previously (Bunker and Nishi, 2002), and Islet-1-positive nuclei (representing all neurons) together with somatostatin-positive cell bodies (representing all choroid neurons) were counted using the Optical Dissector Probe of Stereo Investigator (MBF Biosciences, Williston, VT) in conjunction with a Nikon Optiphot 2 microscope with a Hitachi (Tokyo, Japan) HVC20 camera, Heidenhahn focus encoder, and a motorized, computer-driven X,Y,Z stage (all microscope attachments provided by MBF Biosciences). To avoid inaccuracies caused by cutting artifacts and double counting between adjacent sections, an upper guard of 4 μm and a lower guard of 7 μm were used (Bunker and Nishi, 2002). Spacing between sampling sites (grid size) was set such that 13–15 sampling sites were counted per section, which yielded 100–300 objects per each ciliary ganglion. The number of ciliary neurons was calculated by subtracting the number of somatostatin-positive neurons from the total number of neurons per ciliary ganglion (Bunker and Nishi, 2002).
Immature ciliary ganglion neurons express heterogeneous levels of α7-nAChRs
If cell death in the ciliary ganglion is triggered in some neurons but not others in vivo between E7 and E9 by excessive signaling through α7 subunit-containing nAChRs, then there should be heterogeneity in α7-nAChR expression between individual neurons, in which the neurons with high levels of surface α7-nAChRs would be especially at risk to die from high Ca2+ influx through these receptors. To label surface α7-nAChRs, αbtx–Alexa 488 was bound to live ciliary ganglion neurons at E8. Individual E8 neurons exhibit different intensities of staining with αbtx–Alexa 488 (Fig. 1 A). One example of α7-nAChR heterogeneity is shown in Figure 1 A, where a neuron indicated by arrow labels brightly with αbtx–Alexa 488, whereas neighboring neurons display a low level of αbtx–Alexa 488 binding (Fig. 1 A, arrowhead).
At E13, we identified acutely isolated neurons as choroid or ciliary based on somatostatin immunoreactivity (Epstein et al., 1988; Coulombe et al., 1993). Both ciliary and choroid neurons express high levels of surface α7-nAChRs (Fig. 1 B,C). Moreover, α7-nAChRs on somatostatin-negative ciliary neurons are found tightly clustered on somatic spines at this developmental stage, which is another sign of maturation (Fig. 1 B, arrows) (Shoop et al., 1999). Choroid neurons that express somatostatin do not show α7-nAChR clustering (Fig. 1 C). Qualitatively, the intensity of staining with αbtx–Alexa 488 does not differ between individual E13 neurons of each population.
To survey the number of neurons exhibiting different levels of αbtx–Alexa 488 binding at various stages of development across ganglia, we used flow cytometry (Fig. 1 D–G). Neurons were distinguished from non-neuronal cells by immunoreactivity to Q211, an antibody that recognizes a neuron-specific ganglioside shortly after neural progenitors have commenced differentiation (Rohrer et al., 1985). At E6 and E7, neurons (Q211-positive cells) are very heterogeneous with respect to their surface αbtx binding, as can be seen by the large amount of scatter in αbtx labeling intensities (Fig. 1 D,E). By E9, the Q211-positive cells with little or no αbtx binding have all begun to express α7-nAChRs, and a few neurons still display very high αbtx binding (Fig. 1 F, arrowhead). However, by E10, αbtx binding is much more homogeneous than at E6 and E7 (Fig. 1 G). The loss of Q211 immunoreactivity, as seen by the spread of αbtx-positive cells into the upper left quadrant (Fig. 1 F,G), is a normal progression of Q211 staining that we have observed as neurons mature.
Because α7-nAChRs have a high relative permeability to calcium equal to that of NMDA receptors (Role and Berg, 1996), we tested whether the heterogeneity in surface α7-nAChRs observed at E8 leads to larger increases in intracellular Ca2+ in some neurons versus others. Neurons were imaged using the ratiometric dye fura-2, and 10 μm nicotine was applied by perfusion to activate the maximum number of nAChRs in all the neurons imaged by the microscope. By doing so, we could collect data on a large number of neurons per experiment, which is not possible by single-cell electrophysiology. To ensure that only nicotinic Ca2+ responses were measured, 200 μm CoCl2 and 600 nm TTX were added to the perfusion buffer to inhibit voltage-gated calcium and sodium channels, respectively (see Materials and Methods).
First, we tested whether neurons could recover nAChR sensitivity after nicotine application so that the efficacy of antagonists could be tested. After the first application of nicotine (Nic1), we washed cells for 5 min and then perfused nicotine again (Nic2) (Fig. 2 A). The peak amplitude of the Ca2+ response after the second stimulation was not significantly different from the initial application of nicotine (Fig. 2 B). Therefore, in all subsequent experiments, we washed for at least 5 min after the initial application of nicotine before we applied nicotine together with nicotinic antagonists. The response elicited by nicotine could be significantly reduced by 50 nm MLA, an antagonist known to selectively block α7-nAChRs on ciliary ganglion neurons at this concentration (Alkondon et al., 1992) (Fig. 2 C) (p < 0.0001, one-way ANOVA; n = 34 neurons). The residual response was completely blocked with a mixture containing 50 nm MLA and 5 μm DHβE (Fig. 2 C, compare MLA with DHβE/MLA) (p < 0.0001, one-way ANOVA; n = 30 neurons), which completely blocks transmission through the ciliary ganglion (Bertrand et al., 1992; Liu et al., 2006).
We then examined whether there are differences in α7-nAChR-induced Ca2+ influx between ciliary ganglion neurons isolated at different stages of development. To determine the Ca2+ influx mediated by α7-nAChR, Ca2+ responses obtained in the presence of the α7-nAChR-specific antagonist MLA were subtracted from the total Ca2+ responses to 10 μm nicotine. When E8 neurons are stimulated with nicotine, the elicited Ca2+ responses have a broad distribution (Fig. 2 D), demonstrating that individual E8 neurons exhibit variability in Ca2+ influx through α7-nAChRs. Compared with E8 neurons, α7-specific Ca2+ responses at E9 are skewed to the left with the majority of responses clustered around the mean (Fig. 2 D). The mean of α7-nAChR-specific Ca2+ responses at E9 is significantly smaller compared with E8 neurons (p < 0.0002, Student's t test; E8: 0.09 ± 0.007, n = 41; E9: 0.06 ± 0.004, n = 80). At E13, which is near the end of the cell death period, only half of the ciliary ganglion neurons display increases in intracellular calcium in response to perfused nicotine, despite the fact that all of them respond to high K+, indicating that all cells are equally loaded with fura-2. Of the ones that respond, intracellular Ca2+ decays to baseline at much faster rate than in neurons isolated at E8 (before onset of neuronal loss) (Fig. 2 E). At E8, the rate of Ca2+decay can be fitted with a single exponential (τ = 48.9 ± 0.98 s), whereas the rate of Ca2+ decay at E13 can be fitted with double exponential, with initial fast decay (τ1 = 17.5 ± 0.42 s) and later slower decay (τ2 = 175.6 ± 91.4 s), suggesting that an additional mechanism for sequestering intracellular calcium has developed. Treatment with MLA eliminates differences in the rate of Ca2+ decay between E8 and E13 neurons; at both stages, the rate of Ca2+ decay can be fitted with a double-exponential equation (Fig. 2 F) (E8: τ1 = 27.9 ± 0.0005 s, τ2 = 1014.1 s; E13: τ1 = 20.7 ± 1.56 s, τ2 = 78.8 ± 26.1 s). These results suggest that the activation of α7-nAChRs influences how neurons handle increases in [Ca2+]i.
Membrane-tethered αbtx inhibits α7-nAChR-mediated Ca2+ influx in ciliary ganglion neurons
To block Ca2+ influx via α7-nAChRs cell-autonomously in vivo, we used the avian retroviral vector RCASBP(A) to express an αbtx that was modified at the C terminus to include a linker with a consensus sequence for the GPI linkage from mouse lynx-1 (Fig. 3 A) (Ibanez-Tallon et al., 2004). Ciliary ganglion neurons infected with this construct exhibit bright, punctate αbtx immunoreactivity on their cell surface, indicating efficient expression of tethered αbtx (Fig. 3 C). When the ciliary ganglion cultures are prepared from embryos infected with the control (open) virus, neurons do not label with αbtx antibody (Fig. 3 B). The specificity of αbtx antibody was confirmed by Western blot and ELISA (supplemental Fig. 1, available at www.jneurosci.org as supplemental material).
To test whether GPIαbtx blocks Ca2+ influx through α7-nAChRs, E8 ciliary ganglion neurons infected either with GPIαbtx or open virus were subjected to Ca2+ imaging as described above. GPIαbtx-infected neurons have significantly reduced nicotinic Ca2+ responses compared with the neurons infected with control virus (Fig. 3 D) (open: 1.29 ± 0.13, n = 21; GPIαbtx: 0.36 ± 0.06, n = 23; p < 0.0001, Student's t test). In fact, GPIαbtx inhibits nicotine-induced increases in intracellular Ca2+ to the same extent as exogenously applied αbtx (Fig. 3 D) (open plus αbtx: 0.19 ± 0.04, n = 7; GPIαbtx: 0.36 ± 0.06, n = 23). Moreover, applying exogenous αbtx (50 nm) to GPIαbtx-infected cultures does not cause additional reduction in Ca2+ influx (Fig. 3 D) (GPIαbtx: 0.36 ± 0.06, n = 23; GPIαbtx plus 50 nm αbtx: 0.30 ± 0.05, n = 6). The above findings indicate that tethered αbtx inhibits α7-nAChR-mediated increases in intracellular Ca2+.
Expression of the GPIαbtx does not eliminate all of the calcium influx, leaving behind the residual Ca2+ response (Fig. 3 D), which is very similar to the residual response observed after a treatment with MLA (Fig. 2 C). This residual response is eliminated by the treatment with DHβE (Fig. 2 C), an antagonist that is more selective for heteromeric α3*-nAChRs. Treatment of GPIαbtx-infected neurons with exogenous αbtx does not cause an additional reduction in the residual Ca2+ response (Fig. 3 D).
To determine whether the GPIαbtx can be spontaneously released from the surface of a neuron and bind to α7-nAChRs on nearby neurons, we coincubated uninfected ciliary ganglion neurons overnight with the neurons infected with the GPIαbtx (Fig. 4). The cultures were then live-labeled with αbtx–Alexa 488 to visualize unoccupied surface α7-nAChRs. Uninfected neurons label strongly with αbtx–Alexa 488, whereas neurons infected with GPIαbtx retrovirus do not label with αbtx–Alexa 488 (Fig. 4 B,C). Importantly, uninfected neurons that were coincubated with the GPIαbtx-infected neurons still exhibit intense αbtx–Alexa 488 labeling that is comparable to uninfected controls (Fig. 4 D), demonstrating that their α7-nAChRs are not occupied by αbtx clipped off the surface of GPIαbtx-infected neurons.
GPIαbtx rescues ciliary and choroid neurons from cell death
To block activation of α7-nAChRs on ciliary ganglion neurons in vivo, we infected chicken embryos with RCASBP(A)–GPIαbtx and counted the number of surviving neurons at E14 using design-based stereology (Bunker and Nishi, 2002). If the viral particles are injected into the mesencephalon at stage 8–9 (36 h of development), ciliary ganglion neurons precursors are still present at the neural tube and accessible to retroviral infection (Fig. 5 A–C). However, if viral particles are injected at stage 10–13, the precursors have already migrated away from the neural tube to form a ganglion and are therefore inaccessible to the retrovirus, resulting in uninfected neurons surrounded by infected non-neuronal cells (Fig. 5 D–F).
For counting purposes, ciliary ganglia were labeled with Islet-1 and somatostatin antibodies (Fig. 5 G). Infection with GPIαbtx at 36 h of development rescues 41% of ciliary ganglion neurons from cell death compared with control-infected ganglia (Fig. 5 H) (p < 0.001, ANOVA; open ganglia: 6459 neurons, n = 6; GPIαbtx ganglia: 11,310 neurons, n = 8). Moreover, both ciliary (p < 0.01) and choroid neurons are rescued (Fig. 5 H) (p < 0.001, ANOVA with Tukey's multiple-comparison post hoc). When neurons are not infected, but surrounding non-neuronal cells are with RCASBP(A)–GPIαbtx, then neuronal cell numbers are not significantly different from open-infected control ciliary ganglia (Fig. 5 H) (GPIαbtx ganglia infected at 48 h: 8250 neurons, n = 9). Thus, the GPIαbtx must be expressed in ciliary ganglion neurons to block cell death. These results also show that GPIαbtx is not released from cells in quantities sufficient to block α7-nAChRs on nearby neurons.
The ciliary ganglion receives preganglionic input from the accessory oculomotor nucleus (AON) in the midbrain. The preganglionic neurons also contain presynaptic α7-nAChRs that control the release of acetylcholine by increasing the Ca2+ influx into the presynaptic terminals (Coggan et al., 1997; Rogers and Sargent, 2003). Therefore, inhibition of these receptors with GPIαbtx could also affect survival of ciliary ganglion neurons. However, p27gag immunoreactivity is observed only in a small number of AON neurons (Fig. 6 A–C, arrows), indicating that preganglionic neurons do not express GPIαbtx. Thus, presynaptic α7-nAChRs in terminals from AON are unlikely to affect survival of ciliary ganglion neurons.
The principal findings reported here support the hypothesis that α7-nAChRs facilitate developmental cell death of parasympathetic neurons, possibly by allowing increases in intracellular calcium that exceed the threshold for cell survival. Shortly after all of the neurons in the ciliary ganglion are innervated by preganglionic inputs, neurons express variable levels of α7-nAChRs together with differences in α7-mediated increases in intracellular calcium induced by nicotine. Furthermore, intracellular increases in calcium fall to baseline levels at slower rates in young neurons compared with neurons that have survived the cell death period. Finally, cell-autonomous inhibition of α7-nAChR signaling prevents cell death. Thus, anterograde signals play an important role in sculpting the final number of neurons in the autonomic nervous system.
The conclusion that activation of α7-nAChRs on ciliary ganglion neurons facilitates normal neuronal loss in the ganglion is based on the cell-autonomous action of GPIαbtx. The utility of this reagent in vivo was first demonstrated in zebrafish striated muscle; also in these studies, Xenopus oocytes expressing the GPIαbtx coincubated with oocytes expressing α7-nAChRs did not block α7 responses, demonstrating that the toxin was not cleaved and acting non-cell autonomously (Ibanez-Tallon et al., 2004). We show similar results when neurons expressing GPIαbtx are cocultured with uninfected neurons. Importantly, in vivo, we fail to rescue neurons when the GPIαbtx is expressed only in surrounding non-neuronal cells; the construct must be in neurons to rescue them. However, at the present time we cannot distinguish between α7-nAChRs located in cell soma and the terminals of ciliary ganglion neurons. RCASBP(A) does not efficiently infect early neural progenitors in the neural tube, therefore GPIαbtx is poorly expressed on the preganglionic neurons in the midbrain. Thus, presynaptic α7-nAChRs in GPIαbtx-infected embryos are not affected. This observation excludes α7-nAChRs on neurons in the AON as a significant contributor to cell death of ciliary ganglion neurons.
The hypothesis that α7-nAChR activation facilitates cell death of ciliary ganglion neurons is consistent with the temporal sequence of ganglionic innervation. Afferents from the AON first make contact at E4.5, and the neurons are 100% functionally innervated by E8, before the onset of cell loss (Landmesser and Pilar, 1972, 1974a). Our flow cytometry and calcium imaging data confirm and extend previous observations that ciliary ganglion neurons express many α7 receptors by E8 (Corriveau and Berg, 1993; Blumenthal et al., 1999; Brumwell et al., 2002). Whether the AON is spontaneously active at such early times is not clear; however, spontaneous electrical activity in the brainstem is detected using optical recording techniques at very early stages of embryonic development (Momose-Sato et al., 2003; Sato and Momose-Sato, 2003). In addition, ciliary neurons are enveloped in a preganglionic calyx at E8 that may serve to trap acetylcholine, bringing it to high levels if extracellular esterase activity is low. Finally, the kinetics of α7-nAChR inactivation at E8 versus E13 have yet to be examined. It is plausible that α7-nAChRs at E8 inactivate more slowly than more mature receptors.
The molecular mechanism underlying α7-nAChR effects on survival is likely to involve Ca2+ as an activator of cell death. Ca2+ is an intracellular messenger that operates over a wide temporal and spatial range to regulate many different cellular processes, and Ca2+ overload has been suggested to be the final common pathway of all types of cell death (Berridge et al., 2003; Hajnoczky et al., 2003; Rizzuto et al., 2003). α7-nAChRs have very high relative permeability to Ca2+ that is comparable to that of NMDA receptors. Furthermore, α7-nAChRs are insensitive to Mg2+ blockade, which allows them to mediate Ca2+ influx at rest (Role and Berg, 1996; Berg and Conroy, 2002). Therefore, we propose that activation of α7-nAChRs can increase [Ca2+]i to the levels that can be toxic to immature neurons. In support of this, a naturally occurring mutation in the α7-nAChR homolog induces cell death in Caenorhabditis elegans because of increased Ca2+ influx entering via slowly desensitizing receptors (Treinin and Chalfie, 1995). Transgenic mice homozygous for a similar gain-of-function mutation in α7-nAChRs also exhibit widespread neuronal apoptosis (Orr-Urtreger et al., 2000). Furthermore, Ca2+ influx caused by activation of α7-nAChRs induced apoptotic cell death in undifferentiated and immortalized hippocampal progenitor cells but not in differentiated cells (Berger et al., 1998). These progenitor cells were susceptible to Ca2+ overload because of their lack of buffering capability, whereas more mature neurons upregulated the expression of calbindin and no longer died as a result of α7-nAChR activation. In line with this evidence, ciliary ganglion neurons generate significantly larger increases in [Ca2+]i in response to nicotine at E8 than at E9. In addition, after nicotine stimulation, E8 neurons exhibit a slower rate of intracellular Ca2+ decay than E13 neurons. Thus, E8 neurons with a large response to nicotine are likely to be more vulnerable to Ca2+ influx through α7-nAChRs.
What prevents α7-nAChRs from triggering cell death after the final number of neurons is established? In mature neurons, multiple factors control expression and signaling via α7-nAChRs, thus protecting neurons from large, global increases in [Ca2+]i. For example, mature ciliary neurons cluster α7-nAChRs on somatic spines (Shoop et al., 2001, 2002), narrowing the spatial signaling range of Ca2+ to somatic spines, where it can be handled regionally. Furthermore, α7-nAChRs on mature neurons are excluded from postsynaptic densities and therefore are less likely to be activated by the release of acetylcholine from presynaptic terminals (Jacob and Berg, 1983; Horch and Sargent, 1995). Both types of neurons contain a large number of silent α7-nAChRs, suggesting that the majority of α7-nAChRs are normally prevented from signaling and are recruited only in special circumstances (McNerney et al., 2000). In addition, ciliary neurotrophic factor (CNTF), which is expressed by ciliary ganglion target tissues, downregulates the expression of α7-nAChRs (Halvorsen and Berg, 1989). Finally, endogenous neurotoxin-like molecules such as lynx1 are emerging as important modulators of α7-nAChR function (Miwa et al., 1999; Ibanez-Tallon et al., 2002). In fact, ciliary ganglion neurons turn on expression of the lynx-related gene during the period of cell loss, which reduces the Ca2+ influx through α7-nAChRs receptors (Nishi et al., 2006).
Other studies with pharmacological agents that block nicotinic signaling differ from our conclusion that inhibition of α7-nAChRs prevents cell death in the ciliary ganglion. Pan-nicotinic antagonists such as chlorisondamine, pempidine, or d-tubocurare reduce survival of ciliary ganglion neurons (Wright, 1981; Meriney et al., 1987; Maderdrut et al., 1988). Furthermore, removal of afferent input to the ciliary ganglion exacerbates cell death (Furber et al., 1987). These studies together with some cell culture models indicate that depolarization and increases in [Ca2+]i normally enhance survival of ciliary ganglion neurons (Pugh and Margiotta, 2000). However, increases in intracellular-free calcium can be either survival promoting or death inducing, depending on the Ca2+ set-point of a neuron (Johnson et al., 1992; Ghosh and Greenberg, 1995). Therefore, excessive increases in intracellular Ca2+ via α7-nAChRs could exceed threshold for an optimal Ca2+ set-point and induce cell death, whereas moderate increases in intracellular Ca2+ are survival promoting. To this end, we showed that individual E8 ciliary ganglion neurons express heterogeneous levels of α7-nAChRs on their surface. As a result, some neurons display very large increases in [Ca2+]i, whereas others exhibit a much smaller elevation in [Ca2+]i via α7-nAChRs. Therefore, we propose that the neurons that die as a result of α7-nAChR activation are the ones that express high levels of surface α7-nAChRs and generate large [Ca2+]i that exceed the set-point for cell death. Furthermore, activation of heteromeric α3/β4 nAChRs, which is the second type of nAChRs in the ciliary ganglion, leads to smaller increases in [Ca2+]i that could be survival promoting. Thus, the balance between death-inducing signals via α7-nAChRs and survival-promoting signals via α3*-nAChRs could determine which neurons die and which neurons survive.
Although our findings demonstrate that anterograde signaling via α7-nAChRs influence the survival of ciliary ganglion neurons, they do not invalidate the neurotrophic hypothesis. In fact, inhibition of α7-nAChRs with membrane-tethered αbtx rescues only 41% of neurons from dying, which argues that α7-nAChRs are not solely responsible for cell death of the ciliary ganglion neurons. Actually, ciliary ganglion neurons depend on CNTF for survival in vitro and in vivo, and overexpression of CNTF in chicken embryos rescues 31% of ciliary ganglion neurons that would have normally died in vivo (Nishi and Berg, 1979, 1981; Finn et al., 1998). Therefore, we propose that the balance between death-inducing and survival-promoting factors determines the final numbers of neurons in the ganglion. Initially, death signals predominate, which is evidenced by large programmed cell death at E6, several days before the actual decrement in cell number and synaptogenesis with the target tissues (Lee et al., 2001). As neurons extend processes and synapse with their targets, they become dependent on CNTF, which opposes the deleterious effects of α7-nAChRs by downregulating their expression (Halvorsen and Berg, 1989) or upregulating modulators of nAChR function such as lynx-1. In conclusion, our results highlight the importance of nontraditional roles of nAChRs during neural development.
This work was supported by National Institutes of Health Grants NS25767 and DA017784 (R.N.). Imaging was supported by the Centers of Biomedical Research Excellence Program of the National Center for Research Resources (P20 RR16435), and flow cytometry was supported by the Vermont Cancer Center. We thank Drs. Ines Ibanez-Tallon and Nathanial Heintz for providing us with the GPI-αbtx construct. We are also grateful to Drs. Rodney Parsons, Victor May, Jennifer Straub, and Steven Straub for comments on our manuscript.
- Correspondence should be addressed to Dr. Rae Nishi, Department of Anatomy and Neurobiology, 149 Beaumont Avenue, Burlington, VT 05405.