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Department of Zoology, Stockholm University, SE-10691 Stockholm, Sweden
Submitted 14 July 2003; accepted in final form 7 October 2003
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Zordan et al. 2001b), but the identity and mode of actions of the neurotransmitters mediating photic input onto clock neurons is obscure. Since light input pathways and the neuronal organization of the master clock are less complex in larvae than in adults (Kaneko and Hall 2000), larvae offer good possibilities to understand how neurotransmitters mediate photic input onto Drosophila clock neurons. The molecular clock ticks in larvae (Kaneko et al. 1997) and is entrained by light (Kaneko et al. 2000). Larvae lack obvious behavioral and physiological daily rhythms (Sawin et al. 1994), but light pulses during the larval phase synchronize adult locomotor (Kaneko et al. 2000; Sehgal et al. 1992) and pupal eclosion rhythmicity (Brett 1955) within populations of flies raised under constant darkness, demonstrating the larval time-memory (Zeitgedächtnis).
Larval light entrainment is mediated by the blue-light sensitive protein cryptochrome (CRY), located within the clock neurons (Emery et al. 2000), and by a pathway involving a norpA-encoded phospholipase C. Light pulses entrain clock gene oscillation in the lateral neurons (LNs, the putative pacemaker cells for larval time-memory) both in norpAP41 or cryb single-gene mutants, but not in norpAP41; cryb double-mutant larvae (Kaneko et al. 2000), Thus either the CRY or the norpA-dependent pathway appears to be sufficient to entrain the molecular clock in the larval LNs. NORPA is a phospholipase C involved in rhodopsin-based phototransduction (Bloomquist et al. 1988). It is expressed in the Bolwig organ (BO) (Bolwig 1946; Malpel et al. 2002), the only known rhodopsin-based photoreceptor in larvae, which consequently seems to be responsible for norpA-dependent light entrainment (Kaneko et al. 2000). The BO connects to the brain via Bolwig's nerve (BN) and makes putative contact with the LNs (Helfrich-Förster et al. 2002; Kaneko et al. 1997; Malpel et al. 2002). The LNs appear to be the main circadian pacemaker neurons in the larval brain (Kaneko et al. 2000), and express the neuropeptide pigment-dispersing factor (PDF) (Helfrich-Förster 1997). The BO expresses choline acetyltransferase (ChAT) (Yasuyama et al. 1995), suggesting biosynthesis of acetylcholine (ACh). ACh is thus a major candidate for signal transmission between BO and LNs, or, in other terms, a likely input factor of the larval clock subserving light entrainment.
We wanted to investigate whether the LNs express functional receptors for ACh. For that, we used Ca2+ imaging in short-term cultures of dissociated CNS neurons, since it was known that ACh increases the intracellular Ca2+ concentration ([Ca2+]i) in insect neurons (Bicker and Kreissl 1994; Cayre et al. 1999). LNs in culture were identified by GAL4/UAS-directed expression of green fluorescent protein (GFP) (Brand 1999). Since LNs occur in a low number in the CNS, we first established and validated this approach using a gal4-line with a broader, yet specific expression pattern in peptidergic neurons.
We found that dissociated LNs and many other peptidergic neurons express functional nicotinic ACh receptors (nAChRs). Stimulation of the nAChRs leads to an increase of [Ca2+]i. Our results show that ACh is an input factor of the LNs and imply a role for Ca2+ in the photic input onto larval pacemaker neurons.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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For imaging experiments and immunostainings, flies of a w;pdf-gal4 line specific for PDF-expressing neurons (Park et al. 2000) (kind gift of J. Hall, Brandeis, MA) or the c929-gal4 line specific for a large subpopulation of the peptidergic neurons (Hewes et al. 2003) (kind gift of P. Taghert, St. Louis, MO) were crossed with flies of the w*;P(w+mC = UAS-gfp.S65T)T10 strain (Bloomington stock center, strain 1522, donated by C. Goodman). Crosses of the line per-gal4-3 (Kaneko and Hall 2000) (kind gift of J. Hall) with flies of the UAS-gfp.S65T strain were used for PDF immunostainings in cell cultures. To show the close apposition of the BO with the LN dendrites, PDF immunostaining was performed in larvae of a cross of Cha-gal4 (a driver specific for ChAT-expressing neurons (Kitamoto 2001) (kind gift of T. Kitamoto, Duarte, CA) with flies of the UAS-gfp.S65T strain. Crosses of c929-gal4 flies with either a P{w+mC = UAS-AUG-dsRed}Ea, w*;P{UAS-AUG-dsRed}Eb;P{UAS-AUG-dsRed}Ec strain (Bloomington stock center, strain 6280, donated by J. Kasuya and L. Iverson) or UAS-mCD8-gfp (encoding a GFP-variant that is targeted to the cell membrane) (Lee and Luo 1999) were used to test whether expression of fluorescent marker proteins interferes with Ca2+ measurement in neurons.
Flies were kept under light:dark 12:12 h at 18 or 25°C in an incubator.
Cell dissociation and culture
Cell dissociation and culture was modified from Wu et al. (1983) and Kraft et al. (1998). For each experiment, 6-10 feeding third-instar larvae were collected, washed with ddH2O, and sterilized for 1 min by submerging in 70% ethanol, followed by a further wash with ddH2O. Then, brains (for experiments on the LNs) or whole CNS (for other experiments) were dissected out and cleaned from imaginal disks and other adjacent tissues in standard saline. Tissue was incubated in an Eppendorf tube containing 500 µl dissociation saline containing 0.4 mg/ml dispase II (Roche Diagnostics, Mannheim, Germany) and 0.1 mg/ml collagenase I (Sigma-Aldrich, Stockholm, Sweden) for 2 h at 25°C. Thereafter, the tube was centrifuged for 2.5 min at 0.8 rcf, enzyme solution was pipetted off, and 60 µl medium was added.
For imaging experiments, the enzyme-treated tissue was transferred to a cover glass coated with poly-L-lysin (Sigma-Aldrich) and manually dissociated using fine tungsten needles. Dissociated neurons were allowed to attach to the glass for 1.5-4 h prior to imaging.
For cell culture, all subsequent steps were carried out in a laminar flow box. The enzyme-treated tissue was gently triturated in the tube using siliconized fire-polished Pasteur pipettes. The tube was centrifuged as above, the supernatant was pipetted onto a poly-L-lysincoated cover glass, and the pellet again taken up in 60 µl medium. This trituration procedure was repeated twice.
For both imaging experiments and cell cultures, the cover glass was put into small plastic cell culture dishes. To prevent excess evaporation of the medium, small drops of sterilized ddH2O were placed at the rim of the dishes next to the cover glass. Dishes were kept at a constant temperature of 21°C in darkness in a humidified incubator (MLW, Heiligenstadt, Germany).
Salines and media
Standard saline (Jan and Jan 1976) contained (in mM) 128 NaCl, 2 KCl, 4 MgCl2, 1.8 CaCl2, 36 sucrose, and 5 HEPES; pH 7.10. For Na+-free saline, NaCl was substituted with an equal amount of choline chloride. Dissociation saline contained (in mM) 128 NaCl, 2 KCl, 11.8 sucrose, and 5 HEPES; pH 7.10.
Schneider's modified saline (GibcoBRL, Life Technologies, Täby, Sweden) was mixed with 10% heat-treated fetal calf serum (Gibco-BRL), and 1% of an antibiotic solution (10.000 U/ml penicillin, 10 mg/ml streptomycin, 250 µg amphotericin B, Sigma-Aldrich). The medium was sterile-filtered through a 0.2-µm membrane filter and kept at 4°C. For cell culture purposes, 1 µg/ml 20-hydroxyecdysone and 50 µg/ml bovine insulin (Sigma-Aldrich) were added.
Calcium imaging
The cover glass with cells was mounted in an imaging chamber (RC-26, Warner Imaging, Hamnden, CT) with a volume of 500 µl. Cells were washed with 5 ml standard saline using a gravity flow system and loaded for 40 min at room temperature with 2 µM fura-2 acetoxymethylester (Molecular Probes, Leiden, The Netherlands) in standard saline. Thereafter, cells were washed with 3 ml standard saline and left for 30 min in standard saline containing 0.2 mM Ca2+ to allow for cleavage of residual esters. The imaging system consisted of an Axiovert S100 microscope (Zeiss, Jena, Germany) equipped with a Zeiss 40x fluar oil immersion objective (NA 1.3), conventional rhodamine (Zeiss standard set), FITC and fura-2 filtersets [sets 41001 (exciter HQ480/40x, dichroic Q505LP, emitter HQ 535/50) and 71000 (exciter D380/13 and D340/10, dichroic DLL7, emitter 510/40), Chroma, Brattleboro, VT], an Orbit 1 filterwheel (Improvision, Coventry, UK), and a cooled CCD camera (Hamamatsu 4742-95, Hamamatsu). Excitation light provided by a 75-W xenon lamp (Hamamatsu L2194-01, Hamamatsu) was attenuated by a 10% quartz neutral density filter (Chroma). OpenLab 3 software (Improvision) on a Macintosh G4 PowerPC was used for system control and data acquisition. Ratio images were typically acquired with an intensity resolution of 12 bit at 0.2-1 Hz after background subtraction, with 2 x 2 binning resulting in a pixel size of 0.5 µm. Data processing was performed using Microsoft Excel or GraphPad Prism3 software. Dose-response curves were fitted using a four-parameter equation with variable slope. The intracellular free Ca2+ concentration [Ca2+]i was estimated to be equal to Kd(R - Rmin/Rmax - R)(F380max/F380min) (Grynkiewicz et al. 1985), with R being the ratio measured during the experiment. Rmax, Rmin, F380max, and F380min were determined in vivo after experiments by exposure of the cells to standard saline containing 10 mM Ca2+ and 2 µM ionomycin, and 0 mM Ca2+, 5 mM EGTA, and 2 µM ionomycin, respectively. A Kd of 147 nM was adopted from Berke and Wu (2002).
Drugs
Drugs were pipetted directly into the bath in a volume of 2 ml. (-)Scopolamine was from Fluka; all other drugs were produced by Sigma. All chemicals were purchased from Sigma-Aldrich. Drosophila PDF was custom-synthesized by Åke Engström (Uppsala, Sweden).
Immunocytochemistry
WHOLE MOUNTS OF LARVAL CNS. The CNS of third-instar larvae was dissected and fixed for 2 h at 4°C in 0.1 M sodium phosphate-buffered saline (PBS) containing 4% paraformaldehyde. The fixed tissue was washed three times for 15 min with 0.01 M PBS containing 1% Triton X-100 (PBT) and incubated with primary antibodies diluted in PBT and 10% normal goat serum (DAKO, Glostrup, Denmark) overnight at room temperature on a shaker. Over the next day, the tissue was washed repeated times with PBT before overnight incubation with the secondary antiserum in PBT and 0.5% bovine serum albumin at room temperature on a shaker. The following day, tissue was again washed repeated times with PBT, washed twice with 0.01 M PBS, and mounted in glycerol/PBS (80:20).
CULTURED CELLS. Cultured cells were washed twice with standard saline, fixed at 4°C for 30 min in 0.1 M phosphate buffer containing 4% paraformaldehyde and 4% saccharose, and permeabilized by three 10-min washes with 0.05 M PBT. Blocking was performed with 10% normal goat serum in PBT for 1 h at room temperature, followed by overnight incubation at 4°C with primary antibodies diluted in PBT and 10% normal goat serum. The next day, cells were washed five times for 10 min in PBT and incubated for 1 h at room temperature with the secondary antibodies diluted in PBT + 10% normal goat serum. This was followed by five 10-min washes with PBT, two short washes with 0.1 M Tris-HCl, pH 8.0, and mounting in polyvinylalcohol/glycerol containing 2% of the antibleaching reagent DABCO (Sigma-Aldrich) in 0.2 M Tris-HCl, pH 8.0.
ANTIBODIES. The antibodies against different nAChR subunits were a kind gift of Dr. Eckart Gundelfinger (Magdeburg): rabbit anti-SBD "4596," rabbit anti-ALS "9877," mouse mAb anti-ALS "D4," rabbit anti-D
3 "D 3," mouse mAb anti-D 2 "1f6," and mouse mAb anti-ARD "3d2." Dilutions were 1:250-500 for the rabbit antisera and 1:50-100 for the mouse mAb. The production and characterization of these antibodies have been described previously (Chamaon et al. 2000, 2002; Schuster et al. 1993). In our hands, the antisera "4596" and "D 3" stained either the nucleus or the cytoplasm of many CNS neurons, indicating a lack of specificity for nAChRs. Therefore these antibodies were excluded from further studies. Production and specificity of the rabbit polyclonal antiserum K 9914/13 against Drosophila PDF has been described by Persson et al. (2001). We could also confirm that it specifically stains the PDF neurons in larval and adult Drosophila. In enzyme-linked immunosorbent assays, it recognized both the oxidized and unoxidized form of Drosophila PDF in wildtype brain extracts, and there was an absence of immunoreactivity in extracts from pdf0 mutant flies (unpublished observations). The polyclonal and unabsorbed rabbit anti-Pea-PVK2-serum was a kind gift of Dr. Manfred Eckert (Jena, Germany). It recognizes peptides with the C-terminal sequence PRXamide (Eckert et al. 2002) and specifically stains the pyrokinin- and periviscerokinin-containing neurons in the larval and adult nervous system of Drosophila (unpublished observations). The rabbit anti-FMRFamide-serum 117I was a kind gift of Cornelis J.P. Grimmelikhuijzen (Copenhagen). Its production and specificity are described by Grimmelikhuijzen (1983). All peptide antisera were applied at a dilution of 1:5,000.
As secondary antibodies, Cy3- or Cy2-conjugated AffiniPure goat-anti mouse or goat-anti rabbit IgG (H+L; Jackson ImmuoResearch, West Grove, PA) were used at a dilution of 1:2,000.
All final antibody solutions were centrifuged at 10.000 rcf and 4°C for 10 min before use.
MICROSCOPY. Epifluorescent microscopic images were acquired with a motorized Axioplan 2 microscope (Zeiss) coupled to a cooled CCD-camera (Hamamatsu 4742-95, Hamamatsu) and stored and processed using OpenLab 3 software (Improvision). Images were acquired by focusing through the object in 1-µm steps. The resulting images were corrected for out-of-focus light by nearest-neighbor deconvolution and finally merged. Confocal stacks were acquired in the multi-track mode with a Zeiss LSM 510 confocal microscope based on an Axiovert S100 M microscope (Zeiss) and processed with OpenLab and LSM software. Intensity profiling was performed with the public domain National Institutes of Health Image software (http://rsb.info.nih.gov/nih-image/). Microscopes were equipped with narrow band FITC and Cy3 filter sets. All colors shown are pseudocolors.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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The pattern and morphology of the GAL4-driven GFP expression used to identify cells in culture is shown and described for the intact larval CNS in Fig. 1. The pdf-gal4 driver specifically marked the LNs in the brain. The c929-gal4 driver was well suited to establish and validate the calcium-imaging method described in this investigation, since it marked more neurons than the pdf-gal4 driver and is specific for peptidergic neurons (Hewes et al. 2003). In the following, GFP-expressing brain neurons of the pdf-gal4 crosses will be referred to as LNs and those of the c929-gal4 crosses as c929 neurons. Both will be summarized under the term GFP neurons as opposed to non-GFP neurons, i.e., neurons that do not express GFP.
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5 days in culture (Fig. 2). A typical dissociation yielded around 10-20 c929 neurons and 1-3 LNs. Thus it was possible to routinely obtain identified and isolated GFP neurons (Fig. 2, A-G) even in the case of the LNs, which occur only in small numbers (8 neurons per larval brain). Staining of cultures with antisera against FMRFamide-like or pyrokinin/periviscerokinin-like peptides showed that other specific peptidergic cell types besides the LNs grew out and expressed and transported peptides in vitro (Fig. 2, H-I). A common feature of peptidergic insect neurons in vivo is varicosities along their neurites that show enhanced peptide immunoreactivity. These varicosities could represent storage or release sites for peptide-containing vesicles (see Nässel 2002). Also in vitro, prominent varicosities with strong peptide immunoreactivity were visible along the neurites of the stained peptidergic neurons (Fig. 2, A-E, H, and I).
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Acutely dissociated GFP neurons used in the imaging experiments had an estimated basal [Ca2+]i of 135 ± 21 nM (LNs, n = 20) or 74 ± 11 nM (c929 neurons, n = 20). These [Ca2+]i values are in agreement with those found in other acutely dissociated insect neurons (e.g., Cayre et al. 1999; Grolleau et al. 1996; Messutat et al. 2001). This agreement suggests that our estimates of [Ca2+]i based on in vitro calibration are reasonable. Calibration in vivo has several advantages over that in vitro. Still, inherent difficulties to correctly determine Rmax and Rmin in vivo (Williams and Fay 1990) might have led to an overestimation of the actual [Ca2+]i, at high ratios. Application of saline containing 50 mM K+ typically resulted in a strong rise of [Ca2+]i in GFP as well as non-GFP neurons, demonstrating that dissociated neurons are excitable and actively regulate [Ca2+]i.
To check the specificity of the pdf-gal4-driven GFP expression in acutely dissociated neurons, we fixed the cells after imaging experiments and immunostained them for PDF. All 11 GFP-expressing neurons of pdf-gal4 crosses obtained in five experiments stained positive for PDF, and conversely, all PDF-immunoreactive neurons expressed GFP. Thus we concluded that GFP marker expression is a reliable marker for LNs even in dissociated neurons.
Effect of ACh and other neurotransmitters on [Ca2+]i in dissociated neurons
In a first set of experiments, we tested the effect of various transmitters at fairly high doses on [Ca2+]i of GFP and non-GFP neurons. The c929 reporter gene is expressed in around 200 heterogenous peptidergic neurons (but not the LNs) that express a variety of different neuropeptides (Hewes et al. 2003). However, the response of c929 cells to ACh was remarkably homogeneous. Application of 10 µM ACh induced an increase of [Ca2+]i in 16 of 21 (=76%) tested LNs, 18 of 22 (=82%) tested c929 neurons, but only in 43.3 ± 3.4% of the non-GFP neurons (n = 2,227). Dopamine at 100 µM was able to induce an increase in [Ca2+]i in 4 of 13 tested LNs (=31%), 1 of 12 tested c929 neurons (=8%), and 3.9 ± 1.0% of non-GFP neurons (n = 1,049). Glutamate, 5-HT, octopamine, and histamine, all at 100 µM, increased [Ca2+]i in 1-3% of non-GFP neurons but were ineffective in GFP neurons. An effect of 1 µM Drosophila PDF on the [Ca2+]i of LNs or non-GFP neurons could not be observed in three different experiments.
Since ACh was the only neurotransmitter that consistently increased [Ca2+]i in GFP neurons, we focused on this transmitter and constructed dose-response curves. The ACh doseresponse curves for GFP neurons expressing S65T.GFP showed a threshold concentration of 10-100 nM ACh with an EC50 of 91 ± 11 nM for c929 neurons (n = 8) and 165 ± 100 nM for LNs (n = 5) (Fig. 3A). The EC50 of 1.3 ± 1.1 µM in non-GFP neurons (n = 15) was about one magnitude higher than the EC50 of GFP neurons (Fig. 3A), but still a magnitude lower than the EC50 of ACh on cultured larval Drosophila neurons found by Albert and Lingle (1993). Interestingly, the EC50 was shifted to 452 ± 150 nM in c929 neurons expressing DsRed (n = 6) and 1.3 ± 1.2 µM in c929 neurons expressing CD8-GFP (n = 6, Fig. 3B). Although we did not further characterize this shift to a higher EC50 in neurons expressing these fluorescent marker proteins, we suppose that both DsRed and CD8-GFP might interfere with cell signaling. DsRed is known to oligomerize (Baird et al. 2000; Sacchetti et al. 2002), which might interfere with Ca2+ signaling and cell function. The UAS-DsRed strain used in this investigation has UAS-DsRed inserts on all three chromosomes, which might result in high expression of DsRed. Targeting of GFP to the cell membrane in c929 neurons expressing CD8-GFP might have interfered with signaling molecules (ion channels, receptors) in the plasma membrane.
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). In total, 92% of the 23 tested c929 neurons, and 75% of the 8 tested PDF neurons displayed increases in [Ca2+]i on application of ACh. These percentages obtained from dose-response experiments are in good agreement with those obtained by application of 10 µM ACh only.
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ACh can exert its effect either via ionotropic nicotinic receptors (nAChR) or via metabotropic muscarinic receptors (mAChRs). To test which receptor types are involved, we tested the effect of the nAChR agonist nicotine and the mAChR agonist pilocarpine. The threshold concentration for an effect of nicotine varied between 10 and 100 nM for both GFP and non-GFP neurons, whereas the EC50 was not statistically different (P > 0.05, one-way ANOVA) between GFP and non-GFP neurons [1.6 ± 1.1 µM for LNs (n = 7), 4.2 ± 1.0 µM for c929 neurons (n = 6), and 2.4 ± 1.2 µM for nonGFP neurons (n = 12), Fig. 3C]. Desensitization at high concentrations was much more pronounced for nicotine than for ACh (Fig. 4). Unlike nicotine that increased [Ca2+]i in all GFP neurons tested (n = 7 each for LNs and c929 neurons), pilocarpine was unable to induce an increase in [Ca2+]i in GFP neurons during dose-response experiments (n = 6 for LNs, n = 9 for c929 neurons; Fig. 5, A and B). Two cells were, however, reacting at high concentrations of pilocarpine (0.1 and 1 mM, Fig. 5B). At these high concentrations, pilocarpine might act as a weak agonist of nAChR, which is also suggested by our finding that a single application of 0.1 mM pilocarpine induced [Ca2+]i increases in 8 of 14 (=57%) tested LNs and 9 of 17 (=53%) tested c929 neurons.
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To further test the involvement of nicotinic and muscarinic AChR, we tested the effect of the nAChR antagonist mecamylamine and the mAChR antagonist scopolamine on ACh-induced [Ca2+]i increases. Prior to application of ACh, cells were incubated for 20 min with either 100 µM scopolamine or 10 µM mecamylamine. A blocking effect of scopolamine was not detectable in any of the tested GFP neurons (n = 6 for LNs, n = 2 for c929 neurons). Mecamylamine completely blocked the effect of ACh in 69% of the LNs (n = 13) and in 93% of the c929 neurons (n = 15). Lower mecamylamine concentrations (
1 µM) did not result in a complete block.
ACh-induced Ca2+ increase depends on Ca2+ influx
On application of 10 µM ACh in Ca2+-free saline containing 0.5 mM EGTA, neither GFP nor non-GFP neurons showed an increased [Ca2+]i (n = 5 for LNs, n = 7 for c929 neurons; Fig. 5D). An increase of [Ca2+]i in Ca2+-free saline containing 0.5 mM EGTA could, however, be triggered in some neurons by application of 20 mM caffeine, an agonist of ryanodine receptors at the endoplasmic reticulum (Fig. 5D). This suggests that the intracellular Ca2+ stores were not emptied by incubation in Ca2+-free extracellular saline. Bath application of the nonspecific Ca2+ channel blockers MnCl2 (2 mM) or LaCl3 (100 µM) also completely blocked ACh-induced increases in [Ca2+]i (n = 2 and 4 for c929 neurons and LNs, respectively). ACh was also unable to induce increases of [Ca2+]i in Na+- free saline (n = 3 for LNs, n = 5 for c929 neurons, Fig. 5C).
nAChR subunits locate to the putative contact area of LN dendrites and BN axon terminals
The BN enters the brain and projects to the larval optic neuropil (Fig. 6A) where the BN axon terminals branch (Fig. 6, A-C). Also, the LN dendrites project into the larval optic neuropil, where they are in close apposition to the more dorsal portion of the BN terminals (Fig. 6, A-C).
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-subunits D 2 and ALS as well as the 
-subunit ARD produced distinct immunolabeling in central neuropil areas of the intact larval CNS similar to the staining pattern already described for the ARD and ALS subunits (Schuster et al. 1993). Strongest staining was observed in the neuropil areas of the central brain (Fig. 6D). Only very few cell bodies were immunoreactive (Fig. 6G). Distinct immunolabeling with the different receptor antibodies was also visible in the larval optic neuropil (Fig. 6, D, F, and G), although staining intensity was lower than in the neuropil areas of the central brain (Fig. 6, D and G). The difference in staining intensity between the larval optic neuropil and the neuropil of the central brain is consistent with differences in ChAT-immunostaining intensities between the larval central brain and optic neuropil (Yasuyama et al. 1995). The stained area within the optic neuropile included the region where the LN dendrites are located (Fig. 6, D-I). GFP-signals from the LN dendrites and nAChR immunoreactive material were colocalized in voxels of single confocal sections with a dimension smaller than the LN dendrites (Fig. 6, G-I). In contrast, neither the area including the dorsal branches (the putative PDF release site) nor the neurites and somata of the LNs exhibited nAChR immunoreactivity above the mean staining intensity outside the central neuropil (Fig. 6, D-F).
The finding that nAChR staining within the larval optic neuropil is present in an area slightly larger than the putative contact site of the LNs with the BN axon terminals suggests that cholinergic synapses are distributed throughout the optic neuropil. Since functional impairment of the BO also affects the modulation of locomotion by light (Busto et al. 1999), it is possible that further neurons, additional to the LNs, are postsynaptic to the BO neurons and might have contributed to the nAChR staining in the optic neuropil.
nAChR locate to dissociated LNs
Since our Ca2+ imaging experiments showed that the LNs express functional nAChR, we immunostained acutely dissociated LNs using the different antibodies directed against the subunits D 2, ALS, and ARD. With all four antibodies, immunolabeling was visible in LNs (Fig. 6, J-P). The distribution of the nAChR immunoreactivity was very similar to that found in dissociated vertebrate neurons (Atluri et al. 2001; Zarei et al. 1999) and might at least in part be due to intracellular immunoreactive material.
Only very few neuronal cell bodies could be stained with nAChR antisera in whole mounts of larval brains (Fig. 6G). The discrepancy between the scarceness of cell body staining in situ and its presence in vitro is explainable by assuming that the amount of nAChR molecules in the cell body is too low to result in immunostaining above background levels in the surrounding tissue in situ, whereas staining becomes discernible in isolated neurons where background fluorescence is absent. It is also possible that the freshly dissociated neurons immediately increased their production of nAChRs, which should result in increased cytoplasmic staining. Furthermore, many neurons showed neurite stumps directly after dissociation that often retracted quickly, resulting in rounded-up neurons. These retractions could have resulted in an enriched density of nAChR in the cell body plasma membrane, yielding higher staining intensity.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Using a pharmacological approach, we have demonstrated that nAChRs are functionally expressed by LNs in vitro, and once activated, increase the [Ca2+]i in these clock cells. However, although more than 75% of the tested LNs reacted to ACh, some did not. Those nonresponding cells might have been impaired by the dissociation process. Still, it is also possible that the LNs are heterogenous in their expression of neurotransmitter receptors and that particular LNs do not express functional AChRs. Such heterogeneity could also explain why an effect of dopamine on [Ca2+]i was found in around 30%, but not all, of the tested cells.
The reported effect of ACh on LNs represents the first report on identified transmitter action on a Drosophila clock neuron and lends further support to a possible role of ACh as a transmitter signaling light information from the BO to the LNs (see Introduction). Immunostaining located three different nAChR subunits to the larval optic neuropil in vivo, including the region where the BN axon terminals are in close apposition to the LN dendrites. Furthermore, nAChR immunofluorescence could be found in the same voxels as LN dendrites in single confocal sections. These findings in combination with the absence of measurable nAChR immunoreactivity in the soma and dorsal branch suggest that also in vivo nAChR are expressed by the LNs and are located to their dendrites. However, the resolution of fluorescence confocal microscopy is not sufficient to unequivocally reveal receptor localization on fine dendrites. Thus electron microscopy studies need to be carried out to show the presence of nAChR on LN dendrites. Such studies could also reveal whether BN axon terminals and LN dendrites make synaptic contacts as suggested by their close apposition described in this and other investigations (Helfrich-Förster et al. 2002; Kaneko et al. 1997; Malpel et al. 2002) as well as by a functional study (Kaneko et al. 2000).
A possible role for ACh-induced increases of [Ca2+]i in electrical activity of the LNs
Our experiments show that activation of nAChRs in dissociated larval LNs leads to an influx of extracellular Ca2+ that depends on the presence of extracellular Na+. We therefore infer that activation of the nAChRs results in an influx of Na+. The subsequent membrane depolarization may trigger an influx of extracellular Ca2+ via voltage-dependent Ca2+ channels.
Although freshly dissociated cells have been widely used to demonstrate receptor or channel expression in a variety of insect neurons by patch-clamp electrophysiology, it is not yet clear whether receptors at the synapses have the same properties as those characterized on the isolated cell bodies (see Burrows 1996). Our immunostainings suggest that nAChR are located to the dendrites rather than to the soma or dorsal branch of the LNs. If this is true, it is thus not certain whether ACh in vivo induces an increase in [Ca2+]i comparable to that found in vitro. Still, ACh-induced increases in [Ca2+]i would fit well with a recent model for the role of electrical activity in rhythm generation in adult s-LNvs (Nitabach et al. 2002) that proposed light-driven synaptic inputs to be mediated via an influx of Na+ and Ca2+. The indirect way of increasing [Ca2+]i proposed here would allow fine-tuning of the effect of ACh by other neurotransmitters that affect the membrane potential or Ca2+ channel properties. In mammals, light-induced phase shifts of the pacemaker neurons of the suprachiasmatic nucleus (SCN) are mediated by glutamate and the neuropeptide PACAP (see Reppert and Weaver 2002). The glutamate-induced increase in [Ca2+]i in the SCN pacemaker neurons is mediated in part by membrane depolarization-dependent opening of voltage-dependent L-type Ca2+ channels. PACAP modulates the conductance of these channels, thereby facilitating and potentiating the glutamate-induced increase in [Ca2+]i (Dziema and Obrietan 2002). Some candidate modulators of the LNs can be suggested in insects. In flies, including Drosophila, serotonergic neurons project to the larval optic lobe (Nässel et al. 1987; unpublished observations). GABA and an allatotropin-like neuropeptide seem to be involved in photic entrainment of the cockroach Leucophaea maderae (Petri et al. 2002). It has to be tested whether these or other substances such as PDF modulate the ACh-induced increase in [Ca2+]i.
A possible role for ACh-induced increases of [Ca2+]i in light entrainment of the LNs
The major pathway for light entrainment of the molecular clock seems to be a light-dependent degradation of TIM (Suri et al. 1998; Yang et al. 1998). In peripheral tissue and most adult clock neurons, CRY is the main mediator in this light-dependent TIM degradation (Emery et al. 1998; Stanewsky et al. 1998). In adult cryb mutants, PER/TIM cycling is absent in photoreceptor and glia cells, reduced in most clock neurons, but nearly unchanged in the small ventral LNs (s-LNvs) (Helfrich-Förster et al. 2001; Stanewsky et al. 1998), the counterpart of the larval LNs (Helfrich-Förster et al. 2002). Concomitantly, adult cryb mutants entrain their locomotory rhythms to light (Helfrich-Förster et al. 2001; Stanewsky et al. 1998; Zordan et al. 2001a). Thus light-dependent, but CRY-independent, entrainment mechanisms seem to exist in adult s-LNvs. In larvae, light entrainment is mediated by CRY and a pathway involving norpA (Kaneko et al. 2000) expressed in the BO (Malpel et al. 2002). Since the BO is likely to use ACh as a neurotransmitter to signal to the LNs, it is possible that the ACh-induced increase in [Ca2+]i shown in this investigation is part of the norpA-dependent, but CRY-independent, mechanism for light entrainment of the larval LNs. However, a role of Ca2+ in entrainment of molecular and behavioral rhythms has not been indicated for Drosophila, possibly because it is not trivial to genetically explore the role of a ubiquitous and multifunctional second messenger.
In other animals, Ca2+ plays an important role in light entrainment of biological clocks. In molluscs, light-induced phase shifts of the pacemaker neurons are caused by a light-regulated influx of extracellular Ca2+ (Colwell et al. 1994; McMahon and Block 1987). In mammals, calcium imaging with fura-2 showed that both glutamate and PACAP increase [Ca2+]i in SCN neurons (Dziema and Obrietan 2002; Kopp et al. 1999), triggering expression of per genes via a Ca2+- dependent phosphorylation of CREB involving calcium/calmodulin-dependent protein kinase II (CaMKII) (Akiyama et al. 2001; Schurov et al. 1999). In Drosophila, CaMKII was found to be expressed in the adult s-LNvs (Takamatsu et al. 2002). ACh might also play a role in photic input to the adult s-LNvs. During metamorphosis, rhodopsin 6 - containing photoreceptor cells of the BO redifferentiate into the adult extraretinal eyelet (Helfrich-Förster et al. 2002). This adult BO remnant not only retains a functional role in light entrainment of the LNs (Helfrich-Förster et al. 2001, 2002) but also retains ChAT-expression (Yasuyama and Meinertzhagen 1999).
Nicotinic AChRs might be universal to peptidergic neurons of the larval CNS
Synaptic transmission via nAChR is believed to play a major role in the insect nervous system (Lee and O'Dowd 1999; Yasuyama and Salvaterra 1999). The finding that 43% of the non-GFP neurons react to ACh with an increase in [Ca2+]i lends further support to this hypothesis. It is surprising that 92% of the tested peptidergic c929 neurons reacted to ACh, indicating that ACh has a function common to peptidergic neurons. Anatomical studies have not previously indicated expression of nAChR subunits on peptidergic insect neurons (Hermsen et al. 1998; Jonas et al. 1994; Schuster et al. 1993), and reports on nicotine responsiveness of peptidergic insect neurons are not available with exception for the vasopressin-like immunoreactive interneuron in Locusta migratoria (Baines and Bacon 1994). We can only speculate about the reason for this obviously universal expression of nAChR by peptidergic neurons, which possibly is related to the integrating function of peptides as neuromodulators. Peptidergic neurons are often presumed to receive widespread inputs from different neuron types. Therefore the probability to receive cholinergic presynaptic input should be very high for peptidergic neurons, since ACh seems to be the main transmitter in the insect CNS.
In contrast to the paucity of data for nAChR, pharmacological (Baines and Bacon 1994; Lundquist et al. 1998) and immunocytochemical (Aizono et al. 1997; Shirai et al. 2001) studies in different insect species demonstrated mAChRs on peptidergic neurons. A mAChR could also be immunolocalized to cell bodies in the pars intercerebralis of adult Drosophila (Harrison et al. 1995), an area of the protocerebrum where many peptidergic neurosecretory cells reside. Activation of a cloned Drosophila mAChR expressed in a Drosophila cell line lead to an increase in [Ca2+]i (Millar et al. 1995). Our results do not support the presence of a mAChR-mediated increase in [Ca2+]i in peptidergic neurons. We can, however, not exclude that the enzyme treatment during cell dissociation impaired mAChR function on the isolated neurons. It might also be possible that pilocarpine addition during the dose-response experiments resulted in a complete desensitization already at concentrations too low to evoke increases in [Ca2+]i.
Live cell imaging is a convenient method to study the physiology of dissociated neurons identified by GAL4-driven expression of fluorescent marker proteins
In this study, noninvasive live cell imaging was employed to overcome the "formidable barrier to functional analysis of neuronal properties" offered by the "small size and inherent complexity of the fruit fly CNS " (cf. Rohrbough et al. 2003). Although our results do not contribute to a functional understanding of the role of ACh-induced increases in [Ca2+] in LNs, we provided a first example how imaging techniques can be used to unravel the properties of Drosophila clock neurons. A combination of primary cell cultures with calcium imaging has previously been used to study the effect of neurotransmitters on Kenyon cells of the mushroom bodies of honeybees (Bicker and Kreissl 1994) and crickets (Cayre et al. 1999). Since Kenyon cells are the most numerous neuron type in the insect brain and are densely packed in a tight cluster, they could be obtained as a nearly homogenous population by excision and subsequent dissociation of the mushroom bodies, even in Drosophila (Kraft et al. 1998). However, it seems impossible to identify in vivo and specifically dissect and dissociate most other neuron types. We found that the GAL4/UAS technique can successfully be used to reliably identify neurons in primary cell cultures obtained after a relatively simple dissociation of whole CNS. The finding that LNs can efficiently be brought into culture shows that this in vitro assay is not restricted to Kenyon cells (Kraft et al. 1998), but can be useful even for neuron types that occur in low numbers in the CNS, provided that gal4 lines with sufficient specificity are available. Acutely dissociated GFP-expressing neurons attached, kept their internal [Ca2+]i at normal levels, reacted to depolarization, and responded to application of a neurotransmitter with high sensitivity. The development of UAS constructs of different fluorescent marker proteins covering a broad range of emission/absorption wavelengths (e.g., Halfon et al. 2002) allows the use of a variety of fluorescent dyes to monitor changes in Ca2+, pH, membrane potential, and cAMP (see Yuste et al. 2000). The availability of genetically expressed fluorescent Ca2+- sensitive proteins as UAS constructs (Diegelmann et al. 2002; Wang et al. 2003; Yu et al. 2003) will further facilitate the use of dissociated neurons.
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ACKNOWLEDGMENTS |
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GRANTS
This work was supported by grants from the Human Frontier Science Program and the Karl Trygger's Foundation, both to D. R. Nässel.
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FOOTNOTES |
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Address for reprint requests and other correspondence: C. Wegener, Dept. of Biology, Animal Physiology, Karl-von-Frisch-Str., D-35032 Marburg, Germany (E-mail: wegener{at}staff.uni-marburg.de). D. R. Nässel (E-mail: dnassel{at}zoologi.su.se).
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