Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M306417200 on August 1, 2003

J. Biol. Chem., Vol. 278, Issue 42, 41099-41108, October 17, 2003
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
278/42/41099    most recent
M306417200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rieckhof, G. E.
Right arrow Articles by Littleton, J. T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rieckhof, G. E.
Right arrow Articles by Littleton, J. T.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Presynaptic N-type Calcium Channels Regulate Synaptic Growth*

Gabrielle E. Rieckhof, Motojiro Yoshihara, Zhuo Guan and J. Troy Littleton {ddagger}

From the Picower Center for Learning and Memory, Department of Biology and Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139

Received for publication, June 17, 2003 , and in revised form, July 21, 2003.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Voltage-gated calcium channels couple changes in membrane potential to neuronal functions regulated by calcium, including neurotransmitter release. Here we report that presynaptic N-type calcium channels not only control neurotransmitter release but also regulate synaptic growth at Drosophila neuromuscular junctions. In a screen for behavioral mutants that disrupt synaptic transmission, an allele of the N-type calcium channel locus (Dmca1A) was identified that caused synaptic undergrowth. The underlying molecular defect was identified as a neutralization of a charged residue in the third S4 voltage sensor. RNA interference reduction of N-type calcium channel expression also reduced synaptic growth. Hypomorphic mutations in syntaxin-1A or n-synaptobrevin, which also disrupt neurotransmitter release, did not affect synapse proliferation at the neuromuscular junction, suggesting calcium entry through presynaptic N-type calcium channels, not neurotransmitter release per se, is important for synaptic growth. The reduced synapse proliferation in Dmca1A mutants is not due to increased synapse retraction but instead reflects a role for calcium influx in synaptic growth mechanisms. These results suggest N-type channels participate in synaptic growth through signaling pathways that are distinct from those that mediate neurotransmitter release. Linking presynaptic voltage-gated calcium entry to downstream calcium-sensitive synaptic growth regulators provides an efficient activity-dependent mechanism for modifying synaptic strength.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Axonal sprouting and synaptic rewiring are key regulators of neuronal plasticity in the developing and adult brain (1, 2). Both synapse formation (3) and synapse elimination (4) have been suggested to underlie structural changes in the wiring of the nervous system important for long term modification of brain function. Modulation of synapse formation in Drosophila has been implicated in olfactory learning (5). Likewise, long term habituation and sensitization in Aplysia are associated with short term alterations in synaptic connections between sensory and motor neurons (6). Long term potentiation in the hippocampus is accompanied by changes in the number, size, and structure of synapses within the affected circuit (7, 8). One signaling pathway required for these structural changes is the cyclic AMP cascade, which participates in hippocampal long term potentiation (9), long term facilitation in Aplysia (10), and activity-dependent plasticity in Drosophila (11). Notably, the synaptic structure at neuromuscular junctions (NMJs)1 is dramatically altered in Drosophila mutants defective in cAMP metabolism (12). In addition to cAMP, one synaptic signaling molecule that may function in synaptic development is calcium, which has numerous downstream effectors implicated in behavioral and synaptic plasticity.

Voltage-gated calcium channels are a major source of calcium entry into synapses and are multimeric proteins containing a pore-forming {alpha}-subunit and accessory {beta}- and {alpha}2{delta}-subunits. A large family of these channels has been identified with isoforms classified according to their gating kinetics, single channel conductances, and pharmacological profiles (13). Five high threshold subtypes (L, N, P, Q, and R), together with the low threshold T-type, form the major calcium entry pathways in mammalian excitable cells. Genome sequencing has revealed a conserved set of L-, T-, and N-type calcium channels throughout vertebrate and invertebrate evolution (14, 15). Localized elevation of calcium concentration during presynaptic depolarization triggers the fusion of synaptic vesicles and subsequent neurotransmitter release. However, other roles for presynaptic calcium entry during synaptic transmission are poorly defined.

The Drosophila NMJ provides a useful model system for determining the role of N-type calcium channels in synapse formation. Shortly after motor neurons leave the central nervous system and contact their muscle target, there is a well defined morphological transition from growth cone to pre-varicosity to varicosity that occurs over several hours (16). Following initial synapse formation, there is a second phase of synaptic development that results in an ~10-fold increase in varicosity number on the well studied ventral muscle fibers 6 and 7 during the period between embryogenesis to the third larval instar phase. This growth process is characterized by the addition of new varicosities and additional synaptic branches (17). For new synaptic connections to form, changes in existing terminals must allow for directed synaptic growth and sprouting. These structural changes require temporally controlled and spatially regulated modifications to the nerve terminal cytoskeleton and trans-synaptic cell adhesion complexes. Several calcium-dependent proteins have been implicated in synaptic growth, but the source of calcium and its regulation are unknown. Here we demonstrate that presynaptic N-type calcium channels contribute to the regulation of synaptic growth. The requirement of N-type channels for synaptic growth is not dependent upon its role in triggering neurotransmitter release, but rather reflects a requirement for calcium influx. These findings indicate N-type calcium channels provide an important source of calcium that regulates synaptic growth.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Drosophila Genetics—Drosophila melanogaster was cultured on standard medium at 22 °C. Temperature-sensitive behavioral mutants on the X chromosome were generated by feeding isogenized Canton S males with ethyl methanesulfonate (EMS) for 12 h and mating to C (1)DX females via standard techniques (55). Recombination mapping was performed with a y cho cv v f marked strain. Rescue experiments were performed using a duplication (Dp(1;2)v65b) covering the Dmca1A locus (19). The Dmca1A13 mutant has also been referred to as l(1)L1320-3 and l(1)11Aa13 (19, 42).

Electrophysiology and Morphological Analysis—Electrophysiological analysis of wandering third stage instar larva was performed in Drosophila saline (NaCl, 70 mM;KCl,5mM; MgCl2,4mM; NaHCO3,10mM; Trehalose, 5 mM; sucrose, 115 mM; HEPES-NaOH, 5 mM, pH 7.2, modified from HL3) (56) with the indicated extracellular calcium concentration using an Axoclamp 2B amplifier (Axon Instrument) at 22 °C. Extracellular field potentials were recorded by placing a sharp glass electrode near the longitudinal flight muscles after piercing the cuticle, with a reference electrode in the fly head. Temperature shifts were performed by heating mounting clay encompassing the fly to the desired temperature with a peltier heating device. Immunostaining was performed on third instar larvae at wandering stage after rearing at 22 °C. Larvae were dissected in Drosophila saline and fixed in 4% formaldehyde in HL3 for 45 min. Immunostaining on filleted embryos was performed as described previously (16). Embryos were dissected at 21–24 h (25 °C) after fertilization. Fluorescein isothiocyanate-conjugated IgG against HRP, which labels neuronal cell membranes (57), was purchased from Cappel and used at 1:1000. DSYT2 against synaptotagmin (58) was used at 1:1000. Anti-CSP (59) was used at 1:100 and anti-DLG at 1:1000. Immunoreactive proteins were visualized on a Zeiss Pascal Confocal using fluorescent secondary antibodies (Molecular Probes, Chemicon, The Jackson Laboratory). All error measurements are S.E. unless indicated.

dsRNA Interference—Wild-type embryos were collected at stage 5, dechorionated, mounted on coverslips with double-stick tape, and allowed to desiccate for 6 min at 18 °C. The embryos were covered with halocarbon oil and injected with ~34 picoliters of a 3 µM solution of Dmca1A dsRNA in injection buffer or with injection buffer alone. dsRNA was isolated by PCR amplification of a 725-bp region corresponding to Dmca1A nucleotides 50–772 (following the initiating ATG) using gene-specific PCR primers with the T7 RNA polymerase sequence attached to the 5' end: primer 1, 5'-gaattaatacgactcactatagggagacattgtttattctaaccgag-3'; and primer 2, 5'-gaattaatacgactcactatagggagacgatattatcgaaactcgtg-3'.

Two µg of PCR product was used directly in an in vitro RNA transcription reaction with T7 RNA polymerase using the Megascript Kit (Ambion) and purified with the Mega Clear Kit (Ambion). dsRNA was evaluated by gel electrophoresis prior to use. Following injection, embryos were incubated at 18 °C until hatching, at which time they were transferred to grape juice plates with yeast paste. Larvae were allowed to develop into third instar larvae at 25 °C and processed for immunocytochemistry as described above.

Sequencing—Dmca1ANT27 and CS cDNA was used as template to PCR-amplify the complete ORF in two segments. High fidelity Taq (Takara) was used for these reactions to minimize PCR-introduced errors. Sequencing primers spanning the full ORF were used to sequence in duplicate from either of these two templates in both the forward and reverse direction. Sequences were aligned and compared with the predicted ORF using commercially available software.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
To identify neuronal mechanisms that underlie synaptic function, we conducted large scale behavioral screens in Drosophila for EMS-induced temperature-sensitive (TS) paralytic mutations that result in motor abnormalities at 38 °C. Mutagenized homozygous viable lines were screened for behavioral defects by observing each strain individually in preheated vials in a 38 °C water bath for 5 min. Any strain showing behavioral abnormalities such as paralysis or abnormal seizure-like movements (Fig. 1, A and B) was maintained as a laboratory stock and subjected to two additional screening procedures to detect electrophysiological and morphological defects. First, the mutants were analyzed for temperature-induced seizure activity by screening for recurrent spontaneous neural activity in the adult giant fiber system at the nonpermissive temperature (Fig. 1, C–F). The giant fiber flight circuit is important in escape responses and flight initiation and includes both chemical and electrical synapses. The pathway can be activated by stimulation of the brain, and extracellular recordings can be made from the dorsal longitudinal flight muscles (DLMs). Wild-type Drosophila (Canton S) display little to no spontaneous activity when the temperature is raised to 38 °C (Fig. 1C). In contrast, many of the TS mutants resulted in robust spontaneous spiking in the DLM flight muscles when exposed to the nonpermissive temperature (Fig. 1, D–F). The activation of the DLM flight muscles reflects the firing rate of the innervating motor neurons. To date, 37 seizure-inducing mutations that define 12 complementation groups on the X chromosome have been identified in large scale screens of homozygous viable lines generated in the lab.



View larger version (39K):
[in this window]
[in a new window]
  		FIG. 1.
Temperature-sensitive mutants in Dmca1A exhibit paralysis and conditional seizures. A, the behavioral onset of motor uncoordination is plotted for the newly isolated Dmca1ANT27 and Dmca1ATS3 mutants, as well as the previously isolated Dmca1ATS2 allele (24). Similar to mutations in the sodium channel (para) which cause rapid paralysis (60), Dmca1A mutants exhibit a rapid onset of behavioral dysfunction, with robust bouts of seizure-like activity. Twenty flies of the indicated genotype (Canton S (CS), wild type; para, paraTS1; comt, comt1ST7) were aged 3–5 days and placed in preheated glass vials at 38 °C. The percentage of flies standing was measured in 15-s intervals and plotted. Dmca1ATS4 and Dmca1ATS5 showed more variable behavioral phenotypes and were not analyzed further. Unlike para and Dmca1A mutants, comt mutants, which disrupt NSF function (61), take much longer to paralyze when placed at 38 °C. B, the recovery time following exposure to 38 °C is plotted versus the length of exposure to the nonpermissive temperature. Dmca1ATS2 mutants show the fastest recovery, whereas Dmca1ANT27 mutants are the most severely affected, requiring progressively longer recovery times. C–E, extracellular recordings from the DLM flight pathway from controls (CS), Dmca1ATS3, and Dmca1ANT27, taken at 20 °C (top trace), at the nonpermissive 38 °C (middle trace) and following recovery at 20 °C (bottom trace). Control animals (C) display no seizure activity when the temperature is raised to 38 °C. In contrast, a rapid onset of spontaneous firing is observed inDmca1ATS3 (D) and Dmca1ANT27 (E) at 38 °C. Complete recovery was observed upon return to the permissive temperature. F, an enlarged time scale trace for seizure activity in Dmca1ANT27 at 38 °C. The scale bar for C–E is 5 mV/0.5 s, and for F is 2 mV/20 ms.

 

To examine the link between neuronal activity and synaptic development, we performed a morphological analysis of the seizure mutant collection by visualizing axonal arborizations and terminal morphology with anti-HRP (a general neuronal marker) and anti-synaptotagmin (an antibody against an abundant synaptic vesicle protein) immunocytochemistry at muscle fibers 6 and 7 of third instar larvae. This larval synapse is more accessible to morphological studies than the DLM pathway described above. Abnormal overproliferation of presynaptic varicosities was observed in several of the seizure mutants,2 similar to the excessive synaptic overgrowth seen in Shaker, ether-a-go-go mutants secondary to increased nerve activity (18). Notably in our screen, a single complementation group defined by four EMS alleles resulted in a decrease in axonal terminal branching and synaptic varicosity number (see below), suggesting the mutated gene product defined by this complementation group is likely to function in the control of synaptic growth. The synaptic undergrowth in the mutant, as opposed to the increased synaptic overproliferation observed in several hyperexcitable mutants, prompted us to clone the mutated gene product to characterize further its role in synaptic structure and function.

To identify the gene product defined by this complementation group, we mapped the TS phenotype to 33 centimorgans on the X chromosome via recombination. Subsequent deficiency mapping placed the complementation group near the Dmca1A locus. Complementation analysis with null mutants of Dmca1A (cacophony/nightblind/L(1)L13) demonstrated that the four mutations we isolated (NT27, TS3, TS4, TS5) represented new Dmca1A alleles that disrupt the Drosophila homolog of the N-type calcium channel. Upon exposure to 38 °C, the Dmca1A TS alleles, unlike wild-type CS controls, showed a rapid onset of uncoordination, seizure-like activity, and loss of motor control (Fig. 1A). All of the alleles except one showed a rapid recovery when returned to room temperature, regardless of the length of exposure to 38 °C (Fig. 1B). Dmca1ANT27 was the most severe allele isolated, requiring progressively longer recovery times with increased length of exposure to 38 °C. The temperature-sensitive phenotype of each allele was rescued with a small chromosomal duplication (Dp(1;2)v65b) encompassing the wild-type Dmca1A locus (19), indicating the behavioral defects result from disruption of the N-type calcium channel. Behavioral analysis of the TS mutants as well as complementation testing with other Dmca1A alleles suggest an allelic series of decreasing severity, NT27 > TS3 > TS4 > TS5. The most severe allele, Dmca1ANT27, also shows behavioral defects at room temperature when placed in trans to a null mutant (Dmca1A13) (19). These flies are flightless, exhibit an uncoordinated gait, and have reduced viability and life span. Thus, Dmca1ANT27 mutants disrupt N-type calcium channel function constitutively, with the defect becoming worse at the elevated temperature. Further reduction in channel function at 38 °C triggers the onset of seizure-like activity in the giant fiber circuit (Fig. 1, E and F), accounting for the rapid onset of behavioral dysfunction.

To determine the cause of the behavioral and physiological defects observed in Dmca1ANT27, we sequenced the calcium channel ORF in the mutant. A single bp change (CGT to TGT) at position 2626 of the ORF was detected that converted a charged arginine residue at site 876 to a neutral cysteine residue (Fig. 2A). This amino acid substitution was confirmed by sequencing an independent isolate of the Dmca1ANT27 cDNA. Arginine 876 is a universally conserved residue in all voltage-gated ion channels (Fig. 2B) and corresponds to the third charged residue of the S4 voltage sensor in the third set of six transmembrane domains. We were unable to test directly the effect of the R876C change on the current-voltage relationship of the mutant channel, as Drosophila presynaptic terminals are inaccessible for direct voltage clamp analysis, and experimental attempts to express recombinant Drosophila calcium channels in heterologous systems have not succeeded (20). However, the identical charged arginine residue altered in Dmca1ANT27 has been mutated to a cysteine in recombinant Shaker potassium channels and mammalian L-type calcium channels and expressed in oocytes, mimicking the R876C defect found in Dmca1ANT27 mutants (21, 22). In both cases, the R876C charge neutralization reduces the sensitivity of the channel to voltage, shifting the conductance-voltage relationship and requiring more depolarization for channel activation. Indeed, the third arginine in the S4 voltage domain has been shown to contribute more than any other S4 charged residue in determining total gating charge for voltage-dependent channels (23).



View larger version (44K):
[in this window]
[in a new window]
  		FIG. 2.
Mutations in the N-type calcium channel disrupt the S4 voltage sensor. A, domain diagram of the Drosophila N-type channel showing the localization of known mutations in Dmca1A, including the R876C alteration in the S4 voltage sensor found in Dmca1ANT27, the F1029I change found in cacS (19), and the P1385S change found in Dmca1ATS2 (62). B, the Arg-876 amino acid (circled) is conserved in all voltage-gated ion channels through evolution and forms the third charged residue in the S4 voltage sensor transmembrane helix. Recent crystallization of a bacterial voltage-gated channel suggests the S4 domain forms a paddle within the lipid bilayer that senses voltage by transversing the membrane from inside to outside when the channel opens (63). The R876C change would lead to the loss of a core charge on the voltage-sensor paddle that could alter gating movements required for channel opening. A mutation in the mammalian homolog of Dmca1A has also been mapped to the third transmembrane helix (boxed residue) in the ataxic mouse mutant rolling mouse Nagoya (64). Abbreviations: Na, sodium channel; Ca, calcium channel; H, human; D, Drosophila. The roman numerals in parenthesis refer to the specific set of six transmembrane domains (from 1 to 4) found in sodium and calcium channels. C, dendrogram of the calcium channel subtypes from humans (H), rodents (R), Caenorhabditis elegans (Ce), and Drosophila (Dm). Analysis of the Drosophila genome identifies a single family member of the L-type, N-type, and T-type voltage-gated calcium channels.

 

The Drosophila genome encodes three calcium channel {alpha}-subunit genes (15), including homologs of the mammalian T-type, L-type (Dmca1D), and N-type (Dmca1A) channels (Fig. 2C). Previous physiological analysis in Dmca1A mutants has confirmed that N-type calcium channels are the primary source of extracellular calcium entry into Drosophila presynaptic nerve terminals (24). To determine whether Dmca1ANT27 disrupts calcium channel function at the permissive temperature where we observe the morphological defects in synapse formation, we examined neurotransmitter release at 22 °C. Calcium influx through voltage-dependent calcium channels triggers presynaptic vesicle fusion, allowing us to test the in vivo effects of the R876C change in Dmca1A. Electrophysiological recordings at the neuromuscular junction of third instar Dmca1ANT27 larvae demonstrated a dramatic defect in evoked neurotransmitter release compared with control animals (Fig. 3, A–C). At low extracellular calcium levels, evoked release was undetectable in the mutants upon nerve stimulation, whereas control lines showed substantial synaptic transmission. At higher calcium levels, evoked release was partially restored, but with a greatly reduced quantal content compared with controls. Mini amplitude was unchanged in the mutant (CS, 0.87 ± 0.05 mV; Dmca1ANT27/Dmca1A13, 0.89 ± 0.04 mV), indicating post-synaptic sensitivity was not affected, and the observed defects in evoked release represent presynaptic dysfunction. Likewise, mini frequency was in the normal range in Dmca1ANT27, indicating spontaneous release was largely unaffected. The electrophysiological analysis of neurotransmitter release in Dmca1ANT27 mutants confirms that the R876C change disrupts calcium channel function, calcium entry, and subsequent vesicle fusion.



View larger version (19K):
[in this window]
[in a new window]
  		FIG. 3.
Electrophysiological defects at Dmca1ANT27 neuromuscular junctions. A and B, voltage traces of evoked excitatory junction potentials recorded from muscle fiber 6 of segment A3 in either CS (A) or Dmca1ANT27/Dmca1A13 third instar (B) at the indicated extracellular calcium concentrations (mM). C, plot of evoked excitatory junction potential amplitudes versus the indicated extracellular calcium concentrations. Dmca1ANT27/Dmca1A13 results in a shift to the right, consistent with a presynaptic defect in calcium influx and subsequent neurotransmitter release induced by the R876C amino acid change in Dmca1ANT27. The number of NMJs examined at each calcium concentration (mM) are as follows: CS 0.1 (8), 0.2 (15), 0.4 (20), 0.6 (7); Dmca1ANT27/Dmca1A130.1 (6), 0.2 (15), 0.4 (32), 0.6 (13). D, miniature excitatory junctional potential (mini) frequency recorded at 0.2 mM extracellular calcium in CS and Dmca1ANT27/Dmca1A13 did not show a statistical change. Error bars in C and D are S.E.

 

Our initial observations of abnormal synaptic development in the Dmca1ANT27 mutant suggested that presynaptic calcium entry might not only control neurotransmitter release but also modulate synaptic growth. To explore this possibility, we performed a quantitative analysis of synaptic growth in the mutants by visualizing anti-synaptotagmin staining as a presynaptic marker (Fig. 4, A and B). In Drosophila larvae, each abdominal hemisegment (A2–A7) has a stereotyped musculature with a precise motor neuron innervation pattern. In wild-type third instar larvae, an average of 65 ± 2.5 (S.E.) varicosities are present on muscle fibers 6 and 7 (average of segments A3 to A5). In contrast, both Dmca1ANT27 homozygotes and Dmca1ANT27/Dmca1A13 larvae showed a significant reduction in varicosity number. Dmca1ANT27/Dmca1A13 third instar larvae had on average 42 ± 2.4 (S.E.) varicosities, a 35% reduction in synaptic growth (p < 1 x 10–8, Student's t test). This defect was also manifested at other NMJ synapses, including those at muscle fiber 4, where we observed a 34% reduction in varicosity number in Dmca1ANT27/Dmca1A13 third instar larvae compared with controls. In addition, Dmca1ANT27 mutants exhibited reduced axonal terminal branching and a smaller synaptic innervation domain (Fig. 4B). Because NMJ growth varies in proportion with muscle size (25), we assayed in detail NMJ 6/7s at segment A2 in control and Dmca1ANT27 mutants and normalized measurements to muscle surface area. Normalized varicosity number was also significantly (p < 0.05) reduced in mutant animals (Fig. 4F).



View larger version (30K):
[in this window]
[in a new window]
  		FIG. 4.
Synaptic growth defects in Dmca1ANT27. A–D, immunocytochemical analysis with anti-synaptotagmin antibodies at muscle fibers 6/7 in segment A3 of third instar larvae: A, CS; B, Dmca1ANT27; C, n-syb14/n-syb{Delta}F33B; or D, syxL2/syxL247. Z-series stacks from confocal slices through the entire NMJ 6/7 were generated and used for quantification. Dmca1ANT27 mutants have a reduction in varicosity number, branch number, and branch length compared with controls. In contrast, mutations that disrupt vesicle fusion without affecting presynaptic calcium entry have no defect in synaptic proliferation. On occasion, etopic innervation from the transverse nerve was also observed on muscle fibers 6/7 in Dmca1ANT27, n-syb14/n-syb{Delta}F33B, and syxL2/syxL247, similar to that previously reported in other mutants that reduce synaptic activity (31). E, quantification of varicosity number at muscle fibers 6/7 of segments A3 to A5 of the indicated genotypes. Eight to 20 larvae were examined for each genotype. The total number of 6/7 NMJs examined are as follows: CS (98), Dmca1ANT27/Dmca1A13 (53), Dmca1ANT27 (36), n-syb14/n-syb{Delta}F33B (62), syxL2/syxL247(53). F, quantification of varicosity number of the indicated genotypes at muscle fiber 6/7 of segment A2 normalized to muscle surface area. Ten to 21 larvae were examined for each genotype. The total number of A2 6/7 NMJs examined are as follows: CS (20), Dmca1ANT27/Dmca1A13 (42), Dmca1ANT27 (18). Muscle surface area was quantified using Zeiss Pascal Axiovision Image Analysis software. G, quantification of varicosity number normalized to muscle surface area at muscle fibers 6/7 of segments A3 to A5 for animals injected with either buffer or Dmca1A dsRNA. Ten third instar larvae were examined for each genotype. The total number of 6/7 NMJs examined are as follows: buffer injected (59), Dmca1A dsRNA injected (59). For E–G, error bars are S.E.; statistical significance (* = p < 0.05) was determined by Student's t test.

 

To confirm further that calcium channel dysfunction results in altered synaptic growth, we used RNA interference to generate loss-of-function animals for morphological characterization. We injected double-stranded RNA (dsRNA) prepared from Dmca1A or buffer alone into stage 5 CS embryos. 85% of embryos injected with Dmca1A dsRNA resulted in late stage embryonic lethality (compared with only 28% lethality for bufferonly injections), mimicking the embryonic lethal phase of Dmca1A null mutants. The remaining 15% of animals were able to hatch and proceed through development, suggesting RNA interference generated partial loss-of-function mutants in a small number of cases. We used these animals to quantify the number of varicosities at muscle 6/7 (segments A3 to A5) and normalized measurements to muscle surface area in both buffer and Dmca1A dsRNA-injected third instar larvae. Similar to observations in Dmca1ANT27 mutants, normalized varicosity number in Dmca1A dsRNA-injected animals was reduced by 32% (p < 0.001) compared with buffer only-injected animals (Fig. 4G).

These observations suggest that either presynaptic calcium entry or synaptic transmission is an important modulator of synaptic growth during development. To differentiate between these two possibilities, we examined synaptic development in viable hypomorphic mutations of n-synaptobrevin and syntaxin-1A that disrupt SNARE-mediated vesicle fusion and result in more severe defects in neurotransmitter release than observed in Dmca1ANT27 mutants (26). If defects in neurotransmitter release were the cause of the abnormal synaptic proliferation, we would expect a similar, if not more severe, reduction in synaptic growth in the two SNARE mutants. However, this was not observed. Mutations in syntaxin-1A or nsynaptobrevin did not affect synapse proliferation at the third instar NMJ (Fig. 4, C and D) and showed varicosity numbers identical to controls (Fig. 4E). These observations are in agreement with previous analysis of synaptic growth in synaptotagmin (27) and SNARE mutants (26). Thus, our data suggest that calcium entry through presynaptic N-type calcium channels, not neurotransmitter release per se, is important for normal synaptic growth.

To address whether N-type calcium channels are required for the initial formation of the synapse during embryogenesis, or instead function in the second phase of developmental synaptic proliferation at the NMJ, we analyzed synapse formation in Dmca1A null mutant embryos. Previous genetic analysis of Dmca1A revealed several lethal mutations that map to the locus (19). Out of five embryonic lethal alleles of Dmca1A, we examined Dmca1A6, Dmca1A10, Dmca1A13, Dmca1A20, and Dmca1A24; Dmca1A13 behaves similar to a deficiency in complementation testing with our TS mutants and has been reported previously (19) to act as a null mutant in complementation testing with other cacophany and nightblind alleles of the Dmca1A locus. Therefore, we performed immunocytochemistry on Dmca1A13 embryos in trans to a small deficiency (Df(1)RC29) that removes the Dmca1A locus. Morphological analysis with anti-HRP and anti-synaptotagmin revealed that Dmca1A null mutants and Dmca1ANT27 hypomorphic mutants do not affect neuronal pathfinding or synaptic morphology at muscle fiber 6/7 in fully matured embryos (Fig. 5, A–C). These data rule out a role for presynaptic calcium entry through N-type calcium channels in the initial stages of growth cone guidance and synapse formation. Given the evidence for a role for calcium in axonal pathfinding and neurite extension in neuronal culture systems, we considered the possibility that L-type channels may function in this regard. Mutations in the L-type channel have been generated previously and cause embryonic lethality secondary to a loss of muscle contractions (28). Therefore, we generated double mutants disrupting both N-type and L-type channels. Immunocytochemical analysis of double mutants removing both channels revealed normal axonal pathfinding and initial synapse formation (Fig. 5D). One possibility is that intracellular calcium release channels or presynaptic ligand-gated channels provide calcium signals subserving neurite outgrowth. However, our data indicate that presynaptic calcium entry through voltage-gated calcium channels is not required during embryonic development for neurite outgrowth, axonal pathfinding, or initial synapse formation in vivo. Following synapse formation, presynaptic calcium entry through N-type channels regulates a second phase of activity-dependent synaptic growth during synapse maturation.



View larger version (19K):
[in this window]
[in a new window]
  		FIG. 5.
Defects in initial synapse formation do not account for reduced proliferation in Dmca1ANT27. A–D, immunohistochemical analysis of embryonic muscle fiber 6/7 NMJs stained with anti-HRP and anti-synaptotagmin antibodies at hatching stage: A, CS (n = 25); B, Dmca1ANT27 (n = 23); C, Dmca1A13 (n = 13); and D, Dmca1A13; Dmca1DX10 (n = 7) double mutants. No defect in varicosity number or innervation pattern was observed, indicating presynaptic calcium channels are not required for axonal pathfinding, target recognition, or the initial stages of synapse formation. Scale bar is 5 µm. E, quantification of varicosity number at NMJ 6/7 of embryonic segments A2 to A7 of CS, Dmca1ANT27, Dmca1A13/Df(1)RC29 and Dmca1A13; Dmca1D. Error bars are S.E.

 

Our results suggest that calcium entry through N-type calcium channels contributes to the formation of new varicosities. However, recent data (29) indicate that newly formed varicosities at Drosophila NMJs must be stabilized to prevent synaptic retraction. Thus, calcium entry might be required to stabilize newly formed synapses. We tested these two alternative hypotheses using a recently developed assay for synaptic retraction. We co-stained third instar larvae with antisera against the presynaptic vesicle protein synaptotagmin and the post-synaptic marker DLG. Presynaptic retraction precedes disassembly of the postsynaptic apparatus, leaving behind a long lived postsynaptic footprint (29). This is manifested immunocytochemically as a mismatch between synaptotagmin and DLG staining, with muscle DLG synaptic footprints in the absence of presynaptic synaptotagmin immunoreactivity. This mismatching occurs very rarely in wild-type animals but is common in manipulations that disrupt dynactin function and reduce synapse stability (Fig. 6C) (29). We assayed mismatching in third instar larvae at 78 NMJs on muscle fibers 6/7 in 10 control animals and 70 NMJs on muscle fibers 6/7 in 8 Dmca1ANT27/Dmca1A13 animals. In contrast to mutations that disrupt dynactin and cause retraction (Fig. 6C), no case of a DLG footprint without corresponding synaptotagmin staining was found in either controls or Dmca1ANT27/Dmca1A13 animals (Fig. 6, A and B). These data argue against an increase in synapse retraction in Dmca1ANT27. In summary, we have shown that N-type calcium channels are important in presynaptic proliferation through pathways independent of their role in neurotransmitter release. These findings suggest a link between presynaptic voltage-gated calcium entry through N-type calcium channels and activity-dependent synaptic growth mechanisms at the Drosophila NMJ.



View larger version (21K):
[in this window]
[in a new window]
  		FIG. 6.
Synapse retraction does not account for presynaptic undergrowth of nerve terminals in Dmca1ANT27. Co-staining with anti-synaptotagmin and anti-DLG at muscle fibers 6/7 in segment A3 of third instar larvae: A, CS; B, Dmca1ANT27; and C, Gl1/+. In CS and Dmca1ANT27, DLG post-synaptic staining was co-localized with presynaptic synaptotagmin immunoreactivity. No DLG footprints suggestive of synaptic retraction were observed, indicating the reduced synapse number in Dmca1ANT27 is not due to an increase in synapse retraction. In contrast, Gl1/+ mutants that disrupt dynactin function display DLG footprints without corresponding synaptotagmin staining (arrows), consistent with synaptic retraction as reported previously (29).

 


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The regulation of synaptic connectivity is a dynamic process, with synapse formation, synapse retraction, and synaptic remodeling occurring throughout life. This dynamic plasticity underlies the activity-dependent refinement of developing neuronal circuits and is thought to be essential for plasticity in the adult brain as well. Due to its accessibility and stereotypic development, the Drosophila glutamitergic neuromuscular junction has become a widely studied model system for investigating the molecular mechanisms that underlie synaptic growth (30). The NMJ forms during late embryogenesis and requires outgrowth of the motor neuron growth cone from the central nervous system, recognition, and contact with its target muscle fiber, and formation of enlargements of the growth cone or "prevaricosities" upon contact with the target muscle. Synaptic varicosities develop as a result of constrictions of larger prevaricosities (16). Following the formation of the initial synaptic field, a second phase of synaptic growth occurs that results in a 10-fold increase in varicosity number throughout larval development. This growth process has been shown to be modulated by activity-dependent mechanisms (12, 18, 31). Mutations that lead to increased cAMP levels (dunce) or increased nerve activity (Shaker and ether-a-go-go) result in a developmental increase in varicosity number and terminal branches at the NMJ. Many of these activity-dependent growth mechanisms impinge on the regulation of cell adhesion molecules like FASII and integrins (25, 32). For example, activated CaMKII has been demonstrated to regulate FAS2 localization and modulate synaptic growth (33). Recently, members of the transforming growth factor-{beta} superfamily of ligands and receptors have also been found at the NMJ and shown to modulate synapse number (34, 35). Downstream of these signaling pathways, modulation of the actin and tubulin cytoskeleton is important in synapse maturation (36, 37). Finally, both local protein synthesis and ubiquitin-dependent degradation have been implicated in synaptic growth in Drosophila (38–40). Although a relatively large number of growth pathways have been found, including several that are calcium-dependent, it is largely unknown how they function together and what signals regulate their activity. Cell adhesion proteins and synaptic growth regulators reside in regions adjacent to the active zone where presynaptic calcium channels localize (40, 41). Here we demonstrate that presynaptic calcium channels regulate synaptic growth. These findings suggest calcium diffusion from N-type channels may regulate the properties of calcium-activated synaptic growth regulators present in domains surrounding presynaptic active zones.

Dmca1A is abundantly expressed in the Drosophila nervous system (19) and encodes the presynaptic N-type calcium channel responsible for calcium influx that triggers synaptic vesicle fusion (24). Null mutations in Dmca1A are embryonic lethal (lethal(1)L13), whereas partial loss-of-function mutations disrupt synaptic transmission, leading to defects in various behaviors, including courtship (cacaphony) and phototaxis (nightblind-A) (19, 24, 42). In addition to defects in neurotransmitter release, we now demonstrate that Dmca1A hypomorphic mutants show altered morphology at the mature larval NMJ. There is a decrease in both terminal branching and varicosity number compared with wild-type controls and hypomorphic alleles of syntaxin and n-synaptobrevin. The reduced synaptic proliferation is not secondary to defective synaptic transmission because syntaxin and synaptobrevin show more profound defects in transmitter release but have normal synaptic proliferation. Similar results have also been reported in synaptotagmin mutants as well as SNARE mutants (26, 27). Thus, mutations in Dmca1A affect calcium-regulated synaptic pathways separate from those that regulate transmitter release.

We found no evidence for a role of presynaptic calcium entry through either N- or L-type calcium channels in the early stages of synapse formation during late embryogenesis. We also determined that the morphological defects in Dmca1A mutants are not due to an increase in terminal retraction, suggesting active growth rather than synapse stability is defective. Therefore, the structural defects we observe in Dmca1A mutants occur between the establishment of the initial synaptic field and its final larval maturation. Previous work (17) has demonstrated that the overall shape and branching pattern at the Drosophila NMJ is established early in development. Subsequent growth largely requires the addition of new varicosities to previously formed terminal branches. It is within this second activity-dependent growth phase that we propose presynaptic calcium entry is required to promote synaptic maturation. The 35% reduction we observe in varicosity number in Dmca1ANT27 mutants at the end of larval development is likely an underestimate of the actual contribution of presynaptic calcium entry to synaptic growth regulation. First, the initial activity-independent elaboration of synapses during late embryogenesis does not require calcium channel function, allowing the establishment of the initial synaptic field. Second, the Dmca1ANT27 mutant is a hypomorphic allele, reducing calcium channel function but not eliminating it. Previously isolated alleles of the Dmca1A locus that are more severe than Dmca1ANT27 are embryonic lethal, preventing an analysis of the activity-dependent phase of synaptic growth in more severe alleles. Further studies with mosaic animals will be required to fully characterize the persistence of synaptic growth mechanisms in the complete absence of presynaptic calcium influx.

The opening of presynaptic N-type channels during robust synaptic activity may allow calcium to influence varicosity sprouting mechanisms to locally control synaptic remodeling. Changes in intracellular calcium have been shown to affect growth cone motility (43, 44) and neurite outgrowth (45, 46). Indeed, filopodial protrusions from neuronal growth cones are triggered by altered calcium concentrations (46). Synaptic activity results in calcium-dependent CaMKII activation via binding of calcium/calmodulin and subsequent auto-phosphorylation (47). Activated CaMKII phosphorylates the synaptic MAGUK protein DLG, causing release of FAS2 from its synaptic scaffold and subsequent modulation of synaptic growth in Drosophila (32, 33). In addition, CaMKII activation also regulates the activity of the ether-a-go-go (eag) family of potassium channels in Drosophila, altering aspects of nerve excitability that could contribute to synaptic growth (48, 49). Intracellular calcium levels directly regulate cAMP signaling through the activation of adenylate cyclase by calmodulin (50), enhancing cAMP-dependent pathways implicated in synaptic growth (12). It is likely that disruptions in presynaptic calcium entry in Dmca1ANT27 mutants leads to alterations in several presynaptic signaling cascades that modulate growth. Further genetic analysis should begin to elucidate how the regulation of calcium entry modulates these activity-dependent synaptic growth pathways.

Voltage-gated calcium channels consist of four repeated units (I–IV) containing six {alpha}-helical transmembrane segments (S1–S6). The fourth transmembrane segment (S4) of voltage-gated ion channels has been shown to function as a voltage sensor. It is thought that the S4 sensor, a transmembrane {alpha}-helix in which every third or fourth residue is basic and carries a positive charge, undergoes conformational changes during depolarization that result in channel opening. Sequence analysis of Dmca1ANT27 identified a charge-neutralizing mutation in a highly conserved arginine reside in the S4 voltage sensor, supporting an essential role for S4 helix movement during channel gating and subsequent calcium influx. The altered S4 charged amino acid likely explains the TS phenotype of the mutant, as channel gating requires conformational changes in the S4 helix that are temperature-dependent. Mutations in mammalian Dmca1A homologs have been linked to a variety of neuronal disorders, including episodic and spinocerebellar ataxias, hemiplegic migraine, blindness, hypokalemic periodic paralysis, and epilepsy (51). Although it seems paradoxical that a hyperexcitability phenotype like seizures could arise from a reduction in calcium channel function, the Drosophila giant fiber pathway is extremely sensitive to changes in both inhibition and excitation. The decrease in calcium channel function in the inhibitory pathways may bear more weight in the overall output of the circuit, even though the excitatory outputs have reduced release as well. Similar seizure defects are seen in mammalian N-type calcium channel mutants (tottering, lethargic, and rocker) where overall calcium channel function is also reduced (52). Similar to what we have observed in Drosophila N-type mutants, mutations in the mouse {alpha}1a calcium channel locus (rocker) that disrupt the pore region of the channel and reduce calcium influx cause a profound reduction in Purkinje cell dendritic arborization (53). In addition, pharmacological disruption of calcium channel function in salamander rod photoreceptors has been shown recently (54) to inhibit varicosity formation. The reduced varicosity number in calcium channel mutants suggests an important role for normal levels of calcium entry during synaptic activity for the proper modulation of synaptic growth, in addition to its well established role in neurotransmitter release. Modulation of N-type calcium channel function via alterations in the rates of presynaptic action potential firing could provide an efficient mechanism for the regulation of activity-dependent synaptic growth.


   FOOTNOTES
 
* This work was supported by grants from the National Institutes of Health, the Human Frontiers Science Program Organization, and the Searle Scholars Program. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} Alfred P. Sloan Research Fellow. To whom correspondence should be addressed: Picower Center for Learning and Memory, Massachusetts Institute of Technology, E18-672, 50 Ames St., Cambridge, MA 02139. Tel.: 617-452-2605; Fax: 617-452-2249. E-mail: troy{at}mit.edu.

1 The abbreviations used are: NMJs, neuromuscular junctions; EMS, ethyl methanesulfonate; DLMs, dorsal longitudinal flight muscles; CS, Canton S; HRP, horseradish peroxidase; dsRNA, double-stranded RNA; SNARE, SNARE, soluble N-ethylmaleimide-sensitive factor attachment protein receptor; TS, temperature-sensitive; ORF, open reading frame; CaMKII, calcium/calmodulin-dependent protein kinase II. Back

2 Z. Guan and J. T. Littleton, unpublished observations. Back


   ACKNOWLEDGMENTS
 
We thank Bill Adolfsen, Enrico Montana, Chip Quinn, Sudipta Saraswati, Guosong Liu, and Elly Nedivi for helpful discussions regarding the manuscript; Jeff Hall, Richard Ordway, Graeme Davis, and Brian Stewart for fly strains; and Konrad Zinsmaier for anti-CSP antisera. The anti-DLG antibody developed by Corey Goodman was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD, National Institutes of Health, and maintained by the University of Iowa, Department of Biological Sciences, Iowa City, IA.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Reisman, G. (1969) Brain Res. 14, 25–48[CrossRef][Medline] [Order article via Infotrieve]
  2. Tsukahara, N., Hultborn, H., Murakami, F., and Fujito, Y. (1975) J. Neurophysiol. 38, 1359–1372[Abstract/Free Full Text]
  3. Luscher, C., Nicoll, R. A., Malenka, R. C., and Muller, D. (2000) Nat. Neurosci. 3, 545–550[CrossRef][Medline] [Order article via Infotrieve]
  4. Lichtman, J. W., and Colman, H. (2000) Neuron 25, 269–278[CrossRef][Medline] [Order article via Infotrieve]
  5. Grotewiel, M. S., Beck, C. D., Wu, K. H., Zhu, X. R., and Davis, R. L. (1998) Nature 391, 455–460[CrossRef][Medline] [Order article via Infotrieve]
  6. Bailey, C. H., and Chen, M. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 2373–2377[Abstract/Free Full Text]
  7. Buchs, P. A., and Muller, D. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 8040–8045[Abstract/Free Full Text]
  8. Engert, F., and Bonhoeffer, T. (1999) Nature 399, 66–70[CrossRef][Medline] [Order article via Infotrieve]
  9. Huang, Y. Y., Li, X. C., and Kandel, E. R. (1994) Cell 79, 69–79[CrossRef][Medline] [Order article via Infotrieve]
  10. Schacher, S., Castellucci, V. F., and Kandel, E. R. (1988) Science 240, 1667–1669[Abstract/Free Full Text]
  11. Zhong, Y., and Wu, C. F. (1991) Science 251, 198–201[Abstract/Free Full Text]
  12. Zhong, Y., Budnik, V., and Wu, C. F. (1992) J. Neurosci. 12, 644–651[Abstract]
  13. Catterall, W. A. (2000) Annu. Rev. Cell Dev. Biol. 16, 521–555[CrossRef][Medline] [Order article via Infotrieve]
  14. Bargmann, C. I. (1998) Science 282, 2028–2033[Abstract/Free Full Text]
  15. Littleton, J. T., and Ganetzky, B. (2000) Neuron 26, 35–43[CrossRef][Medline] [Order article via Infotrieve]
  16. Yoshihara, M., Rheuben, M. B., and Kidokoro, Y. (1997) J. Neurosci. 17, 8408–8426[Abstract/Free Full Text]
  17. Zito, K., Parnas, D., Fetter, R. D., Isacoff, E. Y., and Goodman, C. S. (1999) Neuron 22, 719–729[CrossRef][Medline] [Order article via Infotrieve]
  18. Budnik, V., Zhong, Y., and Wu, C. F. (1990) J. Neurosci. 10, 3754–3768[Abstract]
  19. Smith, L. A., Wang, X., Peixoto, A. A., Neumann, E. K., Hall, L. M., and Hall, J. C. (1996) J. Neurosci. 16, 7868–7879[Abstract/Free Full Text]
  20. Ren, D., Xu, H., Eberl, D. F., Chopra, M., and Hall, L. M. (1998) J. Neurosci. 18, 2335–2341[Abstract/Free Full Text]
  21. Garcia, J., Nakai, J., Imoto, K., and Beam, K. G. (1997) Biophys. J. 72, 2515–2523[Medline] [Order article via Infotrieve]
  22. Shao, X. M., and Papazian, M. (1993) Neuron 11, 343–352[CrossRef][Medline] [Order article via Infotrieve]
  23. Tytgat, J., Nakazawa, K., Gross, A., and Hess, P. (1993) J. Biol. Chem. 268, 23777–23779[Abstract/Free Full Text]
  24. Kawasaki, F., Felling, R., and Ordway, R. W. (2000) J. Neurosci. 20, 4885–4889[Abstract/Free Full Text]
  25. Schuster, C. M., Davis, G. W., Fetter, R. D., and Goodman, C. S. (1996) Neuron 17, 655–667[CrossRef][Medline] [Order article via Infotrieve]
  26. Stewart, B. A., Mohtashami, M., Trimble, W. S., and Boulianne, G. L. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 13955–13960[Abstract/Free Full Text]
  27. DiAntonio, A., and Schwarz, T. L. (1994) Neuron 12, 909–920[CrossRef][Medline] [Order article via Infotrieve]
  28. Eberl, D. F., Ren, D., Feng, G., Lorenz, L. J., Van Vactor, D., and Hall, L. M. (1998) Genetics 148, 1159–1169[Abstract/Free Full Text]
  29. Eaton, B. A., Fetter, R. D., and Davis, G. W. (2002) Neuron 34, 729–741[CrossRef][Medline] [Order article via Infotrieve]
  30. Keshishian, H., Broadie, K., Chiba, A., and Bate, M. (1996) Annu. Rev. Neurosci. 19, 545–575[CrossRef][Medline] [Order article via Infotrieve]
  31. Jarecki, J., and Keshishian, H. (1995) J. Neurosci. 15, 8177–8190[Abstract]
  32. Beumer, K., Matthies, H. J., Bradshaw, A., and Broadie, K. (2002) Development 129, 3381–3391[Abstract/Free Full Text]
  33. Koh, Y. H., Popova, E., Thomas, U., Griffith, L. C., and Budnik, V. (1999) Cell 98, 353–363[CrossRef][Medline] [Order article via Infotrieve]
  34. Aberle, H., Haghighi, A. P., Fetter, R. D., McCabe, B. D., Magalhaes, T. R., and Goodman, C. S. (2002) Neuron 33, 545–558[CrossRef][Medline] [Order article via Infotrieve]
  35. Marques, G., Bao, H., Haerry, T. E., Shimell, M. J., Duchek, P., Zhang, B., and O'Connor, M. B. (2002) Neuron 33, 529–543[CrossRef][Medline] [Order article via Infotrieve]
  36. Roos, J., Hummel, T., Ng, N., Klambt, C., and Davis, G. W. (2000) Neuron 26, 371–382[CrossRef][Medline] [Order article via Infotrieve]
  37. Pennetta, G., Hiesinger, P., Fabian-Fine, R., Meinertzhagen, I., and Bellen, H. B. (2002) Neuron 35, 291–306[CrossRef][Medline] [Order article via Infotrieve]
  38. Zhang, N., and Townes-Anderson, E. (2002) J. Neurosci. 22, 7065–7079[Abstract/Free Full Text]
  39. DiAntonio, A., Haghighi, A. P., Portman, S. L., Lee, J. D., Amaranto, A. M., and Goodman, C. S. (2001) Nature 412, 449–452[CrossRef][Medline] [Order article via Infotrieve]
  40. Wan, H. I., DiAntonio, A., Fetter, R. D., Bergstrom, K., Strauss, R., and Goodman, C. S. (2000) Neuron 26, 313–329[CrossRef][Medline] [Order article via Infotrieve]
  41. Uchida, N., Honjo, Y., Johnson, K. R., Wheelock, M. J., and Takeichi, M. (1996) J. Cell Biol. 135, 767–779[Abstract/Free Full Text]
  42. Smith, L. A., Peixoto, A. A., Kramer, E. M., Villella, A., and Hall, J. C. (1998) Genetics 149, 1407–1426[Abstract/Free Full Text]
  43. Kater, S. B., Mattson, M. P., Cohan, C., and Connor, J. (1988) Trends Neurosci. 11, 315–321[CrossRef][Medline] [Order article via Infotrieve]
  44. Gomez, T. M., and Spitzer, N. C. (1999) Nature 397, 350–355[CrossRef][Medline] [Order article via Infotrieve]
  45. Hong, K., Nishiyama, M., Henley, J., Tessier-Lavigne, M., and Poo, M. (2000) Nature 403, 93–98[CrossRef][Medline] [Order article via Infotrieve]
  46. Lankford, K. L., and Letourneau, P. C. (1989) J. Cell Biol. 109, 1229–1243[Abstract/Free Full Text]
  47. Miller, S. G., and Kennedy, M. B. (1986) Cell 44, 861–870[CrossRef][Medline] [Order article via Infotrieve]
  48. Wang, J., Renger, J. J., Griffith, L. C., Greenspan, R. J., and Wu, C. F. (1994) Neuron 13, 1373–1384[CrossRef][Medline] [Order article via Infotrieve]
  49. Wang, Z., Wilson, G. F., and Griffith, L. C. (2002) J. Biol. Chem. 277, 24022–24029[Abstract/Free Full Text]
  50. Schwabe, U., and Daly, J. W. (1977) J. Pharmacol. Exp. Ther. 202, 134–143[Free Full Text]
  51. Jen, J. (1999) Curr. Opin. Neurobiol. 9, 274–280[CrossRef][Medline] [Order article via Infotrieve]
  52. Steinlein, O. K., and Noebels, J. L. (2000) Curr. Opin. Genet. & Dev. 10, 286–291[CrossRef][Medline] [Order article via Infotrieve]
  53. Zwingman, T. A., Neumann, P. E., Noebels, J. L., and Herrup, K. (2001) J. Neurosci. 21, 1169–1178[Abstract/Free Full Text]
  54. Zhang, Y. Q., Bailey, A. M., Matthies, H. J., Renden, R. B., Smith, M. A., Speese, S. D., Rubin, G. M., and Broadie, K. (2001) Cell 107, 591–603[CrossRef][Medline] [Order article via Infotrieve]
  55. Lewis, E. B., and Bacher, F. (1968) Dros. Inf. Serv. 43, 193
  56. Stewart, B. A., Atwood, H. L., Renger, J. J., Wang, J., and Wu, C. F. (1994) J. Comp. Physiol. A Sens. Neural Behav. Physiol. 175, 179–191[CrossRef][Medline] [Order article via Infotrieve]
  57. Jan, L. Y., and Jan, Y. N. (1982) Proc. Natl. Acad. Sci. U. S. A. 79, 2700–2704[Abstract/Free Full Text]
  58. Littleton, J. T., Bellen, H. J., and Perin, M. (1993) Development 118, 1077–1088[Abstract]
  59. Zinsmaier, K. E., Hofbauer, A., Heimbeck, G., Pflugfelder, G. O., Buchner, S., and Buchner, E. (1990) J. Neurogenet. 7, 15–29[Medline] [Order article via Infotrieve]
  60. Loughney, K., Kreber, R., Ganetzky, B. (1989) Cell 58, 1143–1154[CrossRef][Medline] [Order article via Infotrieve]
  61. Littleton, J. T., Barnard, R. J., Titus, S. A., Slind, J., Chapman, E. R., and Ganetzky, B. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 12233–12238[Abstract/Free Full Text]
  62. Kawasaki, F., Collins, S. C., and Ordway, R. W. (2002) J. Neurosci. 22, 5856–5864[Abstract/Free Full Text]
  63. Jiang, Y., Ruta, V., Chen, J., Lee, A., and MacKinnon, R. (2003) Nature 423, 42–48[CrossRef][Medline] [Order article via Infotrieve]
  64. Mori, Y., Wakamori, M., Oda, S., Fletcher, C. F., Sekiguchi, N., Mori, E., Copeland, N. G., Jenkins, N. A., Matsushita, K., Matsuyama, Z., and Imoto, K. (2000) J. Neurosci. 20, 5654–5662[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 DMMHome page
S. Zhang, M. B. Feany, S. Saraswati, J. T. Littleton, and N. Perrimon
Inactivation of Drosophila Huntingtin affects long-term adult functioning and the pathogenesis of a Huntington's disease model
Dis. Model. Mech., May 1, 2009; 2(5-6): 247 - 266.
[Abstract] [Full Text] [PDF]

Home page
 J. Neurophysiol.Home page
H. Gu, S. A. Jiang, J. M. Campusano, J. Iniguez, H. Su, A. A. Hoang, M. Lavian, X. Sun, and D. K. O'Dowd
Cav2-Type Calcium Channels Encoded by cac Regulate AP-Independent Neurotransmitter Release at Cholinergic Synapses in Adult Drosophila Brain
J Neurophysiol, January 1, 2009; 101(1): 42 - 53.
[Abstract] [Full Text] [PDF]

Home page
 J. Neurophysiol.Home page
J. Hou, T. Tamura, and Y. Kidokoro
Delayed Synaptic Transmission in Drosophila cacophonynull Embryos
J Neurophysiol, November 1, 2008; 100(5): 2833 - 2842.
[Abstract] [Full Text] [PDF]

Home page
 J. Neurophysiol.Home page
J. W. Worrell and R. B. Levine
Characterization of Voltage-Dependent Ca2+ Currents in Identified Drosophila Motoneurons In Situ
J Neurophysiol, August 1, 2008; 100(2): 868 - 878.
[Abstract] [Full Text] [PDF]

Home page
 JCBHome page
C. V. Ly, C.-K. Yao, P. Verstreken, T. Ohyama, and H. J. Bellen
straightjacket is required for the synaptic stabilization of cacophony, a voltage-gated calcium channel {alpha}1 subunit
J. Cell Biol., April 3, 2008; 181(1): 157 - 170.
[Abstract] [Full Text] [PDF]

Home page
 J. Neurosci.Home page
D. K. Dickman, P. T. Kurshan, and T. L. Schwarz
Mutations in a Drosophila {alpha}2{delta} Voltage-Gated Calcium Channel Subunit Reveal a Crucial Synaptic Function
J. Neurosci., January 2, 2008; 28(1): 31 - 38.
[Abstract] [Full Text] [PDF]

Home page
 Proc. Natl. Acad. Sci. USAHome page
S. Saraswati, B. Adolfsen, and J. T. Littleton
Characterization of the role of the Synaptotagmin family as calcium sensors in facilitation and asynchronous neurotransmitter release
PNAS, August 28, 2007; 104(35): 14122 - 14127.
[Abstract] [Full Text] [PDF]

Home page
 J. Neurosci.Home page
I-F. Peng and C.-F. Wu
Drosophila cacophony Channels: A Major Mediator of Neuronal Ca2+ Currents and a Trigger for K+ Channel Homeostatic Regulation
J. Neurosci., January 31, 2007; 27(5): 1072 - 1081.
[Abstract] [Full Text] [PDF]

Home page
 J. Neurophysiol.Home page
S. A. Jiang, J. M. Campusano, H. Su, and D. K. O'Dowd
Drosophila Mushroom Body Kenyon Cells Generate Spontaneous Calcium Transients Mediated by PLTX-Sensitive Calcium Channels
J Neurophysiol, July 1, 2005; 94(1): 491 - 500.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
J. D. Spafford, J. van Minnen, P. Larsen, A. B. Smit, N. I. Syed, and G. W. Zamponi
Uncoupling of Calcium Channel {alpha}1 and {beta} Subunits in Developing Neurons
J. Biol. Chem., September 24, 2004; 279(39): 41157 - 41167.
[Abstract] [Full Text] [PDF]

Home page
 JCBHome page
B. Adolfsen, S. Saraswati, M. Yoshihara, and J. T. Littleton
Synaptotagmins are trafficked to distinct subcellular domains including the postsynaptic compartment
J. Cell Biol., July 19, 2004; 166(2): 249 - 260.
[Abstract] [Full Text] [PDF]

Home page
 J. Neurosci.Home page
P. Wang, S. Saraswati, Z. Guan, C. J. Watkins, R. J. Wurtman, and J. T. Littleton
A Drosophila Temperature-Sensitive Seizure Mutant in Phosphoglycerate Kinase Disrupts ATP Generation and Alters Synaptic Function
J. Neurosci., May 12, 2004; 24(19): 4518 - 4529.
[Abstract] [Full Text] [PDF]

Home page
 Proc. Natl. Acad. Sci. USAHome page
W.-C. M. Lee, M. Yoshihara, and J. T. Littleton
Cytoplasmic aggregates trap polyglutamine-containing proteins and block axonal transport in a Drosophila model of Huntington's disease
PNAS, March 2, 2004; 101(9): 3224 - 3229.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
278/42/41099    most recent
M306417200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rieckhof, G. E.
Right arrow Articles by Littleton, J. T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rieckhof, G. E.
Right arrow Articles by Littleton, J. T.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2003 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement