Many mutants modifying the nervous system will alter the communication between neurons and their target cells. To rigorously determine how a particular gene product affects the activity of an excitable cell in vivo, several methods directly monitoring neuronal physiological function are used in Drosophila and described in this section. Elevation of intracellular calcium is the trigger for the exocytosis of synaptic vesicles, and the released transmitter typically leads to a postsynaptic electrical response. Methods that can monitor the neuron at all of these stages of neuronal communication are discussed, i.e., measurements of (1) calcium levels, (2) vesicle dynamics, (3) protein dynamics, and (4) electrical signals.
A. Imaging and Manipulating Calcium Levels in Drosophila Neurons
Calcium is the primary trigger of neuronal activity and as such is the most important ion to be imaged dynamically. In Drosophila, [Ca2+] has been measured in cultured giant neurons (see later; Berke et al., 2002), larval NMJs (Karunanithi et al., 1997; Umbach et al., 1998a,b; Dawson-Scully et al., 2000; Kuromi and Kidokoro, 2002), adult photoreceptors (Ranganathan et al., 1994; Peretz et al., 1994; Hardie, 1996), and kenyon cells of the adult brain mushroom body (MB) (Rosay et al., 2001; Wang et al., 2001). These studies used various methods to generate calcium measurements, including a transgenically expressed calcium-sensitive protein (proaequorin; Rosay et al., 1997, 2001), and either injected (Hardie, 1996) or extracellularly loaded calcium-sensitive fluorescent dyes (Karunanithi et al., 1997; Umbach et al., 1998a,b; Dawson-Scully et al., 2000; Wang et al., 2001; Berke et al., 2002; Kuromi and Kidokoro, 2002). Controlled photo-activation of caged calcium has also been achieved in larval motorneurons (Kuromi and Kidokoro, 2002). In these studies, neuronal activity was stimulated by several means, including application of high [K+] to depolarize cells (Berke et al., 2002), application of a-latrotoxin (Umbach et al., 1998), direct electrical stimulation (Karunanithi et al., 1997; Umbach et al., 1998a; Dawson-Scully et al., 2000; Kuromi and Kidokoro, 2002), light (Peretz et al., 1994; Ranganathan et al., 1994; Hardie, 1996), odors (Wang et al., 2001), or endogenous neuronal activity (Rosay et al., 2001).
Each method has its own advantages and disadvantages. If intact tissues are to be used, none of these methods are useful except for visualizing cells close to the surface because light penetration and reliable, accurate fluorescence capturing is efficient only at the surface. However, an interesting structure, the mushroom body, is close to the surface of the Drosophila brain, and calcium measurements have demonstrated interesting properties of some of the surface cells in this region (Rosay et al., 2001; Wang et al., 2001). Extracellular AM-calcium dyes (ester forms of the dyes allowing membrane permeability) are used frequently in other organisms, but they take 1 h to load and then some time for the endogenous esterase to cleave the side group from the membrane permeable version. If one proceeds without this last maturation step, misleading results could be obtained as the dye is restored to its Ca-binding form only after cleavage. Therefore, this method requires maintaining a preparation healthy for prolonged periods prior to the initiation of the experiment, and some mutant neurons may be less hardy than wild-type neurons. In addition, in order to achieve good sensitivity, high concentrations of intracellular dyes (100 pM) are sometimes required and some damping of the calcium spikes cannot be avoided as these dyes are potent calcium buffers (Requena et al., 1991; Takahashi et al., 1999). A range of dyes with different kinetics and Kd values for calcium exist, and combinations of dyes have been used by vertebrate investigators to extend the dynamic range (Voets, 2000). A distinct advantage of some of these dyes is that they are ratiometric, thereby minimizing the need to obtain equivalent dye loading and eliminating errors due to photoinacti-vation of the dye or dye escaping the cells. Also in live samples that occasionally move, a ratiometric dye will help minimize errors due to movement of the sample. Many of the calcium dyes are measured by sampling at two wavelengths, and typically the measurements at the two wavelengths are inversely proportional to the calcium level. This amplifies the signal and eliminates concerns about dye loss.
Calcium imaging via proaequorin requires loading of the prosthetic group colentrazine (Brini et al., 1995; Rosay et al., 1997, 2001; for review, see Chiesa et al., 2001). Because this can take 1 to 6h and equivalent levels may not necessarily be achieved, loading is done as rapidly as possible to avoid degradation of the sample (Brini et al., 1995; Rosay et al., 1997, 2001). Aequorin gives off luminescence and is therefore relatively insensitive, as each molecule can undergo only one reaction (for review, see Chiesa et al., 2001). Typically, luminometers are used to measure the output, but photon-counting methods can also be used. BioRad confocals have excellent photon-counting abilities and could be used. However, an advantage of aequorin is that it is a poor calcium buffer, and intracellular calcium levels are not perturbed dramatically by this calcium probe (Chiesa et al., 2001). Aequorin has also been fused to different proteins, leading to the targeting of the chimera to different intracellular compartments (Marsault et al., 1997; see review by Chiesa et al., 2001). It has been demonstrated that fusion of aequorin and its native partner in the jellyfish, GFP, allows for fluorescence detection that is more sensitive than luminescence (Waud et al. 2001). In theory, if such a construct were introduced into flies, more sensitive Ca imaging could be assayed by measuring fluorescence. Another advantage of aequorin is its naturally wide dynamic range (0.5-30 pm Ca2+), and a genetically modified version can measure calcium up to the 300-pm range (Llinas et al., 1992). This feature of aequorin has been taken advantage of by Marsault et al., (1997). In this study, they were able to obtain calcium measurements in the range of 100-200 pM calcium close to the membrane using a SNAP-25-aequorin chimera (Marsault et al., 1997).
Our laboratory and others have also attempted FRET using cameleon, a protein fusion of two versions of GFP with different fluorescent properties, fused to calmodulin and to the myosin light chain kinase calmodulin-binding domain
(see Reiff and Schuster, 2000). As has been reported for this fusion in C. elegans (Kerr et al., 2000), this method is quite difficult because the ratio does not change to a great degree, resulting in a poor signal-to-noise ratio. However, better GFP-based calcium-sensing probes are being developed (Truong et al., 2001; Nagai, 2001), which, if introduced into flies, would avoid any incubation prior to initiation of the experiments.
The kinetics of vesicle dynamics may be monitored by members of a family of fluorescent lipophilic stryl dyes (Betz and Bewick, 1992). These dyes are useful tools for studying synaptic vesicle exocytosis and recycling because they reversibly stain membranes, they are membrane impermeant, and they fluoresce. They are a complementary technique to electrophysiological methods in that the distribution of vesicles can be observed directly and rates of endocytosis and exocytosis can be measured. This method also does not depend on measuring a postsynaptic response, which is the typical assay of electrophysiological methods and therefore potentially allowing one to dissect pre- and postsynaptic effects of a mutation. However, dye-loading experiments do not provide the resolution of electrophysi-ology. Several variants have been made with different fluorescent properties (FM1-43, fluorescein-like; FM4-64, rhodamine-like). Modifications of the original dye have yielded products with altered hydrophobic properties, and therefore with different washout kinetics (Betz et al., 1996; Ryan et al., 1996; Klingauf et al., 1998). For example, FM2-10 dissociates much faster from the membrane, allowing one to distinguish populations of vesicles with different recyling properties (Ryan et al., 1996; Klingauf et al., 1998; Schote and Seelig, 1998; Pyle et al., 2000). Vesicle recycling is monitored by labeling the plasma membrane via dye in the extracellular media, and subsequent stimulation of exocytosis. Stimulation of exocytosis causes fusion of the vesicular membrane with the plasma membrane, thereby allowing binding of the dye to the exocytosing membrane. The stained membrane is then internalized. Washout of the dye from the plasma membrane in the absence of synaptic activity is performed using calcium-free buffers. At this point one can either measure the amount of membrane internalized by measuring the total fluorescence or one can restimulate the cell, causing the dye to unload, thereby allowing the measurement of exocytosis. Another useful feature of the dye FM1-43 that has been underutilized by fly neurobiologists is that it can be photoconverted into an electron-dense precipitate, allowing EM detection of the marked vesicle (Henkel et al., 1996; Richards et al., 2000; Schikorski and Stevens, 2001; Harata et al., 2001).
The first use of FM1-43 by fly neurobiologists at the larval NMJ was to demonstrate that a mutation in the gene shibire blocks endocytosis, and that endocytosis requires extracellular calcium (Ramaswami et al., 1994). Since those experiments, FM1-43 has been used to characterize vesicle cycling in wild-type motorneurons (Kuromi et al., 1997; Kuromi and Kidokoro, 1999, 2000, 2002)
and in many synaptic mutants (Gonzales-Gaitan and Jackle, 1997; Ranjan et al., 1998; Kuromi and Kidokoro, 1998; Delgado et al., 2000; Fergestad and Broadie, 2001; Stimson et al., 2001; Verstreken etal., 2002; Guichet etal., 2002; Roche et al., 2002).
To label Drosophila with stryl dyes, embryos or third-instar larvae are dissected (as described in Section IV,D) in calcium-free saline and are loaded with FM1-43. Generally, the dye is loaded for 5 min using elevated extracellular potassium (50-90 mM) containing 0.8-10 ^M FM1-43 (Ramasawami et al., 1994; Gonzalez-Gaitan and Jackle, 1997; Fergstad and Broadie, 2001). A more physiological approach can be achieved via stimulation by a suction electrode leading to labeling of only a subset of neurons. Wash the fillets twice rapidly and then four to six times in Ca-free buffer for 5 min. One can now quantitate the amount of fluorescence retained by the bouton. To measure exocytosis, one can also unload the dye using either elevated potassium or electrical stimulation. By using different staining/destaining protocols, the size and dynamics of subpopulations of vesicles can be determined.
C. Monitoring Protein Dynamics in Living Drosophila Neurons
Various aspects of Drosophila cell biology can be monitored in mutants using available GFP markers of subcellular compartments (Brand, 1995, 1999; Hazel-rigg, 2000). New GFP variants or other fluorescent proteins spanning the visible range have been produced, allowing multicolor labeling and FRET between the appropriate GFP pairs (for reviews, see Tsien, 1998; van Roessel and Brand, 2002), but to date no published reports using Drosophila have taken advantage of these new tools. Many transgenics expressing GFP-tagged proteins are available, including a general membrane marker (mCD8-GFP), a postsynaptic marker (Shaker-GFP), cytoskeletal markers (actin, Verkhusha et al., 1999; moesin, Edwards et al., 1997; Tau, Brand, 1995; and tubulin, Grieder et al., 2000), a mitochondrial marker (cytochrome C-GFP, Pilling and Saxton, 1999), synaptic vesicle markers (synaptobrevin-GFP, Estes et al., 2000; Zhang et al., 2002 and synaptotagmin-GFP, Zhang et al., 2002), and a densecore vesicle marker (ANF-GFP, Rao et al., 2001). These GFP reporters label compartments that are highly dynamic and can be used to assess the effect of a mutation on an organelle and to determine the earliest point of deviation from the wild type. For example, the biogenesis, axonal transport, and localization of synaptic vesicles are regulated by many proteins (Rodesch and Broadie, 2000), and the effect of mutations in these proteins could be studied through the use of GFP markers of the cytoskeleton and vesicles. Bleaching and then observing recovery of the fluorescence (FRAP) has been shown to be feasible at the larval Drosophila NMJ (Zhang et al., 2002), and similar methods should allow one to also look at the effect of mutations on both the dynamics and distribution of vesicles and the underlying cytoskeleton. A word of caution about these markers is that some may not fully rescue null mutants and some markers may even lead to their own phenotypes (Williams et al., 2000).
The process of bouton development has been studied at the neuromuscular junction using a GFP fusion of CD8 with the postsynaptic targeting domain of Shaker (Shaker PDZ domain). The NMJ could be imaged live, and the process of bouton duplication was examined (Zito et al., 1999). New boutons appeared to be added at the end of existing boutons or are "inserted" between existing ones. This division could be symmetric or asymmetric similar to yeast budding. This observation led to a later study testing the idea that yeast budding and bouton division share some machinary in common (Eaton et al., 2002).
All of these GFP markers could also be used for genetic screens and soluble GFP (Kraut et al., 2001) and Shaker-GFP have been used for this purpose. In C. elegans, a GFP-synaptobrevin strain was used to find proteins with roles in synaptogenesis (Jin, 2002) by screening for mutants with an altered distribution of the GFP marker.
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