The authors would like to thank the Developmental Studies Hybridoma Bank at the University of Iowa for sending us antibodies. This work was supported by a grant (No. 29/08) from The Israel Science Foundation. A.S. is also supported by a research grant from the DFG (Deutsche Forschungsgemeinschaft).

Before you start
<ol>
	<li>Grow the desired fly cultures. Keep the vials uncrowded (approximately 30 flies per 50 ml vial). Let the flies lay eggs for only one day in each vial. This will provide the larvae sufficient food supply and allow them to reach maximal size before crawling out of the food. Keep the vials in the appropriate temperature until third instar larvae begin to wander on the vial wall.</li>
	<li>Prepare a fresh stock of phosphate buffered saline (PBS) and PBT (0.1% Tween in PBS). Keep 10 ml of PBS on ice.</li>
	<li>Prepare 1-5 ml of fresh fixative (4% Formaldehyde in PBT) and keep it on ice. We use bottled 37-38% formaldehyde as a stock solution.</li>
</ol>
1. Dissection and Fixation of Third Instar Larvae
<ol>
	<li>Pick a wandering third instar larva from the vial wall and place it in a 50 &mu;l drop of ice-cold PBS on a Sylgard dissection plate (made of Sylgard 184 silicone elastomer kit, DOW Corning Corporation in a tissue culture dish).</li>
	<li>Hold the larva, dorsal side up, near the mouth hooks, using fine forceps (Dumont #5 forceps), and stick an insect pin (minutien, 0.1 mm, stainless steel) through the larva's brain.</li>
	<li>Grab the posterior end of the larva with the forceps, pull gently and stretch the larva gently lengthwise. Stick another insect pin between the two posterior spiracles.</li>
	<li>Using spring scissors cut two horizontal incisions in the body wall (perpendicular to the anterior-posterior axis), close to the anterior and posterior pins.</li>
	<li>Using the spring scissors cut the larval body wall along the dorsal midline from the anterior to the posterior incision.</li>
	<li>Remove internal organs (gut, salivary glands, malpighian tubules etc.) and the trachea using fine forceps. Wash gently once or twice with PBS.</li>
	<li>Grab the two body wall flaps with forceps, stretch them out and pin them to the plate using two insect pins for each side.</li>
	<li>Remove the PBS and add 50 &mu;l of fixing solution (4% formaldehyde in PBT). Incubate 20 minutes at room temperature.</li>
	<li>Remove the fixative. Wash twice with PBS. Pull out the insect pins. Using spring scissors cut the larval head and tail leaving a rectangular fillet. Transfer the fixed tissue to a chilled Eppendorf tube with PBS.</li>
	<li>Once a sufficient number of larvae are fixed, one can continue directly to antibody staining. If immediate staining is not desired, the fixed larvae should be washed 3 times, 5 minutes each, with 95% ethanol and stored in 95% ethanol at -20 &deg;C.</li>
</ol>
2. Dissection and Fixation of Pupae
Part I
<ol>
	<li>Attend to the vials as for larvae fixation, until third instar larvae begin to pupariate. Examine the vials every hour and mark larvae that have pupariated. Allow the marked pupae to develop 30-40 hours at 24 &deg;C, or 24-27 hours at 29 &deg;C.</li>
	<li>Using fine forceps pick 20-30 pupae of the appropriate age and put them in a dark porcelain multi-well dissecting dish. Be careful not to damage the tissues of interest.</li>
	<li>Extract the pupae from the pupal case. Begin by peeling off the operculum and continue until the pupa is free. Put the pupa into a well filled with PBT. Continue extracting all the pupae. Steps 2.4-2.6 should be done in small batches of five pupae at a time.</li>
	<li>Place five pupae on a flat surface between the wells of the dissecting dish. Using a micro dissection knife cut the tip of the head and the posterior tip of the abdomen (alternatively, it is possible to use two pairs of forceps to tear holes at both ends of the pupae).</li>
	<li>Hold the pupa in place using fine forceps and use a 1 ml syringe to wash out the internal organs of the pupa, by injecting PBT through the anterior opening.</li>
	<li>Wash the dissected pupa briefly by dipping it in a well filled with PBS and move it into the cold fixative in an Eppendorf tube kept on ice.</li>
	<li>Incubate overnight (ON) at 4 &deg;C.</li>
</ol>
Part II
<ol start="8">
	<li>Discard the fixative and wash the pupae three times, 5 minutes each, with PBT. Keep the washed pupae on ice.</li>
	<li>Fill two wells of the dissecting dish with PBT. Using a polyethylene Pasteur pipette transfer several pupae to one of the wells.</li>
	<li>Move one pupa at a time into the second well. Using two pairs of sharp forceps with perfectly aligned tips, peel off the transparent pupal cuticle of the wing: secure the pupa to the bottom of the well using one pair of forceps, and gently tear the cuticle using the second pair of forceps. Once the cuticle is torn, peel it off the wing. Be careful not to disconnect the wing from the pupa. After peeling off the cuticle from the wing blade, continue peeling the cuticle of the wing hinge (where the wing ChOs are located).</li>
	<li>After peeling off the wing cuticle, one can attempt peeling off the leg cuticle in a similar manner. Many legs are likely to be lost in the process, but despite the low yield, this step is highly recommended since it greatly improves the staining of leg ChOs. The ChOs of the haltere and abdomen can be visualized without further peeling off the cuticle.</li>
	<li>Place the 'peeled' pupa in an Eppendorf tube filled with methanol and keep at room temperature. Continue peeling off the cuticle of all the pupae, and add them to the same tube. At least 10 nicely peeled pupae should be collected for each staining.</li>
	<li>Remove the methanol and wash three times, 5 minutes each, with 95% Ethanol. Fixed pupae can be kept for long periods of time in 95% Ethanol at -20 &deg;C.</li>
</ol>
3. Immunostaining of Larvae and Pupae
<ol>
	<li>Wash the fixed tissue with PBT three times, 30 minutes each, at room temperature. Larvae can be washed with gentle shaking on a rotating plate. Pupae are washed without shaking.</li>
	<li>Replace the PBT with blocking buffer (PBT + 5% normal goat serum) and incubate ON at 4 &deg;C.</li>
	<li>Remove the Blocking buffer and incubate with primary antibodies (diluted in blocking buffer) ON at 4 &deg;C.</li>
	<li>Wash four times with PBT as described in step 3.1.</li>
	<li>Remove the PBT and incubate with secondary antibodies (diluted in blocking buffer) ON at 4 &deg;C.</li>
	<li>Wash twice with PBT and once with PBS as described in step 3.1.</li>
	<li>Replace the PBS with mounting medium (Dako Fluorescent Mounting Medium (Dako Cytomation, Glostrup, Denmark) and incubate at 4 &deg;C ON.</li>
</ol>
4. Mounting of Larvae
<ol>
	<li>Larvae are mounted on a microscope slide in a drop of mounting media with their cuticle facing down and body wall muscles facing up. Put a small drop of mounting medium on a clean cover slip and use it to cover the preparation. Do not apply any pressure on the mounted larvae.</li>
</ol>
5. Mounting of Pupae
<ol>
	<li>Prepare two microscope slides with a drop of mounting medium, one for dissection and the second for mounting.</li>
	<li>Put several pupae on one of the slides.</li>
	<li>Using spring scissors remove the head and the posterior part of the abdomen.</li>
	<li>Separate the dorsal half of the thorax from the ventral half by cutting between the wings and the legs.</li>
	<li>Place the dissected body parts of the pupa on the second microscope slide in a drop of mounting solution. Ensure that the wings and legs are stretched out and try to minimize overlaying of tissues as much as possible.</li>
	<li>Place a drop of mounting medium on a cover slip and place it over the sample. Do not apply pressure on the mounted sample.</li>
</ol>
6. Representative Results
An example of immuno-stained ChOs of third instar larva is shown in Figure 1. This example shows a nicely stretched abdominal segment in which seven of the eight ChOs are clearly visible. Neurons are labeled with MAb22C10 (1:20, obtained from the Developmental Studies Hybridoma Bank at the University of Iowa); the cap, ligament, cap-attachment and ligament attachment cells are labeled with anti-&alpha;Tub85E (1:10, Klein et al., 2010). Secondary antibodies for fluorescent staining were Cy3, or Cy5-conjugated anti-mouse/rabbit (1:200, Jackson Immunoresearch Laboratories). Samples were viewed using confocal microscopy (LSM 510, Zeiss).
A cluster of wing ChOs at the ventral radial vein of a 35 hr-old pupa is shown in Figure 2.
<img alt="Figure 1" src="/files/ftp_upload/3846/3846fig1.jpg" /> 
Figure 1. (A) Schematic illustration of the six cell types that constitute a single Ch organ: cap attachment (CA), cap (C), scolopale (S), neuron (N), ligament (L) and ligament attachment (LA). The LCh5 organ, in which five ChOs are clustered, is stretched across the group of lateral transverse muscles (LT1-4). The lateral longitudinal muscle (LL1), ventral longitudinal muscles (VL1-4) and lateral oblique muscle (LO1) are also illustrated. Tendon cells are illustrated as brown spheres (Taken from Klein et al., 2010). (B) One abdominal segment of a third instar larva viewed at a 10X magnification. Seven of the eight ChOs present in each abdominal segment are evident: five lateral organs that together form the pentascolopidial organ (LCh5), a single lateral organ (LCh1) and one of two ventral ChOs (VChB). The neurons (N) of the ChOs are labeled with the neuronal marker MAb 22C10 (red). The ligament (L), cap (C) and attachment cells (LA, CA) are labeled with anti-&alpha;Tub-85E (blue). The cap and ligament cells are additionally labeled with a ChO-specific GFP reporter harboring a regulatory element from the dei locus (Nachman et al., unpublished data).
<img alt="Figure 2" src="/files/ftp_upload/3846/3846fig2.jpg" /> 
Figure 2. The wing chordotonal organs at the ventral radial vein are viewed at a 40X magnification. The neurons (N) are marked with the neuronal marker 22C10 (Red, B). The ligament (L) and cap (C) cells are co-labeled with anti-&alpha;Tub85E antibodies (Blue, C) and a ChO-specific GFP reporter transgene (Nachman et al., unpublished data).

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F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Feeling+the+vibes:+chordotonal+mechanisms+in+insect+hearing.">Feeling the vibes: chordotonal mechanisms in insect hearing.</a> Curr. Opin. Neurobiol. 9, 389-393 (1999).</li><li>Eberl, D. F., Boekhoff-Falk, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+Johnston's+organ+in+Drosophila.">Development of Johnston's organ in Drosophila.</a> Int. J. Dev. Biol. 51, 679-687 (2007).</li><li>Eberl, D. F., Hardy, R. W., Kernan, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetically+similar+transduction+mechanisms+for+touch+and+hearing+in+Drosophila.">Genetically similar transduction mechanisms for touch and hearing in Drosophila.</a> J. 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No conflicts of interest declared.

The protocol described in this video provides a way to visualize proprioceptive ChOs during larval and pupal stages. Studies about the structure and development of proprioceptive ChOs have been so far restricted mainly to embryonic stages and to larval imaginal discs. Thus, many aspects of later stages of development, of both larval and adult ChOs, remain largely unknown. The described protocol, combined with common genetic tools in Drosophila, can be utilized to identify and study new genes and pathways that play a role in later stages of ChOs development. This protocol was optimized for the visualization of ChO, but it can be adapted for the staining of other tissues, such as pupal wings 6.

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent/tool</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments </td>
		</tr>
		<tr>
			<td>Dumont #5 (or #55) forceps, biologie tip</td>
			<td>F.S.T</td>
			<td>11252-20 or 11252-10 
			(or similar forceps)</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Austerlitz stainless steel insect pins, minutiens 0.1mm</td>
			<td>Roboz Surgical Instrument Co</td>
			<td>RS-6083-10</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Sylgard 184 silicone elastomer kit</td>
			<td>DOW Corning Corporation</td>
			<td>240.4019862</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Vannas micro scissors (straight, 7.5 cm, blade 3mm)</td>
			<td>AS Mdeizintechnik GmbH</td>
			<td>11-590-00</td>
			<td>Vannas Spring Scissors with identical specifications can be purchased from Roboz Surgical Instrument Co.</td>
		</tr>
		<tr>
			<td>Orbital shaker - Rotamax-120</td>
			<td>Heidolph</td>
			<td>N/A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dako Fluorescent Mounting Medium</td>
			<td>Dako Cytomation, Glostrup, Denmark</td>
			<td>DK-5302392</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>X10 PBS formulation</td>
			<td colspan="2">2 gr/lit KCl, 2 gr/lit KH2PO4, 80 gr/lit NaCl, 
			21.7 gr/lit Na2HPO4.7H2O</td>
			<td>The quality of PBS is critical for the success of this protocol</td>
		</tr>
		<tr>
			<td>PBT</td>
			<td colspan="2">X1 PBS + 0.1% Tween 20</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

visualization of proprioceptors in drosophila larvae and pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Watch this Scientific Journal Video about Visualization of Proprioceptors in Drosophila Larvae and Pupae at JoVE.com

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

visualization-of-proprioceptors-in-drosophila-larvae-and-pupae

Research

JoVE Journal

Neuroscience

11.9K ビュー.  Technion-Israel Institute of Technology. の幼虫や蛹に振動や音響に反応する臓器を免疫染色し、可視化する方法キイロショウジョウバエ記載されています。

ビデオ: Visualization of Proprioceptors in Drosophila Larvae and Pupae

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.
The Drosophila larva is an established model system to study thermal nociception, a sensory response to potentially harmful temperatures that is evolutionarily conserved across species1,2. The advantages of Drosophila for such studies are the relative simplicity of its nervous system and the sophistication of the genetic techniques that can be used to dissect the molecular basis of the underlying biology4-6 In Drosophila, as in all metazoans, the response to noxious thermal stimuli generally involves a "nocifensive" aversive withdrawal to the presented stimulus7. Such stimuli are detected through free nerve endings or nociceptors and the amplitude of the organismal response depends on the number of nociceptors receiving the noxious stimulus8. In Drosophila, it is the class IV dendritic arborization sensory neurons that detect noxious thermal and mechanical stimuli9 in addition to their recently discovered role as photoreceptors10. These neurons, which have been very well studied at the developmental level, arborize over the barrier epidermal sheet and make contacts with nearly all epidermal cells11,12. The single axon of each class IV neuron projects into the ventral nerve cord of the central nervous system11 where they may connect to second-order neurons that project to the brain.
Under baseline conditions, nociceptive sensory neurons will not fire until a relatively high threshold is reached. The assays described here allow the investigator to quantify baseline behavioral responses or, presumably, the sensitization that ensues following tissue damage. Each assay provokes distinct but related locomotory behavioral responses to noxious thermal stimuli and permits the researcher to visualize and quantify various aspects of thermal nociception in Drosophila larvae. The assays can be applied to larvae of desired genotypes or to larvae raised under different environmental conditions that might impact nociception. Since thermal nociception is conserved across species, the findings gleaned from genetic dissection in Drosophila will likely inform our understanding of thermal nociception in other species, including vertebrates.

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of ...

Appetitive Associative Olfactory Learning in Drosophila Larvae

In the following we describe the methodological details of appetitive associative olfactory learning in Drosophila larvae. The setup, in combination with genetic interference, provides a handle to analyze the neuronal and molecular fundamentals of specifically associative learning in a simple larval brain.
Organisms can use past experience to adjust present behavior. Such acquisition of behavioral potential can be defined as learning, and the physical bases of these potentials as memory traces1-4. Neuroscientists try to understand how these processes are organized in terms of molecular and neuronal changes in the brain by using a variety of methods in model organisms ranging from insects to vertebrates5,6. For such endeavors it is helpful to use model systems that are simple and experimentally accessible. The Drosophila larva has turned out to satisfy these demands based on the availability of robust behavioral assays, the existence of a variety of transgenic techniques and the elementary organization of the nervous system comprising only about 10,000 neurons (albeit with some concessions: cognitive limitations, few behavioral options, and richness of experience questionable)7-10.
Drosophila larvae can form associations between odors and appetitive gustatory reinforcement like sugar11-14. In a standard assay, established in the lab of B. Gerber, animals receive a two-odor reciprocal training: A first group of larvae is exposed to an odor A together with a gustatory reinforcer (sugar reward) and is subsequently exposed to an odor B without reinforcement 9. Meanwhile a second group of larvae receives reciprocal training while experiencing odor A without reinforcement and subsequently being exposed to odor B with reinforcement (sugar reward). In the following both groups are tested for their preference between the two odors. Relatively higher preferences for the rewarded odor reflect associative learning - presented as a performance index (PI). The conclusion regarding the associative nature of the performance index is compelling, because apart from the contingency between odors and tastants, other parameters, such as odor and reward exposure, passage of time and handling do not differ between the two groups9.

Appetitive Associative Olfactory Learning in Drosophila Larvae

Drosophila larvae are able to associate odor stimuli with gustatory reward. Here we describe a simple behavioral paradigm that allows the analysis of appetitive associative olfactory learning.

In the following we describe the methodological  details of appetitive associative olfactory learning in Drosophila larvae. The setup, in combination with ...

Light Preference Assay to Study Innate and Circadian Regulated Photobehavior in Drosophila Larvae

Light acts as environmental signal to control animal behavior at various levels. The Drosophila larval nervous system is used as a unique model to answer basic questions on how light information is processed and shared between rapid and circadian behaviors. Drosophila larvae display a stereotypical avoidance behavior when exposed to light. To investigate light dependent behaviors comparably simple light-dark preference tests can be applied. In vertebrates and arthropods the neural pathways involved in sensing and processing visual inputs partially overlap with those processing photic circadian information. The fascinating question of how the light sensing system and the circadian system interact to keep behavioral outputs coordinated remains largely unexplored. Drosophila is an impacting biological model to approach these questions, due to a small number of neurons in the brain and the availability of genetic tools for neuronal manipulation. The presented light-dark preference assay allows the investigation of a range of visual behaviors including circadian control of phototaxis.

Light Preference Assay to Study Innate and Circadian Regulated Photobehavior in Drosophila Larvae

Here we describe a light-dark preference test for Drosophila larva. This assay provides information about innate and circadian regulation of light sensing and processing photobehavior.

Light acts as environmental signal to control animal behavior at various levels. The Drosophila larval nervous system is used as a unique model to answer ...

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the underlying neuronal components in the circuits1-6. Application of optogenetics7,8 in the larval neural circuit allows us to manipulate neuronal activity in spatially and temporally patterned ways9-13. Typically, specimens are broadly illuminated with a mercury lamp or LED, so specificity of the target neurons is controlled by binary gene expression systems such as the Gal4-UAS system14,15. In this work, to improve the spatial resolution to "sub-genetic resolution", we locally illuminated a subset of neurons in the ventral nerve cord using lasers implemented in a conventional confocal microscope. While monitoring the motion of the body wall of the semi-intact larvae, we interactively activated or inhibited neural activity with channelrhodopsin16,17 or halorhodopsin18-20, respectively. By spatially and temporally restricted illumination of the neural tissue, we can manipulate the activity of specific neurons in the circuit at a specific phase of behavior. This method is useful for studying the relationship between the activities of a local neural assembly in the ventral nerve cord and the spatiotemporal pattern of motor output.

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Here we describe a protocol for optogenetic manipulation of motoneuronal activity while monitoring changes in motor output (muscle contraction) in semi-intact Drosophila larvae using lasers within a conventional confocal microscope. This technique enables researchers to achieve local perturbation of neural activity in a few neuromeres to elucidate the dynamics of motor circuits.

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the ...

Foraging Path-length Protocol for Drosophila melanogaster Larvae

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral variation. The larval path-length assay was developed to measure individual differences in foraging behavior that were later linked to the foraging gene. Larval path-length is an easily scored trait that facilitates the collection of large sample sizes, at minimal cost, for genetic screens. Here we provide a detailed description of the current protocol for the larval path-length assay first used by Sokolowski. The protocol details how to reproducibly handle test animals, perform the behavioral assay and analyze the data. An example of how the assay can be used to measure behavioral plasticity in response to environmental change, by manipulating feeding environment prior to performing the assay, is also provided. Finally, appropriate test design as well as environmental factors that can modify larval path-length such as food quality, developmental age and day effects are discussed.

Foraging Path-length Protocol for Drosophila melanogaster Larvae

We provide a detailed protocol for a Drosophila melanogaster foraging path-length assay. We discuss the preparation and handling of test animals, how to perform the assay and analyze the data.

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral ...

Novel Assay for Cold Nociception in Drosophila Larvae

How organisms sense and respond to noxious temperatures is still poorly understood. Further, the mechanisms underlying sensitization of the sensory machinery, such as in patients experiencing peripheral neuropathy or injury-induced sensitization, are not well characterized. The genetically tractable Drosophila model has been used to study the cells and genes required for noxious heat detection, which has yielded multiple conserved genes of interest. Little is known however about the cells and receptors important for noxious cold sensing. Although, Drosophila does not survive prolonged exposure to cold temperatures (&#8804;10 &#186;C), and will avoid cool, preferring warmer temperatures in behavioral preference assays, how they sense and possibly avoid noxious cold stimuli has only recently been investigated.
Here we describe and characterize the first noxious cold (&#8804;10 &#186;C) behavioral assay in Drosophila. Using this tool and assay, we show an investigator how to qualitatively and quantitatively assess cold nociceptive behaviors. This can be done under normal/healthy culture conditions, or presumably in the context of disease, injury or sensitization. Further, this assay can be applied to larvae selected for desired genotypes, which might impact thermosensation, pain, or nociceptive sensitization. Given that pain is a highly conserved process, using this assay to further study&#160;thermal nociception will likely glean important understanding of pain processes in other species, including vertebrates.

Novel Assay for Cold Nociception in Drosophila Larvae

Here we demonstrate a novel assay to study cold nociception in Drosophila larvae. This assay utilizes a custom-built Peltier probe capable of applying a focal noxious cold stimulus and results in quantifiable cold-specific behaviors. This technique will allow further cellular and molecular dissection of cold nociception.

How organisms sense and respond to noxious temperatures is still poorly understood. Further, the mechanisms underlying sensitization of the sensory ...

A Temperature Gradient Assay to Determine Thermal Preferences of Drosophila Larvae

Many animals, including the fruit fly, Drosophila melanogaster, are capable of discriminating minute differences in&#160;environmental temperature, which enables them to seek out their preferred thermal landscape. To define the temperature preferences of larvae over a defined linear range, we developed an assay using a temperature gradient. To establish a single-directional gradient, two aluminum blocks are connected to independent water baths, each of which controls the temperature of individual blocks. The two blocks set the lower and upper limits of the gradient. The temperature gradient is established by placing an agarose-coated aluminum plate over the two water-controlled blocks so that the plate spans the distance between them. The ends of the aluminum plate that is set on the top of the water blocks defines the minimum and maximum temperatures, and the regions in-between the two blocks form a linear temperature gradient. The gradient assay can be applied to&#160;larvae of different ages and can be used to identify mutants that exhibit phenotypes, such as those with mutations affecting genes encoding TRP channels and opsins, which are required for temperature discrimination.

A Temperature Gradient Assay to Determine Thermal Preferences of Drosophila Larvae

Here, we present a protocol to determine the preferred environmental temperature of Drosophila larvae using a continuous thermal gradient.

Many animals, including the fruit fly, Drosophila melanogaster, are capable of discriminating minute differences in&#160;environmental temperature, which ...

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at the Drosophila NMJ enables researchers to acquire synaptic information, encompassing both quantitative data on synapse abundance and detailed insights into their morphology. However, the diffuse distribution and limited visual range of the TEM present challenges for the ultrastructural analysis. This study introduces an innovative and efficient sample preparation method that surpasses the conventional approach. The procedure begins by placing a metal mesh at the base of a flat-bottomed bottle or test tube, followed by positioning fixed larvae samples onto the mesh. An additional mesh is placed over the samples, ensuring that they are positioned between the two meshes. The fixed samples are thoroughly dehydrated and infiltrated before proceeding with the embedding procedure. Then embedding of the samples in epoxy resin is performed in a flat sheet manner, which allows for the preparation of muscles for positioning and sectioning. Benefiting from these steps, all the muscles of Drosophila larvae can be visualized under light microscopy, therey facilitating subsequent positioning and sectioning. Excess resin is removed after locating the 6th and 7th muscles of body segments A2 and A3. Serial ultra-thin sectioning of the 6th or 7th muscle is performed.

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

This report provides a new sample preparation procedure for visualizing neuromuscular junctions in Drosophila Larvae. This method is more effective in preventing the curling of the samples compared to the traditional method and is particularly useful for Drosophila neuromuscular junction ultrastructural analysis.

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at ...

Viewing Microtubule Networks in &lt;em&gt;Drosophila&lt;/em&gt; Neuromuscular Junctions and Muscle Cells

Using Drosophila Larval Neuromuscular Junction and Muscle Cells to Visualize Microtubule Network

The microtubule network is an essential component of the nervous system. Mutations in many microtubules regulatory proteins are associated with neurodevelopmental disorders and neurological diseases, such as microtubule-associated protein Tau to neurodegenerative diseases, microtubule severing protein Spastin and Katanin 60 cause hereditary spastic paraplegia and neurodevelopmental abnormalities, respectively. Detection of microtubule networks in neurons is advantageous for elucidating the pathogenesis of neurological disorders. However, the small size of neurons and the dense arrangement of axonal microtubule bundles make visualizing the microtubule networks challenging. In this study, we describe a method for dissection of the larval neuromuscular junction and muscle cells, as well as immunostaining of &#945;-tubulin and microtubule-associated protein Futsch to visualize microtubule networks in Drosophila melanogaster. The neuromuscular junction permits us to observe both pre-and post-synaptic microtubules, and the large size of muscle cells in Drosophila larva allows for clear visualization of the microtubule network. Here, by mutating and overexpressing Katanin 60 in Drosophila melanogaster,&#160;and then examining the microtubule networks in the neuromuscular junction and muscle cells, we&#160;accurately reveal the regulatory role of Katanin 60 in neurodevelopment. Therefore, combined with the powerful genetic tools of Drosophila&#160;melanogaster, this protocol greatly facilitates genetic screening and microtubule dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

Author Spotlight: Understanding Microtubule Network in Drosophila Neuromuscular Junctions 

Here, we present a detailed protocol to visualize the microtubule networks in neuromuscular junctions and muscle cells. Combined with the powerful genetic tools of Drosophila melanogaster, this protocol greatly facilitates genetic screening and microtubule-dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.