We thank Heidi Miller-Mommerskamp for technical help and Iveth Melissa Guatibonza Arevalo for helpful comments on the manuscript. Initial versions of the protocol were developed in the laboratory of Ralph Greenspan. This work was supported by the German research foundation (DFG), in particular through a grant FOR2705 (TP3) to IGK, and by the Simons foundation (Aimon &#8211; 414701) and the Kavli Institute for Brain and Mind (grant number #2017-954) received by SA.

All steps are performed under a stereomicroscope.
1. Preparing the holder
<ol>
	<li>Print the holder &#8216;FlyholderVJove.stl&#8217; (see Supplementary Material) with a 3D printer or have it printed using online services (Figure 1A). Both SLS (Nylon PA12) and multijet fusion (PA12) are suitable.</li>
	<li>Create the head slot.
	<ol>
		<li>Place a piece of sticky tape rectangularly on a flat surface. Cut a slice of approximately 5 mm x 1 cm. Cut the neck slot (~400 x 400 &#181;m) in the middle of the longer side of the tape using fixed parallel blades (two scalpel blades stuck together) to ensure the same width in every holder (Figure 1B).</li>
		<li>Place the tape over the flatter side of the hole in the holder on the bottom side (see Figure 1C and Figure 1D). The tape can be deformed by pushing it down with forceps ~500 &#181;m around the hole; this will further minimize hindering later fly movements.</li>
		<li>Cover the tape and the holder from the top with black nail polish to prevent the buffer from leaking out. The black nail polish will also protect the fly&#8217;s eyes from the excitation light of the microscope. Let the nail polish dry at least an hour before using the holder.</li>
		<li>Once the nail polish is dry, add ~1 &#181;L of grease in the head slot using a rolled tissue to ensure the glue will not wet the back of the fly&#8217;s head (Figure 2A). Make sure not to put grease outside of the slot that would prevent the glue from sticking.</li>
	</ol>
	</li>
	<li>Create the body slot (Figure 2B).
	<ol>
		<li>Optionally, prepare the tape that will be used to position the fly&#8217;s body (see below) in advance. Use the design shown in Figure 2B&#160;(top) to cut a piece of ~2 cm wide tape in two and cut 1.5 mm wide slices. Cut out 0.3 mm deep shoulders and body slot. Make sure that it fits the holder. 
		NOTE: The variability between fly sizes (in particular with sex, age, genotype or species) can make it necessary to adjust the tape design. If the head tends to be ill-centered, it can be helpful to add a V-shaped piece of tape temporarily over the neck slot (Figure 3). Also apply a few microliters of grease inside the neck slot and on the V-shaped tape.</li>
	</ol>
	</li>
</ol>
2. Placing the fly
NOTE: One to four days old female flies are ideal because the female head is bigger and thus easier to dissect than the male head, and younger flies have softer cuticle. For walking experiments the activity of the fly can be increased by matching the experiments with times of higher circadian activity (ZT0 or ZT11), by using saline containing glucose (such as 103 mM NaCl, 3 mM KCl, 5 mM TES, 8 mM trehalose 2 H2O, 10 mM glucose, 26 mM NaHCO3, 1 mM NaH2PO4, 2.5 mM CaCl2&#183;2 H2O, 4 mM MgCl2&#183;6 H2O), by starving the fly up to 24 h with a water only environment, and by heating the environment to ~28 &#176;C during the experiment. Clipping the wings at least one day&#160;in advance also helps to decrease attempts to fly and thus increase the frequency of walking bouts7,29,30.
<ol>
	<li>Fill a Petri dish or pipette tip box lid with ice, place a laboratory tissue on top of the ice and set the holder upside down on it.</li>
	<li>Transfer a fly on ice to paralyze it by sucking it from its vial into a tube and blowing it onto the ice (make sure the ice is not melted which would drown the fly).</li>
	<li>When the fly stops moving, use dull forceps to slide it into the holder with the neck inside the slot (the fly can be held at the basis of the wing) as in Figure 4 (left). The eyes should be at equal positions in respect to the sides of the slot (Figure 4, middle). If needed, add 1 &#181;L grease on the top of the head to prevent the glue (see next step) from reaching the back of the head and remove later.</li>
	<li>Cover the body with a tissue and some ice to make sure the fly does not move during this step and the next (Figure 4, right). Another option to prevent the legs from reaching the head is to use a piece of tape just below the head (see Figure 6).</li>
</ol>
3. Securing the head
<ol>
	<li>Place the head at a ~20&#176; angle from a fully posterior view (around the lateral axis, see Figure 5A). This is a compromise between decreasing the depth to image and on the other side maintaining the front leg free to move and minimizing the stretch of the neck.</li>
	<li>With a rolled-up tissue (Figure 5B), put UV-glue around the head while avoiding soiling the sensory area of interest (antennae, proboscis and/or eyes). For taste experiments, pull the proboscis out and add glue at its base to prevent movement. If no taste experiment is planned, the proboscis is best pushed into the head and fixed with glue to minimize movement (Figure 6).</li>
	<li>Cure the glue with UV-light for 5 s. Carefully clean the surrounding of the head with a rolled-up tissue to remove remaining liquid glue that could stick to the legs and/or soil sensory areas. 
	NOTE: The tape can be roughed with sandpaper to increase glue adhesion if necessary.</li>
	<li>Use a thin strip of tape or a rolled-up tissue to move the legs to the front (if they are not already), so they will not be damaged by the next step.</li>
</ol>
4. Positioning the body
NOTE: This step needs to be performed fast; before the fly recovers from anesthesia.
<ol>
	<li>Remove the ice container and turn the holder around. Remove the water around the fly with a tissue.</li>
	<li>Place the body slot tape (created in step 1) over the hole and gently push the fly&#8217;s body down (Figure 7). Be careful to not stretch the neck too much.</li>
</ol>
5. Sealing the hole
<ol>
	<li>Cover any remaining large holes with tape.</li>
	<li>Add ~1 &#181;L of grease to the back of the head and in the neck area to make sure no glue will wet there.</li>
	<li>With a rolled-up tissue, paint UV-glue around and on top of the tape and on the thorax (upper dorsal part of the mesonotum) to fix it. Cure the glue with UV-light for ~5 s. 
	NOTE: It is important to minimize the use of UV light as it can strongly affect the fly&#8217;s health.</li>
	<li>Carefully clean grease and uncured glue with a laboratory tissue.</li>
	<li>Put ~1 mL of saline on top of the head. Push air bubbles aside with forceps. Look for leaks by placing a coverslip over the saline and turning the holder around to check for saline on the front side. If there are any leaks, remove the saline and fix the hole (either by adding more glue or more grease). 
	NOTE: This can be a good time to pause if necessary. The fly can be offered a small piece of tissue or Styrofoam ball to walk on to prevent flailing and calm the fly down.</li>
</ol>
6. Dissecting the head
NOTE: Use sharpened forceps for the following steps. Very fine forceps are critical as dull forceps will make it more difficult to open the head cuticle and may lead to additional injuries on the fly&#8217;s head or brain. Strong magnification can help at this stage. To that aim, one can replace the oculars of the binocular microscope with 30x oculars.
<ol>
	<li>Make two cuts at the base of the central dark cuticular triangle on each side of the neck (see crosses in Figure 8A).</li>
	<li>Cut around the dark triangle and remove this part of the cuticle.</li>
	<li>The hole in the brain through which muscle 16 and the esophagus go should now be visible and move rhythmically (Video 1 presents this rhythmic movement in a fly with fluorescent muscles). Carefully pinch the top of this area to cut muscle 16 without puncturing the esophagus. If the rhythmic movement of the brain stopped, muscle 16 was likely removed, however, the movement sometimes pauses and restarts later. It is thus important to pay attention to rhythmic movements and perform this step again if necessary.</li>
	<li>Cut the remaining cuticle in small pieces and remove them carefully. Try not to pull on the cuticle too much. Instead use the forceps like a pair of scissors, to cut pieces of tissue. One can start on the medial edges, where the dark triangle was removed before and work one&#8217;s way to the sides. 
	NOTE: Pieces of cuticle can be used to gently scrape of fat bodies if present.</li>
	<li>Remove the air sacks one piece after the other by grabbing them with the forceps and pulling slowly and steadily.</li>
</ol>

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The authors have nothing to disclose.

Drosophila is one of the rare adult animals where the whole brain can be imaged during complex behaviors. Here, we present a method to prepare the fly and expose its whole brain to image ongoing whole brain activity. Several important points should be noted.
Dissecting a small animal such as D. melanogaster is challenging. The method thus requires a lot of practice and patience to master it. However, after training, the procedure takes less than 30 minutes and produces reproducible results.
The method we presented has additional limitations. First, tilting the fly head from its natural position leads to stretching the neck which could be damaging to connective tissue, nerves or muscle. Second, although the ventral subesophageal zone (SEZ) is optically accessible, it is below the semi-transparent esophagus, which decreases the intensity and resolution in this area. Finally, although the holder is out of reach in most directions, the fly still sometimes realizes its presence and pushes on it to try to escape.
Despite these limitations, the comprehensive data obtained from whole brain imaging during behavior and responses to stimuli will make it possible to decipher brain function at the level of the whole network when the animal interacts with and navigates complex, naturalistic environments.

Imaging brain activity using various techniques have deepened the understanding of brain function. In humans, brain imaging techniques have important limitations: while functional magnetic resonance imaging (fMRI) offers spatio-temporal resolution far below single neuron resolution, fast techniques such as electroencephalography (EEG) only allows indirect and partial access to the brain1. In sufficiently big animal models such as rodents, recording of fluorescent activity sensors (e.g., GCaMP) using head-mounted microscopes permits to observe brain activity while the animal is moving in its environment2. Nevertheless, these techniques currently give access only to a small portion of the brain. Head-fixed animals can be imaged more comprehensively, but coverage is still partial (e.g., the cortex surface3). It is only in small animals, such as the zebrafish larvae, C. elegans and Drosophila that the whole brain can be imaged with temporal and spatial resolution at the level of or close to single neurons4.
D. melanogaster is particularly promising because it has long been used as a genetic model organism5 and powerful genetic tools have been developed6. Complemented by the new large scale anatomical network derived from electron microscopy7, the fly could provide unique opportunities to study complex brain dynamics generated on a large scale network8. Although the cuticle is not transparent, and must thus be removed to image the brain, in vivo functional imaging has become more and more common place since the first study in 20029 and several protocols have already been published. However, these methods involve either separating the fly head from the body10, severely restricting the fly&#8217;s movements and/or responses to stimuli11,12,13,14,15, or only permitting a small part of the brain to be imaged9,16,25,26,27,17,18,19,20,21,22,23,24. To complement these nevertheless powerful approaches, we recently developed a preparation to image the whole brain during behavior and responses to various stimuli28.
Here, we build upon this study to present a method specifically developed to image the whole brain while the fly performs semi-naturalistic behavior (i.e. walking and grooming) and responds to sensory stimuli. This is achieved by using an observation holder designed to give access to the whole brain from the dorsal-posterior side, while leaving the antennae and proboscis intact, and permitting the fly to move its legs to walk (e.g., on an air-cushioned ball). Steps for dissecting the back of the head have been refined for speed, reproducibility, and to minimize their effect on the viability and mobility of the fly.

<table>
	<tbody>
		<tr>
			<td>#5 forceps</td>
			<td>FST by DUMONT</td>
			<td>11252-30</td>
			<td>straight tip 0.05 x 0.02 mm, Dumoxel, 11 cm long</td>
		</tr>
		<tr>
			<td>#55 forceps</td>
			<td>FST by DUMONT</td>
			<td>11255-20</td>
			<td>straight tip 0.05 x 0.02 mm, Inox, 11 cm long</td>
		</tr>
		<tr>
			<td>30x oculars</td>
			<td>yegren</td>
			<td>WF30-9-30-H</td>
			<td>WF30X/9 High Eye-point Eyepiece Wide Field View Ocular Optical Lens for Stereo Microscope or Biological Microscope 30X, 30mm without Reticle</td>
		</tr>
		<tr>
			<td>AHOME/UV flashlight</td>
			<td>Shenzhen Yijiawan Technology Co., Ltd</td>
			<td>B07V2W9543 (ASIN)</td>
			<td>365 nm</td>
		</tr>
		<tr>
			<td>Fotoplast Gel/UV Glue</td>
			<td>Dreve Otoplastik GmbH</td>
			<td>44791</td>
			<td>GHS07, GHS08</td>
		</tr>
		<tr>
			<td>Gloss Finish Transparent Tape</td>
			<td>3M Scotch</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>KIMTECH Science/Precision wipes</td>
			<td>Kimberly-Clark Professional</td>
			<td>7552</td>
			<td>11 x 21 cm</td>
		</tr>
		<tr>
			<td>KL 1500 LCD/Microscope light</td>
			<td>Schott</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Leica MS5 Microscope</td>
			<td>Leica</td>
			<td></td>
			<td>WF30X/9</td>
		</tr>
		<tr>
			<td>Nail Lacqueur</td>
			<td>Opi Products Inc., N. Hollywood</td>
			<td>6306585338</td>
			<td>black</td>
		</tr>
		<tr>
			<td>Saline: Hepes NaH2PO4 NaHCO3 MgCl2 CaCl2 NaCl KCl sucrose threalose</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Werner Dorsch GmbH</td>
			<td>78 621; B07SXCXWFS (ASIN)</td>
			<td>soft handle</td>
		</tr>
		<tr>
			<td>Vacuum grease</td>
			<td>Dow corning</td>
			<td>0020080 /100 gr</td>
			<td>Moly Kote 111 Compound Grease Grease Valve Stamp 100 g</td>
		</tr>
	</tbody>
</table>

preparing adult drosophila melanogaster for whole brain imaging during behavior and stimuli responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

The preparation described above allows observation of the whole brain under a microscope for large scale 3D imaging such as classical 2 photons or confocal microscopy, but also faster techniques such as light sheet31 and other structured illumination microscopy techniques (reviewed in32), or light field microscopy28.
The access to the whole brain while observing the behavior and maintaining functional sensory organs allows to answer several questions.
First, what is the overall brain activity when the fly is at rest, during behavior, and when it responds to stimuli? As an example, we include data obtained with a light field microscope showing brain activation during responses to stimuli and behavior. For example, in Video 2, a calcium probe was expressed in all neurons (nsyb-GAL4 and UAS-syt-GCaMP6s (left) or UAS-GCaMP6M (right)) and a puff of odor was presented. Notice how the preparation allows to get an overview of brain activity during the response to the stimulus. The powerful genetic tools in Drosophila can be used to restrict the expression of these sensors to specific neuronal subtypes. In Video 3, we restricted the expression of a calcium sensor to dopaminergic and serotoninergic neurons (TH-GAL4, DDC-GAL4 and UAS-GCaMP6M). Notice the strong synchronous activity over the brain tightly correlated with the fly walking, allowed by observing the whole brain during behavior. In addition to calcium activity, other physical or chemical signals can be imaged (using for example sensors for voltage28,33, metabolism products34,35 or specific neuromodulators36).
To understand brain activity&#8217;s role more specifically, we can ask what regions are involved in what behavior, response to stimuli or spontaneous patterns of activity. The data can indeed be used as an unbiased screen to extract functional regions using techniques such as principal component analysis and independent component analysis. Figure 9A shows different functional regions in different colors. The shape and localization of the functional regions allow to map them to anatomical templates to identify brain regions and in some cases neuron type. Furthermore, once aligned to the anatomical template, fluorescence values can be averaged in anatomical brain regions for quantitative analysis (see Figure 9B). For example, a hierarchical gaussian model applied to data in Figure 9B shows that regions are more active during walk (with a &#916;F/F median of 0.029, 95% credible interval=[0.017 0.041]) but not during groom (&#916;F/F median = -0.0049 with 95% credible interval=[-0.016 0.0059]).
Adding to the understanding depth, the simultaneous recordings of the different functional regions allowed by the preparation can be used to study the dynamical properties of the functional network. This is important because brain regions in all brains studied so far are highly recurrently interconnected and more and more studies show that even sensory areas respond to the animal&#8217;s behavioral state. Several aspects can be looked at such as functional graph properties (e.g., modules) and spatio-temporal patterns that can be fit with dynamical systems (see8 for examples).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61876/61876fig01.jpg" /> 
Figure 1: Preparing the holder (step 1).&#160;A) Holder design (view from the top). B) Neck slot preparation. top: tape design, bottom: parallel blades used to cut the neck slot. C) View of the holder from below. D) Bottom view indicating where to add the neck slot tape. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61876/61876fig02.jpg" /> 
Figure 2: Further preparations (step 1).&#160;A) Add grease into the neck slot to prevent glue to cover the back of the head and to prevent saline leaks. Scale bars, 1 mm. B) Top: shape of the piece of tape that will be used to push the body down. Bottom: make sure the tape fits the holder. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61876/61876fig03.jpg" /> 
Figure 3: Placement of a v-shaped piece of tape to aid&#160;centering.&#160;(A) Bottom view. (B) Top view. Scale bar, 2 mm. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61876/61876fig04.jpg" /> 
Figure 4: Place the fly (step 2).&#160;Left, use two dull forceps to place the body so that the neck is in the neck slot. Middle, align the head. The eyes lie on both sides on the edges of the slot. The fly&#8217;s head is straight. Right, tuck the fly with a tissue covered with ice to keep it from moving. Scale bar, 1 mm. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61876/61876fig05.jpg" /> 
Figure 5: Fix the head (step 3).&#160;A) Ideal angle of the head. B) Add UV glue around the head, avoiding sensory areas of interest. Scale bar, 1 mm. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61876/61876fig06.jpg" /> 
Figure 6: The proboscis&#8217; tip and the antennae can be kept free from glue (step 3).&#160;The white circle indicates the area where the UV-glue has been applied. The arrow shows which parts are left free from glue for olfactory and gustatory experiments. Note the optional tape that prevents the legs from touching the glue area. Scale bar, 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61876/61876fig07.jpg" /> 
Figure 7: Place the back tape (step 4.2).&#160;The tape created in step 1 (see Figure 2, left) is used to place the body and cover the large hole in the holder. Scale bar, 1 mm. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/61876/61876fig08v2.jpg" /> 
Figure 8: Dissection steps (step 6).&#160;A-B) Cut on each side at the base of the dark triangle (crosses) and remove this part of the cuticle C). D) and E) remove muscle 16. Then gently remove the rest of the cuticle F), the air sacks G), and muscles. H) Dissected head. Scale bar, 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61876/61876fig08v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/61876/61876fig09.jpg" /> 
Figure 9: Representative results. A) Functional regions (extracted using PCA and ICA as in28) for a fly expressing GCaMP6 pan-neuronally imaged with a light field microscope (25x, NA=0.95 with a matching f/12 micro lens array). Scale bar, 100 &#181;m. B) Average calcium activity in large brain regions during responses to stimuli and behavior (reproduced from28). <a href="https://www.jove.com/files/ftp_upload/61876/61876fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Muscle 16 movements (step 6.3). GFP was expressed in muscles. Note the pumping movement that comes from the hole just above the trachea. <a href="https://cloudflare.jove.com/files/ftp_upload/61876/Muscle16_Movie.avi" target="_blank">Please click here to download this video.</a>
Video 2: Pan-neuronal calcium activity during response to odor for two different preparations. Balsamic vinegar was puffed onto the fly. Pan-neuronal calcium activity (nsyb-GAL4 driver, UAS-syt-GCaMP6s left and UAS-GCaMP6M right) was imaged with a light field microscope (25x, NA=0.95 with a matching f/12 microlens array). Light field images were then reconstructed as described in ref.28,37. <a href="https://cloudflare.jove.com/files/ftp_upload/61876/B9_B10CombinedStacks.avi" target="_blank">Please click here to download this video.</a>
Video 3: Calcium activity in a neuron type subset during behavior. GCaMP6 was expressed in dopaminergic and serotoninergic neurons (with TH-GAL4 and DDC-GAL4). In combination with the fly genetic tools, the preparation allows to observe a strong increase in activity in many regions during walk. <a href="https://cloudflare.jove.com/files/ftp_upload/61876/Video3.avi" target="_blank">Please click here to download this video.</a>
Supplementary Material.&#160; <a href="https://cloudflare.jove.com/files/ftp_upload/61876/FlyHolderVJove.stl" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses at JoVE.com

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

preparing-adult-drosophila-melanogaster-for-whole-brain-imaging

Research

JoVE Journal

Neuroscience

5.8K Views.  Technical University of Munich. We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

Video: Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

State-Dependency Effects on TMS:  A Look at Motive Phosphene Behavior

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate cortical excitability (either increasing or decreasing it) via the application of localized magnetic field pulses.1,2 Within the field of TMS, the term state dependency refers to the initial, baseline condition of the particular neural region targeted for stimulation. As can be inferred, the effects of TMS can (and do) vary according to this primary susceptibility and responsiveness of the targeted cortical area.3,4,5

In this experiment, we will examine this concept of state dependency through the elicitation and subjective experience of motive phosphenes. Phosphenes are visually perceived flashes of small lights triggered by electromagnetic pulses to the visual cortex. These small lights can assume varied characteristics depending upon which type of visual cortex is being stimulated. In this particular study, we will be targeting motive phosphenes as elicited through the stimulation of V1/V2 and the V5/MT+ complex visual regions.6

In this article, we examine the effects of visually relevant state dependency on TMS induced motive phosphenic presentations.

Transcranial magnetic stimulation (TMS) is a non-invasive neurostimulatory and neuromodulatory technique that can transiently or lastingly modulate ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To complicate matters even further, the odors detected by the insect nervous system usually are not single compounds but mixtures whose composition and concentration ratios vary. This leads to an almost infinite amount of different olfactory stimuli which have to be evaluated by the nervous system.
To understand which aspects of an odor stimulus determine its evaluation by the fly, it is therefore desirable to efficiently examine odor-guided behavior towards many odorants and odor mixtures. To directly correlate behavior to neuronal activity, behavior should be quantified in a comparable time frame and under identical stimulus conditions as in neurophysiological experiments. However, many currently used olfactory bioassays in Drosophila neuroethology are rather specialized either towards efficiency or towards resolution.
Flywalk, an automated odor delivery and tracking system, bridges the gap between efficiency and resolution. It allows the determination of exactly when an odor packet stimulated a freely walking fly, and to determine the animal&#180;s dynamic behavioral reaction.

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

The automated tracking system Flywalk is used for high-resolution quantification of odor-guided behavior in Drosophila melanogaster.

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To ...

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 ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We ...

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at the synaptic level. The Active Zone (AZ) is the region at the presynaptic membrane that mediates neurotransmitter release and is composed of a dense collection of scaffold proteins. AZs of Drosophila melanogaster (Drosophila) photoreceptors undergo molecular remodeling after prolonged exposure to natural ambient light. Thus the level of neuronal activity can rearrange the molecular composition of the AZ and contribute to the regulation of the functional output.
Starting from the light exposure set-up preparation to the immunohistochemistry, this protocol details how to quantify the number, the spatial distribution, and the delocalization level of synaptic molecules at AZs in Drosophila photoreceptors. Using image analysis software, clusters of the GFP-fused AZ component Bruchpilot were identified for each R8 photoreceptor (R8) axon terminal. Detected Bruchpilot spots were automatically assigned to individual R8 axons. To calculate the distribution of spot frequency along the axon, we implemented a customized software plugin. Each axon&#39;s start-point and end-point were manually defined and the position of each Bruchpilot spot was projected onto the connecting line between start and end-point. Besides the number of Bruchpilot clusters, we also quantified the delocalization level of Bruchpilot-GFP within the clusters. These measurements reflect in detail the spatially resolved synaptic dynamics in a single neuron under different environmental conditions to stimuli.

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

Here we show how to quantify the number and spatial distribution of synaptic active zones in Drosophila melanogaster&#160;photoreceptors, highlighted with a genetically encoded molecular marker, and their modulation after prolonged exposure to light.

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at ...

High-throughput Analysis of Locomotor Behavior in the Drosophila Island Assay

Advances in next-generation sequencing technologies contribute to the identification of (candidate) disease genes for movement disorders and other neurological diseases at an increasing speed. However, little is known about the molecular mechanisms that underlie these disorders. The genetic, molecular, and behavioral toolbox of Drosophila melanogaster makes this model organism particularly useful to characterize new disease genes and mechanisms in a high-throughput manner. Nevertheless, high-throughput screens require efficient and reliable assays that, ideally, are cost-effective and allow for the automatized quantification of traits relevant to these disorders. The island assay is a cost-effective and easily set-up method to evaluate Drosophila locomotor behavior. In this assay, flies are thrown onto a platform from a fixed height. This induces an innate motor response that enables the flies to escape from the platform within seconds. At present, quantitative analyses of filmed island assays are done manually, which is a laborious undertaking, particularly when performing large screens.
This manuscript describes the &#34;Drosophila Island Assay&#34; and &#34;Island Assay Analysis&#34; algorithms for&#160;high-throughput, automated data processing and quantification of island assay data. In the setup, a simple webcam connected to a laptop collects an image series of the platform while the assay is performed. The &#34;Drosophila Island Assay&#34; algorithm developed for the open-source software Fiji processes these image series and quantifies, for each experimental condition, the number of flies on the platform over time. The &#34;Island Assay Analysis&#34; script, compatible with the free software R, was developed to automatically process the obtained data and to calculate whether treatments/genotypes are statistically different. This greatly improves the efficiency of the island assay and makes it a powerful readout for basic locomotion and flight behavior. It can thus be applied to large screens investigating fly locomotor ability, Drosophila models of movement disorders, and drug efficacy.

High-throughput Analysis of Locomotor Behavior in the Drosophila Island Assay

The island assay is a relatively new, cost-effective assay that can be used to evaluate the basic locomotor behavior of Drosophila melanogaster. This manuscript describes algorithms for automatic data processing and objective quantification of island assay data, making this assay a sensitive, high-throughput readout for large genetic or pharmacological screens.