Olfactory experience induces synaptic glomeruli remodeling in the Drosophila brain that
is dose-dependent, temporally-restricted, and reversible only in a short early-life period.
The directionality of this brain circuit remodeling is determined by the odorant acting on
a specific class of olfactory sensory neurons. Ethyl butyrate (EB) odorant induces
synapse elimination in the Or42a receptor neurons. Timed doses of EB exposure with
confocal microscopy imaging allows the investigation of experience-dependent circuit
connectivity pruning in the brain antennal lobe. The sophisticated Drosophila genetic
toolkit enables dissection of the molecular and cellular mechanisms mediating this
olfactory plasticity.

We thank the other Broadie Lab members for their valuable input. Figures were created using BioRender.com. This work was supported by National Institute of Health grants MH084989 and NS131557 to K.B.

1. Odorant exposure
<ol>
	<li>Using a fine paintbrush, sort 40-50 developmentally-staged animals as pharate dark pupae (90+ h post-pupariation at 25 &#176;C) into 25 mm x 95 mm polystyrene Drosophila vials containing standard cornmeal molasses food (Figure 1A).</li>
	<li>Place fine stainless-steel wire mesh over the end of the Drosophila vials to contain the flies while also allowing good airflow. Secure the wire mesh caps with taped transparent film onto the side of the Drosophila vials.</li>
	<li>In a 1.5 mL microcentrifuge tube, place 1 mL of 100% mineral oil (vehicle control) or dissolve the ethyl butyrate (EB) odorant in mineral oil in another tube to produce the desired concentration (e.g., 15% (150 &#181;L) or 25% (250 &#181;L)).</li>
	<li>Place vehicle control or odorant microcentrifuge tubes upright within an air-tight 3700 mL Glasslock container. Using tape, anchor the tubes securely in the center of odorant chambers together with the Drosophila vials (Figure 1B).</li>
	<li>Place the sealed vehicle control and EB odorant chambers containing the staged Drosophila pharate pupae vials into a humidified (70%), temperature-controlled (25 &#176;C) incubator on a 12 h/12 h light/dark cycle.</li>
	<li>Remove the newly-eclosed flies after 4 h of odorant chamber exposure and rapidly transfer 20-25 flies to fresh Drosophila vials in chambers with freshly made odorant (Figure 1A,B). Discard the uneclosed pupae.</li>
	<li>Keep the flies in their sealed odorant chambers in the incubators for a total of 24 h. For longer odorant dosing, rapidly transfer flies to new Drosophila vials in freshly made odorant chambers every 24 h.</li>
	<li>Anesthetize the developmentally-staged flies by submerging them in a dish of 70% ethanol for 1 min in preparation for immediate brain dissection and immunocytochemistry processing.</li>
</ol>
2. Brain dissection
<ol>
	<li>Mark vials as vehicle control (100% mineral oil only) or EB exposed (% EB) by labeling and maintain these designations for the entire duration of brain dissection and subsequent processing. Process 10-20 animals for each genotype/odorant condition.</li>
	<li>Using forceps, transfer a single anesthetized fly into a small dish of freshly made phosphate-buffered saline (PBS)13. Immerse the fly in PBS and continue full brain dissection with the fly fully submerged.</li>
	<li>With fine (#5) sharpened forceps in both hands, place the fly ventral side up and grasp the upper thorax with one forceps and the head under the proboscis with the other forceps. Take care not to penetrate the brain.</li>
	<li>Remove the head from the rest of the body by pulling gently in opposite directions with both hands. The head should easily detach from the thorax; leave the isolated head for dissection (Figure 1C).</li>
	<li>Slide the forceps previously used to grasp the thorax under the opposite side of the proboscis. Begin to gently pull the exoskeleton cuticle in opposite directions so that it tears between the eyes to reveal the brain.</li>
	<li>When pulling, the brain optic lobes may separate along with the exoskeleton. To prevent this, use a slow, steady pull with the two forceps while removing the exoskeleton cuticle (Figure 1C, middle).</li>
	<li>Continue to remove the exoskeleton cuticle from the head. Ensure that all parts of the exoskeleton are removed. Remove any tissues attached to the brain, including the fat body and any projecting trachea.</li>
	<li>To prevent sticking, rinse a P20 pipette tip with PBS + 0.2% Triton-X 100 (PBST). Immediately after dissection, transfer the brains into the fixing solution (4% paraformaldehyde (PFA) + 4% sucrose) in a capped tube (Figure 1C,D).</li>
</ol>
3. Brain immunocytochemistry
<ol>
	<li>Fix the brain for 30 min at room temperature (RT) with end-over-end rotation. While keeping the brains in the tube, pipette off and properly dispose of the hazardous fixative. Quickly wash fixed brains 3x with PBS (Figure 1D).</li>
	<li>Brains can often be found stuck in the cap of the tubes after rotation. To prevent accidental loss of brains, which is a constant hazard in all transfers, use a dissection microscope while pipetting liquids.</li>
	<li>In preparation for antibody labeling, block the fixed brains for 1 h at RT in PBST + 1% bovine serum albumin (BSA) + 0.5% normal goat serum (NGS) with constant end-over-end rotation (Figure 1D).</li>
	<li>Remove the block and incubate brains with the selected primary antibody diluted appropriately in PBST + 0.2% BSA + 0.1% NGS. Incubate the brains overnight (14-16 h) at 4 &#176;C with constant rotation. 
	NOTE: Example antibodies used: rat anti-CadN (dilution 1:50)14 to label synaptic glomeruli and chicken anti-GFP (1:1000)8 to label Or42a-mCD8::GFP (Figure 1D).</li>
	<li>Pipette off the primary antibody. Wash brains 3x for 20 min each with PBST. Incubate brains with secondary antibodies diluted in PBST with 0.2% BSA + 0.1% NGS for 2 h at RT with constant rotation. 
	NOTE: Alternatively, incubate brains overnight (14-16 h) at 4 &#176;C. Dilute antibodies in PBST with 0.2% BSA + 0.1% NGS; secondary antibodies used here: 488 goat anti-chicken (1:250) and 546 goat anti-rat (1:250).</li>
	<li>Pipette off the secondary antibodies. Wash brains 3x for 20 min each with PBST. Finish by washing brains for 20 min with PBS (Figure 1D).</li>
	<li>Prepare glass microscope slides (75 mm x 25 mm) by adding two thin strips of double-sided adhesive tape to the slide ~25 mm apart from one another. This provides a spacer to avoid crushing the brains.</li>
	<li>Pipette ~10-15 &#181;L of mounting medium between the two strips of tape to mount the brains. With a P20 pipette tip pre-rinsed with PBST, transfer the labeled brains onto the microscope slide (Figure 1E).</li>
	<li>Once the brains have been transferred, use a fine paintbrush to align them, ensuring the antennal lobes are facing upwards. Look for the side that has the arched hump, which will contain the antennal lobes.</li>
	<li>Once the brains are properly oriented, cover them with a glass coverslip (No. 1.5H), ensuring the coverslip is secure on the tape. Then, fill in the sides of the coverslip with additional mounting medium (Figure 1E).</li>
	<li>Seal the edges of the coverslip with clear nail polish. Allow the slide to dry thoroughly, and then store it in the refrigerator for subsequent imaging.</li>
</ol>
4. Confocal imaging
<ol>
	<li>Blind all brain slides to both genotype and experience conditions prior to imaging by marking the slide with a coded label for later decoding.</li>
	<li>Use a laser-scanning confocal microscope with a 63x oil-immersion objective. We use a Zeiss 510 META microscope in all images shown here (Figure 1F).</li>
	<li>Use appropriate laser lines for the fluorophores employed. Here, use an Argon 488 and HeNe 543 laser for the antennal lobe synaptic glomeruli and Or42a olfactory sensory neuron imaging (Figure 2).</li>
	<li>Determine the optimal gain and offset for both channels, ideally keeping the gain for both channels &#60;750 and offset &#8776;0. This is done to ensure the signal-to-noise is optimal while limiting the background.</li>
	<li>Position imaging to the center of the brain. When imaging, the VM7 glomeruli reside proximally to the hole left in the middle of the brain following removal of the esophagus during dissection (Figure 2A). 
	NOTE: The VM7 glomeruli will always be close to the opening of the esophagus. However, the exact location and size vary slightly based on brain dissection and mounting differences, so take this into consideration when imaging.</li>
	<li>Select the imaging resolution and optical slice thickness. Here, 1024 x 1024 resolution with a Z-stack slice thickness of 0.37 &#181;m is used.</li>
	<li>Take an entire confocal Z-stack projection through the antennal lobe, ensuring the capture of the full Or42a neuron innervation of the VM7 glomeruli.</li>
</ol>
5. Synaptic measurements
<ol>
	<li>Load the genotype/condition-blinded image into Fiji (1.54f). Split the laser line channels by clicking Image &#62; Color &#62; Split Channels.</li>
	<li>In the Or42a olfactory sensory channel, determine which slices contain the Or42a innervation by scrolling through the entirety of the Z-stack, identifying where the fluorescence begins and ends.</li>
	<li>Create a sum slices projection including only slices containing Or42a neuron innervation by clicking Image &#62; Stacks &#62; Z Project &#62; Sum Slices and entering the desired range.</li>
	<li>In Fiji, click the Lasso tool on the top bar to trace the outline of the Or42a neuron innervation in the VM7 glomerulus, treating each brain side glomerulus independently (Figure 3).</li>
	<li>Multiply the circumference by the number of Z-stack slices and the thickness of each slice to obtain the VM7 synaptic glomerulus innervation volume. To quantify synapse number, the find maxima tool can be used along with the find maxima stacks macro to count synapse puncta in the outlined region8,15.</li>
</ol>

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<li>Stephan, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Drosophila+Psidin+regulates+olfactory+neuron+number+and+axon+targeting+through+two+distinct+molecular+mechanisms%5BTitle%5D)+AND+%22J+Neurosci%22%5BJournal%5D)">Drosophila Psidin regulates olfactory neuron number and axon targeting through two distinct molecular mechanisms.</a> J Neurosci. 32 (46), 16080(2012).</li>
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The authors declare no competing interests.

The odorant exposure and brain imaging protocol presented here can be used to reliably induce and quantify experience-dependent olfactory sensory neuron synaptic glomeruli pruning during an early-life critical period. Earlier studies utilizing this treatment paradigm to explore olfactory circuit remodeling began odorant exposure on the 2nd day after eclosion3,4,5. In contrast, we begin odorant exposure in pharate pupae prior to adult eclosion and find that the vast majority of remodeling occurs within the first 24 h post-eclosion8,9. Thus, an essential step in this odorant exposure protocol is to place staged dark pupae into odor chambers prior to eclosion (Figure 1). This is not for pupal odorant exposure per se but rather to ensure odorant exposure immediately upon adult eclosion. We selected eclosed juvenile adults 4 h following staging to synchronize ages within a tight temporal window and to avoid differences in odorant exposure times. This earlier experience interval is in line with recent studies demonstrating EB olfaction habituation closes after just 2 days post-eclosion10. However, the consequences of odorant exposure timing differences within the critical period (e.g., 0-1 vs. 1-2 days) have not been well characterized. Or42a olfactory sensory neurons selectively innervate the VM7 glomerulus in the antennal lobe, which is easily identified based on the precise glomerular maps and targeted innervation labeling (e.g., Or42a-mCD8::GFP; Figure 2). EB odorant exposure during the critical period does not alter Or42a promoter expression or reporter levels8. Timed odorant exposure causes sequential synapse elimination and axon innervation loss from Or42a receptor neurons. This pruning is completely reversible within the early-life critical period (0-2 dpe) but not at adult maturity (7-9 dpe) when changes in the odorant environment no longer alter innervation. The tight temporal restriction of this sensory experience-dependent mechanism is a major outstanding question in the field. Many attempts are being made to re-open critical period-like plasticity at maturity. The protocol given here is a powerful avenue for investigation.
The ease of the odorant exposure and brain imaging protocol described here allows for wide accessibility and broad adaptability. The odorant exposure experience onset, termination, and duration are all completely at the investigator&#39;s discretion to allow targeted studies of critical period properties, remodeling reversibility, and the re-opening of critical period-like plasticity. Likewise, the strength of the odorant experience can be easily manipulated for the study of dose-dependent synaptic glomeruli pruning (Figure 3). Importantly, studies that use lower EB concentrations (e.g., 15% EB) can be used to test manipulations that could enhance the rate or extent of synaptic glomeruli remodeling. For example, results could show synaptic pruning at lower EB levels (e.g., 15%) to be indistinguishable from higher EB levels (e.g., 25%). Across different manipulations, outcomes can be compared to consistent innervation of odorant vehicle controls and the robust synaptic glomeruli pruning occurring at 25% EB (Figure 3). One limitation of this method is that the odorant exposure entirely relies upon the volatility of the dissolved odorant. Variable factors such as temperature and relative humidity affect the odorant exposure effectiveness8. Fly density in the odorant chambers could also potentially impact the odorant exposure, for example, through the production of fly-derived odorants. A density of no more than 20-25 flies per vial is recommended. The odorant concentration required to induce very strong synaptic glomeruli pruning is likely higher than what is typically experienced in nature10,17. Much higher resolution imaging techniques would be required to assay synapse elimination at lower odorant concentrations. However, the current protocol allows consistent and reliable studies for a range of manipulations.
There are immense benefits to this odorant exposure and brain imaging protocol. Drosophila is an enormously advantageous animal model founded on an extensive and sophisticated genetic toolkit, which can be systematically exploited for the broad study of the gene-environment interactions in critical period olfactory circuit remodeling. Examples shown here include the Or42a-Gal4 transgenic driver line, UAS-mCD8::GFP membrane marker, and Or42a-mCD8::GFP direct fusion. There are no distinguishable differences between synaptic glomeruli pruning with the Or42a direct fusion versus Gal4/UAS indirect labeling approaches8,9. The Or42a-Gal4 driver provides the ability to both manipulate the targeted olfactory receptor neurons and visualize experience-dependent changes in VM7 innervation, whereas the direct fusion line frees up the binary UAS-Gal4 transgenic system for other cell-targeted manipulations. Many other mutant and transgenic lines from the extensive Drosophila toolkit can be employed to test temporal-restriction and experience-dependent mechanisms, including identifying cells mediating synaptic pruning, the intercellular signaling pathways relaying olfactory experience information, and the intracellular machinery required for targeted synapse elimination. Overall, this protocol allows the systematic interrogation of many mechanisms driving olfactory experience-dependent brain circuit remodeling. This odorant exposure and brain imaging protocol can also be used to investigate critical period-specific olfactory experience behavioral modifications, genetic and pharmaceutical manipulations that may enable the re-opening of critical period-like plasticity at maturity and other aspects of olfactory circuit plasticity beyond synaptic glomeruli pruning.

The remodeling of juvenile brain circuits during early life represents the last chance for large-scale synaptic connectivity changes to match the highly variable, unpredictable environment into which an animal is born. As the most abundant group of animals, insects share this evolutionarily conserved, foundational critical period remodeling mechanism1. Critical periods open with the onset of sensory input, exhibit reversible circuit changes to optimize connectivity, and then close when stabilization forces resist further remodeling2. Insects are particularly reliant on olfactory sensory information and show a well-defined olfactory critical period. Drosophila provides an excellent genetic model to investigate this experience-dependent critical period in the juvenile brain. Odorant experience during the first few days following eclosion drives striking circuit connectivity changes in individually identified synaptic glomeruli3,4. The direction of remodeling is dependent on the specific input odorant experience. Some odorants cause an increase in the synaptic glomerulus volume for a couple of days post-eclosion (dpe)3,5,6,7, whereas other odorants cause a rapid elimination of synapses during the 0-2 dpe critical period, resulting in decreased innervation volume8,9,10. Specifically, ethyl butyrate (EB) odorant experience drives dose-dependent synaptic pruning of the Or42a olfactory receptor neurons only during this early-life critical period8. The synapse elimination is completely reversible by modulating EB odorant input within the critical period but becomes permanent following the closure of the critical period. This olfactory experience-dependent synaptic pruning provides a valuable experimental system to elucidate the temporally restricted mechanisms underlying juvenile brain circuit remodeling.
Here, we present a detailed protocol used to induce and analyze EB experience-dependent synaptic pruning of Or42a receptor olfactory sensory neurons during the early-life critical period. We show that Or42a synaptic terminals in the antennal lobe VM7 glomerulus can be specifically labeled by transgenically driving a membrane-tethered mCD8::GFP marker, either directly fused to the Or42a promoter (Or42a-mCD8::GFP)11 or using the Gal4/UAS binary expression system (Or42a-Gal4 driving UAS-mCD8::GFP)12. Individual Or42a neuron synapses can be similarly labeled using targeted transgenic expression of presynaptic active zone markers fused to an array of fluorescent tags (e.g., Bruchpilot::RFP)8 or an electron-dense signal for ultrastructural synapse analyses (e.g., miniSOG-mCherry)8. Or42a synaptic terminals can be imaged with a combination of laser-scanning confocal microscopy and transmission electron microscopy. We show that Or42a synaptic glomeruli pruning is EB dose-dependent, scaling to the concentration of the timed odorant experience. The percentage of EB odorant dissolved in mineral oil used as a vehicle can be varied, as can the timed duration of the odorant exposure in developmentally staged animals. Finally, we show the methods used to analyze the extent of synaptic glomeruli pruning by measuring the VM7 innervation fluorescence intensity and volume. Synapse number can also be quantified by counting labeled synaptic puncta and by measuring synaptic ultrastructure parameters using transmission electron microscopy8. Overall, the protocol shown here is a powerful approach that enables the systematic dissection of both cellular and molecular mechanisms mediating Drosophila olfactory circuit synaptic connectivity pruning during a juvenile critical period. The general odor exposure setup described in this study has been utilized in previous studies using other odors and assaying other glomeruli3,7.

<table><tbody><tr><td>&lt;strong&gt;For Odor Exposure&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;em&gt;Drosophila &lt;/em&gt;vials</td><td>Genesee Scientific</td><td>32-110</td><td></td></tr><tr><td>Ethyl butyrate</td><td>Sigma Aldrich</td><td>E15701</td><td></td></tr><tr><td>Microcentrifuge tubes&amp;nbsp;</td><td>Fisher Scientific&amp;nbsp;</td><td>05-408-129</td><td></td></tr><tr><td>Mineral oil</td><td>Sigma Aldrich</td><td>M3516</td><td></td></tr><tr><td>Odor chambers</td><td>Glasslock</td><td></td><td></td></tr><tr><td>Paint brushes</td><td>Winsor &amp;amp; Newton</td><td>Series 233</td><td></td></tr><tr><td>Parafilm</td><td>Thermofisher</td><td>S37440</td><td></td></tr><tr><td>Wire mesh</td><td>Scienceware</td><td>378460000</td><td></td></tr><tr><td>&lt;strong&gt;Brain Dissection&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Ethanol, 190 proof</td><td>Decon Labs</td><td>2801</td><td>Diluted to 70%</td></tr><tr><td>Forceps</td><td>Fine Science Tools</td><td>11251-30</td><td>Dumont #5</td></tr><tr><td>Paraformaldehyde&amp;nbsp;</td><td>Electron Microscope Sciences</td><td>157-8</td><td>Diluted to 4%</td></tr><tr><td>Petri dishes</td><td>Fisher Scientific&amp;nbsp;</td><td>08-757-100B</td><td></td></tr><tr><td>Phosphate-buffered saline</td><td>Thermo Fisher Scientific</td><td>70011-044</td><td>Diluted to 1x</td></tr><tr><td>Sucrose</td><td>Fisher Scientific&amp;nbsp;</td><td>BP220-1</td><td></td></tr><tr><td>Sylgard</td><td>Electron Microscope Sciences</td><td>24236-10</td><td></td></tr><tr><td>Triton-X 100</td><td>Fisher Scientific&amp;nbsp;</td><td>BP151-100</td><td></td></tr><tr><td>&lt;strong&gt;Brain Immunocytochemistry&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>488 goat anti-chicken</td><td>Invitrogen</td><td>A11039</td><td></td></tr><tr><td>546 goat anti-rat</td><td>Invitrogen</td><td>A11081</td><td></td></tr><tr><td>Bovine serum albumin&amp;nbsp;</td><td>Sigma Aldrich</td><td>A9647</td><td></td></tr><tr><td>Chicken anti-GFP</td><td>Abcam</td><td>13970</td><td></td></tr><tr><td>Coverslips</td><td>Avantor</td><td>48366-067</td><td>25 x 25 mm</td></tr><tr><td>Double-sided tape</td><td>Scotch</td><td>34-8724-5228-8</td><td></td></tr><tr><td>Fluoromount-G&amp;nbsp;</td><td>Electron Microscope Sciences</td><td>17984-25</td><td></td></tr><tr><td>Microscope slides</td><td>Fisher Scientific</td><td>12-544-2</td><td>75 x 25 mm</td></tr><tr><td>Nail polish</td><td>Sally Hansen</td><td>109</td><td>Xtreme Wear, Invisible</td></tr><tr><td>Normal goat serum</td><td>Sigma Aldrich</td><td>G9023</td><td></td></tr><tr><td>Rat anti-CadN</td><td>Developmental Studies Hybridoma Bank</td><td>AB_528121</td><td></td></tr><tr><td>&lt;strong&gt;Confocal/Analysis&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Any computer/laptop</td><td></td><td></td><td></td></tr><tr><td>Confocal microscope</td><td>Carl Zeiss</td><td></td><td>Zeiss 510 META&amp;nbsp;</td></tr><tr><td>Fiji software</td><td>Fiji</td><td></td><td>Version 2.14.0/1.54f</td></tr></tbody></table>

Early-life olfactory sensory experience induces dramatic synaptic glomeruli remodeling in the Drosophila juvenile brain, which is experientially dose-dependent, temporally restricted, and transiently reversible only in a short, well-defined critical period. The directionality of brain circuit synaptic connectivity remodeling is determined by the specific odorant acting on the respondent receptor class of olfactory sensory neurons. In general, each neuron class expresses only a single odorant receptor and innervates a single olfactory synaptic glomerulus. In the Drosophila genetic model, the full array of olfactory glomeruli has been precisely mapped by odorant responsiveness and behavioral output. Ethyl butyrate (EB) odorant activates Or42a receptor neurons innervating the VM7 glomerulus. During the early-life critical period, EB experience drives dose-dependent synapse elimination in the Or42a olfactory sensory neurons. Timed periods of dosed EB odorant exposure allow investigation of experience-dependent circuit connectivity pruning in juvenile brain. Confocal microscopy imaging of antennal lobe synaptic glomeruli is done with Or42a receptor-driven transgenic markers that provide quantification of synapse number and innervation volume. The sophisticated Drosophila genetic toolkit enables the systematic dissection of the cellular and molecular mechanisms mediating brain circuit remodeling.

Figure 1 shows the workflow for the olfactory experience-dependent critical period odorant exposure and brain imaging methods. The protocol starts with the age-matching of pharate dark pupae immediately prior to eclosion (Figure 1A). The pupae are placed into odorant chambers for 4 h, and then newly-eclosed adults are flipped into fresh vials in either the vehicle control or dosed EB odorant chambers (Figure 1B). We typically expose to the odorant for 24 h, although any duration is possible. After odorant exposure, the brain dissection begins by removing the head from the thorax and then removing the cuticle exoskeleton with fine forceps (Figure 1C). The isolated brain is cleaned of any associated tissues that could interfere with imaging and then immediately fixed in 4% paraformaldehyde and 4% sucrose in PBS. A short fix of 30 min is sufficient for good integrity of the brain tissue. The immunocytochemical protocol begins with a 1 h block (e.g., BSA, NGS), overnight incubation in primary antibodies (e.g., anti-GFP, -CadN), and then a brief 2 h incubation in secondary antibodies (Figure 1D). The labeled brains are mounted on glass slides under sealed coverslips with the brain dorsal surface up for best imaging of the antennal lobes (Figure 1E). Note that all the samples are blinded to both genotype and experience conditions by marking the slide with a coded designation for later decoding. A laser-scanning confocal microscope with a 63x oil-immersion objective is used to image the central brain antennal lobes (Figure 1F). Laser lines optimal for the fluorophores used are selected (e.g., Argon 488 and HeNe 543) to image the Or42a olfactory sensory neuron innervation of VM7 in the context of surrounding glomeruli.
It is important to select the optimal gain, offset, imaging resolution, and optical slice thickness to capture the entire Z-stack projection of the Or42a neuron innervation in the VM7 glomeruli. All genotypes and experience conditions to be compared are imaged with identical microscope settings. The Z-stacks are processed using ImageJ software with experience-dependent changes assessed primarily by innervation volume7,14. There are two ways in which volumetric measurements have been computed. One calculates the innervation area slice-by-slice using the ImageJ lasso function and then multiplies each value by the thickness of each slice (µm)8,9. The other uses sum slice Z-projections and multiplies the number of Z-stack slices by slice thickness to determine the average volume. Both methods give consistent results. In addition to volume measurements, fluorescence intensity can also be assessed with maximum intensity projection of VM7 glomeruli16. The weighted sum of all pixels is assessed via ImageJ sum slice max projection, with the pixel number at each intensity level summed to calculate the overall intensity8,9. The background can be subtracted by quantifying an extra-glomerular region of identical size. This subtraction method is more time-consuming but more accurate, as it accounts for varying levels of background antibody labeling in each brain sample to produce a more representative quantification of the innervation fluorescence intensity values. Intensity measurements are made in a typical range of 18-30 optical slices (0.37 µm/slice) within a Z-stack, averaged for each brain16. In nearly all cases, the innervation fluorescence intensity and volume measurements give consistent results.
Across all measurement modalities, the representative results remain unwavering; early-life EB odorant experience results in a significant loss of Or42a neuron innervation. The VM7 glomerulus can be reliably identified by size, shape, and location among other precisely mapped antennal lobe olfactory glomeruli (Figure 2A) and by targeted expression of the Or42a-mCD8::GFP membrane label coupled to glomerular labeling with anti-neural cadherin (CadN; Figure 2B). Timed exposure to the odorant vehicle control (mineral oil) results in Or42a neuron innervation indistinguishable from an untreated, normal animal. Still, EB odorant experience in the first few days post-eclosion (dpe) causes synaptic glomeruli pruning (Figure 2C). This pruning mechanism is entirely restricted to the brief early-life critical period8,10. We typically now use 24 h odorant exposures to assess VM7 glomeruli synaptic pruning and the resulting decrease in innervation volume in the juvenile Drosophila brain. A 24 h exposure (0-1 dpe) to the odorant vehicle control (mineral oil) maintains the normal, dense Or42a-mCD8::GFP innervation (green) in the VM7 synaptic glomeruli (white dashed circles) in both brain hemisphere antennal lobes (Figure 2D, left). In striking contrast, 24 h exposure (0-1 dpe) to 25% EB odorant dissolved in the mineral oil results in strong pruning of Or42a-mCD8::GFP innervation (green) and a loss of the synaptic glomerular volume (Figure 2D, right). These results are both visually obvious and consistently clear. One of the greatest attributes of this odorant experience protocol is the clarity of the brain imaging results, which are also extremely amenable to straightforward innervation fluorescence intensity and volume quantifications.
The major advantage of this odorant experience protocol is the ability to harness it for the study of temporal restriction and odorant dose-dependent mechanisms during critical period synaptic pruning. For EB-driven Or42a neuron synapse elimination in the VM7 glomerulus, the remodeling mechanism is primarily restricted to the early juvenile (0-2 dpe), with no detectable experience-dependent synaptic pruning at maturity (7-9 dpe) except minor modulation at the highest EB exposure levels8. Such temporal restriction defines a critical period, and the understanding of this timing mechanism is an important objective. Another attribute of this protocol is that experience-dependent synaptic pruning is EB dose-dependent (Figure 3). Timed exposure to the odorant vehicle control (mineral oil) with 0% EB odorant results in normal Or42a neuron innervation of the VM7 synaptic glomerulus, which is indistinguishable from an untreated animal. Calculation of the Or42a neuron innervation 3-dimensional volume shows a size typical of a normal juvenile brain (2260 µm3; Figure 3A, left). Representative results show that increasing EB odorant concentration causes progressively greater synaptic glomeruli pruning. For example, a 24 h exposure (0-1 dpe) to 15% EB odorant results in clear and consistent innervation loss (Figure 3A, middle). Quantification of the 3D innervation volume shows a striking reduction compared to vehicle controls (760 µm3 vs. 2260 µm3). A 24 h exposure (0-1 dpe) to 25% EB odorant causes even more innervation loss (281 µm3; Figure 3A, right). Representative quantifications for this range of EB odorant concentrations show consistent results for fluorescence intensity (Figure 3B) and innervation volume (Figure 3C). This dose dependence allows for the study of the means to prevent synaptic pruning as well as enhance pruning with lower odorant exposures to demonstrate increased synaptic glomeruli remodeling. Some prior studies have shown the opposite effect of increased glomerular volume using other odorants with this protocol10. EB odorant exposure in the critical period always results in striking and easily detectable synaptic glomeruli pruning.
<img alt="Drosophila brain dissection with antibody staining and confocal imaging; includes setup and methods." class="xfigimg" src="/files/ftp_upload/66629/66629fig01.jpg" data-altupdatedat="1788304762000" data-alt="Figure 1"> 
Figure 1: Flowchart of odorant exposure studies in the Drosophila juvenile brain. (A) Pharate dark pupae (90+ h post-pupariation at 25 °C) are collected and placed in odorant chambers. Under odorant conditions, 4 h are allowed for eclosion, and then juvenile flies are transferred to a fresh vial. (B) Age-matched flies are divided into either the vehicle control (100% mineral oil) or the ethyl butyrate (EB; % v/v in mineral oil) experience conditions. A fine wire mesh (inset) allows odorant access. Flies are typically exposed for an additional 20 h for a total of 24 h of exposure, 0-1 day post-eclosion (dpe). (C) Flies are dissected by first removing the head, and then fine forceps are used to delicately remove the cuticle exoskeleton and isolate the juvenile brain. The isolated brain is cleared of all surrounding tissues and protruding trachea. (D) Isolated brains are fixed, blocked, and then sequentially incubated in primary and secondary antibodies. The range of antibodies used typically includes a general membrane probe to reveal the antennal lobe synaptic glomeruli and a specific membrane probe for the Or42a olfactory sensory neurons. (E) Labeled brains are placed in a mounting medium on a glass microscope slide with tape spacers under a sealed coverslip. Correct brain orientation (dorsal surface up) is critical for optimal imaging of the antennal lobes. (F) The brains are imaged using a laser-scanning confocal fluorescent microscope. All samples are blinded to genotype and experience conditions prior to imaging and subsequent quantified analyses. <a href="https://www.jove.com/files/ftp_upload/66629/66629fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Antennal lobe diagram and synaptic pruning analysis; GFP labeling, olfactory neuron study." class="xfigimg" src="/files/ftp_upload/66629/66629fig02.jpg" data-altupdatedat="1788304762000" data-alt="Figure 2"> 
Figure 2: Odorant experience-dependent critical period synaptic glomeruli pruning. (A) The antennal lobe (AL) olfactory glomeruli are precisely mapped. A few representative synaptic glomeruli are shown. The Or42a receptor olfactory sensory neuron (OSN) class specifically innervates the VM7 glomeruli. (B) Or42a-mCD8::GFP tagged neurons (green) densely innervate the paired VM7 glomeruli on the left (L) and right (R) sides of the brain, with N-Cadherin (CadN) antibody labeling of all the AL synaptic glomeruli (magenta). The fly line is w1118; Or42a-mCD8::GFP/+. (C) Schematic graphical representation of the Or42a OSN innervation of the VM7 synaptic glomeruli in the vehicle control condition (100% mineral oil; left) and ethyl butyrate exposure (% EB v/v in mineral oil; right) during the juvenile critical period. Experience-dependent innervation pruning only occurs with early-life EB exposure, with synapses eliminated and axons lost, resulting in reduced innervation volume. (D) Representative confocal images of Or42a OSN innervation of the paired VM7 synaptic glomeruli (dashed white outline circles; Or42a-mCD8::GFP, green) following oil vehicle control (left) and 25% EB exposure (right) for 24 h (0-1 dpe). The EB odorant experience drives synaptic pruning and the loss of innervation volume. This experience-dependent mechanism is temporally restricted and transiently reversible only within a well-defined olfactory critical period in the first few days post-eclosion. <a href="https://www.jove.com/files/ftp_upload/66629/66629fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Fluorescence intensity and innervation volume diagram; GFP expression; microscopy result; EB effect." class="xfigimg" src="/files/ftp_upload/66629/66629fig03.jpg" data-altupdatedat="1788304762000" data-alt="Figure 3"> 
Figure 3: Quantifying EB dose-dependent synaptic glomeruli innervation fluorescence intensity and volume. (A) Or42a-mCD8::GFP labeled innervation in a VM7 synaptic glomerulus (green) exposed from 0-1 dpe to the odorant vehicle mineral oil control (0% EB, left), 15% EB (middle), or 25% EB (right). Below: The same images with glomerular innervation outlined (white circles) and the 3-dimensional innervation volume calculated. The vehicle control has VM7 synaptic glomerulus innervation by Or42a neurons that is indistinguishable from the untreated animals. Critical period experience with 15% EB odorant exposure results in obvious synaptic glomerulus pruning, with a 66% decrease in the 3-dimensional innervation volume in this case. Exposure to the higher level of 25% EB results in greater pruning, with a nearly 90% loss of synaptic glomerulus volume in this case. Thus, one can image dose-dependent pruning as the odorant experience increases. (B) Representative results of the Or42a-mCD8::GFP labeled fluorescence intensity within VM7 synaptic glomeruli exposed from 0-1 dpe to the odorant vehicle control (oil, green), 15% EB (middle, orange), or 25% EB (right, black). (C) Representative results of the VM7 glomeruli 3-dimensional innervation volumes under the same critical period odorant exposure conditions. The scatterplots show all the individual data points with the mean ± SEM. The significance is indicated as p≤0.05 (*), p ≤ 0.01 (**), p ≤ 0.001 (***), and p ≤ 0.0001 (****). <a href="https://www.jove.com/files/ftp_upload/66629/66629fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Read this Scientific Article Protocol about Author Spotlight: Exploring Glial Influence in Experience-Dependent Synaptic Pruning During Critical Periods at JoVE.com

Author Spotlight: Exploring Glial Influence in Experience-Dependent Synaptic Pruning During Critical Periods

We describe here methods for inducing and analyzing olfactory experience-dependent remodeling of antennal lobe synaptic glomeruli in the Drosophila juvenile brain.

experience-dependent-remodeling-juvenile-brain-olfactory-sensory

Experience-Dependent Remodeling of Juvenile Brain Olfactory Sensory Neuron Synaptic Connectivity in an Early-Life Critical Period

Early-life olfactory sensory experience induces dramatic synaptic glomeruli remodeling in the Drosophila juvenile brain, which is experientially ...

Early-life experience-dependent remodeling of Drosophila olfactory sensory neurons

	Insects taste the external world through taste hairs, or sensilla, that have pores at their tips. When a sensillum comes into contact with a potential food source, compounds from the food source enter through the pore and activate neurons within. For over 50 years, these responses have been recorded using a technique called tip recording. However, this method has major limitations, including the inability to measure neural activity before or after stimulus contact, and the requirement for tastants to be soluble in aqueous solutions.  We describe here a technique that we call base recording, which addresses these limitations. Base recording allows the measurement of taste neuron activity before, during, and after the stimulus.  Thus it allows extensive analysis of OFF responses that occur after a taste stimulus.  It can be used to study hydrophobic compounds such as long-chain pheromones that have very low solubility in water.   In summary, base recording offers the advantages of single-sensillum electrophysiology as a means of measuring neuronal activity--high spatial and temporal resolution, without the need for genetic tools--and overcomes key limitations of the traditional tip recording technique.

We thank Zina Berman for support, Lisa Baik for comments on the manuscript, and other members of the Carlson laboratory for discussion. This work was supported NIH grant K01 DC020145 to H.K.M.D; and NIH grants R01 DC02174, R01 DC04729, and R01 DC011697 to J.R.C.

The following protocol complies with all the animal care guidelines of Yale University.
1. Flies
<ol>
	<li>Place 10-15 newly emerged flies in fresh standard culture vials at 25 &#176;C and 60% relative humidity in a 12:12 h light-dark cycle.</li>
	<li>Use flies when 3-7 days old.</li>
</ol>
2. Chemosensory stimuli
<ol>
	<li>Obtain chemosensory stimuli of the highest available purity. Store them as recommended by the vendor until use.</li>
	<li>Dissolve chemosensory stimuli and dilute to the desired concentrations in water or another desired, non-toxic solvent such as paraffin oil. Stir the prepared solutions for a minimum of 1 h in the case of dissolved solid compounds.</li>
</ol>
3. Glass stimulus capillary
<ol>
	<li>Pull a glass capillary to hold the stimulus from a borosilicate glass capillary (100 mm length, 1 mm outer diameter, 0.58 mm inner diameter) using a pipette puller instrument. Aim to achieve a tip diameter between 3 &#181;m and 10 &#181;m.</li>
	<li>Fill the glass capillary with the preferred stimulus solution using a microloader pipette tip. Take care to avoid bubbles, which can be removed by gentle tapping.</li>
	<li>If the stimulus crystallizes at the tip, clean or replace the glass stimulus capillary.</li>
</ol>
4. Reference and recording electrodes
<ol>
	<li>Use tungsten rods (127 &#181;m diameter and 76.2 mm length) for both reference and recording electrodes. Sharpen the reference and recording electrodes to approximately 1 &#181;m diameter at the tip (the shapes of these electrodes are depicted in Delventhal et al.36) by dipping them repeatedly for several seconds in either a 10% KNO3 (~1 M) solution or a 10% KOH (1.8 M) solution. 
	NOTE: This solution requires current (0.3-3 mA) to facilitate this process.</li>
</ol>
5. Preparation of the fly for base recording
<ol>
	<li>Draw a single fly from the vial into an aspirator. Withdraw the aspirator and trap the animal by placing a finger over the end.</li>
	<li>Expel the fly into a 200 &#181;L plastic pipette tip. Keeping the end of the aspirator in the pipette tip, use the end to push the fly forward, headfirst, toward the narrow end of the pipette tip.</li>
	<li>Trim the pipette tip at each end (i.e., anterior and posterior to the animal) using a razor blade.</li>
	<li>Use either clay or a small piece of cotton to push the fly forward further, until half of the head protrudes from the end of the trimmed pipette tip. Use forceps to gently push until the labellum at the front of the head is exposed.</li>
	<li>Fix the trimmed pipette tip onto a glass microscope slide using clay (Figure 1).</li>
	<li>Under the stereomicroscope, position the labellum laterally on a cover slip so that one lobe, along with its 31 taste sensilla, is exposed (Figure 1). The cover slip keeps the labellum in place.</li>
</ol>
6. Electrophysiology rig
<ol>
	<li>Select a room for the rig setup that has stable temperature and relative humidity (&#60;70%) and is isolated from sources of electrical and mechanical noise, such as refrigerators and centrifuges.</li>
	<li>Place the microscope on the center of an antivibration table.</li>
	<li>Secure a manual micromanipulator to the antivibration table (Figure 2).</li>
	<li>Attach a stainless-steel shaft that holds the tungsten reference electrode to the manual micromanipulator (Figure 2).</li>
	<li>Connect motorized manipulators-one with a holder for the recording electrode probe and a second one with a holder connected to a stainless-steel shaft for the glass stimulus capillary-to the same table using stands (Figure 2).</li>
	<li>Connect the recording electrode probe to an Intelligent Data Acquisition Controller (IDAC) system or another amplifier/digitizer system.</li>
	<li>Link this IDAC system to the computer at the workstation.</li>
	<li>Ground the manual and motorized manipulators to the same location within the rig.</li>
	<li>Install appropriate acquisition software for the IDAC system on the computer. Ensure that the digital acquisition drivers are compatible with the operating system (e.g., Windows XP-7, -8, or -10) on the computer.</li>
</ol>
7. Recording from taste sensilla
<ol>
	<li>Place the preparation slide on the microscope stage with a low magnification (e.g.,&#160;10x) objective in position. Move the stage until the labellum is in focus at the center of the field of view at both low-magnification and high-magnification (e.g., 50x) objectives.</li>
	<li>Insert the reference electrode into the eye using the low magnification objective. To insert the reference electrode, target the eye on the side of the fly opposite the side with the recording electrode, for example, if the recording electrode is approaching from the right, place the reference electrode in the left eye. Utilize a manual micromanipulator for precise insertion.</li>
	<li>Bring the tip of the glass stimulus capillary into focus at the center of the field of view of both low-magnification and high-magnification objectives using a motorized micromanipulator (Figure 3).</li>
	<li>Under low magnification, bring the recording electrode close to the labellum using a second motorized micromanipulator.</li>
	<li>Under high magnification, insert the recording electrode into the base of a taste sensillum using the motorized micromanipulator until the sound of neuronal firing activity from the audio output of the IDAC system is heard.</li>
	<li>Once a stable signal has been established, start recording the signal using the software that comes with the IDAC system (Figure 4A-D). To start recording, press the Start recording button.</li>
	<li>Bring the tip of the stimulus glass capillary to cover the tip of the taste sensillum using the motorized manipulator.</li>
	<li>To end the stimulus, remove the glass stimulus capillary from the sensillum using the motorized manipulator.</li>
	<li>Mark the start and end of the stimulation manually using a pedal. The pedal is connected to the IDAC, and its communication with the software is facilitated through the IDAC to mark the start/end of the stimulus.</li>
</ol>
8. Analysis
<ol>
	<li>Use the different functions of the software that comes with the IDAC system to sort spike populations by amplitude (when possible) and analyze response dynamics.
	<ol>
		<li>To count spikes, left click on the recording of interest, bringing up a window to select from. Choose To Spikes, initiating another window named Convert waves to spikes. Enter a name in the New field and press the OK button.</li>
		<li>Entering the name in the New field in step 8.1.1 leads to the Amplitude Histogram view. Choose the amplitude to count, then close this view. Left click to add a counter.</li>
		<li>Examine the spikes manually to confirm conclusions based on analysis with software. 
		NOTE: The software also allows the exportation of data in different formats for further analysis.</li>
	</ol>
	</li>
</ol>

<ol><li>Joseph, R. M., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Drosophila+chemoreceptors%3A+a+molecular+interface+between+the+chemical+world+and+the+brain%5BTitle%5D)+AND+%22Trends+Genet%22%5BJournal%5D)">Drosophila chemoreceptors: a molecular interface between the chemical world and the brain.</a> Trends Genet. 31 (12), 683-695 (2015).</li>
<li>Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Drosophila+sensory+receptors-a+set+of+molecular+Swiss+Army+Knives%5BTitle%5D)+AND+%22Genetics%22%5BJournal%5D)">Drosophila sensory receptors-a set of molecular Swiss Army Knives.</a> Genetics. 217 (1), 1-34 (2021).</li>
<li>Dweck, H. K. M., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Molecular+logic+and+evolution+of+bitter+taste+in+Drosophila%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">Molecular logic and evolution of bitter taste in Drosophila.</a> Curr Biol. 30 (1), 17-30 (2020).</li>
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The authors have no conflicts of interest to disclose.

In recordings from some types of sensilla, it can be challenging to differentiate the spikes of different neurons. For example, the sugar neurons and mechanosensory neurons of S and I sensilla produce spikes of similar amplitudes, making it difficult to distinguish them4,14. We find that the use of a very sharp tungsten recording electrode reduces the firing of the mechanosensory neuron, as does the judicious placement of the recording electrode. Insertion of the recording electrode into the collar of the sensillum socket (i.e. into the base but not deep) often results in reduced mechanosensory stimulation. Additionally, we find that if a recording has a high level of firing of the mechanosensory neuron, withdrawing the recording electrode and placing it in a different position relative to the sensillum often leads to a lower level of firing.
Ensuring the stability of the recording electrode throughout experiments is another critical issue (see Figure 4E). Mechanical disturbances or shifts in the electrode position can adversely impact the quality of recordings. Investing time and effort in thorough preparation, as specified in the protocol described above, is key to preventing frustration that may result from labellum movement.
We note one other issue we have observed with certain compounds that produce both ON and OFF responses from I sensilla of the I-a class14. Sometimes, a second delivery of a stimulus elicits the OFF but not the ON response. The reason for this reduction in ON response is unknown.
Another challenge encountered in base recording is that not all sensilla are easily accessible for recording because of their position on the labellum. Of the 31 labellar sensilla, only 26 are convenient to record from, presenting a limitation to the technique, as illustrated in Figure 3B. However, in principle, these sensilla (I8, I9, I10, L9, and S10) can be made accessible by rotation of the Olympus microscope mechanical XY stage.
Finally, we emphasize the importance of investing effort into fly preparation. A well-prepared fly is much more likely to yield reliable and high-quality recordings throughout the course of the experiment. Additionally, sucrose can be used as a positive control to ensure the quality of recording; it elicits a response from all labellar sensilla.

Insects, including drosophilid flies, are endowed with a sophisticated taste system that enables them to extract complex chemical information from their surroundings. This system allows them to discern the chemical composition of various substances, distinguishing between those that are nutritious and those that are harmful1,2.
At the core of this system are specialized structures known as taste hairs or sensilla, strategically located on various body parts. In drosophilid flies, these sensilla are located on the labellum, which is the major taste organ of the fly head1,2,3,4, as well as on the legs and wings1,2,5,6. The labellum is located at the tip of the proboscis and contains two lobes4,7,8. Each lobe is covered with 31 taste sensilla categorized as short, long, and intermediate4,7,8. These sensilla each house 2-4 taste neurons1,2,9,10. These taste neurons express members of at least four different gene families, namely the Gustatory receptor (Gr), Ionotropic receptor (Ir), Pickpocket (Ppk), and Transient receptor potential (Trp) genes1,2,11,12,13. This diversity of receptors and channels equips insects with the ability to recognize a wide array of chemical compounds, including both nonvolatile and volatile cues1,2,14.
For over 50 years, scientists have quantified the response of taste neurons and their receptors using a technique called tip recording3,4,6,8,13,15,16,17,18,19,20,21,22,23,24,25,26,27,28, 
29,30,31,32,33,34,35. However, this method has major limitations. First, neural activity can be measured only during contact with the stimulus, and not before or after contact. This limitation precludes the measurement of spontaneous spiking activity and prevents the measurement of OFF responses. Second, only tastants that are soluble in aqueous solutions can be tested.
These limitations can be overcome by a rarely used alternative electrophysiological technique called &#34;base recording.&#34; Here we describe this technique, which we have adapted from a method used by Marion-Poll and colleagues24, and show the crucial taste coding features that it can now conveniently measure14.

<table><tbody><tr><td>Microscope</td><td>Olympus</td><td>BX51WI</td><td>equipped with a 50X objective (LMPLFLN 50X, Olympus) and 10X eyepieces.&amp;nbsp;</td></tr><tr><td>Antivibration Table</td><td>TMC</td><td>63-7590E</td><td></td></tr><tr><td>motorized Micromanipulators</td><td>Harvard Apparatus and M&amp;auml;rzh&amp;auml;user Micromanipulators</td><td>Micromanipulator PM 10 Piezo Micromanipulator</td><td></td></tr><tr><td>manual Micromanipulators</td><td>M&amp;auml;rzh&amp;auml;user Micromanipulators</td><td>MM33 Micromanipulator</td><td></td></tr><tr><td>Magnetic stands</td><td>ENCO</td><td>Model #625-0930</td><td></td></tr><tr><td>Reference&amp;nbsp; and recording Electrode Holder</td><td>Ockenfels Syntech GmbH</td><td></td><td></td></tr><tr><td>Stimulus glass capillary Holder</td><td>Ockenfels Syntech GmbH</td><td></td><td></td></tr><tr><td>Universal Single Ended Probe</td><td>Ockenfels Syntech GmbH</td><td></td><td></td></tr><tr><td>4-CHANNEL USB ACQUISITION CONTROLLER , IDAC-4</td><td>Ockenfels Syntech GmbH</td><td></td><td></td></tr><tr><td>Stimulus Controllers</td><td>Ockenfels Syntech GmbH</td><td>Stimulus Controller CS 55</td><td></td></tr><tr><td>Personal Computer</td><td>Dell</td><td>Vostro</td><td>Check for compatibility with digital acquisition system and software</td></tr><tr><td>Tungsten Rod</td><td>A-M Systems</td><td>Cat#716000</td><td></td></tr><tr><td>Aluminum Foil and/or Faraday Cage</td><td></td><td></td><td>Electromagnetic noise shielding</td></tr><tr><td>Borosilicate Glass Capillaries</td><td>World Precision Instruments</td><td>1B100F-4</td><td></td></tr><tr><td>Pipette Puller</td><td>Sutter Instrument Company</td><td>Model P-97 Flaming/Brown Micropipette Puller</td><td></td></tr><tr><td>Stereomicroscope</td><td>Olympus</td><td>VMZ 1x-4x</td><td>For fly preparation</td></tr><tr><td>p200 Pipette Tips</td><td></td><td></td><td>Generic</td></tr><tr><td>Microloader tips&amp;nbsp;</td><td>Eppendorf</td><td>E5242956003</td><td></td></tr><tr><td>1 ml Syringe</td><td></td><td></td><td>Generic</td></tr><tr><td>Crocodile clips</td><td></td><td></td><td></td></tr><tr><td>Power Transformers</td><td>STACO ENERGY PRODUCTS</td><td>STACO 3PN221B</td><td>Assembled from P1000 pipette tips, flexible plastic tubing, and mesh</td></tr><tr><td>Modeling Clay</td><td></td><td></td><td>Generic</td></tr><tr><td>Forceps</td><td></td><td></td><td>Generic</td></tr><tr><td>Plastic Tubing</td><td>Saint Gobain</td><td>Tygon S3&amp;trade; E-3603</td><td></td></tr><tr><td>Standard culture vials</td><td>Archon Scientific</td><td></td><td>Narrow 1-oz polystyrene vails, each with 10 mL of glucose medium, preloaded with cellulose acetate plugs</td></tr><tr><td>Berberine chloride (BER)</td><td>Sigma-Aldrich</td><td>Cat# Y0001149</td><td></td></tr><tr><td>Denatonium benzoate (DEN)</td><td>Sigma-Aldrich</td><td>Cat# D5765</td><td></td></tr><tr><td>N,N-Diethyl-m- toluamide (DEET)</td><td>Sigma-Aldrich</td><td>Cat# 36542</td><td></td></tr></tbody></table>

Insects taste the external world through taste hairs, or sensilla, that have pores at their tips. When a sensillum comes into contact with a potential food source, compounds from the food source enter through the pore and activate neurons within. For over 50 years, these responses have been recorded using a technique called tip recording. However, this method has major limitations, including the inability to measure neural activity before or after stimulus contact and the requirement for tastants to be soluble in aqueous solutions. We describe here a technique that we call base recording, which overcomes these limitations. Base recording allows the measurement of taste neuron activity before, during, and after the stimulus. Thus, it allows extensive analysis of OFF responses that occur after a taste stimulus. It can be used to study hydrophobic compounds such as long-chain pheromones that have very low solubility in water. In summary, base recording offers the advantages of single-sensillum electrophysiology as a means of measuring neuronal activity - high spatial and temporal resolution, without the need for genetic tools - and overcomes key limitations of the traditional tip recording technique.

Figure 4A shows spontaneous spikes that arise from a sensillum. They fall into two classes based on amplitude, with the larger spikes deriving from the neuron that is sensitive to bitter compounds and the smaller spikes from the neuron that responds to sugars. The relationship between spike amplitude and functional specificity has been corroborated by genetic experiments4,14,37,38,39 .
Figure 4B shows the response of the bitter-sensitive neuron of the S5 sensillum to the odor of DEET; this response occurred without any contact between the DEET solution and the sensillum. One spike appears to be of intermediate amplitude and may result from the superposition of two smaller spikes, one from the sugar neuron and one from the mechanosensory neuron.
Figure 4C shows the ON response that occurs following contact between a bitter stimulus and the I1 sensillum, followed by an OFF response that occurs following the termination of contact. Not that the magnitude of the OFF response, measured in spikes/s, is greater than the ON response.
Figure 4D shows an ON and an OFF response to the bitter compound berberine from an I1 sensillum of another species of fly, Drosophila virilis. One of the advantages of this technique is that it can be performed on other species, including mosquitoes, without the need to introduce any transgenes into them.
Figure 4E illustrates two problems that can occur during recording. First, if the labellum is not properly secured it can move, producing spikes from the mechanosensory neuron of the sensillum. Second, the recording electrode can become dislodged from the sensillum, often as a result of movement of the labellum. In this case, contact is lost and the electrode must be re-inserted to record spikes.
<img alt="Fly immobilization setup using clay and a coverslip, shown in a microscopy experiment diagram." class="xfigimg" src="/files/ftp_upload/66665/66665fig01v2.jpg" data-altupdatedat="1788304762000" data-alt="Figure 1"> 
Figure 1: Fly preparation for base recording. A trimmed pipette tip containing a female fly with the labellum securely placed on a glass coverslip. (A) Both the pipette tip and the coverslip are held securely on mounds of clay. (B) Higher magnification of labellum resting on a coverslip. <a href="https://www.jove.com/files/ftp_upload/66665/66665fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Micromanipulator setup for electrode positioning in electrophysiology, shown in labeled diagram." class="xfigimg" src="/files/ftp_upload/66665/66665fig02v2.jpg" data-altupdatedat="1788304762000" data-alt="Figure 2"> 
Figure 2: Electrophysiology rig setup. (A) An overview showing the positions of a manual micromanipulator for the reference electrode, a motorized micromanipulator for the glass stimulus capillary, and a motorized micromanipulator for the recording electrode. (B) Overview showing holders for the reference electrode, glass stimulus capillary, and recording electrode. <a href="https://www.jove.com/files/ftp_upload/66665/66665fig02largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Electrophysiology setup for insect sensillum recording using glass stimulus capillary and electrode." class="xfigimg" src="/files/ftp_upload/66665/66665fig03v5.jpg" data-altupdatedat="1788304762000" data-alt="Figure 3"> 
Figure 3: The labellum and sensilla. (A) The labellum under an inverted microscope. Close-up of the labellum showing the taste sensilla of one lobe of the labellum. It also shows the stimulus glass capillary close to one of the large sensilla known as L2 (Large type 2) and a recording electrode in the base of this sensillum. (B) Accessibility of labellar sensilla. The labellum showing sensilla that are easily accessible to base recording and sensilla that are less accessible due to their position in the preparation that we generally use. Abbreviations: A = anterior; M = medial; P = posterior; L = lateral. <a href="https://www.jove.com/files/ftp_upload/66665/66665fig03largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Electrophysiology spike traces; action potentials; sensilla neuron recordings; diagram; stimulus response." class="xfigimg" src="/files/ftp_upload/66665/66665fig04v2.jpg" data-altupdatedat="1788304762000" data-alt="Figure 4"> 
Figure 4: Example traces of spontaneous spikes, bitter neuron responses, and suboptimal recording. (A) Example trace of spontaneous spikes. The spikes are from a bitter-sensitive neuron (red dots) and a sugar-sensitive neuron (green dots) in an I1 sensillum. In the drawing of the sensillum, the bitter, sugar, and mechanosensory neurons are colored red, green, and black, respectively. (B) Example trace of the response of the bitter neuron (red dots) in an S5 sensillum to the vapor of 1 mM DEET. Note that the small amplitude spikes are from the sugar neuron and mechanosensory neurons, which in this example trace are difficult to distinguish by amplitude. Abbreviation: DEET = N,N-diethyl-meta-toluamide. (C) Example trace of ON and OFF responses from a bitter neuron in I1 sensilla to 1 mM denatonium benzoate. Spikes are observed before (spontaneous firing), during (ON response), and after (OFF response) contact with the stimulus. Abbreviation: DEN = denatonium benzoate. (D) Example trace of ON and OFF responses from a bitter neuron in I1 sensilla of D. virilis to 0.5 mM berberine chloride. Spikes are observed before, during, and after contact. Abbreviation: BER = berberine chloride. (E) Example trace of a suboptimal recording. The contact was lost in the middle of the experiment due to the movement of the labellum. <a href="https://www.jove.com/files/ftp_upload/66665/66665fig04largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Author Spotlight: Advances in Chemoreception &#8211; From Insect Odor Receptors to Non-Coding RNAs

A rarely used method of electrophysiological recording, base recording, allows analysis of features of taste coding that cannot be examined by conventional recording methods. Base recording also allows the analysis of taste responses to hydrophobic stimuli that cannot be studied using traditional electrophysiological methods.

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Base Recording: A Technique for Analyzing Responses of Taste Neurons in Drosophila

Insects taste the external world through taste hairs, or sensilla, that have pores at their tips. When a sensillum comes into contact with a potential ...

Base recording:  a technique for analyzing responses of taste neurons in Drosophila

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Today, researchers are studying insect olfaction to classify semiochemical-guided behaviors. The requirement for insect olfaction research is a promising area of research for developing an attractant or repellent for pest control. Methods developed by researchers in the lab are not publicly available. Therefore, we invite researchers in this field to publish their work and to benefit other researchers in the field through this Methods Collection.

JoVE Bioengineering merges both physical and life sciences to understand and predict biological processes. Applying physical science tools to life science questions allow for the discovery of better technologies to measure, diagnose, and clinically treat disease.

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Insect Olfaction

Today, researchers are studying insect olfaction to classify semiochemical-guided behaviors. Curated by Zhao Liu. Explore Bioengineering techniques on JoVE.

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Base Recording: A Technique for Analyzing Responses of Taste Neurons in Drosophila