Stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018637) were used in this study. The research reported in this publication was supported by the Institutional Development Award (IDeA) Network for Biomedical Research Excellence from the National Institute of General Medical Sciences of the National Institutes of Health under grant number P20GM103430, and by the Rhode Island Foundation. Authors thank J. Waters and P. Snodgrass-Belt for their support in developing this protocol.

1. Assay Preparation (Day 1)
<ol>
	<li>Place the metabolic analyzer (Table of Materials) in an incubator or a temperature-controlled room set to 11 &#176;C. 
	NOTE: This temperature is ideal for measurements taken at 25 &#176;C, as it allows for the ~14 &#176;C temperature fluctuation that occurs while running the assay. This is caused by the heat generated by the instrument during the run.</li>
	<li>Open the appropriate software on the computer attached to the metabolic analyzer. Click on the numerical temperature in the bottom left side of the screen. In the new window that opens, click on the box next to the &#34;Heater on&#34; command and ensure a check mark appears. Adjust the temperature to 25 &#176;C and adjust the tolerance range to 0.2 &#176;C using the arrows. 
	NOTE: Several hours are required to achieve stable temperatures.</li>
	<li>Open the cartridge container (Table of Materials) and remove the cartridge from the utility plate without touching the probes. 
	NOTE: The utility plate is used during the calibration of the cartridge and is not used while the metabolic assay is run.</li>
	<li>Add 200 &#956;L of a calibrant solution (Table of Materials) to the utility plate and place the cartridge back on top of the utility plate to rehydrate the sensor probes on the cartridge. Do not touch the probes and protect them from light.</li>
	<li>Seal the cartridge with the paraffin film.</li>
	<li>Incubate the cartridge overnight at 25 &#176;C.</li>
</ol>
2. Assay Media Preparation (Day 2)
<ol>
	<li>Add 10 mM glucose and 10 mM sodium pyruvate to the assay medium.</li>
	<li>Incubate the medium at 25 &#176;C until the liquid has equilibrated to this temperature.</li>
	<li>Adjust the pH to 7.4 using a pH meter.</li>
</ol>
3. Setup of the Software for the Metabolic Analysis of the Drosophila melanogaster Brains (Day 2)
<ol>
	<li>Set up the metabolic software on the metabolic analyzer used in step 1.2.</li>
	<li>Select &#34;Template&#34; on the software home screen.</li>
	<li>Double-click on &#34;Blank&#34; to start a new assay design.</li>
	<li>Click on the &#34;Plate Map&#34; button in the top toolbar to view the design of the cell assay plate. 
	NOTE: The cell assay plate will contain the brain samples and will be paired with the cartridge before beginning the metabolic assay run.</li>
	<li>Click the Add groups button 3x.
	<ol>
		<li>Double-click on the &#34;Group 1&#34; label and rename it to &#34;Experimental&#34;. 
		NOTE: This group will contain a fly brain and a micro-tissue restraint treated with drugs.</li>
		<li>Double-click on the &#34;Group 2&#34; label and rename it to &#34;Control&#34;. 
		NOTE: This group will contain a fly brain and a micro-tissue restraint with no drug treatment.</li>
		<li>Double-click on the &#34;Group 3&#34; label and rename it to &#34;Restraint only&#34;. 
		Note: This group will contain a micro-tissue restraint without any brains or drug treatment.</li>
	</ol>
	</li>
	<li>Assign the wells to each group based on where samples in the cell plate are intended to be placed.
	<ol>
		<li>Click on the &#34;Experimental&#34; label and then highlight wells B2 - B8 on the plate map.</li>
		<li>Click on the &#34;Control&#34; label and then highlight wells C2-C8 on the plate map.</li>
		<li>Click on the &#34;Restraint only&#34; label and then highlight wells D2-D8 on the plate map.</li>
		<li>Check that all four corners (A1, A12, H1, and H12) are designated as background. 
		Note: Wells along the perimeter of the plate are not used to reduce any edge effects. This is a default setting in the software.</li>
	</ol>
	</li>
	<li>Click on the &#34;Protocol&#34;&#160;button in the top toolbar to set up the protocol.
	<ol>
		<li>Click on the &#34;Edit measurement details&#34; in the baseline measurement box.</li>
		<li>Adjust the time that the metabolic analyzer will measure the brain by changing the basal measure cycle time to &#34;3:00&#34; by clicking the up and down arrows.</li>
		<li>Adjust the time that the metabolic analyzer will wait between brain measurements by changing the basal wait cycle time to &#34;0:00&#34; by clicking the up and down arrows.</li>
		<li>Adjust the time that the metabolic analyzer will mix the media prior to measuring the brain by changing the basal mix cycle time to &#34;1:00&#34; by clicking the up and down arrows.</li>
		<li>Adjust the total number of basal cycles, which includes the measure, wait, and mix cycles, to &#34;7&#34; by clicking the up and down arrows. 
		NOTE: Stable Oxygen Consumption Rate measurements are obtained after ~25 min from the start of the assay.</li>
		<li>Click on the &#34;Add injection&#34; button, located in the upper left toolbar.</li>
		<li>Click on the &#34;Injection label&#34; and change it to &#34;Oligomycin&#34;.</li>
		<li>Adjust the injection measure, wait, and mix times to match those for the basal.</li>
		<li>Adjust the total number of injection cycles to &#34;6&#34; by clicking the up and down arrows.</li>
		<li>Repeat steps 3.7.6 - 3.7.8 but change the label to &#34;Rot/AA&#34; and adjust the total number of injection cycles to 7 by clicking the up and down arrows.</li>
		<li>Click on the &#34;Run Assay&#34; button in the top toolbar.</li>
		<li>&#34;Save&#34;&#160;the assay and begin setting up the assay cartridge (Table of Materials).</li>
	</ol>
	</li>
</ol>
4. Preparation of the Cartridge for the Calibration (Day 2)
<ol>
	<li>Remove the cartridge from the 25 &#176;C incubator and take off the cover.
	<ol>
		<li>Add 20 &#956;L of 100 &#956;M oligomycin to the small upper left circular injection port, port A, in the cartridge for all corresponding experimental wells on the cell plate and add 20 &#956;L of assay media to port A in all other wells, including the control and background wells. 
		NOTE: The cartridge contains 4 ports (A-D) for each of the 96 wells corresponding to the cell plate what will contain the samples. This step is done prior to adding the samples. Adding 20 &#956;L of 100 &#956;M oligomycin results in a final oligomycin concentration of 10 &#956;M in the cell plate well once the drug is injected by the metabolic analyzer. Oligomycin is an inhibitor of ATP synthase. The resulting oxygen consumption value will reflect the amount of ATP-linked respiration in the brain. In order to properly inject the drugs into the specified wells, all ports of the same letter must be filled with the same volume of liquid, even if not in use.</li>
		<li>Add 22 &#956;L of 50 &#956;M rotenone/antimycin A to the small upper right circular port, port B, in the cartridge for all corresponding experimental wells on the cell plate, and add 22 &#956;L of assay media to port B of all other wells, including the control and background wells. 
		NOTE: This results in a final rotenone/Antimycin A concentration of 5 &#956;M in the cell plate well once the drug is injected by the metabolic analyzer. Rotenone and antimycin A are inhibitors of electron transport chain complexes I and III, respectively. The resulting oxygen consumption value will reflect non-mitochondrial respiration in the brain.</li>
		<li>Click &#34;Start run&#34; to place the loaded cartridge with the utility plate into the metabolic analyzer with well A1 positioned in the upper left corner when facing the instrument, with the plate loader extending from the right side of the machine.</li>
	</ol>
	</li>
	<li>Select &#34;I&#8217;m Ready&#34; in the software to begin the equilibration and calibration of the cartridge prior to beginning the metabolic assay with the cell plate containing the samples. 
	Note: The program will ask to save the file and then begin equilibrating and calibrating. These steps can take up to 1 h. Steps 5&#8211;8 are conducted during the equilibration and calibration time.</li>
</ol>
5. Preparation of the Micro-tissue Restraints (Day 2)
<ol>
	<li>Select 1 micro-tissue restraint per brain that is being measured, plus at least 3 for use as restraint-only controls, from the storage container (contains 70% ethanol).</li>
	<li>Rinse the restraints with fresh 70% ethanol and wash them with deionized water 3x for 2 min each, using a mesh basket and 6-well plate (Table of Materials). Add additional water washes if the restraints still give off an alcohol odor.</li>
	<li>Wash the micro-tissue restraints in assay media and leave them in this solution until they are ready for use.</li>
</ol>
6. Dissection of the Drosophila melanogaster Larval Brains (Day 2)
<ol>
	<li>Select late (wandering) third instar larvae from a vial of cultured Oregon-R flies using high precision, style 5 (0.10 x 0.06 mm2) tweezers.</li>
	<li>Place the larva in the well of a clean dissecting spot plate (Table of Materials) containing 500 &#956;L of 1x PBS. 
	NOTE: A spot plate is a glass plate with depressions, used to dissect small tissues and organisms.</li>
	<li>Wash the larva by gently shaking it in the well, using tweezers.</li>
	<li>Move the larva to the clean well of a spot plate containing 500 &#956;L of 1x PBS.</li>
	<li>Place the spot plate under the dissecting microscope.</li>
	<li>Grasp the larva at its midsection with one pair of tweezers, while grasping the eye hooks with a second pair of tweezers.</li>
	<li>Gently and smoothly pull the larva in opposite directions using the two sets of tweezers.</li>
	<li>Visualize the brain, which typically stays attached to the eye hooks and will commonly have eye-antennal discs attached to it.</li>
	<li>Carefully remove additional tissues from the brain, using the eye hooks to hold the brain in place. Lastly, separate the eye hooks from the brain.</li>
	<li>Use tweezers to move the dissected brain to a new well on a spot plate containing 1x PBS.</li>
	<li>Repeat steps 6.1&#8211;6.10 until 14 brains are dissected. 
	NOTE: The total time from the start of the dissection to the assay run should not exceed 30 min.</li>
</ol>
7. Addition of the Dissected Brains to the 96-well Metabolic Assay Cell Plate (Day 2)
<ol>
	<li>Add 50 &#956;L of assay media to the wells of the 96-well metabolic assay plate (Table of Materials) being used in the experiment, including the experimental, control, restraint-only, and background wells.</li>
	<li>Carefully place one brain in each well from A2-A8 and B2-B8 of the cell plate, using tweezers, a spatula, or a pipette. 
	NOTE: This may be done away from the dissecting microscope.</li>
	<li>Under the dissection microscope, use a bent needle micro-probe to sink the brains to the bottom of the well.</li>
	<li>Gently position the brain in the middle of the three raised spheres using the probe.</li>
</ol>
8. Addition of the Micro-tissue Restraints to the 96-well Metabolic Assay Cell Plate (Day 2)
<ol>
	<li>Using tweezers, place the micro-tissue restraint on the edge of the assay plate well, and use the microscope to inspect that it is oriented with the plastic ring facing down and the mesh on top.</li>
	<li>Remove the plate from under the microscope and use tweezers to grasp the restraint on both sides and gently drop it into the well.</li>
	<li>Use a bent needle micro-probe to push the restraint down into the well.</li>
	<li>Under the microscope, verify that the brain can be seen through the tissue restraint and that it is centered in the well.</li>
	<li>Repeat steps 8.1&#8211;8.4 for all the wells that will be in the assay &#8212; A2-A8, B2-B8, and C2-C8.</li>
	<li>Carefully add 130 &#956;L of assay media to each of the experimental, control, and restraint-only wells.</li>
	<li>Verify that the brains and micro-tissue restraints have not moved while adding the media by visualizing them under the microscope.</li>
	<li>Add 180 &#956;L of assay media to the four corner wells for use as background control.</li>
</ol>
9. Addition of the Cell Plate to the Metabolic Analyzer and the Start of the Assay (Day 2)
<ol>
	<li>Verify that the equilibration and calibration are complete by waiting for the software to prompt for the cell plate containing the sample.</li>
	<li>Select &#34;Open Tray&#34; and wait for the instrument to eject the utility plate.</li>
	<li>Remove the utility plate and add the cell plate, without the lid, in the same orientation.</li>
	<li>Select &#34;Load Cell Plate&#34;&#160;for the instrument to take in the cell plate and close the tray, and then begin the assay.</li>
	<li>View the measurements in real-time with error bars on the computer screen running the software.</li>
	<li>Remove the ejected cell plate and cartridge when the assay is complete. 
	NOTE: Paired cartridges and cell plates can be re-used 5x.</li>
</ol>
10. Post-measurement Analysis (Day 2)
<ol>
	<li>Use a dissecting microscope to verify that the brains and the micro-tissue restraints are still positioned properly in the well after the assay has completed. Exclude any wells with abnormalities from the analysis.</li>
	<li>Export the Oxygen Consumption Rate (OCR) and the Extracellular Acidification Rate (ECAR) data for further analysis.</li>
</ol>

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M. Tipping holds a provisional patent for the micro-tissue restraints described in this protocol17.

Here, a novel method for the metabolic analysis of D. melanogaster larval and adult brains ex vivo is described. Under basal conditions, oxygen consumption and extracellular acidification rates indicate how metabolically active the brain is and can determine whether a tissue has become more dependent on mitochondrial versus glycolytic respiration, respectively11.
Treating the brains with inhibitors or therapeutic drugs can provide further information about the metabolic status of the tissue. Metabolic substrates can also be added to determine dependencies on specific metabolites, such as glucose to test for the Warburg effect22 or glutamine to investigate glutaminolysis23&#8212;both common in metabolic reprogramming. For long-term studies, larvae or flies could also be fed drugs instead of using injection ports, and brains could be assayed at intervals or at the end-point, depending on the experimental design. This method can be optimized for numerous applications.
This method has been optimized for brains but can be used to analyze other larval18 and adult tissues. Additionally, other small model systems can utilize this method to measure whole organisms18 or tissues. The only limitation to optimizing this method for other systems is the size of the organ, tissue, or organism. The chamber created by the micro-tissue restraint is the size barrier to running the assay. If the metabolic output is too low for the brain or tissue being tested, and the sample size is not a constraint, pooling samples can be used to increase the assay measurements and sensitivity. Modifying this protocol for other applications should only require minor optimizations, including 1) choosing the appropriate measure and mix cycle numbers and timing, and 2) determining effective treatment concentrations for the tissue. The measure and mixing times were determined by the monitoring oxygen concentration measured by the metabolic analyzer during each cycle. This can be viewed by selecting O2 in the Y2 axis button in the overview window after the run is complete. During each cycle, there should be a drop-in oxygen concentration reflecting the tissue&#39;s oxygen consumption during the measured time, followed by a return to the initial oxygen concentration during the mixing cycle. The measure and mix times should be tested until this pattern is observed. Using this method, other insect brains have been studied. These brains were larger than the fly brains used here but still fit within the chamber generated by the tissue restraint. The OCR readings from these brains were as high as 400 pmol/min (data not shown). The results from these studies, as well as OCR rates that align with other studies measuring whole-fly oxygen consumption18,24, suggest that the D. melanogaster brains were not oxygen-limited. For both D. melanogaster brains and tissues from other organisms, applying this method requires special attention to four critical steps.
To successfully achieve reproducible and biologically relevant measurements, there are critical steps of the protocol that must be followed. First, the use of a micro-tissue restraint is required for this method. Manufacturing these restraints using the specifications and materials listed is essential. The materials were selected due to their inert chemical composition, and their ability to allow proper media exchange during the mixing steps without the creation of air bubbles. Second, a timely dissection and plate preparation is required to maximize the metabolic output of the tissue. Short dissection times should not significantly impair the brain. Other studies have shown that fly brains can be kept alive for hours to days after dissection if properly stored in a perfusion chamber25. However, as seen in Figure 3B and 3D, if brains are left sitting in assay media for long periods of time prior to the assay, the oxygen consumption rates are decreased. During the assay, samples undergo a mixing step during every cycle. This mixing not only aids with the injections of chemical compounds but also acts to recirculate the media and provide oxygen. The brains are able to stay metabolically active for longer periods during these measure-mix cycles than they would be incubating in media on the bench top. Therefore, the dissection of larval brains should be mastered before beginning to set up this assay. When setting up this assay, analyze a small number of genotypes at a time, to minimize the number of brains needed and wells utilized. This will prevent delays between the dissecting and the assay measurements. Third, maintaining a stable temperature is required to analyze the metabolic rates in physiologically relevant conditions. Increased assay temperatures can cause tissue death, observed by lower oxygen consumption rates (Figure 3C). If flies are reared at a different temperature for experimental purposes, the measurements should also be performed at this temperature. While dissections will most likely be performed at room temperature, if the assay will be conducted at a different temperature, the plated and restrained brains should be incubated at the required temperature for 15 min prior to running the assay. Lastly, observing the wells post-assay is critical to determine if tissue positioning affected the measurements. Occasionally, there will be one outlier well that, after observing the plate under the dissecting microscope, can be eliminated from the data analysis due to either loss of tissue or mispositioning, where the brain has slipped out of the center of the well and is stuck under the polymer ring of the restraint. Tissue can be lost if the restraint was not secured properly to the well and was disturbed during the mixing cycles. While neither of these events happens often, if they are not excluded from the analysis, they will lower the rate values substantially.
The measurement of whole-tissue metabolic flux by monitoring oxygen consumption and extracellular acidification provides a biologically relevant method for understanding how metabolism is altered by the genetic landscape or other experimental conditions. This method is sensitive enough to detect metabolic changes in a single D. melanogaster brain, which is not possible using stop-flow respirometry or indirect calorimetry. It is also less technically challenging than using a Clark electrode to measure oxygen consumption. An additional advantage of this protocol is the ability to analyze one whole brain per well. The 96-well format allows for more sensitive readings, and thus, more than one genotype can be assayed at a time due to the smaller number of samples required per assay. While measuring metabolism in tissue is still challenging and requires carefully reared and synchronized animals, this protocol describes a fast, relatively simplistic method, and has the potential to analyze numerous metabolic sensitivities in D. melanogaster larval and adult brains.

Metabolic reprogramming has been identified in many neurological diseases, including Glioblastoma Multiforme (GBM), Huntington&#39;s disease, and Major Depressive Disorder (MDD)1,2,3. As metabolism becomes the focus of therapeutic strategies, tools for basic metabolic research have advanced. However, most of these methods were designed to study cell lines and primary cells or to analyze larger tissues post-fixation or -freezing. Some approaches have relied on kits to simplistically measure specific metabolites, while others have utilized more costly and complex analyses utilizing chromatography in combination with mass spectrometry4 for the same goal. To understand the larger metabolic landscape, metabolic profiling5,6 and metabolic flux analysis (MFA)7 emerged to complement the large-scale proteomic and genomic studies. Profiling provides a quantitative representation of metabolites in a cell or tissue at one point in time, while MFA expands upon this by allowing the tracking of labeled metabolites over time. The latter has been useful in revealing how energy sources are utilized differently in a cell or tissue when in a disease state8. However, these methods do not include the measurement of the overall metabolic rate.
To interrogate the overall metabolic state of the small model systems, traditional methods, such as the Clark electrode9 and indirect calorimetry10, can be used to measure oxygen consumption or configured for stop flow respirometry, to measure oxygen and carbon dioxide concentration, respectively. These techniques, while accurately providing insight into the readout of metabolic reprogramming at the organism level, have limitations. The use of the Clark electrode can be technically challenging and is not designed for high-throughput studies. The stop flow respirometer cannot function with the sensitivity required to assay cells or tissues. Several years ago, new technology was developed specifically for these smaller applications11. These instruments were initially designed to measure the oxygen consumption and extracellular acidification of cell lines and primary cells in a 24- or 96-well format. The simplicity of the set-up and data output established this method as an alternative to the traditional approaches. This methodology is a powerful tool for cells, and recent advances have allowed for the measurement of tissue punches12,13,14. However, the methods utilized in these assays do not allow for the measurement of whole organs from small model systems.
The metabolic analysis of disease models often involves the interaction between cells of specialized function residing within the same tissue. For example, glial cells produce metabolites utilized by neurons. These metabolic interactions are required for the neuronal survival15. Small model systems are advantageous for investigating questions such as these. Studies to measure the metabolism of a whole organ, containing a variety of cell types, will add to the understanding of the energy utilization in vivo. Recently, Becker et al. reported a method of measuring the oxygen consumption in whole fly heads16. By pooling a number of heads together in one well of a 24-well cell plate, oxygen consumption readings were obtained using a metabolic analyzer. While this works well for heads, which are easily separated from the body, it is much more difficult to use for organs such as larval brains, because a large quantity needs to be dissected for this method. Therefore, a method utilizing a 96-well cell plate, which increases the sensitivity of the oxygen consumption and extracellular acidification readings, was developed to assay a single whole larval brain.
The metabolic analyzer used in this study requires the sample being measured to remain at the bottom of the well of a 96-well cell plate. For cells and relatively flat tissues, this is not a challenge; however, for D. melanogaster larval and adult brains, it is not possible using traditional plate coating protocols. The spherical three-dimensional shape of the brains will not reliably adhere to the plate surface, and those that do are often damaged while attempting to place them properly in the well. A critical part of this new protocol is the design and development of micro-tissue restraints17 that provide a small chamber for the brain to reside in without interfering with the metabolic measurements. The chamber generated is approximately 0.016 in in height and provides enough space to easily hold many D. melanogaster brains. The restraints consist of nylon mesh attached to an inert polymer ring and will be discussed further in the Representative Results. Using these restraints minimizes the time required to prepare dissected brains for the metabolic measurement. Optimizing the measuring procedure generates steady, reproducible oxygen consumption and extracellular acidification rates after six measurement cycles (of 25 min). The brains remain metabolically active under these conditions for a minimum of 2 h, which enables injections of therapeutics, inhibitors, or other treatments via the metabolic analyzer cartridge drug delivery ports. This protocol can also be easily adapted for other tissues and small model systems18.
The protocol detailed below describes how to assay the oxygen consumption in a whole Drosophila larval brain when challenged with mitochondrial stress. To stress the brain, oligomycin is added to inhibit the ATP synthase19. The decrease in oxygen consumption from basal readings shows a measurement of the amount of ATP-depending respiration in the brain. Further decreases in the oxygen consumption after treatments with rotenone20 and antimycin A21, inhibitors of electron transport chain complexes I and III, respectively, indicate the amount of non-mitochondrial oxygen consumption in the brain. This assay is one example of how mitochondrial respiration in a fly brain might be compared and how this protocol can be used to compare the metabolism in varying genotypes. However, this protocol can be adapted to measure simply basal oxygen consumption and extracellular acidification rates, as well as to design more elaborate studies investigating specific energy utilization or substrate utilization hypotheses.

<table>
	<tbody>
		<tr>
			<td>Assay medium</td>
			<td>Agilent</td>
			<td>102365-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Calibrant solution</td>
			<td>Agilent</td>
			<td>100840-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Cell-Tak Cell Tissue Adhesive</td>
			<td>Sigma Aldrich</td>
			<td>DLW354240</td>
			<td></td>
		</tr>
		<tr>
			<td>delrin</td>
			<td>Grainger</td>
			<td>2XMK6</td>
			<td></td>
		</tr>
		<tr>
			<td>Drosophila Schneider Media</td>
			<td>Thermo Fisher</td>
			<td>21720-024</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont High Precision Tweezers (style 5)</td>
			<td>Ted Pella</td>
			<td>5622</td>
			<td></td>
		</tr>
		<tr>
			<td>Loctite 409</td>
			<td>Amazon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mitostress kit&#160;</td>
			<td>Agilent</td>
			<td>103015-100</td>
			<td>oligomycin, rotenone, and antimycin A included</td>
		</tr>
		<tr>
			<td>Netwell inserts (6-well)</td>
			<td>VWR</td>
			<td>29442-136</td>
			<td></td>
		</tr>
		<tr>
			<td>nylon mesh</td>
			<td>Component Supply</td>
			<td>U-CMN-64</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm M wrapping film</td>
			<td>Fisher Scientific</td>
			<td>S37440</td>
			<td></td>
		</tr>
		<tr>
			<td>PYREX spot plate</td>
			<td>Fisher Scientific</td>
			<td>13-748B</td>
			<td></td>
		</tr>
		<tr>
			<td>Seahorse XFe96 Analyzer</td>
			<td>Agilent</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Wave v2.4 XFe96 analyzer software</td>
			<td>Agilent</td>
			<td>https://www.agilent.com/en/products/cell-analysis/software-download-for-wave-desktop</td>
			<td>free download to mac or windows</td>
		</tr>
		<tr>
			<td>XFe96 catridge and cell plates</td>
			<td>Agilent</td>
			<td>102416-100</td>
			<td></td>
		</tr>
	</tbody>
</table>

metabolic analysis of drosophila melanogaster larval and adult brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

The protocol presented here requires the use of micro-tissue restraints that meet the specifications described below. Using other methods to hold the brains in place at the bottom of the 96-well cell plate were unsuccessful (Figure 2A and 2B). First, various agents to increase the tissue adherence to the plate were tested. A commercially available tissue adhesive (Table of Materials) is recommended for the use of cells and organoids with cell plates and spheroid plates, respectively. Initially, brains appeared to adhere to the well surface; however, the oxygen consumption (OCR) readings were low, and observing the wells after the assay revealed that the brains were no longer attached to the well surface (Figure 2A). The same results occurred with super glue (Figure 2A). Next, manufacturing small screens to hold the brains within a small chamber at the bottom of the well was attempted (Figure 2B).
Metal screens produced high OCR readings alone, as did metal screens with a polymer ring holding the screen in place (Figure 2B). Plastic mesh netting was used with the same polymer ring. This device caused a negative OCR reading to be obtained. Finally, micro-tissue restraints were designed using an inert polymer for the ring measuring an outer diameter of 0.146 in, an inner diameter of 0.1335 in, a thickness of 0.0125 in, and a height of 0.016 in (Table of Materials). Super glue (Table of Materials) was used to attach the ring to a nylon mesh with a specific pore size of 0.0039 in in diameter (Table of Materials). The pore size is critical, as it allows the media and metabolite exchange without the formation of air bubbles, yet still acts as a barrier to the brains. These restraints alone result in little to no OCR reading, and when used with brains, allow for reproducible readings (Figure 2A and 2B). The verification after the assay showed that the brains were still positioned properly in the wells. These tissue restraints are currently not commercially available but can be manufactured by following the specification above. While this requires precise manufacturing, most machine shops will be able to reproduce this restraint using the materials referenced above.
The protocol was further optimized to account for the assay media, the time allowed before the assay run, and the temperature (Figure 3). Initially, assays were set up in the Schneider medium to model the larval in vivo environment. However, the media used cannot contain buffering agents since pH is used to measure the ECAR. This media, thus, has to be custom made, which is costly. To optimize this, a commercially available assay media designed for metabolic analysis (Table of Materials) was used in place of the Schneider media. The OCR levels were unchanged, even when increasing the glucose levels slightly to model assay media conditions (Figure 3A). All assays from this point on were conducted in the assay medium. The supplements to the media were optimized by observing whether the oxygen consumption and the extracellular acidification rates showed a response when treated with known mitochondrial inhibitors. Next, the waiting time between the dissections and the assay runs was analyzed. An incubation of 3 h significantly dropped the OCR levels (Figure 3B). Figure 3D demonstrates the difference at the sixth time point, which is when the OCR and ECAR levels stabilize for both adult and larval brains. Thus, the protocol indicates to begin the assay immediately following the dissections, unless the experiment requires a temperature equilibration, in which case an incubation for less than 30 min is suggested. The metabolic analyzer is stored in an incubator set to 11 &#176;C to allow for a 25 &#176;C temperature throughout the assay. If the temperatures are elevated due to the ambient temperature and the instrument operation, reduced OCR levels are observed (Figure 3C). This is hypothesized to be due to tissue death.
When the optimized conditions are followed, the assays with the larval and adult brains result in readings of OCR slightly exceeding 150 pmol/min at the stabilized sixth time point, ~25 min into the assay (Figure 4B). This rate is maintained for at least 30 min (Figure 4A) and up to 2 h (data not shown). The ECAR is slightly lower in adult brains than in larval brains at the sixth time point (Figure 4D) and is maintained for at least 30 min (Figure 4C). This finding corresponds to an increase in glycolysis during the larval stages to support growth. An injection with a mitochondrial inhibitor, such as oligomycin, lowers the OCR readings to reveal the ATP-dependent respiration, and further treatment with rotenone and antimycin A lowers the OCR to reveal non-mitochondrial oxygen consumption (Figure 5A). This data demonstrates that these inhibitors are able to penetrate the brain tissue and can be used to compare mitochondrial respiration in brains of varying genotypes. For this assay, the mix, wait, and measure times were optimized by observing the oxygen levels, not the consumption rate, and ensuring that after the mixing, the oxygen level returned to the level prior to the measurement.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58007/58007fig1.jpg" /> 
Figure 1: A schematic of larval brain OCR and ECAR measurements in the metabolic analyzer. The cartridge is prepared a day prior to running the assay. The next day, drugs are added to the injection ports of the cartridge. D. melanogaster larval brains are dissected, and micro-tissue restraints are added to secure the brain to the bottom of the well. The cell plate with the brains is assayed in the metabolic analyzer and the data is analyzed. <a href="https://www.jove.com/files/ftp_upload/58007/58007fig1large.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/58007/58007fig2.jpg" /> 
Figure 2: Micro-tissue restraints are not metabolically active and hold the brain in a chamber at the bottom of a plate well. (A) The OCR of Oregon-R D. melanogaster larval brains is measured in wells coated with tissue adhesive (Table of Materials) or when the brains are super glued to the bottom of the well. The results are compared to those of brains positioned at the bottom of the well using micro-tissue restraints. (B) The OCR is measured in wells containing assay media and metal, metal with a polymer ring, plastic mesh netting and a polymer ring, or micro-tissue restraints. * p-value &#60;0.05, ** p-value &#60;0.01, *** p-value &#60;0.001, **** p-value &#60;0.0001. <a href="https://www.jove.com/files/ftp_upload/58007/58007fig2large.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/58007/58007fig3.jpg" /> 
Figure 3: Optimization of the media, incubation time, and temperature. (A) The OCR is measured in Oregon-R D. melanogaster larval brains using Schneider media (S2) with sodium pyruvate added, S2 with glucose and sodium pyruvate added, and assay media with glucose and sodium pyruvate added. (B) The OCR is measured in larval brains after 3 h of incubation prior to the assay, or after 30 min of incubation prior to the assay. (C) The OCR is measured in larval brains at assay run temperatures of 33 &#176;C and 25 &#176;C. The sixth time point is graphed. (D) The OCR is measured in larval brains after 3 h of incubation prior to the assay, or after 30 min of incubation prior to the assay. The data is reported for the sixth time point. * p-value &#60;0.05, ** p-value &#60;0.01, *** p-value &#60;0.001, **** p-value &#60;0.0001. <a href="https://www.jove.com/files/ftp_upload/58007/58007fig3large.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/58007/58007fig4.jpg" /> 
Figure 4: The OCR and ECAR are reproducibly measured in adult and larval D. melanogaster brains. (A) The OCR is measured in Oregon-R D. melanogaster adult and larval brains for 30 min. (B) The OCR data of the sixth time point from panel A is graphed. This is the time point at which the OCR readings stabilize. (C) The ECAR is measured in D. melanogaster adult and larval brains for 30 min. (D) The ECAR data of the sixth time point from panel A is graphed. This is the time point at which the ECAR readings stabilize. * p-value &#60;0.05, ** p-value &#60;0.01, *** p-value &#60;0.001, **** p-value &#60;0.0001. <a href="https://www.jove.com/files/ftp_upload/58007/58007fig4large.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/58007/58007fig5.jpg" /> 
Figure 5: Oligomycin lowers the D. melanogaster larval brain OCR levels to reveal ATP-dependent respiration, and rotenone/antimycin A further lowers the OCR to reveal non-mitochondrial oxygen consumption. (A) The OCR is measured in Oregon-RD. melanogaster larval brains after a treatment with a 20 &#181;M oligomycin and 5 &#181;M rotenone/antimycin A mixture. The control and restraint-only wells were injected with assay media. <a href="https://www.jove.com/files/ftp_upload/58007/58007fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains at JoVE.com

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...

metabolic-analysis-of-drosophila-melanogaster-larval-and-adult-brains

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JoVE Journal

Neuroscience

9.2K Views.  Providence College. We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

Video: Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

Ex vivo Culturing of Whole, Developing Drosophila Brains

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for investigating the cell biology and development of central brain structures by allowing acute pharmacological interventions and live imaging of cellular processes. As an example of the technique, prior work from our lab1 has shown that a previously unrecognized subcellular compartment lies between the axonal and somatodendritic compartments of axons of the Drosophila central brain. The development of this compartment, referred to as the axon initial segment (AIS)2, was shown genetically to depend on the neuron-specific cyclin-dependent kinase, Cdk5. We show here that ex vivo treatment of wild-type Drosophila larval brains with the Cdk5-specific pharmacological inhibitors roscovitine and olomoucine3 causes acute changes in actin organization, and in localization of the cell-surface protein Fasciclin 2, that mimic the changes seen in mutants that lack Cdk5 activity genetically.
A second example of the ex vivo culture technique is provided for remodeling of the connections of embryonic mushroom body (MB) gamma neurons during metamorphosis from larva to adult. The mushroom body is the center of olfactory learning and memory in the fly4, and these gamma neurons prune their axonal and dendritic branches during pupal development and then re-extend branches at a later timepoint to establish the adult innervation pattern5. Pruning of these neurons of the MB has been shown to occur via local degeneration of neurite branches6, by a mechanism that is triggered by ecdysone, a steroid hormone, acting at the ecdysone receptor B17, and that is dependent on the activity of the ubiquitin-proteasome system6. Our method of ex vivo culturing can be used to interrogate further the mechanism of developmental remodeling. We found that in the ex vivo culture setting, gamma neurons of the MB recapitulated the process of developmental pruning with a time course similar to that in vivo. It was essential, however, to wait until 1.5 hours after puparium formation before explanting the tissue in order for the cells to commit irreversibly to metamorphosis; dissection of animals at the onset of pupariation led to little or no metamorphosis in culture. Thus, with appropriate modification, the ex vivo culture approach can be applied to study dynamic as well as steady state aspects of central brain biology.

Ex vivo Culturing of Whole, Developing Drosophila Brains

This article describes a method by which one can mimic in vivo development of the Drosophila mushroom body in an ex vivo culture system.

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for ...

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of the nervous system often produce viable larvae and adults that have locomotion defective phenotypes that are difficult to adequately describe with text or completely represent with a single photographic image. Current modes of scientific publishing, however, support the submission of digital video media as supplemental material to accompany a manuscript. Here we describe a simple and widely accessible microscopy technique for acquiring high-quality digital video of both Drosophila larval and adult phenotypes from a lateral perspective. Video of larval and adult locomotion from a side-view is advantageous because it allows the observation and analysis of subtle distinctions and variations in aberrant locomotive behaviors. We have successfully used the technique to visualize and quantify aberrant crawling behaviors in third instar larvae, in addition to adult mutant phenotypes and behaviors including grooming.

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Here we describe a simple and widely accessible microscopy technique to acquire high-quality digital video of Drosophila adult and larval mutant phenotypes from a lateral perspective.

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of ...

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 ...

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in communication with the epidermal cells they innervate and thus their analysis benefits from in situ visualization of both neuronally and epidermally expressed proteins by immunofluorescence. Traditional methods of preparing larval fillets for immunofluorescence experiments leave intact the muscle tissue that covers most of the body wall, presenting several challenges to imaging neuronal and epidermal proteins. Here we describe a method for removing muscle tissue from Drosophila larval fillets. This protocol enables imaging of proteins that are otherwise obscured by muscle tissue, improves signal to noise ratio, and facilitates the use of super-resolution microscopy to study da neuron development.

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Studies of neuronal morphogenesis using Drosophila larval dendritic arborization (da) neurons benefit from in situ visualization of neuronal and epidermal proteins by immunofluorescence. We describe a procedure that improves immunofluorescence analysis of da neurons and surrounding epidermal cells by removing muscle tissue from the larval body wall.

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in ...

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

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Chapters in this video

Visualization of Larval Segmental Nerves in 3^rd Instar Drosophila Larval Preparations

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Ex vivo Culturing of Whole, Developing Drosophila Brains

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

Functional Analysis of the Larval Feeding Circuit in Drosophila

Live Imaging of Drosophila Larval Neuroblasts

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Foraging Path-length Protocol for Drosophila melanogaster Larvae

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells