We thank all the members of the Sierralta Lab. This work was supported by FONDECYT-Iniciaci&#243;n 11200477 (to AGG) and FONDECYT Regular 1210586 (to JS). UAS-FLII12Pglu700&#181;&#948;6 (glucose sensor) was kindly donated by Pierre-Yves Pla&#231;ais and Thomas Preat, CNRS-Paris.

1. Fly strain maintenance and larval synchronization
<ol>
	<li>To perform these experiments, use fly cultures raised at 25 &#176;C on standard Drosophila food composed of 10% yeast, 8% glucose, 5% wheat flour, 1.1% agar, 0.6% propionic acid, and 1.5% methylparaben.</li>
	<li>To follow this protocol, use the following lines: w1118 (experimental control background), OK6-GAL4 (driver for motor neurons), repo-GAL4 (driver for all glial cells), CG-GAL4 (driver for fat bodies), UAS-Pyronic (pyruvate sensor), UAS-FLII12Pglu700&#181;&#948;6 (glucose sensor), UAS-Laconic (lactate sensor), UAS-GCaMP6f (calcium sensor), UAS-AT1.03NL (ATP sensor) and UAS-Chk RNAi GD1829. All the lines expressing sensors or RNAi are in the w1118 genetic background.</li>
	<li>To obtain synchronized third instar wandering larvae, place 300 flies (3-5 days old, 100 males, 200 virgin females) of the desired genetic cross in the egg-laying chamber, which contains a 60 mm Petri dish covered with 1% agarose in phosphate-buffered saline solution (PBS). Place a drop of liquid/creamy yeast (1.5 cm diameter) in the center of the plaque to encourage flies to lay eggs (Figure 1).</li>
	<li>Keep the flies at 25 &#176;C for 3 days replacing the agarose Petri dish with a fresh one and freshly dissolved yeast twice daily.</li>
	<li>Allow the flies to lay eggs for 4 h on the fourth day before changing the Petri dish. Remove this plaque. Then, for 3 h, allow the flies to lay eggs in a new agarose plaque with freshly dissolved yeast. Use these larvae in the experiments.</li>
	<li>After 3 h, remove the plaque containing the eggs and place it in a 25 &#176;C incubator for 24 h.</li>
	<li>Collect 50 to 100 newly hatched larvae from this plaque (0 h after larval hatching) and place them in a plastic vial containing standard food. To allow the larvae to feed properly, ensure that the food is ground and soft. Use the larvae 96 h after the transfer.</li>
</ol>
2. Make the glass coverslips with poly-L-lysine
<ol>
	<li>Perform this step 1 h before the larval dissection. In a 6-well cell culture plate, place 25 mm glass coverslips (the diameter of the covers to be used depends on the recording chamber available in the microscope).</li>
	<li>Place a drop (300 &#181;L) of poly-L-lysine in the center of each coverslip for 30 min at room temperature.</li>
	<li>Wash each coverslip 3x with distilled water, then 2x with a saline solution devoid of Ca2+ (the same solution used to dissect the larvae, see step 3.2). Install the covers in the recording chamber and fill it with Ca2+-free saline solution. 
	&#8203;NOTE: Although the larval brain can adhere directly to the glass coverslips, the adhesion is weak, and the brain occasionally moves or displaces during the experiment due to the continuous flow of saline solution. The addition of the poly-L-lysine step reduces the risk of movements or even brain loss as a result of the washes.</li>
</ol>
3. Dissect the ventral nerve cord (VNC) and fat bodies (FBs)
<ol>
	<li>Gather the wandering third-instar larvae from the desired genetic cross (from step 1.7) and thoroughly wash them 3x with distilled water.</li>
	<li>Place the larvae in a glass dissection dish well containing 750 &#181;L of nominally zero Ca2+ ice-cold saline solution composed of 128 mM NaCl, 2 mM KCl, 4 mM MgCl2, 5 mM trehalose, 5 mM HEPES, and 35 mM sucrose (pH = 6.7, measured with a pH meter and adjusted with 1 M NaOH and HCl 37% v/v). 
	NOTE: Setting the pH to 6.7 is critical because, as previously observed, monocarboxylate transport is highly dependent on the pH of the solution. In addition, 6.7 corresponds to the pH in the hemolymph of a third instar larva17,18.</li>
	<li>Place the larva under a stereomicroscope and make a transverse cut across the back of the abdomen with a pair of forceps.</li>
	<li>Push the jaw with the forceps while turning the larvae inside out.</li>
	<li>Observe the ventral nerve cord (VNC) next to the jaw. Carefully remove the imaginal discs and remaining brain-caudal tissues.</li>
	<li>Separate the VNC with the central brain and optic lobes from the rest of the tissue by cutting the nerves. Transfer the VNC, picking it up with forceps from the remaining nerves, and placing it in the recording chamber containing the Ca2+-free recording solution (same solution from step 3.2). The VNCs will immediately adhere to the coverslips.</li>
	<li>To avoid interference from the remaining nerves, attach them at the bottom of the coverslip using forceps (the nerves will also be fluorescent depending on the driver line used) (Figure 2).</li>
	<li>To perform experiments in fat bodies (FBs) measuring glucose or monocarboxylate transport in another set of experiments, proceed to isolate the tissues following steps 3.1-3.4. Once the larvae are turned inside out, observe that the FB is a white, bilateral, flat tissue.</li>
	<li>Place the isolated FBs expressing the FRET sensors (obtained from a genetic cross using the appropriate drivers) in the recording chamber containing the recording solution without Ca2+. 
	NOTE: The FB is highly hydrophobic (it tends to float on the surface of the solution) and extremely vulnerable to manipulation with forceps, it must be handled with care. Any rough handling of this tissue will result in dead cells later (with a decreased fluorescence of the sensors).</li>
	<li>Place the recording chamber containing VNCs/FBs on the microscope stage.</li>
</ol>
4. Live cell imaging
<ol>
	<li>To acquire images of the tissue expressing sensors, use an upright fluorescence microscope coupled to an emission splitting system and a CCD camera. Turn on the illumination system of the microscope 30 min before starting any experiment. 
	NOTE: Here we use a Spinning Disk fluorescence microscope equipped with a 20x/0.5 water immersion objective to observe both VNCs and FBs. Figure 2 also shows the use of a 40x/0.8 water immersion objective.</li>
	<li>To visualize GCaMP6f fluorescence, set the excitation and emission wavelengths at 488 nm and 540 nm, respectively. For Laconic/Pyronic/ATP/glucose sensors, set the excitation wavelength at 440 nm and the emission wavelengths at 488 nm (mTFP, CFP) and 540 nm (Venus, YFP).</li>
	<li>Acquire images of 512 x 512 pixels size every 2 s for GCaMP6f and every 10-30 s for Laconic/Pyronic/ATP/glucose sensors. Determine the optimal exposure to the light source observing the quality of the image obtained. To follow this protocol, ensure that the images are of 300 ms exposure for Laconic and Pyronic sensors and 100-150 ms for the glucose and ATP sensors. 
	NOTE: The exposure can vary depending on the use of a confocal or normal fluorescence microscope.</li>
	<li>Once the recording chamber is installed in the stage of the microscope, carefully place the water immersion objective and be sure that the objective remains submerged throughout the experiment. Keep the room temperature at 25 &#176;C.</li>
	<li>Connect the recording chamber to a perfusion system and keep the tissues bathed in the recording solution containing 128 mM NaCl, 2 mM KCl, 1.5 mM CaCl2, 4 mM MgCl2, 5 mM trehalose, 5 mM HEPES, and 35 mM sucrose, pH 6.7 (measured with a pH meter and adjusted with NaOH and HCl).</li>
	<li>Maintain a constant flow of 3 mL/min of recording solution through the tissue using gravity, coupled to a low flux peristaltic pump to extract the liquid from the chamber while keeping the volume constant. To achieve the required flow, position the solution-containing tubes (50 mL plastic tubes) 25 cm above the microscope stage. 
	NOTE: This gravity-driven flow is used to prevent tissue movement caused by peristaltic pumps, allowing for more accurate image analysis later on.</li>
	<li>Before any stimulation, maintain the recording solution flowing for 5-10 min to obtain a stable baseline of fluorescence from the sensors. Change the duration as necessary to obtain this baseline.</li>
	<li>Replace the flowing solution with the stimulation solution for 5 min to stimulate the VNC/FBs (with glucose, pyruvate, or lactate at the concentration described for each experiment dissolved in saline solution; see each figure). 
	NOTE: The duration of stimulation can be varied according to the needs of the experiment (for example, glucose pulses can be increased to 10 min to reach a plateau in neurons, as previously reported16).</li>
	<li>To maintain osmolarity in the stimulation solution, rectify the sucrose concentration (a carbohydrate nonmetabolizable by the Drosophila brain) according to the concentration of the monocarboxylate or glucose added.</li>
	<li>Change the solutions using a simple system of valves. Be careful not to move the microscope stage.</li>
	<li>Expose the VNC to 80 &#181;M picrotoxin (PTX, continuously flowing) to stimulate neuronal activity. This procedure increases the frequency of Ca2+ oscillations, making it possible to observe changes in metabolically relevant molecules (e.g., intracellular ATP).</li>
	<li>At the end of any experiment, thoroughly wash the complete flow system, including the tubes and the recording chamber, for at least 10 min with the saline solution and another 10 min with distilled water to eliminate any trace of the solutions used.</li>
</ol>
5. Image processing and data analysis
<ol>
	<li>To process the images obtained, proceed with the steps shown in Figure 3.</li>
	<li>Import the image (Laconic) to the ImageJ software. The image has two views of the sample: the left one is the mTFP signal and the one on the right is the Venus signal. Select a region that includes the cells to be analyzed and separate these images (mTFP and Venus) in a new window. Ensure that these images contain exactly the same area.</li>
	<li>Open the registration plugin and correct the small drift of the tissue that is normally observed in the experiment with the function rigid body. Perform this in both images (mTFP and Venus).</li>
	<li>Select at least 10 regions of interest (ROIs) in the cytosol of the corresponding 10 cells in the mTFP signal (488 nm) and transfer the selections to the Venus (540 nm) image.</li>
	<li>Select two or three ROIs close to the cells being analyzed but with no discernible signal (always within the VNC or tissue analyzed, see Figure 3). These are the background ROIs.</li>
	<li>With the ROIs selected in both images, obtain the mean grey value of each signal using the function measure. Transfer the data to a data sheet and subtract the average background value for each signal.</li>
	<li>Determine the mTFP fluorescence ratio over Venus (for Laconic and Pyronic). This value is computed differently than the other FRET sensors described here (where the ratio is usually YFP/CFP) because when bound to their respective substrates, both Laconic and Pyronic reduce their FRET efficiencies.</li>
	<li>Normalize the data dividing each recorded value by the baseline. In a separate data sheet, calculate the baseline by taking the mean value of the mTFP/Venus ratio during the 2 min preceding the stimulus.</li>
	<li>For the glucose and ATP measurements using FRET-based sensors, calculate the YFP/CFP ratio, and normalize the data as described in step 5.8.</li>
</ol>

Monitoring Brain Metabolite Transport in Drosophila Using FRET Sensors

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</ol>

The authors declare no competing or financial interests.

The use of the Drosophila model for the study of brain metabolism is relatively new26, and it has been shown to share more characteristics with mammalian metabolism than expected, which has primarily been studied in vitro in primary neuron cultures or brain slices. Drosophila excels at in vivo experiments thanks to the battery of genetic tools and genetically encoded sensors available that allows researchers to visualize in real time the metabolic changes caused...

The brain has high energy requirements due to the high cost of restoring ion gradients in neurons caused by neuronal electric signal generation and transmission, as well as synaptic transmission1,2. This high energy demand has long been thought to be met by the continuous oxidation of glucose to produce ATP3. Specific transporters at the blood-brain barrier transfer the glucose in the blood to the brain. Constant glycemic levels ensure that the brain receives a steady supply of glucose4. Interestingly, growing experimental evidence suggests that molecules derived from glucose metabolism, such as lactate and pyruvate, play an important role in the brain cells&#39; energy production5,6. However, there is still some debate about how important these molecules are for energy production and which cells in the brain produce or use them7,8. The lack of appropriate molecular tools with the high temporal and spatial resolution required for this task is a significant issue that has prevented this controversy from being completely resolved.
The development and application of several engineered fluorescent metabolic sensors have resulted in a remarkable increase in our understanding of where and how metabolites are produced and used, as well as how the metabolic fluxes occur during basal and high neuronal activity9. Genetically encoded metabolic sensors based on F&#246;rster resonance energy transfer (FRET) microscopy, such as ATeam (ATP), FLII12Pglu700&#181;&#948;6 (glucose), Laconic (lactate), and Pyronic (pyruvate), have contributed to our understanding of brain energy metabolism10,11,12,13. However, due to the high costs and sophisticated equipment required to conduct experiments on live animals or tissues, results in vertebrate models are still primarily limited to cell cultures (glial cells and neurons).
The emerging use of the Drosophila model to express these sensors has revealed that key metabolic features are conserved across species and their function can be easily addressed with this tool. More importantly, the Drosophila model has shed light on how glucose and lactate/pyruvate are transported and metabolized in the fly brain, the link between monocarboxylate consumption and memory formation, and the remarkable demonstration of how increases in neural activity and metabolic flux overlap14,15,16,17. The method presented here for measuring monocarboxylate, glucose, and ATP levels using genetically encoded FRET sensors expressed in the larval brain allows researchers to learn more about how the brain of Drosophila uses energy, which can be applied to the brains of other animals.
We show that this method is effective for detecting lactate and glucose in glial cells and neurons, and that a monocarboxylate transporter (Chaski) is involved in lactate import into glial cells. We also demonstrate a simple method for studying metabolite changes during increased neuronal activity, which can be easily induced by bath application of a GABAA receptor antagonist. Finally, we show that this methodology can be used to measure monocarboxylate and glucose transport in other metabolically significant tissues, such as fat bodies.

<table><tbody><tr><td>Agarose</td><td>Sigma</td><td>A9539</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>C3881</td><td></td></tr><tr><td>CCD Camera ORCA-R2</td><td>Hamamatsu</td><td>-</td><td></td></tr><tr><td>Cell-R Software</td><td>Olympus</td><td>-</td><td></td></tr><tr><td>CG-GAL4</td><td>Bloomington Drosophila Stock Center</td><td>7011</td><td>Fat body driver</td></tr><tr><td>Dumont # 5 Forceps</td><td>Fine Science Tools</td><td>11252-30</td><td></td></tr><tr><td>DV2-emission splitting system</td><td>Photometrics</td><td>-</td><td></td></tr><tr><td>Glass coverslips (25 mm diameter)</td><td>Marienfeld</td><td>111650</td><td>Germany</td></tr><tr><td>Glucose</td><td>Sigma</td><td>G8270</td><td></td></tr><tr><td>GraphPad Prism</td><td>GraphPad Software</td><td>Version 8,0,2</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td></td></tr><tr><td>ImageJ software</td><td>National Institues of Health</td><td>Version 1,53t</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P9541</td><td></td></tr><tr><td>LUMPlanFl 40x/0.8 water immersion objective</td><td>Olympus</td><td>-</td><td></td></tr><tr><td>Methylparaben</td><td>Sigma</td><td>H5501</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>M1028</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653</td><td></td></tr><tr><td>OK6-GAL4</td><td>Bloomington Drosophila Stock Center</td><td></td><td>Motor neuron driver</td></tr><tr><td>Picrotoxin</td><td>Sigma</td><td>P1675S</td><td>&lt;strong&gt;CAUTION-Fatal if swallowed&lt;/strong&gt;</td></tr><tr><td>Poly-L-lysine</td><td>Sigma</td><td>P4707</td><td></td></tr><tr><td>Propionic Acid</td><td>Sigma</td><td>P1386</td><td></td></tr><tr><td>Repo-GAL4</td><td>Bloomington Drosophila Stock Center</td><td>7415</td><td>Glial cell driver (all)</td></tr><tr><td>Sodium Lactate</td><td>Sigma</td><td>71718</td><td></td></tr><tr><td>Sodium pyruvate</td><td>Sigma</td><td>P2256</td><td></td></tr><tr><td>Spinning Disk fluorescence Microscope BX61WI</td><td>Olympus</td><td>-</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S0389</td><td></td></tr><tr><td>Trehalose</td><td>US Biological</td><td>T8270</td><td></td></tr><tr><td>UAS-AT1.03NL&amp;nbsp;</td><td>Kyoto Drosophila Stock Center</td><td>117012</td><td>ATP sensor</td></tr><tr><td>UAS-Chk RNAi GD1829</td><td>Vienna Drosophila Resource Center</td><td>v37139</td><td>Chk RNAi line</td></tr><tr><td>UAS-FLII&lt;sup&gt;12&lt;/sup&gt;Pglu700md6&amp;nbsp;</td><td>Bloomington Drosophila Stock Center</td><td>93452</td><td>Glucose sensor</td></tr><tr><td>UAS-GCaMP6f&amp;nbsp;</td><td>Bloomington Drosophila Stock Center</td><td>42747</td><td>Calcium sensor</td></tr><tr><td>UAS-Laconic</td><td>Sierralta Lab</td><td>-</td><td>Lactate sensor</td></tr><tr><td>UAS-Pyronic</td><td>Pierre Yves Placais/Thomas Preat</td><td>-</td><td>CNRS-Paris</td></tr><tr><td>UMPlanFl 20x/0.5 water immersion objective</td><td>Olympus</td><td>-</td><td></td></tr></tbody></table>

visualizing monocarboxylates and other relevant metabolites in the ex vivo drosophila larval brain using genetically encoded sensors

The high energy requirements of brains due to electrical activity are one of their most distinguishing features. These requirements are met by the production of ATP from glucose and its metabolites, such as the monocarboxylates lactate and pyruvate. It is still unclear how this process is regulated or who the key players are, particularly in Drosophila.
Using genetically encoded F&#246;rster resonance energy transfer-based sensors, we present a simple method for measuring the transport of monocarboxylates and glucose in glial cells and neurons in an ex-vivo Drosophila larval brain preparation. The protocol describes how to dissect and adhere a larval brain expressing one of the sensors to a glass coverslip. 
We present the results of an entire experiment in which lactate transport was measured in larval brains by knocking down previously identified monocarboxylate transporters in glial cells. Furthermore, we demonstrate how to rapidly increase neuronal activity and track metabolite changes in the active brain. The described method, which provides all necessary information, can be used to analyze other Drosophila living tissues.

For up to 1 h, this procedure allows for easy measurement of intracellular changes in the fluorescence of monocarboxylate and glucose sensors. As shown in Figure 4, Laconic sensors in both glial cells and motor neurons respond to 1 mM lactate at a similar rate at the start of the pulse, but motor neurons reach a higher increase over the baseline during the 5 min pulse, as previously demonstrated17. This lactate concentration was chosen because it is comparable to the ...

Watch this Scientific Journal Video about Author Spotlight: Advances in Brain Energy Metabolism Research Using the Drosophila Model at JoVE.com

Author Spotlight: Advances in Brain Energy Metabolism Research Using the Drosophila Model

Here we present a protocol to visualize the transport of monocarboxylates, glucose, and ATP in glial cells and neurons using genetically encoded F&#246;rster resonance energy transfer-based sensors in an ex-vivo Drosophila larval brain preparation.

Visualizing Monocarboxylates and Other Relevant Metabolites in the Ex Vivo Drosophila Larval Brain Using Genetically Encoded Sensors

The high energy requirements of brains due to electrical activity are one of their most distinguishing features. These requirements are met by the ...

visualizing-monocarboxylates-other-relevant-metabolites-ex-vivo

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Neuroscience

701 Views.  Universidad de Chile. Here we present a protocol to visualize the transport of monocarboxylates, glucose, and ATP in glial cells and neurons using genetically encoded Förster resonance energy transfer-based sensors in an ex-vivo Drosophila larval brain preparation.

Video: Author Spotlight: Advances in Brain Energy Metabolism Research Using the Drosophila Model

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated. Thus, simultaneous recordings of neuronal activity and behavior are essential to correlate brain activity to behavior. For such behavioral analyses, the fruit fly, Drosophila melanogaster, allows us to incorporate genetically encoded calcium indicators such as GCaMP1, to monitor neuronal activity, and to use sophisticated genetic manipulations for optogenetic or thermogenetic techniques to specifically activate identified neurons2-5. Use of a thermogenetic technique has led us to find critical neurons for feeding behavior (Flood et al., under revision). As a main part of feeding behavior, a Drosophila adult extends its proboscis for feeding6 (proboscis extension response; PER), responding to a sweet stimulus from sensory cells on its proboscis or tarsi. Combining the protocol for PER7 with a calcium imaging technique8 using GCaMP3.01, 9, I have established an experimental system, where we can monitor activity of neurons in the feeding center &#x2013; the suboesophageal ganglion (SOG), simultaneously with behavioral observation of the proboscis. I have designed an apparatus ("Fly brain Live Imaging and Electrophysiology Stage": "FLIES") to accommodate a Drosophila adult, allowing its proboscis to freely move while its brain is exposed to the bath for Ca2+ imaging through a water immersion lens. The FLIES is also appropriate for many types of live experiments on fly brains such as electrophysiological recording or time lapse imaging of synaptic morphology. Because the results from live imaging can be directly correlated with the simultaneous PER behavior, this methodology can provide an excellent experimental system to study information processing of neuronal networks, and how this cellular activity is coupled to plastic processes and memory.

Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

The fruit fly, Drosophila melanogaster, extends its proboscis for feeding, responding to a sugar stimulus from its proboscis or tarsus. I have combined observations of the proboscis extension response (PER) with a calcium imaging technique, allowing us to monitor the activity of neurons in the brain, simultaneously with behavioral observation.

To study neuronal networks in terms of their function in behavior, we must analyze how neurons operate when each behavioral pattern is generated.  Thus, ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model animal for endocrine research, Drosophila melanogaster, which has sophisticated genetic tools and genome information, is being increasingly used. This article describes a method for the calcium imaging of Drosophila brain explants. This method enables the detection of the direct signaling of a hormone to the brain. It is well known that many peptide hormones act through G-protein-coupled receptors (GPCRs), whose activation causes an increase in the intracellular Ca2+concentration. Neural activation also elevates intracellular Ca2+ levels, from both Ca2+ influx and the release of Ca2+ stored in the endoplasmic reticulum (ER). A calcium sensor, GCaMP, can monitor these Ca2+ changes. In this method, GCaMP is expressed in the neurons of interest, and the GCaMP-expressing larval brain is dissected and cultured ex vivo. The test peptide is then applied to the brain explant, and the fluorescent changes in GCaMP are detected using a spinning disc confocal microscope equipped with a CCD camera. Using this method, any water-soluble molecule can be tested, and various cellular events associated with neural activation can be imaged using the appropriate fluorescent indicators. Moreover, by modifying the imaging chamber, this method can be used to image other Drosophila organs or the organs of other animals.

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

This paper describes a protocol for ex vivo calcium imaging of the Drosophila brain. In this method, natural or synthetic compounds can be applied to the buffer to test their ability to activate particular neurons in the brain.

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model ...

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.

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.

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

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. ...

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

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

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

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

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

Monitoring Epileptiform Events in Drosophila via Calcium Imaging

Ex Vivo Calcium Imaging for Drosophila Model of Epilepsy

Epilepsy is a neurological disorder characterized by recurrent seizures, partially correlated with genetic origin, affecting over 70 million individuals worldwide. Despite the clinical importance of epilepsy, the functional analysis of neural activity in the central nervous system is still to be developed. Recent advancements in imaging technology, in combination with stable expression of genetically encoded calcium indicators, such as GCaMP6, have revolutionized the study of epilepsy at both brain-wide and single-cell resolution levels. Drosophila melanogaster has emerged as a tool for investigating the molecular and cellular mechanisms underlying epilepsy due to its sophisticated molecular genetics and behavioral assays. In this study, we present a novel and efficient protocol for ex vivo calcium imaging in GCaMP6-expressing adult Drosophila to monitor epileptiform activities. The whole brain is prepared from cac, a well-known epilepsy gene, knockdown flies for calcium imaging with a confocal microscope to identify the neural activity as a follow-up to the bang-sensitive seizure-like behavior assay. The cac knockdown flies showed a higher rate of seizure-like behavior and abnormal calcium activities, including more large spikes and fewer small spikes than wild-type flies. The calcium activities were correlated to seizure-like behavior. This methodology serves as an efficient methodology in screening the pathogenic genes for epilepsy and exploring the potential mechanism of epilepsy at the cellular level.

Author Spotlight: Insights into the Techniques and Findings of Recent Advancements in Epilepsy Research 

Here, we present a protocol for ex vivo calcium imaging in GCaMP6-expressing adult Drosophila to monitor epileptiform activities. The protocol provides a valuable tool for investigating ictal events in adult Drosophila through ex vivo calcium imaging, allowing for exploration of the potential mechanisms of epilepsy at the cellular levels.

Epilepsy is a neurological disorder characterized by recurrent seizures, partially correlated with genetic origin, affecting over 70 million individuals ...

Multiplex Detection of Gene Expression in the Intact Drosophila Brain Using Expansion-Assisted Iterative Fluorescence In Situ Hybridization

Understanding gene expression is essential for deciphering cellular functions. However, methods for analyzing the expression of numerous genes in situ within a given tissue remain limited. The EASI-FISH protocol described here has been adapted to detect the expression of dozens of genes in the intact adult Drosophila central nervous system (CNS) using commercially available reagents. This protocol includes a new gel formulation that enhances gel robustness, enabling multiple rounds of hybridization and allowing the embedding of multiple brains per gel. This improvement increases throughput, facilitates optimal comparison of experimental conditions, and reduces reagent costs. Additionally, by employing the GAL4-UAS system for co-detection of green fluorescent protein (GFP), gene expression can be visualized in specific neuronal or glial cell types. Notably, the high resolution achieved through expansion microscopy, combined with the sensitivity of the method, allows for the detection of single RNA transcripts. This approach effectively integrates high image quality with high throughput, making it a powerful tool for studying gene expression throughout the intact fly brain.

Expansion-Assisted Iterative Fluorescence In Situ Hybridization (EASI-FISH) is a robust technique that combines expansion microscopy with fluorescence in situ hybridization (FISH) to detect the expression of multiple genes in thick tissue. This protocol outlines recent advancements in the method and its adaptation for labeling the intact Drosophila central nervous system..

Understanding gene expression is essential for deciphering cellular functions. However, methods for analyzing the expression of numerous genes in situ ...

Ex Vivo Calcium Imaging to Study Drosophila Brain Responses to Neuropeptides

Source:&#160;<a style='text-decoration:none;' href='https://app.jove.com/v/57701/ex-vivo-calcium-imaging-for-visualizing-brain-responses-to-endocrine-signaling-in-drosophila'><span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:12pt;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'>Ishimoto, H.&#160;et. al.,<span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:12pt;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'> Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila.&#160;J. Vis. Exp.<span style='-webkit-text-decoration-skip:none;font-family:Arial,sans-serif;font-size:12pt;font-style:normal;font-variant:normal;font-weight:400;text-decoration-skip-ink:none;vertical-align:baseline;white-space:pre-wrap;'> (2018)</a> &#160;This video demonstrates calcium imaging to study&#160;Drosophila brain responses to neuropeptides. Neurons expressing a fluorescent calcium indicator reveal intracellular calcium dynamics and functional responses to signaling molecules.

Source:&#160;Ishimoto, H.&#160;et. al., Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila.&#160;J. Vis. Exp. ...

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Neuroimaging

Live Imaging & In Vivo Tracking

In Vivo Calcium Imaging Using Synaptically Localized Calcium Sensors in Drosophila melanogaster

Source: Hancock, C. E. et al., <a href="https://app.jove.com/v/60288">In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster.</a>&#160;J. Vis. Exp. (2019).
This video demonstrates a method for in vivo calcium imaging using synaptically localized calcium sensors in flies. It outlines the steps for preparing the flies for microscopy, conducting olfactory conditioning, and visualizing odor-induced calcium influx.

In vivo (in vivo) Obrazowanie wapnia za pomocą synaptycznie zlokalizowanych czujników wapnia u Drosophila melanogaster

1. Transgenic fruit flies,&#160;Drosophila melanogaster

	Cross female virgin and male flies (raised at 25 &#176;C in 60% relative humidity on a 12 h ...

Visualizing Monocarboxylates and Other Relevant Metabolites in the Ex Vivo Drosophila Larval Brain Using Genetically Encoded Sensors

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Simultaneous Recording of Calcium Signals from Identified Neurons and Feeding Behavior of Drosophila melanogaster

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

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

Ex Vivo Calcium Imaging for Drosophila Model of Epilepsy

Multiplex Detection of Gene Expression in the Intact Drosophila Brain Using Expansion-Assisted Iterative Fluorescence In Situ Hybridization

Ex Vivo Calcium Imaging to Study Drosophila Brain Responses to Neuropeptides

In vivo (in vivo) Obrazowanie wapnia za pomocą synaptycznie zlokalizowanych czujników wapnia u Drosophila melanogaster