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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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agarose</td>
			<td>Sigma</td>
			<td>A9539</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</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>MgCl2</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>CAUTION-Fatal if swallowed</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&#160;</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-FLII12Pglu700md6&#160;</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>93452</td>
			<td>Glucose sensor</td>
		</tr>
		<tr>
			<td>UAS-GCaMP6f&#160;</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

Research

JoVE Journal

Neuroscience

595 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

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

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 ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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 ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with enhanced [Ca2+]i signals. It has been proposed that astrocytes in turn regulate neurons and blood vessels through calcium-dependent mechanisms, such as the release of signaling molecules. However, [Ca2+]i imaging in entire astrocytes has only recently become feasible with genetically encoded calcium indicators (GECIs) such as the GCaMP series. The use of GECIs in astrocytes now provides opportunities to study astrocyte [Ca2+]i signals in detail within model microcircuits such as the striatum, which is the largest nucleus of the basal ganglia. In the present report, detailed surgical methods to express GECIs in astrocytes in vivo, and confocal imaging approaches to record [Ca2+]i signals in striatal astrocytes in situ, are described. We highlight precautions, necessary controls and tests to determine if GECI expression is selective for astrocytes and to evaluate signs of overt astrocyte reactivity. We also describe brain slice and imaging conditions in detail that permit reliable [Ca2+]i imaging in striatal astrocytes in situ. The use of these approaches revealed the entire territories of single striatal astrocytes and spontaneous [Ca2+]i signals within their somata, branches and branchlets. The further use and expansion of these approaches in the striatum will allow for the detailed study of astrocyte [Ca2+]i signals in the striatal microcircuitry.

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ.

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with ...

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.