We thank Michael Rosbash for his mentorship and support during the initial phase of the development of this method. This work was funded by the JST PRESTO program, the Swiss National Science Foundation (31003A_149893 and 31003A_169548), the European Research Council (ERC-StG-311194), Novartis Foundation for Medical-Biomedical Research (13A39) and the University of Geneva.

1. Preparation of Stock Solutions Under a Culture Hood
<ol>
	<li>Prepare 400 mL of 1x Schneider Active Medium (SAM) optimized for ex vivo culture of larval brains (modified from reference24,25) (Table 1). Aliquot to 5 mL and flash freeze in liquid nitrogen (LN2), store at - 80 &#176;C.</li>
	<li>Prepare 1x Dissecting Saline Solution (DSS) by diluting 10x stock solution (modified from reference26) (Table 2).</li>
	<li>Aliquot fibrinogen to 10 mg and store at 4 &#176;C for a single use. 
	Caution: Weigh fibrinogen under laminar flow, wearing gloves, as it is harmful.</li>
	<li>Prepare thrombin stock solution in SAM (1500 NIH unit/mg) and aliquot to 10 &#181;L, flash freeze in LN2 and keep at -80 &#176;C. 
	Caution: Wear gloves when handling thrombin as it is harmful.</li>
	<li>Aliquot stock solution of 100x Penicillin-Streptomycin. Keep at -20 &#176;C.</li>
	<li>Cut circles of polytetrafluoroethylene (PTFE) membrane (see Materials Table) to the size that fits the inside of a glass bottom dish. Rinse them in 70% EtOH and then in autoclaved dH2O. Sterilize and dry under laminar flow with UV. Keep in a sterile Petri dish. Single use. 
	NOTE: PTFE is a porous flexible membrane that allows gas exchange and prevents medium from evaporating during long-term recording.</li>
</ol>
2. Time-lapse Microscopy Setup
NOTE: The following setup is for a tandem scanner inverted confocal microscope (see Materials Table) equipped with a resonant scanner (8,000 Hz). The system includes a highly sensitive photo-multiplier detector, which together with the resonant scanner prevents phototoxicity and photobleaching. Temperature and humidity are controlled within a microscope environmental chamber (see Materials Table) that covers most of the microscope.
<ol>
	<li>Place a stage-top humidity chamber.</li>
	<li>Turn on the temperature and humidity controllers to equilibrate the microscope chamber to 25 &#176;C and 80% humidity.</li>
	<li>To perform experiments in constant darkness (DD), cover the microscope environmental chamber with an opaque cloth (unless the microscope chamber is made of an opaque material).</li>
	<li>If using a water-immersion objective, position a Water Immersion Micro Dispenser on top of the objective and program a protocol with the Autoimmersion Objective Controller software to dispense water at a constant rate during time-lapse imaging. 
	NOTE: Water-immersion objectives together with a water dispenser or dry objective lens are recommended for long-term live imaging, as other type of immersion medium would dry out.</li>
	<li>Place a Petri dish holder on the stage, inside of the humidity chamber.</li>
	<li>Launch the microscope software and select the resonant scanner mode from the first dialogue box. Select OK to initialize the stage when prompted. Go to Configuration tab and to Hardware configuration to switch on the relevant laser lines.</li>
	<li>Go to Acquire tab and to XY panel to select bidirectional mode and set image acquisition parameters. 
	NOTE: The described imaging setup presented here (Figure 2, Figure 3 and Movie 1) is optimized to observe circadian expression of specific fluorescent reporters. The following parameters were chosen to best fit our specifications: 40X water immersion objective, 8X line average with 512 &#215; 512 image resolution. For multicolor imaging, sequential image acquisition is required when the emission wavelength of one fluorophore and the excitation wavelength of another overlap (here, wavelengths of Yellow Fluorescent Protein (YFP) emission and Tomato excitation overlap). Choose the sequential mode &#34;in between stack&#34; as it is the fastest and the movement of clock neurons is negligible during the acquisition of a Z-stack. Microscopy settings should be adapted to suit the aim of each experiment.</li>
</ol>
3. Setup of Larval Brain Explant Culture
NOTE: The whole procedure from dissection to embedding of brain explants takes ~30 min.
<ol>
	<li>Preparation of reagents under a culture hood.
	<ol>
		<li>Thaw an aliquot of thrombin and keep at room temperature until the embedding step (step 3.3). Single use.</li>
		<li>Thaw quickly an aliquot of SAM at 37 &#176;C and keep on ice for a single use.</li>
		<li>Add SAM to a 10 mg aliquot of fibrinogen to 10 mg/mL, flick gently and incubate at 37 &#176;C for at least 30 min and during the embedding step (step 3.3). Single use. 
		NOTE: Do not incubate the fibrinogen solution more than 2 h as it starts to polymerize.</li>
		<li>Thaw an aliquot of 100x stock Penicillin-Streptomycin. Prepare 2.5 mL of SAM containing 1x Penicillin-Streptomycin (SAM/antibiotics). Keep at room temperature. 
		NOTE: Store the remaining 100x stock Penicillin-Streptomycin at 4 &#176;C up to 1 month.</li>
		<li>Prepare an aliquot of 500 &#181;L from the remaining SAM. Keep on ice.</li>
		<li>For long-term imaging, apply autoclaved vacuum grease to the plastic part of the bottom of the glass bottom dish (Figure 1A) and sterilize under laminar flow with UV.</li>
	</ol>
	</li>
	<li>Dissection of the central nervous system and ventral nerve cord.
	<ol>
		<li>Clean the forceps and 2 x three-well glass dissection dishes with 70% EtOH. Pour 400 &#181;L of DSS into 4 wells and 400 &#181;L of SAM in one well. Keep on ice.</li>
		<li>Scoop out fly medium containing larvae and pick non-wandering 3rd instar larvae (L3) with forceps. Rinse them successively in 2 DSS-filled wells. Keep them in the 3rd DSS-filled well on ice that anesthetizes the larvae. 
		NOTE: L3 lasts approximately 2 days at 25 &#176;C. To observe the brain in physiologically relevant timeframe we chose to use non-wandering L3 larvae. Other subgroups of clock neurons, such as l-LNvs and DN3s, start to be formed at wandering L3 stage but they are not necessarily cycling yet (data not shown). Therefore, although it is possible to image cycling clock neurons at this stage (data not shown), it is important to carefully distinguish the already functional larval clock neurons from developing ones for data analysis. Non-wandering L3 appear approximately 4 to 4.5 days after egg laying at 25 &#176;C. To use this protocol for studying circadian rhythms, synchronize (entrain) the developing larvae in 12 h/12 h LD cycles at 25 &#176;C for 3 days before collecting L3 larvae.</li>
		<li>Dissect larval brains with fine forceps in the 4th DSS-filled well, which has not been used for rinsing larvae, using the inside-out method27.
		<ol>
			<li>Briefly, cut the larva in two, gently pinch the mouth hooks with a pair of forceps and roll up inside-out the body wall using the other pair of forceps, thus exposing the brain. Free the brain by cutting out all the trachea and nerves that attach it to the rest of the body.</li>
			<li>Aspirate the brain in about 3 &#181;L of DSS with a 20 &#181;L pipette (pre-rinsed with DSS) and dispense in the SAM-filled well. Repeat the procedure for up to 7 brains. 
			NOTE: Dissection should be performed on a cold surface to keep the larvae anesthetized and the brain chilled. For example, place the dissection dish on a cooling block connected to a pump to circulate ice-chilled water. The dissection procedure takes ~30 s per brain. It is important to keep the eye/antennal imaginal discs attached to the brains and the ventral nerve cord intact. Tearing off these parts will damage the brain and reduce the quality of the culture. Also, avoid exposing the brains to the air. Before aspirating the first brain, pipette up and down the DSS containing parts of the dissected larvae, as it prevents the brains from sticking to the wall of the tip.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Embedding of brain explants in a 3D matrix and mounting (~20 min).
	<ol>
		<li>Submerge a dissected brain in fibrinogen working solution (fibrinogen final concentration ~3.3 mg/mL, 10.5 &#181;L per brain) as follows:</li>
		<li>Aspirate 3.5 &#181;L of SAM/antibiotics (with the same tip used in section 3.2.3). Aspirate a dissected brain from the dissection well into the same pipette tip without dispensing the SAM/antibiotics medium in the tip.</li>
		<li>Dispense the brain and the medium on the lid of a sterile Petri dish. Add another 3.5 &#181;L of SAM/antibiotics and 3.5 &#181;L of warm fibrinogen/SAM 10 mg/mL solution on the drop containing the brain. Mix the solution around the brain gently by pipetting with a 20 &#181;L pipette tip. 
		NOTE: Using a pipette rather than forceps in this step avoids stressing the brain.</li>
		<li>Take 2 &#181;L of the fibrinogen working solution from the drop and apply it on the coverslip of the glass bottom dish as a small circle. Add 0.8 &#181;L of thrombin and mix very quickly with a pipette tip. The fibrinogen is converted to fibrin and starts to polymerize.</li>
		<li>Take the brain that sits in the fibrinogen working solution (step 3.3.1) between a pair of forceps and position it on the matrix once a white matrix of fibrin is visible. Orient the brain anterior side down (for imaging clock neurons). Push it down as close as possible to the cover glass. Fibrin clot is composed of layers of fibrin. Pick one layer and fold it on top of the brain with small and gentle movements without detaching the matrix.
		<ol>
			<li>Repeat 2 to 3 times from different parts of the clot to stabilize the brain. 
			NOTE: When handling the brain with forceps, be careful not to dry it or impose it a physical stress.</li>
		</ol>
		</li>
		<li>Add 20 &#181;L of SAM/antibiotics on top of the embedded brain to stop the action of thrombin.</li>
		<li>Repeat the procedure to mount the remaining brains. It is possible to embed up to 5 to 6 brains on a glass bottom dish.</li>
		<li>For long-term live imaging, add 600 &#181;L of SAM/antibiotics in the inner well (the glass part). Position a pre-cut PTFE membrane on top of the medium and stick it to the vacuum grease applied on the plastic part of the bottom (3.1.5) (Figure 1A).</li>
		<li>For pharmacological assays, fill the dish with 2 mL of SAM/antibiotics (Figure 1B). 
		NOTE: It is very important to avoid evaporation of the medium, as the osmolality of the medium is critical to the viability of neurons. Therefore, for long-term imaging experiments, it is necessary to seal the culture dish with PTFE membrane. Whereas for short-term experiments, sealing membrane is not required as long as the dish is filled with 2 mL of medium and monitored in a humidified chamber.</li>
	</ol>
	</li>
</ol>
4. Time-lapse Image Acquisition
<ol>
	<li>Place the glass bottom dish on the Petri dish holder inside the stage-top humidity chamber.</li>
	<li>Go to z-stack panel in the Acquire tab and set the z-section steps from the microscope software (2 &#181;m &#215; ~40 in the experiments shown in Figure 2 and Figure 3, and Movie 1). Set the beginning of a z-stack at the plane farthest from the coverslip and the end at the surface of the brain explant, close to the coverslip. Though brain movements are negligible, add extra z-sections at the beginning and end of the z-stack.</li>
	<li>Go to Mark &#38; Find panel and set-up multi-position image acquisition. With a 40X objective, 1 brain hemisphere can be captured within a field of view</li>
	<li>Go to Acquisition mode panel and select xyzt (z-stack time-lapse). Go to Time Acquisition panel and set the preferred time interval and frequency parameters. 
	NOTE: Acquire time-lapse images with desired frequency. Note that long-term live imaging (24-72 h) with time lapse interval shorter than 2 h tends to results in fewer cycling neurons, probably because it reduces the viability of neurons. The data volume expands as the rate of image acquisition increases, which makes data analysis and storage more challenging.</li>
</ol>
5. Pharmacological Treatment of brain Explants with Short-term Live Imaging
NOTE: The following procedure is essentially the same as that for long-term imaging but does not require a PTFE membrane. To avoid exposing the brains to blue light when the chemical is added to the culture, the microscope should be placed in the dark room with red light.
<ol>
	<li>Go to Time Acquisition panel and set the desired time interval and number of scan to obtain the baseline data before drug application. Start Scan.</li>
	<li>After the baseline scan is completed, lift the scan head of the microscope system. Remove the lid of the stage-top humidity controller and very carefully remove the lid of the glass bottom dish without moving it.</li>
	<li>Remove the appropriate amount of culture medium if necessary. Add the experimental solution in the medium and, carefully and very gently, mix with a sterile Pasteur pipette without touching the brain explants. 
	NOTE: For bath application of PDF, use 2 &#181;M PDF stock solution (in 10% DMSO).</li>
	<li>Replace the lids of the glass bottom dish and the humid chamber, and the scan head. If using, reposition the black opaque cloth around the microscope chamber.</li>
	<li>Check if the brains are at their original positions in live scan mode. Adjust if necessary.</li>
	<li>Go to Time Acquisition panel and set the desired time interval and number of scan. Start Scan. 
	NOTE: Here we tested time-lapse images acquired every hour for up to 10 h.</li>
</ol>

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytochalasin+D+and+latrunculin+affect+chromosome+behaviour+during+meiosis+in+crane-fly+spermatocytes.">Cytochalasin D and latrunculin affect chromosome behaviour during meiosis in crane-fly spermatocytes.</a> Chromosome Res. 6 (7), 533-549 (1998).</li><li>Lukinavi&#269;ius, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SiR-Hoechst+is+a+far-red+DNA+stain+for+live-cell+nanoscopy.">SiR-Hoechst is a far-red DNA stain for live-cell nanoscopy.</a> Nature Communications. 6, 8497 (2015).</li><li>Medioni, C., Ephrussi, A., Besse, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+axonal+transport+in+Drosophila+pupal+brain+explants.">Live imaging of axonal transport in Drosophila pupal brain explants.</a> Nat Protoc. 10 (4), 574-584 (2015).</li><li>Enoki, R., Ono, D., Hasan, M. T., Honma, S., Honma, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+resolution+fluorescence+imaging+of+circadian+rhythms+detected+with+a+Nipkow+spinning+disk+confocal+system.">Single-cell resolution fluorescence imaging of circadian rhythms detected with a Nipkow spinning disk confocal system.</a> J Neurosci Methods. 207 (1), 72-79 (2012).</li><li>Ozel, M. N., Langen, M., Hassan, B. A., Hiesinger, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Filopodial+dynamics+and+growth+cone+stabilization+in+Drosophila+visual+circuit+development.">Filopodial dynamics and growth cone stabilization in Drosophila visual circuit development.</a> Elife. 4, (2015).</li><li>Yao, Z., Macara, A. M., Lelito, K. R., Minosyan, T. Y., Shafer, O. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+functional+neuronal+connectivity+in+the+Drosophila+brain.">Analysis of functional neuronal connectivity in the Drosophila brain.</a> J Neurophysiol. 108 (2), 684-696 (2012).</li></ol>

The authors have nothing to disclose.

Here we described the method for long-term fluorescence time-lapse microscopy of cultured larval brains. The success of this type of experiments depends on multiple factors, such as the health of the culture, method for immobilization of the brain explant, fluorescence intensity and signal-to-noise ratio of the reporter, temporal and spatial resolution, and accessibility to the explant. These factors can be mutually exclusive. For example, increasing time-lapse frequency affects the health of the culture, and the immobil...

Circadian clocks help organisms to adapt to the periodic environmental changes generated by the 24 h rotation of the Earth. The interlocked transcriptional-translational feedback loops commonly underlie the molecular machinery of circadian clocks across species1. The circadian pacemaker circuit composed of clock-containing neurons integrates time-of-day information conveyed by the environmental cues, such as light/dark (LD) and temperature cycles, to orchestrate the rhythms of a plethora of daily physiological and behavioral processes2,3. The coordination of molecular rhythms with neuronal inputs and outputs is critically important for the operation of the circadian circuit but remains only partially understood.
In Drosophila, at the core of the molecular clock, the CLOCK/CYCLE (CLK/CYC) heterodimer activates the transcription of period (per) and timeless (tim). PER and TIM form a complex and enter the nucleus, where they inhibit transcriptional activity of CLK/CYC and consequently their own transcription. Post-transcriptional and post-translational regulations cause delays between CLK/CYC-mediated transcription and repression by PER/TIM, ensuring the generation of circa 24-h molecular oscillations1,3,4. About 150 neurons containing these molecular clocks form a circuit to control circadian behavior of adult flies5. A much simpler yet fully functional circadian circuit consisting of 3 groups of clock neurons - 5 ventral Lateral Neurons (LNvs; 4 PDF-positive LNvs and one PDF-negative LNv, see below), 2 Dorsal Neuron 1s (DN1s) and 2 Dorsal Neuron 2s (DN2s) - is present in the larval brain6,7.
The simple larval circadian circuit offers an excellent model to study the interactions between inter-neuronal communication and the molecular clockwork. Using our newly developed fluorescent reporter PER-TDT, which mimics the levels of PER protein and its subcellular location, we sought to characterize the dynamics of the molecular clockwork in different clock neuron subgroups in the larval circadian circuit8. Furthermore, knowing the key role of the neuropeptide pigment-dispersing factor (PDF) produced by 4 LNvs in regulating circadian rhythms at the neuronal level9,10,11, we wanted to examine the direct effect of PDF on the molecular clocks. To this end, we developed a method to monitor circadian gene expression rhythms in larval brain explant over multiple days by time-lapse confocal microscopy. The protocol was also adapted for pharmacological assays to test the effect of PDF or other compounds on the level of PER-TDT. Thus, the adaptation consists of making the brain explant culture accessible to drug application, increasing temporal resolution and imaging for a shorter duration.
Ex vivo culture of Drosophila brains of different developmental stages have been previously established12,13,14,15,16,17,18. Whereas these protocols have been used for imaging various biological phenomena, some of them are not compatible with imaging at single-cell resolution or do not support the culture for more than several hours. Alternative methods to perform long-term live imaging of circadian neurons in Drosophila include bioluminescence imaging of molecular rhythms19,20,21 and fluorescence imaging of calcium indicator with light sheet microscopy22,23. Although bioluminescence imaging can achieve higher temporal resolution and light sheet microscopy can be adaptable for in vivo imaging, they are limited on the spatial resolution and require specialized microscope systems.
The method described here is tailored to visualize fluorescent signals in the whole brain culture at single-cell resolution over multiple days. This facile and versatile method could be adapted to image cultured adult fly brains and for pharmacological experiments to study many different problems in Drosophila neurobiology.

<table><tbody><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>P5655</td><td>I am not sure they are exactly the same ones we have in the lab. I chose &quot;suitable for insect cell culture&quot; whenever available</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>C7902</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;.7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>230391</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S5886</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S5761</td><td></td></tr><tr><td>D-(+) Glucose</td><td>Sigma-Aldrich</td><td>G7021</td><td></td></tr><tr><td>Yeast extract</td><td>Sigma-Aldrich</td><td>Y1000</td><td></td></tr><tr><td>Insulin</td><td>Sigma-Aldrich</td><td>I0516-5ML</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Sigma-Aldrich</td><td>P4333</td><td></td></tr><tr><td>BIS-TRIS</td><td>Sigma-Aldrich</td><td>B4429</td><td></td></tr><tr><td>L-(&amp;minus;)-Malic acid</td><td>Sigma-Aldrich</td><td>M7397</td><td></td></tr><tr><td>D-(+)-Trehalose dihydrate</td><td>Sigma-Aldrich</td><td>T0167</td><td></td></tr><tr><td>Succinic acid</td><td>Sigma-Aldrich</td><td>S9512</td><td></td></tr><tr><td>Fumaric acid</td><td>Sigma-Aldrich</td><td>F8509</td><td></td></tr><tr><td>&amp;alpha;-Ketoglutaric acid</td><td>Sigma-Aldrich</td><td>K1128</td><td></td></tr><tr><td>Non-heat-inactivated, Foetal Calf Serum (FCS) Mycoplasma and Virus screened</td><td>BioConcept Ltd. Amimed</td><td>2-01F30-I</td><td></td></tr><tr><td>HEPES-KOH, pH 7.4</td><td>E&amp;amp;K Scientific Products</td><td>EK-654011</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P5405</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S5011</td><td></td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>S7903</td><td></td></tr><tr><td>Fibrinogen from bovine plasma</td><td>Calbiochem (Merck)</td><td>341573-1GM</td><td>CAUTION: Harmful by inhalation, in contact with skin and if swallowed. Manipulate under laminar flow</td></tr><tr><td>Thrombin from bovine plasma</td><td>Sigma-Aldrich</td><td>T9549</td><td>CAUTION: Health Hazard, use gloves</td></tr><tr><td>PDF, NH&lt;sub&gt;2&lt;/sub&gt;-NSELINSLLSLPKNMNDA-OH</td><td>Chi Scientific</td><td>custom made</td><td></td></tr><tr><td>Vaccum grease</td><td>Sigma-Aldrich</td><td>18405</td><td></td></tr><tr><td>35 mm Dish, No. 1.5 Coverslip, 20 mm Glass Diameter, Uncoated</td><td>MatTek</td><td>P35G-1.5-20-C</td><td></td></tr><tr><td>Corning Falcon Easy-Grip Tissue Culture Dishes</td><td>fisherscientific</td><td>08-772A</td><td></td></tr><tr><td>Sterile 500 mL Steritop-GP 33 mm threaded bottle top filter, 0.22 &amp;mu;m</td><td>Millipore</td><td>SCGPS05RE</td><td></td></tr><tr><td>Polytetrafluoroethylene (PTFE) film</td><td>Dupont</td><td>200A Teflon FEP Film</td><td></td></tr><tr><td>Millex-HV Syringe Filter Unit, 0.45&amp;nbsp;&amp;micro;m, PVDF, 33&amp;nbsp;mm, gamma sterilized</td><td>Millipore</td><td>SLHV033RS</td><td></td></tr><tr><td>Millex-GV Syringe Filter Unit, 0.22&amp;nbsp;&amp;micro;m, PVDF, 33&amp;nbsp;mm, gamma sterilized</td><td>Millipore</td><td>SLGV033RS</td><td></td></tr><tr><td>Three-well glass dissection dish</td><td>Any company</td><td></td><td></td></tr><tr><td>Fine forceps, size 5, Dumont</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>Tandem scanner inverted TCS SP5 confocal microscope, with resonant scanner and HyD photo-multiplier detectors</td><td>Leica microsystem CMS GmbH</td><td></td><td></td></tr><tr><td>Temperature control chamber</td><td>Life Imaging Services</td><td>The CUBE &amp;amp; BOX temperature control system, custom designed</td><td></td></tr><tr><td>Stage-top humidity controller</td><td>Life Imaging Services</td><td>custom made</td><td></td></tr><tr><td>Water Immersion Micro Dispenser: dispenser, extended micro-pump MP6 series and Autoimmersion Objective Controller software</td><td>Leica microsystem CMS GmbH</td><td></td><td></td></tr><tr><td>SUM-stack creation and 3D correction drift plugin</td><td>ImageJ software</td><td></td><td></td></tr><tr><td>10x iterative deconvolution</td><td>AutoQuant and Imaris software</td><td></td><td></td></tr></tbody></table>

single-cell resolution fluorescence live imaging of drosophila circadian clocks in larval brain culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Here, we show the representative results of the long-term recording of a circadian fluorescent reporter in ex vivo larval brain culture, and the live imaging results of PDF bath application on reporter expression.
Non-wandering L3 larvae expressing the molecular clock reporter PER-TDT and UAS-mCD8::YFP driven by a clock neuron driver Clk(856)-gal4 (Figure 1C) were entrained in LD. ...

Watch this Scientific Journal Video about Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture at JoVE.com

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

single-cell-resolution-fluorescence-live-imaging-drosophila-circadian

Research

JoVE Journal

Neuroscience

7.5K Views.  University of Geneva. Het doel van dit protocol is om de ex vivo Drosophila larvale hersenen cultuur, geoptimaliseerd voor het controleren van de circadiane moleculaire ritmes met langdurige fluorescentie time-lapse beeldvorming. De toepassing van deze methode op farmacologische testen wordt ook besproken.

Video: Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Developmental Biology

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 ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

Biology

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Imaging Live Brain Explants from Drosophila melanogaster Third Instar Larvae

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a differentiating ganglion mother cell (GMC), which will undergo one additional division to give rise to two neurons or glia. Studies in NBs have uncovered the molecular mechanisms underlying cell polarity, spindle orientation, neural stem cell self-renewal, and differentiation. These asymmetric cell divisions are readily observable via live-cell imaging, making larval NBs ideally suited for investigating the spatiotemporal dynamics of asymmetric cell division in living tissue. When properly dissected and imaged in nutrient-supplemented medium, NBs in explant brains robustly divide for 12-20 h. Previously described methods are technically difficult and may be challenging to those new to the field. Here, a protocol is described for the preparation, dissection, mounting, and imaging of live third-instar larval brain explants using fat body supplements. Potential problems are also discussed, and examples are provided for how this technique can be used.

Author Spotlight: Investigating Asymmetric Cell Division Dynamics: A Protocol for Live-Imaging of Drosophila Larval Brain Explants 

Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological conditions.

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a ...

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 ...

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding environment and initiate contacts with potential synaptic partners. Although the connection between dendritic branch dynamics and synaptogenesis is well established, how developmental and activity-dependent processes regulate dendritic branch dynamics is not well understood. This is partly due to the technical difficulties associated with the live imaging and quantitative analyses of these fine structures using an in vivo system. We established a method to study dendrite dynamics using Drosophila larval ventral lateral neurons (LNvs), which can be individually labeled using genetic approaches and are accessible for live imaging. Taking advantage of this system, we developed protocols to capture branch dynamics of the whole dendritic arbor of a single labeled LNv through time-lapse live imaging. We then performed post-processing to improve image quality through drift correction and deconvolution, followed by analyzing branch dynamics at the single-branch level by annotating spatial positions of all branch terminals. Lastly, we developed R scripts (Supplementary File) and specific parameters to quantify branch dynamics using the coordinate information generated by the terminal tracing. Collectively, this protocol allows us to achieve a detailed quantitative description of branch dynamics of the neuronal dendritic arbor with high temporal and spatial resolution. The methods we developed are generally applicable to sparsely labeled neurons in both in vitro and in vivo conditions.

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Here, we describe the method we employed to image highly motile dendritic filopodia in a live preparation of the Drosophila larval brain, and the protocol we developed to quantify time-lapse 3D imaging datasets for quantitative assessments of dendrite dynamics in developing neurons.

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Circadian Entrainment of Drosophila Melanogaster

Nearly universal among organisms, circadian rhythms coordinate biological activity to earth&#39;s orbit around the sun. To identify factors creating this rhythm and to understand the resulting outputs, entrainment of model organisms to defined circadian time-points is required. Here we detail a procedure to entrain many Drosophila to a defined circadian rhythm. Furthermore, we detail post-entrainment steps to prepare samples for immunofluorescence, nucleic acid, or protein extraction-based analysis.

Here, we detail how to synchronize Drosophila to a circadian day. This is the first, and most important step necessary for studying biological rhythms and chronobiology.

Nearly universal among organisms, circadian rhythms coordinate biological activity to earth&#39;s orbit around the sun. To identify factors creating this ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

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