Name: light spot-based assay for analysis of drosophila larval phototaxis
Uploaded: 2019-09-27T14:45:00
Description: Watch this Scientific Journal Video about Light Spot-Based Assay for Analysis of Drosophila Larval Phototaxis at JoVE.com

This work is supported by natural science foundation of China (31671074) and Fundamental Research Funds for the Zhejiang Provincial Universities (2019XZZX003-12).

1. Preparation of Drosophila larvae
<ol>
	<li>Prepare standard medium consisting of boiled corn meal (73 g), agar (5.6 g), soybean meal (10 g), yeast (17.3 g), syrup (76 mL) and water (1000 mL).</li>
	<li>Raise all flies at 25 &#176;C on standard medium in a room with a 12 h/12 h light/dark cycle.</li>
</ol>
2. Preparation of agar plates
<ol>
	<li>Prepare 1.0% agar solution. Weigh 3 g of agar in a 500 mL beaker with a balance, then add 300 mL of distilled water. Place a foil paper ove...

<ol>
	<li>Grossfield, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Geographic+distribution+and+light-dependent+behavior+in+Drosophila.">Geographic distribution and light-dependent behavior in Drosophila.</a> Proceedings of the National Academy of Sciences of the United States of America. 68, 2669 (1971).</li><li>Godoy-Herrera, R. C. L. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+behaviour+of+Drosophila+melanogaster+larvae+during+pupation.">The behaviour of Drosophila melanogaster larvae during pupation.</a> Animal Behaviour. 37, (1989).</li><li>Busto, M., Iyengar, B., Campos, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+dissection+of+behavior:+modulation+of+locomotion+by+light+in+the+Drosophila+melanogaster+larva+requires+genetically+distinct+visual+system+functions.">Genetic dissection of behavior: modulation of locomotion by light in the Drosophila melanogaster larva requires genetically distinct visual system functions.</a> Journal of Neuroscience. 19, 3337 (1999).</li><li>Mazzoni, E. O., Desplan, C., Blau, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circadian+pacemaker+neurons+transmit+and+modulate+visual+information+to+control+a+rapid+behavioral+response.">Circadian pacemaker neurons transmit and modulate visual information to control a rapid behavioral response.</a> Neuron. 45, 293 (2005).</li><li>Keene, A. C., 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=Distinct+visual+pathways+mediate+Drosophila+larval+light+avoidance+and+circadian+clock+entrainment.">Distinct visual pathways mediate Drosophila larval light avoidance and circadian clock entrainment.</a> Journal of Neuroscience. 31, 6527 (2011).</li><li>Keene, A. C., Sprecher, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Seeing+the+light:+photobehavior+in+fruit+fly+larvae.">Seeing the light: photobehavior in fruit fly larvae.</a> Trends in Neurosciences. 35, (2012).</li><li>Kane, E. A., 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=Sensorimotor+structure+of+Drosophila+larva+phototaxis.">Sensorimotor structure of Drosophila larva phototaxis.</a> Proceedings of the National Academy of Sciences of the United States of America. 110, E3868 (2013).</li><li>Humberg, T. H., 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=Dedicated+photoreceptor+pathways+in+Drosophila+larvae+mediate+navigation+by+processing+either+spatial+or+temporal+cues.">Dedicated photoreceptor pathways in Drosophila larvae mediate navigation by processing either spatial or temporal cues.</a> Nature Communications. 9. 1260, (2018).</li><li>Gong, Z., 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=Two+pairs+of+neurons+in+the+central+brain+control+Drosophila+innate+light+preference.">Two pairs of neurons in the central brain control Drosophila innate light preference.</a> Science. 330, (2010).</li><li>Yamanaka, N., 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=Neuroendocrine+Control+of+Drosophila+Larval+Light+Preference.">Neuroendocrine Control of Drosophila Larval Light Preference.</a> Science. 341, 1113 (2013).</li><li>Zhao, W., 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=A+disinhibitory+mechanism+biases+Drosophila+innate+light+preference.">A disinhibitory mechanism biases Drosophila innate light preference.</a> Nature Communications. 10, (2019).</li><li>Gong, C., 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=A+Neuronal+Pathway+that+Commands+Deceleration+in+Drosophila+Larval+Light-Avoidance.">A Neuronal Pathway that Commands Deceleration in Drosophila Larval Light-Avoidance.</a> Neuroscience Bulletin. Feb. 27, (2019).</li><li>Larderet, I., 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=Organization+of+the+Drosophila+larval+visual+circuit.">Organization of the Drosophila larval visual circuit.</a> eLife. 6, (2017).</li><li>Sawin-McCormack, E. P., Sokolowski, M. B., Campos, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+and+genetic+analysis+of+Drosophila+melanogaster+photobehavior+during+larval+development.">Characterization and genetic analysis of Drosophila melanogaster photobehavior during larval development.</a> Journal of Neurogenetics. 10, (1995).</li><li>Farca, L. A., von Essen, A. M., Widmer, Y. F., Sprecher, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light+preference+assay+to+study+innate+and+circadian+regulated+photobehavior+in+Drosophila+larvae.">Light preference assay to study innate and circadian regulated photobehavior in Drosophila larvae.</a> Journal of Visualized Experiments. 74 (74), e50237 (2013).</li><li>Xiang, Y., 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=Light-avoidance-mediating+photoreceptors+tile+the+Drosophila+larval+body+wall.">Light-avoidance-mediating photoreceptors tile the Drosophila larval body wall.</a> Nature. 468, 921 (2010).</li><li>Gomez-Marin, A., Partoune, N., Stephens, G. J., Louis, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+tracking+of+animal+posture+and+movement+during+exploration+and+sensory+orientation+behaviors.">Automated tracking of animal posture and movement during exploration and sensory orientation behaviors.</a> PLoS ONE. 7, e41642 (2012).</li></ol>

This protocol presents the light spot assay to test the ability of Drosophila larvae to escape from light. This assay allows tracking of the behavior of larvae before entering, during encountering, and after leaving a light spot. Details of larval movement can be captured and analyzed. The light spot assay is very simple and possesses strong practicability. The cost of the whole device is not high. In the experiment, LED light is used as the light source. It can be replaced with light sources of different wavele...

The larvae of Drosophila melanogaster show obvious light-avoiding behavior during the foraging stage. Drosophila larval phototaxis has been under investigation, especially in the past 50 years1,2,3,4,5,6,7,8. In recent years, despite the fact that 1) many neurons mediating larval light avoidance have been identified4,5,9,10,11,12 and 2) the complete connectome of larval visual system at the resolution of synapses has been established13, the neural mechanisms underlying larval phototaxis remain largely unclear.
A number of behavioral assays have been used in studying larval phototaxis. They can be largely divided into two classes: one involving spatial light gradients and the other involving temporal light gradients. For spatial light gradient assays, the arena is divided into equal number of sections in light and dark. The arena can be divided into light and dark halves2,4 or light and dark quadrants14,15, or can even be separated into alternate light and dark squares like on a checkerboard7. Usually, agar plates are used for spatial light gradient assay, but tubes that are divided into alternate light and dark sections can also be used10,14.
In older version of assays, light illumination generally originates from below the larvae. However, illumination in newer versions largely originates from above, since larval eyes (e.g., the Bolwig&#39;s organs that are sensitive to low or medium light intensities16) are contained in the opaque cephalopharyngeal skeleton with openings towards the upper front. This makes larvae more sensitive to light from upper front directions than from below behind directions7. For temporal light gradient assays, the light intensity is spatially uniform in the arena, but the intensity changes over time. In addition to temporal square wave light (i.e., flashing on/off or strong/weak light3,7), temporally varying light that conforms to a linear ramp in intensity is also used8 to measure the sensitivity of larvae to a temporally changing light stimulus.
A third type of phototaxis assay is the directional light scape navigation, which involves illumination from above at an angle of 45&#176;7. Before the work of Kane et al.7, only coarse parameters such as the number of larvae in light and dark regions, frequency of turning, and trail length were calculated in larval phototaxis assays. Since the work of this same group, with the analysis of high temporal resolution video record for larval phototaxis, detailed dynamics of larval movement during phototaxis (i.e., instant speeds of different parts of larval body, heading direction, turning angle and corresponding angular velocity) have been analyzed7. Thus, more details of larval phototaxis behavior have been able to be discovered. In these assays, larvae are tested in groups so that group effects are not excluded.
This protocol introduces a light-spot assay for the investigation of larval behavioral responses to individual light stimulation. The main experimental set-up consists of a visual stimulation system and infrared light-based imaging system. In the visual stimulation system, an LED light source generates a round 2 cm-diameter light spot on an agar plate, where the larva is tested. The light intensity can be adjusted using an LED driver. The imaging system includes an infrared camera that captures the behavior of the larva in addition to three 850 nm infrared LEDs that provide illumination for the camera. The lens of the camera is covered by an 850 nm band-pass filter to block light from the visual stimulation system from entering the camera, while the infrared light is allowed to enter the camera. Thus, interference of visual stimulation on imaging is prevented. In this assay, the behavioral details of the fast responses of individual larvae within a period including before, during, and after entering light are recorded and analyzed.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>850 nm &#177; 3 nm infrared-light-generating LED</td>
			<td>Thorlabs, USA</td>
			<td>PM100A</td>
			<td>Compatible Sensors: Photodiode and Thermal 
			Optical Power Rangea: 100 pW to 200 W 
			Available Sensor Wavelength Rangea: 185 nm-25 &#956;m Display Refresh Rate: 20 Hz 
			Bandwidtha: DC-100 kHz 
			Photodiode Sensor Rangeb: 50 nA-5 mA 
			Thermopile Sensor Rangeb: 1 mV-1 V</td>
		</tr>
		<tr>
			<td>AC to DC converter</td>
			<td>Thorlabs, USA</td>
			<td>S120VC</td>
			<td>Aperture Size: &#216;9.5 mm 
			Wavelength Range: 200-1100 nm 
			Power Range: 50 nW-50 mW 
			Detector Type: Si Photodiode (UV Extended) 
			Linearity: &#177;0.5% 
			Measurement Uncertaintyc: &#177;3% (440-980 nm), &#177;5% (280-439 nm), &#177;7% (200-279 nm, 981-1100 nm)</td>
		</tr>
		<tr>
			<td>band-pass filter</td>
			<td>Thorlabs, USA</td>
			<td>DC2100</td>
			<td>LED Current Range: 0-2 A 
			LED Current Resolution: 1 mA 
			LED Current Accuracy: &#177;20 mA 
			LED Forward Voltage: 24 V 
			Modulation Frequency Range: 0-100 kHz Sine Wave 
			Modulation: Arbitrary</td>
		</tr>
		<tr>
			<td>Collimated LED blue light&#160;</td>
			<td>ELP, China</td>
			<td>USBFHD01M</td>
			<td>Max. Resolution: 1920X1080 
			F6.0 mm 
			Sensor: 1/2.7&#34; CMOS OV2710</td>
		</tr>
		<tr>
			<td>Compact power meter console&#160;</td>
			<td>Ocean Optics, USA</td>
			<td>USB2000+(RAD)</td>
			<td>Dimensions: 89.1 mm x 63.3 mm x 34.4 mm 
			Weight: 190 g 
			Detector: Sony ILX511B (2048-element linear silicon CCD array) 
			Wavelength range: 200-850 nm 
			Integration time: 1 ms &#8211; 65 seconds (20 seconds typical) 
			Dynamic range: 8.5 x 10^7 (system); 1300:1 for a single acquisition 
			Signal-to-noise ratio: 250:1 (full signal) 
			Dark noise: 50 RMS counts 
			Grating: 2 (250 &#8211; 800 nm) 
			Slit: SLIT-50 
			Detector collection lens: L2 
			Order-sorting: OFLV-200-850 
			Optical resolution: ~2.0 nm FWHM 
			Stray light: &#60;0.05% at 600 nm; &#60;0.10% at 435 nm 
			Fiber optic connector: SMA 905 to 0.22 numerical aperture single-strand fiber</td>
		</tr>
		<tr>
			<td>High-Power LED Driver</td>
			<td>Minhongshi, China</td>
			<td>MHS-48XY</td>
			<td>Working voltage: DC12V 
			Central wavelength: 850nm</td>
		</tr>
		<tr>
			<td>high-resolution web camera</td>
			<td>Thorlabs, USA</td>
			<td>MWWHL4</td>
			<td>Color: Warm White 
			Correlated Color Temperature: 3000 K 
			Test Current for Typical LED Power: 1000 mA 
			Maximum Current (CW): 1000 mA 
			Bandwidth (FWHM): N/A 
			Electrical Power: 3000 mW 
			Viewing Angle (Full Angle): 120&#730; 
			Emitter Size: 1 mm x 1 mm 
			Typical Lifetime: &#62;50 000 h 
			Operating Temperature (Non-Condensing): 0 to 40 &#176;C 
			Storage Temperature: -40 to 70 &#176;C 
			Risk Groupa: RG1 &#8211; Low Risk Group</td>
		</tr>
		<tr>
			<td>LED Warm White</td>
			<td>Mega-9, China</td>
			<td>BP850/22K</td>
			<td>&#216;25.4(+0~-0.1) mm 
			Bandwidth: 22&#177;3nm 
			Peak transmittance:80% 
			Central wavelength: 850nm&#177;3nm&#160;</td>
		</tr>
		<tr>
			<td>Spectrometer&#160;</td>
			<td>Noel Danjou</td>
			<td>Amcap9.22</td>
			<td>AMCap&#160;is a still and video capture application with advanced preview and recording features. It is a Desktop application designed for computers running Windows 7 SP1 or later. Most&#160;Video-for-Windowsand&#160;DirectShow-compatible devices are supported whether they are cheap webcams or advanced video capture cards.</td>
		</tr>
		<tr>
			<td>Standard photodiode power sensor&#160;</td>
			<td>Super Dragon, China</td>
			<td>YGY-122000</td>
			<td>Input: AC 100-240V~50/60Hz 0.8A 
			Output: DC 12V 2A</td>
		</tr>
		<tr>
			<td>Thermal power sensor&#160;</td>
			<td>Thorlabs, USA</td>
			<td>M470L3-C1</td>
			<td>Color: Blue 
			Nominal Wavelengtha: 470 nm 
			Bandwidth (FWHM): 25 nm 
			Maximum Current (CW): 1000 mA 
			Forward Voltage: 3.2 V 
			Electrical Power (Max): 3200 mW 
			Emitter Size: 1 mm x 1 mm 
			Typical Lifetime: 100 000 h 
			Operating Temperature (Non-Condensing): 0 to 40 &#176;C 
			Storage Temperature: -40 to 70 &#176;C 
			Risk Groupb: RG2 &#8211; Moderate Risk Group</td>
		</tr>
		<tr>
			<td>Thermal power sensor&#160;</td>
			<td>Thorlabs, USA</td>
			<td>S401C</td>
			<td>Wavelength range: 190 nm-20 &#956;m 
			Optical power range:10 &#956;W-1 W(3 Wb) 
			Input aperture size: &#216;10 mm 
			Active detector area: 10 mm x 10 mm 
			Max optical power density: 500 W/cm2 (Avg.) 
			Linearity: &#177;0.5%</td>
		</tr>
	</tbody>
</table>

light spot-based assay for analysis of drosophila larval phototaxis

The larvae of Drosophila melanogaster show obvious light-avoiding behavior during the foraging stage. Drosophila larval phototaxis can be used as a model to study animal avoidance behavior. This protocol introduces a light-spot assay to investigate larval phototactic behavior. The experimental set-up includes two main parts: a visual stimulation system that generates the light spot, and an infrared light-based imaging system that records the process of larval light avoidance. This assay allows tracking of the behavior of larva before entering, during encountering, and after leaving the light spot. Details of larval movement including deceleration, pause, head casting, and turning can be captured and analyzed using this method.

According to the protocol, the light spot assay was used to investigate light avoidance behavior of third instar larva that were raised at 25 &#176;C on standard medium in a room with a 12 h/12 h light/dark cycle. A single w1118 larva was tested using the light spot assay at 25.5 &#176;C. The average light intensity of the light spot generated by a 460 nm LED was 0.59 &#181;W/cm2. The whole process of larval entering and exiting the light spot was recorded and analyzed using SOS software an...

Watch this Scientific Journal Video about Light Spot-Based Assay for Analysis of Drosophila Larval Phototaxis at JoVE.com

Light Spot-Based Assay for Analysis of Drosophila Larval Phototaxis

This protocol introduces a light-spot assay to investigate Drosophila larval phototactic behavior. In this assay, a light spot is generated as light stimulation, and the process of larval light avoidance is recorded by an infrared light-based imaging system.

Light Spot-Based Assay for Analysis of Drosophila Larval Phototaxis

The larvae of Drosophila melanogaster show obvious light-avoiding behavior during the foraging stage. Drosophila larval phototaxis can be used as a model ...

Using the light spot assay, the behavior for larvae can be tracked before, during, and after entering the light area allowing the detail of the larval movement to be analyzed. This assay is simple and easy to perform. The light response is tested only one time, so that any possible effect of light adaptation can be excluded.To prepare the agar plates, slowly and evenly cover the bottom of a 15 centimeter Petri dish with an approximately four millimeter thick layer of hot 1%agar. Allow the plate to cool at room temperature until the agar has solidified. To set up the visual stimulation system, clip a prepared LED light source onto an iron frame, taking care that the light projects down toward the desktop.Tilt the cylinder slightly until the angle between the cylinder plane and the vertical plane is about 10 degrees. Connect the 470 nanometer blue LED light to the LED one plug of the high power LED driver and turn on the driver. Turn the knob in the upper right corner of the driver to select channel 470 nanometers and click LED.When the screen displays a check, a blue light spot will appear on the desktop. Click OK and turn the knob to adjust the intensity of the light. Move the position of the light source up and down to adjust the diameter of the light spot to two centimeters.Rotate the knob to select the light intensity according to the experimental requirements. Use a compact power meter console with a standard photo diode power sensor to measure the maximal and minimal light power in the spot. The light power needs to measured in the dark in the actual experiment.Then record the light power, measure it three times and use the average value. To set up the imagining system, clamp a high resolution web camera with an iron clip about 10 centimeters above the light spot on the desktop and adjust the orientation of the camera lens toward the desktop. Connect the camera to a computer running Windows 7 through a USB interface and place the agar plate on the desktop right below the camera.Open the AMCap 9.22 software, the light spot will automatically be shown in the AMCap window. Move the camera slightly left or right to ensure that the light spot is near the center of the window. Use a clip to fix an 850 plus or minus three nanometer bandpass filter at five to seven millimeters immediately below the camera.Place three infrared light generating LEDs with a central wavelength of 850 nanometers evenly around the agar plate about five centimeters away from the edge of the plate and with LED lenses at a 70 degree angle facing toward the plate. Then connect the LEDs to a power source through their AC to DC converters. In the menu of the AMCap software, select options, video device, and capture format and set the pixel size of the captured video to 800 by 600 and the frame rate to 60 frames per second.Remove the filter and place a ruler under the camera. Adjust the camera focus until the scale line is clear and parallel to the width of the video field of view and click capture, setup, and video capture to select the save path. Then click start recording to record the actual distance corresponding to 600 pixels and calculate the ratio of each pixel to the actual distance.Before introducing the larva to the imaging setup, acquire a short video of the light position and name the unfiltered control background video light area one. Place the filter back on the camera lens and turn on a light far from the experimental device at as low a setting as possible while still allowing the larvae to be clearly observed. Use a spoon to remove some larvae from the culture medium and gently select a third instar larva.Wash the larva in distilled water and place the larva in the center of the agar plate. Gently remove any excess water from the larva and turn off the room light. Allow the larva to acclimate for two minutes in the dark environment before turning on the infrared LED light.The following light avoidance behavior experiment needs to be carried out in the dark. For the convenience of demonstration we filmed in the bright light. Gently brush the larva to the center of the plate.When the larva begins to crawl straight, rotate the plate to make the larva head towards the light spot. Click capture, setup, and video capture to select the save path and click start recording to record. Allow the larva crawl toward, enter, and leave the light spot until it is nearly out of the field of view, and click stop recording.Move the filter away from the camera and acquire a short video of the position of the light spot. Then name the post assay unfiltered video light area two and compare this video with the light area one video to ensure that the light spot position is not changed. Here, time curves of the tail speed, body bending angle and angular speed of body bending of a representative larva can be observed.The size of the larval head cast and, therefore, maximal body bending angle is significantly reduced in third instar larvae with octopaminergic neurons inhibited by tetanus toxic expression compared to parental controls indicating that Tdc2-Gal4 neurons are necessary for a normal larval light response. For successful image processing, make sure that the camera doesn't block the light and that the light spot is round and near the center of the window. Following this procedure, optogenetics can be performed to observe how the larvae react when specific neurons are activated.Using this assay, the details of larval movement can be analyzed allowing the far impact of environmental and of internal factors on larval light avoidance behavior to be studied.

light-spot-based-assay-for-analysis-of-drosophila-larval-phototaxis

Research

JoVE Journal

Biology

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Drosophila Larval NMJ Immunohistochemistry

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 31 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. Immunohistochemistry can be used to clearly image the components of the NMJ. In this article, we demonstrate how to use antibody staining to visualize the Drosophila larval NMJ.

Drosophila Larval NMJ Immunohistochemistry

This protocol demonstrates how to perform immunohistochemistry on dissected Drosophila larva.

A Neuronal and Astrocyte Co-Culture Assay for High Content Analysis of Neurotoxicity

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing measurement of multiple cellular phenotypes within a single assay. In this study, we utilized HCA to develop a novel assay for neurotoxicity. Neurotoxicity assessment represents an important part of drug safety evaluation, as well as being a significant focus of environmental protection efforts. Additionally, neurotoxicity is also a well-accepted in vitro marker of the development of neurodegenerative diseases such as Alzheimer's and Parkinson's diseases. Recently, the application of HCA to neuronal screening has been reported. By labeling neuronal cells with &beta;III-tubulin, HCA assays can provide high-throughput, non-subjective, quantitative measurements of parameters such as neuronal number, neurite count and neurite length, all of which can indicate neurotoxic effects. However, the role of astrocytes remains unexplored in these models. Astrocytes have an integral role in the maintenance of central nervous system (CNS) homeostasis, and are associated with both neuroprotection and neurodegradation when they are activated in response to toxic substances or disease states. GFAP is an intermediate filament protein expressed predominantly in the astrocytes of the CNS. Astrocytic activation (gliosis) leads to the upregulation of GFAP, commonly accompanied by astrocyte proliferation and hypertrophy. This process of reactive gliosis has been proposed as an early marker of damage to the nervous system. The traditional method for GFAP quantitation is by immunoassay. This approach is limited by an inability to provide information on cellular localization, morphology and cell number. We determined that HCA could be used to overcome these limitations and to simultaneously measure multiple features associated with gliosis - changes in GFAP expression, astrocyte hypertrophy, and astrocyte proliferation - within a single assay. In co-culture studies, astrocytes have been shown to protect neurons against several types of toxic insult and to critically influence neuronal survival. Recent studies have suggested that the use of astrocytes in an in vitro neurotoxicity test system may prove more relevant to human CNS structure and function than neuronal cells alone. Accordingly, we have developed an HCA assay for co-culture of neurons and astrocytes, comprised of protocols and validated, target-specific detection reagents for profiling &beta;III-tubulin and glial fibrillary acidic protein (GFAP). This assay enables simultaneous analysis of neurotoxicity, neurite outgrowth, gliosis, neuronal and astrocytic morphology and neuronal and astrocytic development in a wide variety of cellular models, representing a novel, non-subjective, high-throughput assay for neurotoxicity assessment. The assay holds great potential for enhanced detection of neurotoxicity and improved productivity in neuroscience research and drug discovery.

This article describes a novel protocol and reagent set designed for sensitive measurement of neurotoxic effects of compounds and treatments on co-cultures of neurons and astrocytes using high content analysis. Results demonstrate that high content analysis represents an exciting novel technology for neurotoxicity assessment.

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Examination of Drosophila Larval Tracheal Terminal Cells by Light Microscopy

Cell shape is critical for cell function. However, despite the importance of cell morphology, little is known about how individual cells generate specific shapes. Drosophila tracheal terminal cells have become a powerful genetic model to identify and elucidate the roles of genes required for generating cellular morphologies. Terminal cells are a component of a branched tubular network, the tracheal system that functions to supply oxygen to internal tissues. Terminal cells are an excellent model for investigating questions of cell shape as they possess two distinct cellular architectures. First, terminal cells have an elaborate branched morphology, similar to complex neurons; second, terminal cell branches are formed as thin tubes and contain a membrane-bound intracellular lumen. Quantitative analysis of terminal cell branch number, branch organization and individual branch shape, can be used to provide information about the role of specific genetic mechanisms in the making of a branched cell. Analysis of tube formation in these cells can reveal conserved mechanisms of tubulogenesis common to other tubular networks, such as the vertebrate vasculature. Here we describe techniques that can be used to rapidly fix, image, and analyze both branching patterns and tube formation in terminal cells within Drosophila larvae. These techniques can be used to analyze terminal cells in wild-type and mutant animals, or genetic mosaics. Because of the high efficiency of this protocol, it is also well suited for genetic, RNAi-based, or drug screens in the Drosophila tracheal system.

Examination of Drosophila Larval Tracheal Terminal Cells by Light Microscopy

Here, we present a method for light microscopy analysis of tracheal terminal cells in Drosophila larvae. This method allows for quick examination of branch and lumen morphology in whole animals and would be useful for analysis of individual mutants or screens for mutations affecting terminal cell development.

Cell shape is critical for cell function. However, despite the importance of cell morphology, little is known about how individual cells generate specific ...

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.