The authors would like to thank members of the Grillo-Hill pHly lab for discussions and support. We thank Tim Andriese, Randy Kirschner, Kitty (Ngoc-Huong) Nguyen, Marco Parent, Jonny Shaloub, and Librado Veliz for excellent technical support. This work was supported by NIH SC3GM132049 and 1R16GM153640 awards (BKGH), a CSU Biotechnology Faculty-Student Research Award (LM and BKGH), and start-up funds from the College of Science and the Department of Biological Sciences at San Jos&#233; State University (FJL). Special mention goes to Bernd Becker for their resourcefulness and assistance during this process. We thank the BioIcons (https://bioicons.com/) community for providing high-quality icons for our figures and especially to Serviere for the pipet icon, and DBCLS for the&#160;Drosophila, forceps, and desktop electron microscope icons used in Figure 1 and Figure 2, which are licensed under CC-BY 4.0 Unported. We also thank the SciDraw (https://scidraw.io/) community for providing high-quality icons for our figures, especially Diogo Losch De Oliveira (doi.org/10.5281/zenodo.3925953), which are licensed under Creative Commons 4.0 license (CC-BY).&#160;

1. Adult Drosophila collection and fixation
<ol>
	<li>Set up Drosophila crosses or select strains and place them in vials containing fly food. Incubate the vials at the desired temperature (usually 25 &#176;C) until the flies have developed&#160;and adults eclose (~10-14 days at 25 &#176;C).</li>
	<li>Anesthetize the flies with CO2 and place them on a CO2 pad.</li>
	<li>Sort the flies with a feather and select the individuals with the desired phenotype (e.g., straight wings). Make a feather fly sorter by trimming a goose feather to fit in the tapered end of a 1 mL serological pipette.</li>
	<li>Prepare a 1.7 mL microcentrifuge tube with 1 mL of 70% ethanol. Place the selected flies in the microcentrifuge tube and put on ice. Store the microcentrifuge tubes at 4 &#176;C overnight (Figure 2A). 
	NOTE: Do not preserve flies in ethanol for more than 24 h. Long-term storage of flies in 70% ethanol will result in loss of eye and body pigment.</li>
</ol>
2. Sample preparation by point-mounting
NOTE: Drosophila are soft-bodied insects that become brittle and collapse when air-dried; therefore, this protocol requires samples to be imaged the same day as they are mounted. Work in small sets of ~5 flies at a time to prevent sample loss. Increase the number of samples in a set based on efficiency. Specimens that require more time before imaging can be dehydrated through an increasing concentration series of hexamethyldisilazane (HMDS)14.
<ol>
	<li>Cut small triangular points (7.1 mm x 2.7 mm) from archival 65 lb cardstock using a specialized point punch. Prepare points by bending the tip (narrowest 25%) to a 90&#176; angle with Dumont #5 fine-tip forceps (Figure 2B).</li>
	<li>Using Dumont #5 fine-tip forceps, remove the flies from the microcentrifuge tubes (step 1.4). Gently blot the flies with lint-free lab tissue to remove excess ethanol. Position each fly on its left side on an index card under a dissecting microscope. 
	NOTE: Remove the flies from the tube by holding onto an anatomical structure that is not the area of interest-when imaging the head, we hold samples by the wing or leg. Do not hold samples by the abdomen, as that structure is used to glue the fly onto the card point.</li>
	<li>Prepare the hide glue, adjusting its consistency to the desired viscosity. Mix 1-2 drops of hide glue with 1-2 drops of deionized (DI) water, and mix with a transfer pipet on an index card. Pick up a prepared card point at the broad end with forceps, and place a small amount of the diluted glue on the bent tip of the point by dabbing it into the glue-water mix (Figure 2C). 
	NOTE: The glue should be spreadable but not runny.</li>
	<li>Apply the bent tip of the point to the anterior side of the right abdomen around abdominal segments 2-3 (Figure 2C). Before the glue dries, make slight adjustments to the fly such that the anterior-posterior axis of the fly is perpendicular to the bent tip of the point.</li>
	<li>Insert a No. 3 mounting pin into the wide end of the card-point (Figure 2D) and secure it to an insect pinning block (Figure 2E). Label each pin or row of pins with the corresponding genotype.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67485/67485fig02.jpg" /> 
Figure 2: Sample preparation. (A) Adult Drosophila are sorted based on phenotypic markers and collected into labeled microcentrifuge tubes containing 70% ethanol on ice. Flies are stored at 4&#176; C overnight.&#160;(B) Paper card points are prepared by bending the narrow end 90&#176; from the rest of the card using a pair of #5 forceps. (C) Flies are recovered from tubes and briefly allowed to air dry. Hide glue is applied to the small, folded end of the prepared card point and glued to the adult fly at abdominal segments 2-3. (D) Specimens are mounted, with an identification label, onto a #3 stainless steel insect pin. (E) Pinned specimens are stored on a sample board until ready for image acquisition. <a href="https://www.jove.com/files/ftp_upload/67485/67485fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. High-resolution focus stacking imaging acquisition
<ol>
	<li>Acquire high-resolution photographs of fly eyes with an assembled and customized focus stacking imaging system assembled and customized.
	<ol>
		<li>Capture photographs with a DSLR Camera Body with a 70-200 mm telephoto lens connected to a 20x Apo Microscope Objective via a 77 mm lens adapter.</li>
		<li>Ensure that the specimen is illuminated with a flash through a diffuser (Figure 3).</li>
		<li>Control the Z positioning using a Stackshot Controller and Macro Rail.</li>
		<li>Connect the camera, flash, and motorized stage to a heavy-duty anodized aluminum tripod.</li>
	</ol>
	</li>
	<li>Position each point-mounted sample on a universal stage gimbal with the head oriented so that the eye is facing toward the lens. Make head position adjustments by gently moving the head with the forceps. 
	CAUTION: Large and fast adjustments can result in accidental decapitation.</li>
	<li>With the camera tethered to a laptop computer, adjust the acquisition settings in the software. Photograph specimens at 20x magnification with these settings: Flash Power 1/32, Shutter Speed 1/200, Aperture F2.8, and ISO 400. Ensure that any Autofocus and Image Stabilization features are turned off. 
	NOTE: These settings balance optimal flash illumination, shutter speed, and depth of field. They would need to be adjusted for other magnifications and/or lens combinations.</li>
	<li>Set the location for saving the resulting image stack (10-50 images) to the desired file folder. Ensure enough storage capacity for the images (~8.5 MB per image).</li>
	<li>Adjust the focus stack settings on the Stackshot control unit in Auto Distance mode.
	<ol>
		<li>Set the step size to 5 &#181;m and calculate the number of steps by setting the start and stop positions of the focus stack.</li>
		<li>View the specimen in the LiveView Mode and with the camera in Auto Shoot Mode to identify the start and stop positions.</li>
		<li>Move the rail so that the closest part of the specimen is in focus (set start position), then move to where the furthest feature of interest is in focus (set end position).</li>
	</ol>
	</li>
	<li>Return the camera to the Manual Shoot Mode and start the image acquisition from the Stackshot control unit. 
	NOTE: Image acquisition time depends on the size of the specimen. The greater the depth of field necessary for large specimens, the more slices are included in the image stack, which will extend the overall acquisition time.</li>
	<li>Open files in the referenced focus stacking software. Generate a stacked image by clicking Stack | Align &#38; Stack All (PMax).</li>
	<li>Save the final image to the computer&#39;s hard drive as a .tif file by clicking File | Save Output Image. 
	&#8203;NOTE: Depending on the resolution of the stacked image file and the number of specimens imaged, large external hard drives (1 TB) may be necessary for image backup. In this protocol, stacked images are approximately 100 MB each before being compressed.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67485/67485fig03.jpg" /> 
Figure 3: Image acquisition. (A) Imaging apparatus with parts labeled as follows: a) DSLR Camera Body; b) telephoto lens; c) 20x Apo Microscope Objective and Adapter; d) Flash; e) Lens and Dome Diffusers; f) Stackshot Controller, Macro Rail, and Rotary Stage; g) Universal Stage Gimbal; h) Tripod. (B) Imaging apparatus with light diffuser&#160;in place. (C) Close-up of mounted specimen in position for imaging. <a href="https://www.jove.com/files/ftp_upload/67485/67485fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. FIJI analysis workflow to calculate adult eye area
<ol>
	<li>For image analysis, obtain the FIJI software15 from the ImageJ.net website.</li>
	<li>Choose images for analysis where the eye is centered and aligned with adequate lighting and minimal peripheral blurring, indicating proper alignment with the camera.</li>
	<li>Calibrate the image scale.
	<ol>
		<li>Download the scale bar image for 20x magnification that correlates to 500 &#181;m. Alternately, at the time of image acquisition, photograph a ruler using the same settings. Open the image in FIJI Software.</li>
		<li>Measure the length of scale bar (Figure 4A). Use the Straight Line Tool to exactly trace the line. Click Analyze | Measure. This pixel distance is equivalent to 500 &#181;m (Figure 4B).</li>
		<li>Calculate pixels per micron. Use this to convert pixel measurements into micrometer measurements.</li>
	</ol>
	</li>
	<li>Open the stacked image file in FIJI (Figure 4C).</li>
	<li>Select Magnifying Glass from the toolbar to enlarge the area of focus. Try to fill the screen with the eye and immediate surrounding head cuticle (Figure 4D).</li>
	<li>Select Freehand Select Tool from the toolbar. Outline the retinal area as closely as possible, following the contours of the outermost row of ommatidia (Figure 4E). To remove part of the selection, hold down the option button and select the pixels to remove. To add to the selection, hold down the option and shift buttons and select the pixels to add.</li>
	<li>To calculate the area, select Analyze | Measure from the top menu (Figure 4F). A new window will appear with area, mean, minimum, and maximum parameters. Copy and paste these data into a spreadsheet for documentation and conversion from pixels to micrometer measurements.</li>
	<li>Perform appropriate statistical analyses.</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67485/67485fig04.jpg" /> 
Figure 4: Image analysis in FIJI. (A) Scale the original image. Download the calibration image and measure the length of the 500 &#181;m bar. (B) Adjust the scaling using the Set Scale function. (C) Open the stacked image. (D) Magnify the image so that the eye is centered and nearly full screen. (E) Use the Freehand Select tool to outline the eye at the border between the outermost row of ommatidia and the surrounding cuticle. (F) Measure the area within the selected region is calculated by clicking Analyze | Measure | Area. <a href="https://www.jove.com/files/ftp_upload/67485/67485fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

High-Resolution Imaging and Analysis of Drosophila Eye Defects

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+larval+models+of+invasive+tumorigenesis+for+in+vivo+studies+on+tumour/peripheral+host+tssue+interactions+during+cancer+cachexia.">Drosophila larval models of invasive tumorigenesis for in vivo studies on tumour/peripheral host tssue interactions during cancer cachexia.</a> Int J Mol Sci. 22 (15), 8317 (2021).</li><li>Lam Wong, K. K., Verheyen, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metabolic+reprogramming+in+cancer:+mechanistic+insights+from+Drosophila.">Metabolic reprogramming in cancer: mechanistic insights from Drosophila.</a> Dis Model Mech. 14 (7), dmm048934 (2021).</li><li>Bonini, N. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+electron+microscopy+of+Drosophila+mutant+and+wild+type+eyes.">Scanning electron microscopy of Drosophila mutant and wild type eyes.</a> Trans Am Microsc Soc. 91 (4), 600-602 (1972).</li><li>Gibb, T. J., Oseto, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Insect collection and identification: Techniques for the field and laboratory. , (2019).</li><li>Mertens, J., Roie, M. 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The authors have no conflicts of interest to disclose.

Here we describe a method for sample preparation, high-resolution imaging, and analysis of adult Drosophila structures. The Drosophila eye is a genetically tractable model system that has yielded critical insights into molecular mechanisms underlying diseases including cancer19, neurodegeneration20 and metabolic diseases21. In particular, cancer patient &#34;avatars&#34; are generated where transgenic Drosophila carrying on...

Drosophila melanogaster is a powerful genetic model organism that has been used for decades to elucidate molecular signaling pathways and cellular behaviors. Many of the evolutionarily conserved signaling pathways that are essential for multicellular development were first identified and their mechanism of action defined in Drosophila. About 65-75% of all human disease-associated genes have orthologs in Drosophila1,2. The adult Drosophila eye is an important model that has allowed for unbiased genetic screens that facilitated the discovery of important conserved genes implicated in human diseases, including cancer3,4, neurodegeneration5, and metabolic disorders6.
The Drosophila eye is composed of ~800 unit eyes, termed ommatidia, that are precisely arrayed in a hexagonal pattern across the surface of the adult eye7. Each ommatidium is composed of eight photoreceptor neurons that occupy a distinct location within an asymmetrical trapezoid. These are supported by four non-neural cone cells and two primary pigment cells that secrete lens and pseudo-cone to focus light onto the light-sensing rhabdomeres of the photoreceptor neurons. Neighboring ommatidia are separated by a single row of interommatidial lattice cells, comprised of secondary pigment cells, tertiary pigment cells, and mechanosensory bristle complexes8,9,10.
Perturbations in eye development are visible in adult eyes as increased or decreased eye size, abnormal abundance or structure of lenses or bristles, or as a &#34;rough eye&#34; where the normally invariant hexagonal patterning is disrupted such that a row of ommatidia can no longer be followed across the surface of the eye. These phenotypes can be scored at the gross tissue level using dissecting microscopes. Detailed analysis of phenotypes traditionally includes scanning electron microscopy followed by quantitative image analysis11. However, scanning electron microscopy requires expensive instrumentation, costly reagents, sample preparation that spans days, and often, full-time staff to maintain.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67485/67485fig01v2.jpg" /> 
Figure 1: Workflow for imaging adult Drosophila structures. (A) Collect and fix adult Drosophila in 70% ethanol. (B) Prepare samples for imaging by point mounting and affixing to pins. (C) Acquire high-resolution images through focus stacking and integration. (D) Quantify images using FIJI. <a href="https://www.jove.com/files/ftp_upload/67485/67485fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
This paper presents a workflow that is relatively inexpensive, has a short sample preparation time, can easily be set up on a 3-foot lab bench, does not require hazardous materials, and could be a long-lived addition to Drosophila research labs (Figure 1). Point mounting is an entomological technique used to air dry and preserve small, soft-bodied insects, such as Drosophila12. This method relies on combining microscope objectives with high-resolution DSLR cameras for effective magnifications of 10x to 1,000x. The limited depth of field inherent to macrophotography is overcome by focus stacking: stitching together a series of images with the focal plane moving through the specimen of interest13. This method yields high-resolution images suitable for the quantification of phenotypes and could easily be adapted for other structures of interest, such as the wing, leg, thorax, and abdomen. The image analysis workflow uses the free image analysis program FIJI (NIH ImageJ). This methodology makes sample preparation, high-resolution imaging, and analysis accessible for undergraduate students and scientists at under-resourced institutions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mL serological pipette</td>
			<td>ThermoFisher Scientific</td>
			<td>170353N</td>
			<td></td>
		</tr>
		<tr>
			<td>1.7 mL microcentrifuge tubes</td>
			<td>Genesee Scientific&#160;</td>
			<td>24-282LR</td>
			<td></td>
		</tr>
		<tr>
			<td>20x Apo Microscope Objective</td>
			<td>Mitutoyo Corp.</td>
			<td>378-804-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Archival 65 lb cardstock&#160;</td>
			<td>Neenah, Inc.</td>
			<td>91901</td>
			<td></td>
		</tr>
		<tr>
			<td>Canon EF 70-200 mm USM II telephoto lens</td>
			<td>Canon</td>
			<td>3044C002</td>
			<td></td>
		</tr>
		<tr>
			<td>Canon EOS 6D Mark II DSLR Camera Body</td>
			<td>Canon</td>
			<td>1897C002</td>
			<td></td>
		</tr>
		<tr>
			<td>Diffuser Dome</td>
			<td>Macroscopic Solutions&#160;</td>
			<td>PA-DIF-GIM-SM</td>
			<td></td>
		</tr>
		<tr>
			<td>Diffuser for Mitutoyo M Plan APO Objectives</td>
			<td>Macroscopic Solutions&#160;</td>
			<td>mitutoyo-diffusers</td>
			<td></td>
		</tr>
		<tr>
			<td>Drosophila vials and plugs</td>
			<td>Genesee Scientific&#160;</td>
			<td>32-117BF</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 fine-tip forceps</td>
			<td>Fisher Scientific</td>
			<td>NC9889584</td>
			<td></td>
		</tr>
		<tr>
			<td>Goose feathers</td>
			<td>Amazon</td>
			<td>B01CMMJI6U</td>
			<td></td>
		</tr>
		<tr>
			<td>Heavy-Duty Anodized Aluminum Tripod</td>
			<td>Really Right Stuff, LLC</td>
			<td>TFA-32G</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimwipes</td>
			<td>Fisher Scientific</td>
			<td>06-666A</td>
			<td>lint-free lab tissue</td>
		</tr>
		<tr>
			<td>Levenhuk M1000 Plus Digital Camera</td>
			<td>Levenhuk</td>
			<td>70358</td>
			<td></td>
		</tr>
		<tr>
			<td>No. 3 mounting pin</td>
			<td>Indigo Instruments</td>
			<td>33414-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Nutri-Fly Bloomington Drosophila media</td>
			<td>Genesee Scientific&#160;</td>
			<td>66-113</td>
			<td>fly food</td>
		</tr>
		<tr>
			<td>Point-Punch</td>
			<td>M.C. Mieth Manufacturing, Inc.</td>
			<td>448Detail</td>
			<td></td>
		</tr>
		<tr>
			<td>Screwknob Clamp</td>
			<td>Really Right Stuff, LLC</td>
			<td>SK-Clamp</td>
			<td>For attaching the macro rail to the tripod</td>
		</tr>
		<tr>
			<td>Stackshot Controller and Macro Rail</td>
			<td>Cognisys Inc.</td>
			<td>ST3X_100_BUNDLE</td>
			<td></td>
		</tr>
		<tr>
			<td>Step-down Ring Adapter&#160;</td>
			<td>RAF Camera</td>
			<td>763461174207</td>
			<td>Lens adapter to connect the microscope objective to the camera lens</td>
		</tr>
		<tr>
			<td>Titebond Glue</td>
			<td>Franklin International</td>
			<td>5013</td>
			<td></td>
		</tr>
		<tr>
			<td>Yongnuo YN-24-EX Macro Twin Lite Flash</td>
			<td>Shenzhen Yongnuo Photographic Equipment Co.</td>
			<td>YN-24EX</td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Canon EOS Utility (v. 3.16.1).&#160;</td>
			<td>Canon</td>
			<td></td>
			<td>acquisition software</td>
		</tr>
		<tr>
			<td>FIJI</td>
			<td>National Institutes of Health</td>
			<td></td>
			<td>Fiji is released as open source under the GNU General Public License.&#160; FIJI Version 2.14.0/1.54f&#160;</td>
		</tr>
		<tr>
			<td>GraphPad Prism</td>
			<td>GraphPad Software, Boston, Massachusetts USA</td>
			<td></td>
			<td>Prism Version 10.3.1</td>
		</tr>
		<tr>
			<td>Zerene Stacker (v.1.04)</td>
			<td>Zerene Systems, LLC</td>
			<td></td>
			<td>Focus Stacking Software</td>
		</tr>
	</tbody>
</table>

a rapid, simple workflow for quantification of external adult drosophila structures

The Drosophila compound eye is a precisely patterned tissue that has revealed molecular mechanisms and biological processes that drive morphogenesis. It is a simple structure of repeating unit eyes, termed ommatidia, that is used to characterize genetic interactions and gene functions. Mutations that affect eye architecture can be easily detected and analyzed; hence, this system is frequently used in under-resourced institutions. Further phenotypic analysis often includes a Scanning Electron Microscope (SEM) to generate high-magnification images suitable for quantitative analysis. However, SEMs are expensive and require costly reagents; sample preparation spans days; and, often, they need full-time staff for sample preparation and instrument maintenance. This limits their utility at under-resourced institutions or during budgetary austerity. In entomology, the use of high-resolution digital imaging technology is a common practice for the identification and characterization of species. This paper describes a method that combines strategies and allows for high-resolution digital imaging of adult Drosophila structures and quantitative analysis using the open software ImageJ. The workflow is a rapid and student-friendly alternative that remedies the limitations of underfunded and under-resourced research facilities with a cost-effective and rapid approach to quantitative phenotypic analysis.

The Drosophila eye is an excellent model system for studying tissue patterning, growth control, and cell death. We recently published a study investigating how intracellular pH (pHi) influences tissue growth. First, we established a genetic system where overexpression of the sodium-proton exchanger DNhe2 (the ortholog of mammalian NHE1) in the developing eye causes patterning defects and increased proliferation16. Increased proliferation with higher pHi is observed across species...

A Rapid, Simple Workflow for Quantification of External Adult Drosophila Structures

Here, we present a rapid, low-cost, workflow for high-resolution imaging of adult Drosophila eyes to quantify patterning and growth defects. We describe our protocol for sample preparation by point-mounting, high-resolution image acquisition, and image analysis.

The Drosophila compound eye is a precisely patterned tissue that has revealed molecular mechanisms and biological processes that drive morphogenesis. It ...

a-rapid-simple-workflow-for-quantification-external-adult-drosophila

Research

JoVE Journal

Biology

264 Views.  San José State University. Here, we present a rapid, low-cost, workflow for high-resolution imaging of adult Drosophila eyes to quantify patterning and growth defects. We describe our protocol for sample preparation by point-mounting, high-resolution image acquisition, and image analysis.

Video: A Rapid, Simple Workflow for Quantification of External Adult Drosophila Structures

Whole Cell Recordings from Brain of Adult Drosophila

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by describing the dissecting solution and capture of the adult females used in our studies. The procedure for removing the whole brain intact, including both optic lobes, is illustrated. Dissection of the overlying trachea is also shown.  The isolated brain is not only small but needs special care in handling at this stage to prevent damage to the neurons, many of which are close to the outer surface of the tissue. We show how a special holder we developed is used to stabilize the brain in the recording chamber. A standard electrophysiology set up is used for recording from single neurons or pairs of neurons. A fluorescent image, viewed through the recording microscope, from a GAL4 line driving GFP expression (GH146) illustrates how projection neurons (PNs) are identified in the live brain. A high power Nomarski image shows a view of a single neuron that is being targeted for whole cell recording. When the brain is successfully removed without damage, the majority of the neurons are spontaneously active, firing action potentials and/or exhibiting spontaneous synaptic input. This in situ preparation, in which whole cell recording of identified neurons in the whole brain can be combined with genetic and pharmacological manipulations, is a useful model for exploring cellular physiology and plasticity in the adult CNS.

This video demonstrates the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons using standard whole cell technology. It includes images of GFP labeled cells and neurons viewed during recording.

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

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 ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

A Quantitative Fitness Analysis Workflow

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. QFA can be applied to focused observations of single cultures but is most useful for genome-wide genetic interaction or drug screens investigating up to thousands of independent cultures. The central experimental method is the inoculation of independent, dilute liquid microbial cultures onto solid agar plates which are incubated and regularly photographed. Photographs from each time-point are analyzed, producing quantitative cell density estimates, which are used to construct growth curves, allowing quantitative fitness measures to be derived. Culture fitnesses can be compared to quantify and rank genetic interaction strengths or drug sensitivities. The effect on culture fitness of any treatments added into substrate agar (e.g. small molecules, antibiotics or nutrients) or applied to plates externally (e.g. UV irradiation, temperature) can be quantified by QFA.
The QFA workflow produces growth rate estimates analogous to those obtained by spectrophotometric measurement of parallel liquid cultures in 96-well or 200-well plate readers. Importantly, QFA has significantly higher throughput compared with such methods. QFA cultures grow on a solid agar surface and are therefore well aerated during growth without the need for stirring or shaking.
QFA throughput is not as high as that of some Synthetic Genetic Array (SGA) screening methods5,6. However, since QFA cultures are heavily diluted before being inoculated onto agar, QFA can capture more complete growth curves, including exponential and saturation phases3. For example, growth curve observations allow culture doubling times to be estimated directly with high precision, as discussed previously1.
Here we present a specific QFA protocol applied to thousands of S. cerevisiae cultures which are automatically handled by robots during inoculation, incubation and imaging. Any of these automated steps can be replaced by an equivalent, manual procedure, with an associated reduction in throughput, and we also present a lower throughput manual protocol. The same QFA software tools can be applied to images captured in either workflow.
We have extensive experience applying QFA to cultures of the budding yeast S. cerevisiae but we expect that QFA will prove equally useful for examining cultures of the fission yeast S. pombe and bacterial cultures.

Quantitative Fitness Analysis (QFA) is a complementary series of experimental and computational methods for estimating microbial culture fitnesses. QFA estimates the effect of genetic mutations, drugs or other applied treatments on microbe growth. Experiments scaling from focussed analysis of single cultures to thousands of parallel cultures can be designed.

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. ...

An Improved Method for Accurate and Rapid Measurement of Flight Performance in Drosophila

Drosophila has proven to be a useful model system for analysis of behavior, including flight. The initial flight tester involved dropping flies into an oil-coated graduated cylinder; landing height provided a measure of flight performance by assessing how far flies will fall before producing enough thrust to make contact with the wall of the cylinder. Here we describe an updated version of the flight tester with four major improvements. First, we added a &#34;drop tube&#34; to ensure that all flies enter the flight cylinder at a similar velocity between trials, eliminating variability between users. Second, we replaced the oil coating with removable plastic sheets coated in Tangle-Trap, an adhesive designed to capture live insects. Third, we use a longer cylinder to enable more accurate discrimination of flight ability. Fourth we use a digital camera and imaging software to automate the scoring of flight performance. These improvements allow for the rapid, quantitative assessment of flight behavior, useful for large datasets and large-scale genetic screens.

An Improved Method for Accurate and Rapid Measurement of Flight Performance in Drosophila

Here we describe a method for rapid and accurate measurement of flight performance in Drosophila, enabling high-throughput screening.

Drosophila has proven to be a useful model system for analysis of behavior, including flight. The initial flight tester involved dropping flies into an ...

Simple Bulk Readout of Digital Nucleic Acid Quantification Assays

Digital assays are powerful methods that enable detection of rare cells and counting of individual nucleic acid molecules. However, digital assays are still not routinely applied, due to the cost and specific equipment associated with commercially available methods. Here we present a simplified method for readout of digital droplet assays using a conventional real-time PCR instrument to measure bulk fluorescence of droplet-based digital assays.
We characterize the performance of the bulk readout assay using synthetic droplet mixtures and a droplet digital multiple displacement amplification (MDA) assay. Quantitative MDA particularly benefits from a digital reaction format, but our new method&#160;applies to any digital assay. For established digital assay protocols such as digital PCR, this method serves to speed up and simplify assay readout.
Our bulk readout methodology brings the advantages of partitioned assays without the need for specialized readout instrumentation. The principal limitations of the bulk readout methodology are reduced dynamic range compared with droplet-counting platforms and the need for a standard sample, although the requirements for this standard are less demanding than for a conventional real-time experiment. Quantitative whole genome amplification (WGA) is used to test for contaminants in WGA reactions and is the most sensitive way to detect the presence of DNA fragments with unknown sequences, giving the method great promise in diverse application areas including pharmaceutical quality control and astrobiology.

We describe an endpoint digital assay for quantifying nucleic acids with a simplified (analog) readout. We measure bulk fluorescence of droplet-based digital assays using a standard qPCR machine rather than specialized instrumentation and confirm our results by microscopy.

Digital assays are powerful methods that enable detection of rare cells and counting of individual nucleic acid molecules. However, digital assays are ...