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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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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E., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+Drosophila+retina,+a+neurocrystalline+lattice.">Development of the Drosophila retina, a neurocrystalline lattice.</a> Dev Biol. 53 (2), 217-240 (1976).</li><li>Johnson, R. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hexagonal+patterning+of+the+Drosophila+eye.">Hexagonal patterning of the Drosophila eye.</a> Dev Biol. 478, 173-182 (2021).</li><li>Weasner, B. P., Kumar, J. 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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 oncogenic mutations are generated and can be used for high-throughput drug screening22. Although we focus on the adult eye, this protocol can be adapted to analyze other structures of interest such as wing, leg, thorax, and abdomen.
Fine motor skills are required at several critical steps of this protocol. Sample preparation of the adult Drosophila by point-mounting requires patience and practice. We advise beginners to practice point-mounting before preparing rare or precious samples. There are several critical steps that will require the practice of fine motor skills and careful attention.
The first critical steps in the protocol are the preparation of the card points and the application of the glue to adhere the card point to the abdomen (protocol step 2.4-2.5). It is important to apply just the right amount of glue because target anatomical structures (i.e., the head) may be covered by glue if too much is applied or it is not the right consistency. The sample must also be glued in an appropriate position, with the anterior-posterior axis perpendicular to the mounting pin. The second critical step is positioning the head for high-resolution imaging (protocol step 3.2). It is crucial that the head is oriented with the eye facing the camera lens. To adjust the head, small adjustments are made using forceps. Gentle and incremental movements are crucial as accidental decapitation may occur with large or forceful adjustments. An additional step where fine motor skills are required is when using the Freehand Select Tool from the FIJI toolbar&#160;(protocol step 4.6). Here, it is crucial to trace as closely to the outer row of ommatidia of the eye as possible. This precision is key for rigorous and reproducible area measurements of the eye (Figure 4E).
Imaging methods such as scanning electron microscopy (SEM) remain the primary approach to visualize and analyze external structures. However, SEM protocols require the user to handle hazardous chemicals and operate multiple instruments, including critical point dryers, sputter coating apparatuses, and the SEM itself. Sample preparation takes several days and requires intensive training and skill.&#160;A significant challenge academic institutions face&#160;is the financial investment and commitment to maintaining the instrument and, in our experience, personnel with expertise in maintaining and operating this equipment are generally not supported at an institutional level. Our protocol mitigates cost and eliminates hazardous chemical exposure at the cost of lower-resolution imaging compared to SEM.
The system described in this study was a preassembled package, but its modular nature permits lowering the cost by substituting individual components with equipment fabricated in-house (e.g., sample stages, flash diffusers), alternative brands or models, or equipment that might already be present in the lab. While connecting microscope objectives to macro cameras is increasing in popularity among hobbyists and professional photographers, some trial and error may be required for different combinations of objectives and lenses. To replicate the system described here, an infinity-corrected microscope objective must be connected to a tube lens (telephoto lens here) with an appropriately threaded adapter. Another consideration when exchanging components is to ensure that the working distance of the lens is compatible with the dimensions of the camera mount, specimen mount, and macrofocusing rail. Finally, the focus stacking software selected here is commercial, but many low-cost or free post-processing focus stacking solutions exist, including ImageJ plugins.
While this protocol requires practice to master sample handling techniques, it is designed to acquire and analyze high-resolution images for detailed phenotypic analysis. The procedures presented here constitute a rapid, low-cost alternative to traditional imaging methodologies. By using freeware-based digital imaging techniques for quantitative analysis, our methodology is accessible and will not require costly software purchases or updates. The imaging setup requires some cost and effort but once established, should be a long-lived addition to Drosophila labs. 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. This methodology makes sample preparation, high-resolution imaging, and analysis accessible for undergraduate students and scientists at under-resourced institutions.

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.remotexs.ntu.edu.sg/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><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&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</td><td>PA-DIF-GIM-SM</td><td></td></tr><tr><td>Diffuser for Mitutoyo M Plan APO Objectives</td><td>Macroscopic Solutions&amp;nbsp;</td><td>mitutoyo-diffusers</td><td></td></tr><tr><td>&lt;em&gt;Drosophila &lt;/em&gt;vials and plugs</td><td>Genesee Scientific&amp;nbsp;</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 &lt;em&gt;Drosophila &lt;/em&gt;media</td><td>Genesee Scientific&amp;nbsp;</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&amp;nbsp;</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>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Canon EOS Utility (v. 3.16.1).&amp;nbsp;</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.&amp;nbsp; FIJI Version 2.14.0/1.54f&amp;nbsp;</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 from yeasts to mammals17. However, we also observed that with increased pHi, the adult eye appeared smaller in size compared to controls.
Quantitative analysis of adult eye area was done to determine how elevated DNhe2 expression decreases tissue size in the Drosophila eye. We first acquired low-resolution images using an ocular insert microscope (Figure 5A), but these were of insufficient quality to perform quantitative measurements. Specifically, the edges of the eye are not in focus, thus preventing an accurate measurement of the eye area.
We measured the area of each eye in high-resolution images of adult Drosophila eyes (Figure 5B). GMR&#62;DNhe2-expressing flies had significantly smaller eyes (mean 74.28 &#181;m2) than the genetic background control w1118 flies (mean 125.83 &#181;m2) or the driver control GMRGAL4 heterozygous flies (mean 142.49 &#181;m2), which is a 41%&#160;and 48% reduction in area, respectively (Figure 5C). This decrease in eye size, in spite of increased proliferation, could be attributed to elevated levels of cell death.
Known autophagy genes, including Atg1, which induces phagophore formation, were used in genetic interaction studies (Figure 5). Effects on adult eye size were quantified using high-resolution photography and area analysis (Figure 5B). Eye size was restored (129.9 &#181;m2 vs 74.28 &#181;m2) in GMR&#62;DNhe2 flies heterozygous for Atg13&#160;compared to GMR&#62;DNhe2 alone and was&#160;similar to&#160;controls (Figure 5C). Combined with molecular marker analysis, our results support increased autophagic cell death at higher pHi.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/67485/67485fig05.jpg" /> 
Figure 5: Rescue of decreased retinal size associated with increased intracellular pH with decreased autophagy.&#160;(A) Low-resolution images of adult fly eyes taken with an ocular insert microscope. (B) High-resolution images of adult fly eyes used for eye size quantification for the following genotypes: w1118; GMRGAL4; GMR&#62;DNhe2; GMRGAL4; Atg13; GMR&#62;DNhe2; Atg13. (C) Mean measured adult eye sizes were: w1118 (125.83 &#181;m2, N = 7); GMRGAL4 (142.49 &#181;m2, N = 9), and GMR&#62;DNhe2 (74.28 &#181;m2, N = 6), GMRGAL4; Atg13 (146.4 &#181;m2, N = 15); GMR&#62;DNhe2; Atg13 (129.9 &#181;m2, N = 19) and indicated by showing median values with interquartile ranges. Statistical significance was determined by unpaired t-tests with Welch&#39;s correction. The data in B and C were previously published18. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67485/67485fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Our conclusions are strengthened by the sample preparation, high-resolution imaging, and the analysis described here. These methods allowed for a precise and quantitative description of the size of adult Drosophila eyes with various genotypes. We are continuing to use this approach to assess other genetic interactions that will elucidate mechanisms of pH-dependent growth control.

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

Nanyang Technological University

Research

JoVE Journal

Biology

355 Views.  San José State University. Ons laboratorium is geïnteresseerd in het begrijpen hoe intracellulaire pH-dynamiek cellulair gedrag moduleert. We zijn geïnteresseerd in gedrag dat bijdraagt aan de progressie van kanker, omdat we weten dat de intracellulaire pH bij de meeste vormen van kanker verhoogd is. We doen veel aan microscopie en ontwikkelen protocollen om gegevens van deze beelden te kwantificeren.We leren meer over cellulaire routes en processen die worden gereguleerd door...

Video: Een snelle, eenvoudige workflow voor het kwantificeren van externe volwassen Drosophila-structuren

Quantifying Abdominal Pigmentation in Drosophila melanogaster

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for understanding the development and evolution of morphological phenotypes. Abdominal pigmentation in Drosophila melanogaster has been particularly useful, allowing researchers to identify the loci that underlie inter- and intraspecific variations in morphology. Hitherto, however, D. melanogaster abdominal pigmentation has been largely assayed qualitatively, through scoring, rather than quantitatively, which limits the forms of statistical analysis that can be applied to pigmentation data. This work describes a new methodology that allows for the quantification of various aspects of the abdominal pigmentation pattern of adult D. melanogaster. The protocol includes specimen mounting, image capture, data extraction, and analysis. All the software used for image capture and analysis feature macros written for open-source image analysis. The advantage of this approach is the ability to precisely measure pigmentation traits using a methodology that is highly reproducible across different imaging systems. While the technique has been used to measure variation in the tergal pigmentation patterns of adult D. melanogaster, the methodology is flexible and broadly applicable to pigmentation patterns in myriad different organisms.

Quantifying Abdominal Pigmentation in Drosophila melanogaster

This work presents a method to quickly and precisely quantify the abdominal pigmentation of Drosophila melanogaster using digital image analysis. This method streamlines the procedures between phenotype acquisition and data analysis and includes specimen mounting, image acquisition, pixel value extraction, and trait measurement.

Pigmentation is a morphologically simple but highly variable trait that often has adaptive significance. It has served extensively as a model for ...

Genetics

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography

Biomedical imaging tools permit investigation of molecular mechanisms across spatial scales, from genes to organisms. Drosophila melanogaster, a well-characterized model organism, has benefited from the use of light and electron microscopy to understand gene function at the level of cells and tissues. The application of imaging platforms that allow for an understanding of gene function at the level of the entire intact organism would further enhance our knowledge of genetic mechanisms. Here a whole animal imaging method is presented that outlines the steps needed to visualize Drosophila at any developmental stage using microcomputed tomography (&#181;-CT). The advantages of &#181;-CT include commercially available instrumentation and minimal hands-on time to produce accurate 3D information at micron-level resolution without the need for tissue dissection or clearing methods. Paired with software that accelerate image analysis and 3D rendering, detailed morphometric analysis of any tissue or organ system can be performed to better understand mechanisms of development, physiology, and anatomy for both descriptive and hypothesis testing studies. By utilizing an imaging workflow that incorporates the use of electron microscopy, light microscopy, and &#181;-CT, a thorough evaluation of gene function can be performed, thus furthering the usefulness of this powerful model organism.

Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography

A protocol is presented that allows for the visualization of intact Drosophila melanogaster at any stage of development using microcomputed tomography.

Biomedical imaging tools permit investigation of molecular mechanisms across spatial scales, from genes to organisms. Drosophila melanogaster, a ...

Developmental Biology

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

Neuroscience

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Two Algorithms for High-throughput and Multi-parametric Quantification of Drosophila Neuromuscular Junction Morphology

Synaptic morphology is tightly related to synaptic efficacy, and in many cases morphological synapse defects ultimately lead to synaptic malfunction. The Drosophila larval neuromuscular junction (NMJ), a well-established model for glutamatergic synapses, has been extensively studied for decades. Identification of mutations causing NMJ morphological defects revealed a repertoire of genes that regulate synapse development and function. Many of these were identified in large-scale studies that focused on qualitative approaches to detect morphological abnormalities of the Drosophila NMJ. A drawback of qualitative analyses is that many subtle players contributing to NMJ morphology likely remain unnoticed. Whereas quantitative analyses are required to detect the subtler morphological differences, such analyses are not yet commonly performed because they are laborious. This protocol describes in detail two image analysis algorithms "Drosophila NMJ Morphometrics" and "Drosophila NMJ Bouton Morphometrics", available as Fiji-compatible macros, for quantitative, accurate and objective morphometric analysis of the Drosophila NMJ. This methodology is developed to analyze NMJ terminals immunolabeled with the commonly used markers Dlg-1 and Brp. Additionally, its wider application to other markers such as Hrp, Csp and Syt is presented in this protocol. The macros are able to assess nine morphological NMJ features: NMJ area, NMJ perimeter, number of boutons, NMJ length, NMJ longest branch length, number of islands, number of branches, number of branching points and number of active zones in the NMJ terminal.

Two Algorithms for High-throughput and Multi-parametric Quantification of Drosophila Neuromuscular Junction Morphology

Two image analysis algorithms, "Drosophila NMJ Morphometrics" and "Drosophila NMJ Bouton Morphometrics" were created, to automatically quantify nine morphological features of the Drosophila neuromuscular junction (NMJ).

Synaptic morphology is tightly related to synaptic efficacy, and in many cases morphological synapse defects ultimately lead to synaptic malfunction. The ...

Unbiased Quantification of Ommatidia in Drosophila Eye Models

Quantifying Drosophila melanogaster Eye Phenotypes: A Computational Approach Integrating ilastik and Flynotyper

The Drosophila melanogaster compound eye is a well-structured and comprehensive array of around 800 ommatidia, exhibiting a symmetrical and hexagonal pattern. This regularity and ease of observation make the Drosophila eye system a powerful tool to model various human neurodegenerative diseases. However, ways of quantifying abnormal phenotypes, such as manual ranking of eye severity scores, have limitations, especially when ranking weak alterations in eye morphology. To overcome these limitations, computational approaches have been developed such as Flynotyper. The use of a ring light allows for better qualitative images accessing the intactness of individual ommatidia. However, these images cannot be analyzed by Flynotyper directly due to shadows on ommatidia introduced by the ring light. Here, we describe an unbiased way to quantify rough eye phenotypes observed in Drosophila disease models by combining two software, ilastik and Flynotyper. By preprocessing the images with ilastik, successful quantification of the rough eye phenotype can be achieved with Flynotyper.

Author Spotlight: Quantifying Rough Eye Phenotypes in Drosophila Models of Amyotrophic Lateral Sclerosis with Frontotemporal Dementia

The Drosophila eye system is a useful tool for studying various biological processes, particularly human neurodegenerative diseases. However, manual quantification of rough eye phenotypes can be biased and unreliable. Here we describe a method by which ilastik and Flynotyper are used to quantify eye phenotype in an unbiased way.

The Drosophila melanogaster compound eye is a well-structured and comprehensive array of around 800 ommatidia, exhibiting a symmetrical and hexagonal ...

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. Drosophila larval neuronal dendritic arborization (da) is an ideal model for studying morphogenesis of neural dendrites and gene function in the development of nervous system. There are four classes of da neurons. Class IV is the most complex with a branching pattern that covers almost the entire area of the larval body wall. We have previously characterized the effect of silencing the Drosophila ortholog of SOX5 on class IV neuronal dendritic arborization complexity (NDAC) using four parameters: the length of dendrites, the surface area of dendrite coverage, the total number of branches, and the branching structure. This protocol presents the workflow of NDAC quantitative analysis, consisting of larval dissection, confocal microscopy, and image analysis procedures using ImageJ software. Further insight into da neuronal development and its underlying mechanisms will improve the understanding of neuronal function and provide clues about the fundamental causes of neurological and neurodevelopmental disorders.

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

This protocol focuses on quantitative analysis of neuronal dendritic arborization complexity (NDAC) in Drosophila, which can be used for studies of dendritic morphogenesis.

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. ...

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 ...

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to recognize a wide range of particles and destroy microbial pathogens. As organisms age, this process begins to decline, yet little is known about the underlying mechanisms or the genetic basis of immunosenescence. Here, an injection based in vivo phagocytosis assay is used to assess age related changes in different aspects of phagocytosis, such as binding, engulfment, and degradation of internalized particles, by quantifying phagocytic events in hemocytes in adult Drosophila. Drosophila melanogaster has become an ideal model to investigate age related changes in innate immune function for many reasons. For one, many genetic components and functions of the innate immune response, including phagocytosis, are evolutionarily conserved between Drosophila and mammals. Because of that, results obtained from using this protocol are likely to be widely relevant to understanding the age related changes in immune function in a variety of organisms. Additionally, we note that this method provides quantitative estimates of hemocyte phagocytic ability, which could be useful for a variety of research topics, and need not be limited to studies of aging.

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

This protocol describes an in vivo assay of phagocytosis used to assess and quantify the ability of young and aged Drosophila melanogaster hemocytes to phagocytose bacteria.

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to ...

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

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