None of this would have been possible without the support of Nasser Rusan. I would like to thank H. Doug Morris, Danielle Donahue, and Brenda Klaunberg of the NIH Mouse Imaging Facility and Ben Ache of Micro Photonics for training and helpful discussion. I also thank Mansoureh Norouzi Rad of Zeiss for scanning abdomen samples on the Xradia 520 Versa. Lauren Smith, Samantha Smith, and Rachel Ng also helped with scanning. Mike Marsh of Object Research Systems provided Dragonfly technical support. I am also grateful for support from the National Heart, Lung, and Blood Institute (1K22HL137902-01) and Start Up Funds from the University of Wyoming. I also thank the anonymous reviewers for their helpful suggestions and comments.

1. Sample selection and cuticle preparation
<ol>
	<li>Choose the appropriate developmental timepoint (embryo, larva, pupa, or adult) for imaging.
	<ol>
		<li>For embryonic stages, cage females on grape juice agar plates smeared with yeast paste and collect eggs every 30-60 min. Leave these embryos to develop at 25 &#176;C until the proper stage is reached.</li>
		<li>For larval stages, collect first and second instars from timed embryo collection experiments. Directly pick wandering 3rd instars from the side of the food vial under non-crowding conditions.</li>
		<li>For pupal timing, collect white pre-pupae (inverted spiracles) and make note of the time when the cuticle starts to brown. Animals will progress through 15 stages of pupal development at defined timepoints following cuticle browning and can be collected accordingly42.</li>
		<li>Collect adults as virgins following eclosion and allow to age in a food vial for a required amount of time (e.g., 5 days for complete gut development).</li>
	</ol>
	</li>
	<li>Transfer 5-50 animals to a 1.5 mL microcentrifuge tube containing 1 mL of 0.5% Triton X-100 in 1x Phosphate Buffered Saline (0.5% PBST). While tapping the tube periodically on the benchtop, incubate for 5 min at room temperature (RT) to assist in the removal of the hydrophobic coating and allow animals to become fully submerged.
	<ol>
		<li>For embryonic, larval, and pupal stages, arrest further development by placing the tube in a heat block set to 100 &#176;C for 20 s followed by cooling at RT for 5 min. 
		NOTE: Animals can also be flash frozen in liquid nitrogen37.</li>
	</ol>
	</li>
</ol>
2. Fixation and staining
<ol>
	<li>Remove 0.5% PBST and add 1 mL of Bouin&#8217;s Solution. Tap tube to ensure animals are fully submerged. 
	CAUTION: Bouin&#8217;s Solution contains formaldehyde. It may cause acute toxicity to skin and eyes if spilled and can be fatal if swallowed. Wear gloves, safety glasses and a lab coat when handling. 
	NOTE: Additional fixation techniques can also be employed, such as the use of ethanol. The merits of other fixatives are described in detail in ref.1,30.
	<ol>
		<li>For embryo and adult samples, incubate on the benchtop for 16-20 h at RT.</li>
		<li>For larval and pupal samples, fix animals for 2 h at RT. Discard Bouin&#8217;s solution and wash with 1x PBS for 5 min thrice. Transfer to a multi-well dissecting dish containing 1x PBS and use a small minutien pin attached to a holder to poke a hole in the anterior and posterior cuticle being careful to not disrupt any underlying soft tissue.</li>
		<li>Transfer larval and pupal samples back to a microfuge tube and add 1 mL of fresh Bouin&#8217;s solution. Incubate on benchtop for 24 h at RT.</li>
	</ol>
	</li>
	<li>Remove Bouin&#8217;s Solution and wash sample for 30 min with 1 mL of &#956;-CT Wash Buffer (0.1 M Na2HPO4, 1.8% Sucrose) or 1x PBS thrice.</li>
	<li>Add 1 mL of the appropriate staining solution and incubate on benchtop for 2-7 days. 
	NOTE: Stain choice will depend on many factors, but is generally a balance between penetration, incubation time, and resolution. In general, phosphotungstic acid (PTA) provides superior contrast and resolution of soft tissue but requires mechanical disruption of the cuticle and longer incubation times. The merits of different stain types are described in detail in ref.1.
	<ol>
		<li>For iodine staining, use 1 mL of 0.1 N I2KI (Lugol Solution). Incubate for 2 days. No further disruption of the adult cuticle is necessary.</li>
		<li>For phosphotungstic acid (PTA) staining, use 1 mL of a 0.5% (w/v) solution diluted in water. Disrupt the adult cuticle, either by removing the mouthparts or poking holes in the thorax or abdominal cuticle with a small minutien pin attached to a holder. Incubate for 5-7 days, or longer if tissue staining appears non-homogenous.</li>
	</ol>
	</li>
	<li>Wash with 1 mL of ultrapure water or 1x PBS for 30 min. Repeat wash step. Store animals at RT in ultrapure water or 1x PBS for up to one month.</li>
	<li>If animals are to be scanned while hydrated, proceed directly to section 4. If longer preservation of the sample is needed, proceed to section 3 of the protocol.</li>
</ol>
3. Critical point drying (Optional)
<ol>
	<li>Perform an ethanol (EtOH) dehydration series on the sample: 10%, 25%, 50%, 75%, 100%, 100%. Add 1 mL of the EtOH solution and incubate on the benchtop for 1 h for each concentration in the order stated.</li>
	<li>After the final 100% EtOH soak, replace with fresh 100% EtOH and let sample incubate on the benchtop overnight.</li>
	<li>Perform critical point drying of samples following the manufacturer&#8217;s instructions. 
	NOTE: Electron microscopy facilities generally have a critical point drying machine (see Table of Materials) that will perform the drying of the sample following EtOH dehydration.</li>
</ol>
4. Sample mounting
<ol>
	<li>For critically point dried samples, hot glue samples to an insect pin or other holder designed to fit within the chuck of the rotating stage or place&#160;it in a plastic or glass capillary tube&#160;(~1.0-1.25 mm inner diameter).</li>
	<li>For hydrated samples, fill a P10 pipette tip with water and secure the narrow end by either heat sealing or paraffin film to prevent leakage. 
	CAUTION: Be sure to wrap any connections tightly so that water does not leak into the scanner and cause damage.
	<ol>
		<li>Transfer a single specimen to the pipette tip using forceps. Using a long, slender object, such as a dulled 20 G needle or another pipette tip, carefully push the specimen down the tip until it just contacts the wall of the pipet tip to hold it in place.</li>
		<li>Cover the opening of the pipette tip with paraffin film to prevent evaporation of water during long scans.</li>
		<li>Mount the P10 pipette using a holder designed to fit within the chuck of the rotating stage (Figure 1A-C).</li>
	</ol>
	</li>
</ol>
5. Scanning
<ol>
	<li>Perform any necessary calibrations of the machine prior to imaging for the day. 
	NOTE: These will vary by manufacturer and it is recommended to consult with an application specialist to determine the proper steps. Please see the Table of Materials for specific information on the setup and software used in this protocol. Generally, calibrations such as a stage axis alignment to remove any &#8216;wobble&#8217; associated with the chuck being off-axis relative to the rotating stage and performing flat-field corrections to ensure uniform background pixel intensities on the camera provide optimal imaging conditions for the best resolution and datasets that are comparable across multiple scans.</li>
	<li>Open the scanner door to gain access to the rotating stage chuck by clicking on the &#8216;Open Door&#8217; icon in the upper left corner of the software. Attach the sample by tightening the collar around the base of the sample holder. 
	NOTE: Use gentle pressure when attaching the sample to the rotating chuck to maintain scanner alignment.</li>
	<li>Set scanning parameters in the software controlling the scanner for optimal resolution and contrast. 
	NOTE: These will need to be determined empirically as the X-ray source, camera/detector, and geometry of each scanner will vary by manufacturer. Additional information for selecting the best parameters can also be found here43, as well as from the manufacturer&#8217;s application specialist.
	<ol>
		<li>Open the X-Ray source power control (Options | X-ray Source). Use the slider bars to set X-ray voltage to 30-40 kV and current to 100-110 &#956;A to produce an X-ray beam with 3-4W of power and a small spot size for enhanced resolution.</li>
		<li>Use the Acquisition Modes dialog (Options | Acquisition Modes) to set camera exposure time to 500-800 ms.</li>
		<li>Use the slider bar located next to the magnifying glass icon at the bottom of the software to set the desired image pixel size (~700 nm to 4 &#956;m), depending on camera settings and position. This determines the number of projection images that are acquired during the scan, with more projections leading to enhanced resolution but longer scan times (see Representative Results).</li>
		<li>Click and drag the slider bar located next to the rotating arrow at the bottom of the software to move the sample along its 360&#730; rotation path. Ensure that the sample stays within the field of view.</li>
	</ol>
	</li>
	<li>Click on the &#8216;Start Acquisition&#8217; icon (blue circle arrow icon) at the top of the software. A dialog box appears that allows additional scanning parameters to be set, and the file and output folder where the scan will be saved to be named.
	<ol>
		<li>Set the random movement to 10 and average 4-6 frames. The rotation step is automatically calculated depending on the camera settings used.</li>
	</ol>
	</li>
	<li>Begin scan by clicking &#8216;OK&#8217; on the Acquisition dialog box. A second progress bar dialog appears that shows the scan time. The scanner will now acquire a series of projection images (Figure 1D) of the specimen along the rotation path and does not need to be monitored.</li>
</ol>
6. Reconstruction
<ol>
	<li>To generate the tomograms, perform image reconstruction using the projection images. 
	NOTE: While almost all reconstruction of images from cone beam geometry scanners rely on the Feldkamp algorithm16, individual parameters will vary depending on the software implementation and should be determined empirically. The following settings, specific for a commercially available software (see Table of Materials) can be used as a guide. Parameters such as misalignment compensation, ring artifact reduction, and beam hardening correction (0% for most fly samples) are performed in an iterative fashion to choose the best values for the final reconstruction. For advanced users who would like more control over the reconstruction process, see the MATLAB interface here44.</li>
	<li>Perform an initial image alignment by utilizing a shift-correction algorithm based on reference scans45 (Actions | X/Y alignment with a reference scan).</li>
	<li>Fine tune each reconstruction parameter (Start | Fine tuning). This activates a &#8216;Parameter fine-tuning&#8217; dialog box.
	<ol>
		<li>Fine tune the alignment by clicking &#8216;Next&#8217; to &#8216;Post-Alignment&#8217;. Set the number of iterations to five and the parameter step to 1.0. Click the Start button to generate a series of previews. Select the image that is properly aligned. 
		NOTE: A properly aligned image will not be blurry or display &#8216;shearing&#8217; artifacts where an otherwise continuous structure (such as the cuticle) appears split.</li>
		<li>Fine tune the ring artifact reduction by clicking next to it. Set the number of iterations to five and the parameter step to 1.0. Click the Start button to generate a series of previews. Select the image that contains the fewest number of rings.</li>
	</ol>
	</li>
	<li>Once the above parameters have been optimized to give the best image, ensure that the selected values are properly represented in the Settings tab (Settings).</li>
	<li>Adjust any final brightness and contrast values using the histogram, file parameters such as bit depth and type, and utilize a Region of Interest (ROI) encompassing only the structures of interest (e.g., whole fly or head only) to reduce computational time (Output).</li>
	<li>Begin the reconstruction (Start | Start). If multiple reconstructions are required, add the current image to the batch manager (Start | Add to batch) and repeat steps 6.2-6.5 for the remaining images.</li>
</ol>
7. Image analysis
<ol>
	<li>Visualize tomograms in two and three dimensions and perform further morphometric analysis with freeware or commercial software. 
	NOTE: Details of the software used in this protocol is available in the Table of Materials. Other software packages that are capable of evaluating &#181;-CT datasets include freeware options such as FIJI46, Seg3D (www.seg3D.org)47, and ITK-SNAP48, plus commercial software (e.g., AMIRA). Other applications that employ machine-learning based algorithms to semi-automate the segmentation process can help speed workflows and reduce human bias (e.g., Biomedisa49).</li>
	<li>Import the tomogram file into the software (File | Import Image Files). The metadata associated with the file should automatically load into the window but can also be set manually to match the image properties.</li>
	<li>To segment a structure of interest, click on the Segment tab on the left-hand side of the screen. Create a new Region of Interest (ROI) (Basic | New), give it a name and select an appropriate color.</li>
	<li>Define the threshold range that encompasses the structure of interest (Range | Define Range) using the slider bar.</li>
	<li>Select an appropriate 2D or 3D ROI painter tool mode (ROI Painter | Paintbrush option).</li>
	<li>To paint an area defined by the threshold; press and hold the Ctrl key while holding the left mouse key. To remove an area, press and hold the Shift key while holding the left mouse key. 
	NOTE: To change the size of circular or square paintbrush, simply scroll the mouse wheel while holding either the Ctrl or Shift keys.</li>
	<li>Continue to paint the structure of interest throughout its entire Z-volume.</li>
	<li>Convert the ROI into a Mesh (Export | To a Mesh | Normal).</li>
	<li>Highlight the mesh overlay by clicking on it in the Data Properties and Settings panel in the upper right-hand corner.</li>
	<li>Smooth the mesh using an appropriate number of iterations (Smooth Mesh | Iterations). 
	NOTE: Use the same mesh smoothing iteration value for all images that are to be compared.</li>
	<li>Ensure that the measurements of the mesh ROI (Surface Area, Volume and Feret Diameter) are displayed in the Information panel on the right-hand side of the screen for basic morphometric analysis. Additional analysis can be performed using the Object Analysis Module.</li>
	<li>Render the mesh object and the entire tomogram image and visualize in 3D. Use the built-in Movie Maker to generate a video of the object (Right Mouse Click | Show Movie Maker) using individual frames from the viewer (Add Key). 
	NOTE: Several visual enhancements can also be applied to the movie using the visual effects tab in the right-hand panel to highlight certain features, etc.</li>
	<li>Export the video for viewing using the Export Animation button in the Movie Maker.</li>
</ol>

The author declares no competing or financial interests.

Visualizing intact Drosophila melanogaster at all developmental stages has remained a challenge, primarily due to the incompatibility of light microscopy with the thick, pigmented cuticle found in this animal. While other whole animal imaging methods, such as Magnetic Resonance Imaging (MRI), Optical Coherence Tomography (OCT), and ultramicroscopy coupled with tissue clearing have been used with success in flies50,51,52,53,54, &#956;-CT presents a number of advantages that make it ideal for whole animal imaging of this organism13,15,30. X-rays easily penetrate the pigmented cuticle and their small wavelength allows for sub-micron imaging. Labeling requires minimal investment in widely available chemicals and no specialized bench skills13. &#956;-CT scanners are also commercially available, and costs are comparable to light microscopy platforms, while also being more attractive to wider range of disciplines (Geology, Paleontology, Engineering, etc.) that can also benefit from its availability at an institution. Synchrotron X-Ray sources can also be used for high resolution &#956;-CT imaging of fixed and living insects31,55,56, but are less accessible than commercial benchtop scanners.
This protocol provides an efficient way to obtain &#956;-CT images of fly adults, pupa, larva and cellularized embryos. Note that for many of the steps outlined above, alternative methods can also be applied to prepare samples for imaging. Other studies have provided a detailed comparison of different fixation, labeling, and drying steps for use in insects and those interested in adopting this technique are encouraged to evaluate the merits of each approach1,4,13,29,30,57. While this protocol is relatively straightforward, a few helpful suggestions are presented.
First, care should be taken when disrupting the cuticle of intact specimens such that underlying soft tissues are not significantly disrupted. It is important to let larval and early pupal stages undergo fixation for 2 hours in Bouin&#8217;s solution before poking. This will stiffen the tissue and limit the amount of hemolymph that will ooze out of the cuticle holes, which can alter organ architecture. Individual body segments (head, thorax and abdomen) of the adult can be separated if the structure(s) of interest are located there. It is recommended to use a scalpel to cleanly slice through these segments rather than pulling them apart with forceps, which could disrupt the 3D architecture of the gut or central nervous system, for example. As for timing, adults generally need only 16 hrs. for complete fixation, whereas larval and pupal stages need 24 h. Also, if iodine or PTA staining appears uneven, the sample can be placed back in solution to incubate longer until even staining is achieved. Finally, hydrated samples should not be placed at 4 &#176;C, as this seems to induce the formation of air bubbles within the body cavity after warming to room temperature.
Second, sample mounting will vary by instrument, stage type and whether the sample needs to remain hydrated or has been critical point dried. If hydrated, ensure the sample does not leak and possibly destroy the scanner. When mounting the sample inside a pipette tip, be sure to push gently with a dulled object until the specimens encounters slight resistance and can&#8217;t move. Pushing too hard can lead to cuticle deformation and underlying structural defects. Also, be sure that the sample is aligned in the holder as close to the axis of rotation as possible. Any wobble will increase scan times due to the larger field of view and reduce the resolution of the final tomogram following reconstruction.
Third, scanner settings for acquiring projection images will also vary by instrument. To maximize the resolution capabilities of the scanner, the X-ray beam spot size should be as small as possible (5-10 &#956;m). This can be achieved by balancing X-Ray voltage and current settings such that the total power is 3-4 W. With these settings and the appropriate exposure time on the camera, proper X-ray beam attenuation by the sample and optimal image contrast can be achieved. The use of aluminum or copper filters between the object and the X-ray source can be used to fine-tune the optimal X-ray energy settings for the best image contrast or attenuate the beam sufficiently for higher powered sources to be used. As for image resolution, this will depend on many different variables, including stain type, number of projection images, image pixel size, camera position, sample movement, scanner vibrations and reconstruction parameters. A bar pattern phantom (QRM GmbH) containing known size markers can help evaluate spatial resolution for a given scanner and camera setting.
It is also worth evaluating the merits of imaging critical point dried or hydrated samples. Sombke et al. performed a comparative assessment of the two methods and found critical point drying to be superior for &#956;-CT applications involving arthropods30. However, benefits of hydrated samples are that animals are subjected to less chemical and mechanical exposure that could lead to both quantitative and morphological artifacts. This also tends to preserve delicate tissues better than CPD. However, hydrated samples have a much shorter shelf life and should be imaged no later than one month after fixation since tissue degradation and reduced image quality becomes obvious at that point. Also, the resolution of hydrated samples will be slightly less than a critical point dried sample, because X-rays must also penetrate through both a plastic pipette tip and the surrounding liquid (water or buffer). Critical Point Dried samples can be preserved for much longer periods of time, especially when kept on Drierite. They also can be placed directly in the X-ray beam path by simply gluing the wings or legs to an insect pin and placing it in the stage chuck, simplifying the mounting process. However, the extensive ethanol dehydration of these samples can lead to tissue shrinkage and loss of delicate tissue architecture, which is why it is important to perform a range of increasing EtOH concentrations to minimize these effects. Nonetheless, it should be noted that all forms of chemical treatment, including paraformaldehyde fixation and even iodine staining can cause tissue shrinkage58,59. While neither method will provide measurements of &#8216;actual organ size&#8217; in a living fly, morphometric measurements are still valid when comparing mutant and wildtype animals so long as the fixation, staining, and drying steps are carried out identically for both sets of samples&#8212;preferably in parallel.
In conclusion, &#956;-CT provides a useful whole animal imaging tool for Drosophila33,34,35,36,37,38,39,40,41. Many other studies have showcased the power of this technology for understanding various aspects of insect taxonomy, ecology, physiology, development, and anatomy that can help inform future studies in flies1,28,30,31,32,55,56,57. Combined with the genetic and light microscopy tools already widely used in this organism, &#956;-CT can position itself within an experimental pipeline that allows for a deeper understanding between genotype and phenotype.

Imaging methods that allow for the detailed investigation of interior structures of an object without destroying its overall 3D architecture have proven to be widely beneficial to a number of different disciplines, including physics, engineering, materials science, archaeology, paleontology, geology, and biology1,2,3,4,5,6,7,8,9. Among these nondestructive imaging methods, X-ray based platforms are especially useful due to the ability of high energy X-rays to penetrate many different sample types and materials with minimal scattering compared to visible light waves. Computed Tomography (CT), Microcomputed Tomography (&#181;-CT), Nanocomputed Tomography (Nano-CT), and Synchrotron Microtomography have, therefore, emerged as the primary technologies for X-ray based imaging of samples ranging from meters to microns, with millimeter to sub-micron resolution capabilities10,11,12,13,14.
While these platforms differ in their design, X-ray geometry, and components in order to balance sample size and resolution, they all rely on the same basic principle for image capture: a source of X-rays that travel through the object and are captured by a detector. Differential attenuation of the X-ray beam as it passes through varying densities within the object generates image contrast. 3D data is obtained by rotating either the sample or the detector, collecting a series of 2D projection images that are then reconstructed using algorithms into tomograms containing 3D information whose resolution is isotropic in x,y,z15. For many benchtop &#181;-CT scanners that utilize a cone-beam X-ray geometry to project X-rays at the object being imaged, the Feldkamp algorithm is used to accurately reconstruct the object with minimal errors16.
Resolution of a given platform is determined primarily by system parameters such as the size of the X-ray beam (spot size), scanner geometry (distance from object to X-ray source), size of the pixels on the detector, and the reconstruction algorithm employed. Additional factors, such as scanner vibrations, X-ray beam fluctuations, sample movement, and material type or chemical stain used to visualize the object can also significantly influence spatial resolution under real world imaging condidtions15.
For biomedical applications, CT and &#181;-CT have played a key role in advancing our understanding of anatomy, physiology, development, and disease mechanisms, serving as a tool for both human patient diagnoses and as a preclinical imaging platform for model organisms17,18. For example, the Mouse International Phenotyping Consortium, whose goal is to identify the function of every gene in the mouse genome, utilizes &#956;-CT as part of their phenotyping pipeline19. Their results have been critical for understanding genes involved in development and disease processes, while also serving as an atlas for mouse anatomy and development20. Other model organisms, such as zebrafish and rats, have also fully embraced the use of &#956;-CT for performing whole animal phenotyping of a number of gene mutants17,21,22,23.
The advantage of combining whole animal imaging with model organisms is that a mechanistic understanding of gene function for a given biological process can be fully explored. This is possible because of the well-characterized genomes and many genetic tools available in model organisms that allow for precise manipulation of gene function at distinct developmental timepoints, specific tissues, individual cells, and even subcellular organelles. These include binary expression systems such as the UAS/GAL4 system (and its many derivatives), CRISPR/Cas9, and RNAi24,25,26. When these genetic tools are used in conjunction with a powerful imaging pipeline consisting of electron microscopy, light microscopy (fluorescent and non-fluorescent), and whole animal imaging such as &#956;-CT, a thorough evaluation of molecules, cells, tissues, organs, and the entire organism can be achieved, allowing for a much deeper understanding of gene function.
This protocol focuses on the use of &#181;-CT in the non-mammalian model organism Drosophila melanogaster, whose myriad genetic tools have helped elucidate numerous molecular mechanisms26,27. It was adopted from previous protocols in non-model insects1,28,29,30,31,32, and builds off of previous &#181;-CT studies in Drosophila to establish a standardized protocol for its use in this animal33,34,35,36,37,38,39,40,41. The steps for successful sample preparation, imaging and analysis of fly &#956;-CT datasets using commercially available scanners are outlined. With this protocol, all developmental stages of the fly can be visualized at high resolution for both descriptive and hypothesis-testing studies, including taxonomy, anatomy, development, physiology, and disease27. This protocol will also be useful for imaging virtually any insect and even non-living materials that require chemical staining for image contrast to enhance visualization by &#956;-CT.

<table><tbody><tr><td>100% Ethanol</td><td></td><td></td><td>For critical point drying</td></tr><tr><td>Bouin&#039;s Solution</td><td>Sigma-Aldrich</td><td>HT10132</td><td>For animal fixation</td></tr><tr><td>Critical Point Dryer</td><td></td><td></td><td>Dries samples using the critical point method; multiple options available (Balzers CPD 020 or Leica EMCPD300)</td></tr><tr><td>Dragonfly Software</td><td>Object Research Systems</td><td></td><td>For visualization and segmentation of micro-CT datasets; https://www.theobjects.com/dragonfly/index.html</td></tr><tr><td>Heat Block</td><td></td><td></td><td>For microfuge tubes</td></tr><tr><td>Image Analysis Workstation</td><td></td><td></td><td>Should contain sufficient RAM and quality graphics card for 3D rendering</td></tr><tr><td>Iodine Solution (I&lt;sub&gt;2&lt;/sub&gt;KI)</td><td>Fisher Scientific</td><td>SI86-1</td><td>For staining</td></tr><tr><td>Microcomputed Tomography Scanner</td><td>Bruker</td><td>Skyscan 1172</td><td>Cone-beam X-Ray geometry; detector is a Hamamatsu 10 MP camera with 11.54 &amp;micro;m pixel size.</td></tr><tr><td>Microcomputed Tomography Scanner Software</td><td>Bruker</td><td></td><td>For controling the scanner itself (e.g., performing flat field corrections, X-ray tube power, camera expsoure times, acquisition, etc.)</td></tr><tr><td>Minutien Pins</td><td>Fine Science Tools</td><td>26002-15</td><td>For poking hole in cuticle</td></tr><tr><td>NRecon Image Reconstruction Software</td><td>Bruker</td><td></td><td>Used to reconstruct cross-section images from 2D projection images taken with cone-beam X-Ray geometry</td></tr><tr><td>P10 pipet tips</td><td>Genesee Scientific</td><td>24-120</td><td>Sample mounting</td></tr><tr><td>Phosphate Buffered Saline</td><td>Resarch Products International</td><td>P32060-4000.0</td><td>Dilute to 1X with water before use</td></tr><tr><td>Phosphotungstic Acid Hydrate</td><td>Sigma-Aldrich</td><td>79690-25g</td><td>For staining</td></tr><tr><td>Pin Holder</td><td>Fine Science Tools</td><td>26018-17</td><td>For Minutien Pins</td></tr><tr><td>Triton X-100</td><td>Research Products International</td><td>111036</td><td>To remove waxy coating from adult flies (as 0.5% PBST)</td></tr><tr><td>X-Ray Microscope</td><td>Zeiss</td><td>Xradia 520 Versa</td><td>Cone-beam X-Ray geometry featuring Fresnel zone plate objective lenses for Resoluton at a Distance (RaaD&amp;trade;)</td></tr></tbody></table>

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.

Figure 2 shows images of an embryo, 3rd instar larvae, pupae at the pharate adult stage (P7), and an adult female fly stained with iodine and imaged hydrated in water using a commercial benchtop scanner. Excellent preservation and even staining of delicate tissue are apparent, allowing all major organs to be readily identified and used for morphometric analysis and 3D visualization.
Typically, scans that acquire fewer projection images of the specimen provide lower resolution than scans that acquire more projection images, with the tradeoff being time spent scanning. Although scan times will vary by instrument and other scanning parameters, scans that acquire a few hundred projections (~3 &#181;m image pixel size) takes roughly 30 min per specimen, whereas scans consisting of thousands of projection images (700 nm-1.25 &#181;m image pixel size) can take 8-16 h. A comparison of the same adult fly headcase taken in both &#8216;slow&#8217; (thousands of projections) and &#8216;fast&#8217; (hundreds of projections) scan mode is shown in Figure 3. Importantly, morphometric analyses do not differ between &#8216;slow&#8217; and &#8216;fast&#8217; scans (Figure 3C)40. Our imaging pipeline, therefore, utilizes fast scans to generate a sufficiently large sample size for morphometric analysis, and slow scans to visualize any morphological or anatomical defects at higher resolution. Using the software (Step 7), any tissue or organ system of interest can be segmented and used for morphometric analysis and visualized in 3D using the movie maker (Movie 1).
Figure 4 shows an example of the fly abdomen stained with PTA and imaged hydrated (water) or following critical point drying (CPD) on an X-ray microscope (Table of Materials). The detail afforded by a combination of the PTA and the capabilities of this platform is readily evident in these images, such that individual epithelia cells of the midgut and sperm bundles within the testes are easily resolved. While the CPD image shows marginally increased resolution compared to the hydrated sample, better preservation of the ultrastructure of delicate tissues (such as the fat cells near the cuticle) is achieved with hydrated samples (Movie 2).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61515/61515fig01.jpg" /> 
Figure 1: Overview of scanner design and sample mounting for &#181;-CT.&#160;(A) A commercial benchtop &#181;-CT scanner. (B) View inside the scanner. The X-ray source (right) emits X-Rays that pass through the sample located on a rotating stage (yellow arrow). Attenuation of these X-rays generate image contrast as they pass through the sample and onto the detector, which consists of a scintillation screen that converts X-rays to photons and a standard CCD camera (left). (C) Mounting an adult fruit fly for hydrated imaging in water. The connection points between the pipette tip and the brass holder are wrapped in paraffin film to prevent leakage and potential damage to the scanner. The stage chuck is also highlighted. Note that the pipette tip was positioned slightly off-axis, which led to a longer scan time and reduced resolution in the final reconstruction. (D) A single 2D projection image of an adult female fly; hundreds to thousands of these projections are acquired during a scan along the rotation axis and are used for reconstruction to generate tomograms containing isotropic resolution and accurate 3D information. Scale Bars (C) = 2 mm. P, Posterior; V, Ventral. This figure has been modified from Schoborg et al.40. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61515/61515fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61515/61515fig02.jpg" /> 
Figure 2: All Drosophila melanogaster life cycle stages, imaged by &#181;-CT.&#160;Samples stained with iodine and imaged hydrated in water. Shown is a single 2D slice. (A) An embryo that has completed the early stages of gastrulation (asterisk). (B) A third instar larva. (C) A P7 pharate adult during metamorphosis. (D) An adult female. Various organs are highlighted: BWM, body wall muscles; Br, brain; Cd, cardia; Cr, crop; DLMs, dorsal longitudinal muscles; DVM, dorsal ventral muscles; E-AD, eye-antennal disc; Em, embryo; FB, fat bodies; FBCs, fat body cells; H, heart; Hg, hindgut; La, lamina; L, leg; Mg, midgut; OL, brain optic lobe; Ov, ovipositor; PC, pupal cuticle; SG, salivary glands; VNC, ventral nerve cord; W, wing; WD, wing disc. Scale Bars (A) = 100 &#181;m; (B)-(D) = 500 &#181;m. D, Dorsal; A, Anterior; L, Left. Scanning parameters: Source to Sample Distance (mm): (A, D) 36.5, (B) 48.8, (C) 40.3. Source to Camera Distance (mm): (A, D) 350, (B, C) 285. Camera Pixel Size (&#181;m): (A-D) 11.6. Image Pixel Size (&#181;m): (A, D) 1.2, (B) 1.9, (C) 1.7. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61515/61515fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61515/61515fig03.jpg" /> 
Figure 3: Scanning parameters and image resolution do not alter morphometric analyses.&#160;An adult head scanned using both (A, A&#8217;) &#8216;fast&#8217; scanner settings (hundreds of projections) and (B, B&#8217;) &#8216;slow&#8217; scanner settings (thousands of projections). The brain is outlined in yellow. (C) Brain volume measurements from slow and fast scans. Highlighted structures: AL; antennal lobe; CB, central brain; FB, fan shaped body; FCs, fat cells; La, lamina; Lo, lobula; LoP, lobula plate; Me, medulla; Re, retina. n = 5, Welch&#8217;s t-test. ns = not significant. Scale bars = 100 &#181;m. Scanning parameters: Source to Sample Distance (mm): (A) 44.4, (B) 36.5. Source to Camera Distance (mm): (A) 348 (B) 350. Camera Pixel Size (&#181;m): (A-B) 11.6. Image Pixel Size (&#181;m): (A) 2.95, (B) 1.2. This figure has been modified from Schoborg et al.40. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61515/61515fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61515/61515fig04.jpg" /> 
Figure 4: Drosophila melanogaster abdomen imaged by X-ray Microscopy.&#160;Abdomens were stained with 0.5% PTA and imaged hydrated (water) or following critical point drying (CPD). (A) Critical Point Dried abdomen, shown from the YZ perspective and (A&#8217;) XZ perspective. (B) Hydrated abdomen, shown from the YZ perspective and (B&#8217;) XZ perspective. Various organs are highlighted: FC, fat cells; Hg, hindgut; Mg, Midgut; SP, Sperm Pump; Te, Testes. Scale Bars (A) = 250 &#181;m. D, Dorsal; A, Anterior; L, Left. Scanning parameters: Source to Sample Distance (mm): (A) 6.7, (B) 7. Source to Camera Distance (mm): (A) 28 (B) 29.5. Objective: (A-B) 4X. Image Pixel Size (&#181;m): (A-B) 0.65. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61515/61515fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Movie 1: A third instar larva, rendered in 3D using the Movie Maker in Dragonfly. Highlighted organs include the brain (yellow), eye-antennal imaginal discs (red), fat body (blue) and the body wall muscles (green). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61515/Movie_1_Revised.avi" target="_blank">Please click here to download this movie.</a>
Movie 2: Comparison of samples imaged in water or follow critical point drying. Abdomens stained with 0.5% PTA are shown. Both abdomens were scanned with identical image pixel size settings (0.65 &#181;m). A series of 2D slices are shown in a &#8216;Z-stack&#8217; format (YZ) starting at the dorsal surface and ending at the ventral surface of the abdomen. Organs highlighted: FC, fat cells; Mg, Midgut; SP, Sperm Pump; Te, Testes. Scanning parameters: Source to Sample Distance (mm): (A) 6.7, (B) 7. Source to Camera Distance (mm): (A) 28 (B) 29.5. Objective: (A-B) 4X. Image Pixel Size (&#181;m): (A-B) 0.65. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61515/Movie_2_Revised.avi" target="_blank">Please click here to download this movie.</a>

Watch this Scientific Journal Video about Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography at JoVE.com

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.

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 ...

whole-animal-imaging-drosophila-melanogaster-using-microcomputed

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

4.9K Views.  University of Wyoming. Er wordt een protocol gepresenteerd dat de visualisatie van intacte Drosophila melanogaster in elk stadium van ontwikkeling mogelijk maakt met behulp van microcomputed tomografie.

Video: Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Biology

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 ...

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

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

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

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

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

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

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in every tissue to be identified and characterized. Our laboratory is actively working to make technological advances in X-ray micro-computed tomography (micro-CT) that will bring the diagnostic power of histology to the study of full tissue volumes at cellular resolution (i.e., an X-ray Histo-tomography modality). Toward this end, we have made targeted improvements to the sample preparation pipeline. One key optimization, and the focus of the present work, is a straightforward method for rigid embedding of fixed and stained millimeter-scale samples. Many of the published methods for sample immobilization and correlative micro-CT imaging rely on placing the samples in paraffin wax, agarose, or liquids such as alcohol. Our approach extends this work with custom procedures and the design of a 3-dimensional printable apparatus to embed the samples in an acrylic resin directly into polyimide tubing, which is relatively transparent to X-rays. Herein, sample preparation procedures are described for the samples from 0.5 to 10 mm in diameter, which would be suitable for whole zebrafish larvae and juveniles, or other animals and tissue samples of similar dimensions. As proof of concept, we have embedded the specimens from Danio, Drosophila, Daphnia, and a mouse embryo; representative images from 3-dimensional scans for three of these samples are shown. Importantly, our methodology leads to multiple benefits including rigid immobilization, long-term preservation of laboriously-created resources, and the ability to re-interrogate samples.

We developed protocols and designed a custom apparatus to enable embedding of millimeter-scale specimens. We present sample preparation procedures with an emphasis on embedding in acrylic resin and polyimide tubing to achieve rigid immobilization and long-term storage of specimens for the interrogation of tissue architecture and cell morphology by micro-CT.

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in ...

Bioengineering

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Neuroscience

Imaging Through the Pupal Case of Drosophila melanogaster

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

Imaging Through the Pupal Case of Drosophila melanogaster

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

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

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy (OCM)

Longitudinal study of the heartbeat in small animals contributes to understanding structural and functional changes during heart development. Optical coherence microscopy (OCM) has been demonstrated to be capable of imaging small animal hearts with high spatial resolution and ultrahigh imaging speed. The high image contrast and noninvasive properties make OCM ideal for performing longitudinal studies without requiring tissue dissections or staining. Drosophila has been widely used as a model organism in cardiac developmental studies due to its high number of orthologous human disease genes, its similarity of molecular mechanisms and genetic pathways with vertebrates, its short life cycle, and its low culture cost. Here, the experimental protocols are described for the preparation of Drosophila and optical imaging of the heartbeat with a custom OCM system throughout the life cycle of the specimen. By following the steps provided in this report, transverse M-mode and 3D OCM images can be acquired to conduct longitudinal studies of the Drosophila cardiac morphology and function. The en face and axial sectional OCM images and the heart rate (HR) and cardiac activity period (CAP) histograms, were also shown to analyze the heart structural changes and to quantify the heart dynamics during Drosophila metamorphosis, combined with the videos constructed with M-mode images to trace cardiac activity intuitively. Due to the genetic similarity between Drosophila and vertebrates, longitudinal study of heart morphology and dynamics in fruit flies could help reveal the origins of human heart diseases. The protocol here would provide an effective method to perform a wide range of studies to understand the mechanisms of cardiac diseases in humans.

Drosophila Preparation and Longitudinal Imaging of Heart Function In Vivo Using Optical Coherence Microscopy OCM

Here, the experimental protocols are described for preparing Drosophila at different developmental stages and performing longitudinal optical imaging of Drosophila heartbeats using a custom optical coherence microscopy (OCM) system. The cardiac morphological and dynamical changes can be quantitatively characterized by analyzing the heart structural and functional parameters from OCM images.

Longitudinal study of the heartbeat in small animals contributes to understanding structural and functional changes during heart development. Optical ...

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique cristae-rich double membrane ultrastructure of this organelle is elegantly arranged to carry out multiple functions by partitioning biomolecules. Mitochondrial ultrastructure is intimately linked with various functions; however, the fine details of these structure-function relationships are only beginning to be described. Here, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle. Serial-section electron tomography may be adapted to study any cellular structure in three-dimensions.

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

In this protocol, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle.

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique ...

Whole Animal Imaging of Drosophila melanogaster using Microcomputed Tomography

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