We thank members of the Melkani lab for their help with valuable feedback for developing the protocol. This work was supported by National Institutes of Health (NIH) grants AG065992 and RF1NS133378 to G.C.M. This work is also supported by UAB Startup funds 3123226 and 3123227 to G.C.M.

1. Preparation of equipment
<ol>
	<li>Ensure that the cryostat is powered on and set to -20 &#176;C. Allow for adequate time to elapse for the temperature to reach this point. If the machine is at ambient temperature, cooling to optimal cutting temperature can take around 5 h.</li>
	<li>Power on the slide warmer or incubator, ensuring it is set to 37 &#176;C. Ensure that slides have enough time to reach the desired temperature. 
	NOTE: At this stage, labeled slides can be placed on the warmer or incubator and left indefinitely until sectioning.</li>
	<li>Ensure that dry ice is ready for use. Ice can be removed from storage at &#8211; 80 &#176;C once step 5 has been reached.</li>
</ol>
2. Preparation of solutions
<ol>
	<li>Prepare 50 mL of PBS. Prepare 10 mL of 10 mM Ethelyne glycol tetraacetic acid in PBS. Prepare a solution of 4% Paraformaldehyde in PBS where the final volume is equal to 100 &#181;L times the number of experimental groups.</li>
	<li>Prepare a cleaning solution of 70% ethanol in water in a&#160;spray bottle. Prepare all fluorescent staining solutions. 
	NOTE: This protocol will detail how to perform simple&#160;staining using simple fluorescent stains or dyes. In this&#160;case, we will describe the use of 1x Phalloidin 594 at&#160;a 1:250 ratio in PBS. Fluorescent stain concentrations&#160;will vary depending on the reagent and experimental&#160;goals. Preliminary testing should be done to find the&#160;appropriate concentration. Antibody staining will not be&#160;detailed here but should be applicable using conventional&#160;staining methods.</li>
</ol>
3. Collection of tissue
<ol>
	<li>Anesthetize 5-10 flies using a CO2 fly mat and position the flies within view under the microscope at 1x-2x magnification. The CO2 regulator should register between 1 and 5 kPa to the CO2 mat. 
	NOTE: Other methods for anesthetizing are available (e.g., FlyNap2), and these methods do not significantly affect results.</li>
	<li>Using spring scissors, decapitate, clip the wings, and remove the legs from 5 to 10 flies. What remains is the thorax and abdomen, together will be referred to as the body. 
	NOTE: To align fly bodies appropriately in step 5, any remaining wing or leg lengths must not interfere with a flat alignment along the bottom of the mold. Accomplish this by clipping the wings and legs as close to the thorax as possible.</li>
	<li>Using a brush, gently place the bodies into labeled 1.5 mL tubes on ice until all experimental groups have been collected.</li>
</ol>
4. Fixation of whole tissue
<ol>
	<li>After removing the tubes from the ice, add 100 &#181;L of 10 mM EGTA, pH 8, into each tube to relax the heart tissue within the abdomen. Ensure that all bodies are submerged for 10 min and placed on the orbital shaker.&#160;Table-top centrifuge can be used to facilitate the bodies to submerge.</li>
	<li>Remove and discard the solution in the tube. Pipette 100 &#181;L of PBS in each tube to wash tissue for 10 min, placing on the orbital shaker. Remove and discard after the time has elapsed.</li>
	<li>Pipette 100 &#181;L of 4% Paraformaldehyde in PBS solution into each tube. Ensure that all bodies are submerged in the solution.</li>
	<li>Place on an orbital shaker for 15 min, setting the speed to around 100 units. Ensure that the setting does not agitate the tissues out of contact with the solution.</li>
	<li>After this time has elapsed, discard the PFA and replace it with 1x PBS for 10 min, again ensuring that all bodies are contacting the solution. Return the tubes to the orbital shaker at a similar speed as in step 4.4 for each remaining wash.</li>
	<li>Discarding the previous solution each time, wash the tissue with PBS 2x for 10 min each, for a total of 3 washes.</li>
	<li>After the final wash is discarded, transfer bodies to a 10% sucrose in PBS solution, ensuring that they contact the solution within the tube. For optimal saturation, leave them overnight or long enough such that the bodies sink to the bottom. 
	NOTE: Overnight cryoprotection is preferred, but if same-day imaging is necessary, allow for as long of time as possible for the cryoprotectant to infiltrate the tissue.</li>
</ol>
5. Mold preparation
<ol>
	<li>At this point, retrieve the dry ice from storage and select a few pieces for placement in a small metal or plastic tray. The dimensions of the tray should be such that the mold can be inserted, placing it directly on top of a single, large, and preferably flat, piece of dry ice.</li>
	<li>Fill a labeled mold approximately 50% with Optimal Cutting Temperature (OCT) compound. Try to reduce the introduction of bubbles by letting the compound flow slowly.</li>
	<li>Using a brush, carefully place the bodies on the surface of the OCT within the mold. Multiple experimental groups should be distinctly separated at this stage. 
	NOTE: It is important to prevent dilution of the OCT compound through contamination with the sucrose solution by the brush. In addition, placement deep into the OCT using the brush can create many air bubbles. Avoid these mistakes to prevent sectioning issues later.</li>
	<li>Using forceps, push each body to the bottom of the mold with the dorsal wall of the thorax and abdomen, generally along the bottom. Then, adjust the body such that the abdomen is prioritized as being flat along the bottom of the mold. Align all bodies in the X, Y, and Z dimensions such that each section will contain all subjects simultaneously.</li>
	<li>There may be a curvature of the abdomen, particularly in the males; flatten this out to the best of the technician&#39;s ability. This is essential for longer sections to be collected in the coronal plane. Avoid puncturing or crushing the bodies against the bottom of the mold during this process. Work methodically to align each subject correctly.</li>
	<li>Once all subjects are in place, take the mold and place the bottom directly on top of a piece of dry ice. Check to see if the OCT begins to freeze after a few moments post-contact. Allow this to continue until all subjects are enveloped in the frozen portion of the mold.</li>
	<li>Once this stage is reached, transfer the mold to -20 &#176;C to freeze throughout. Fill the remaining portion of the mold when the first portion is completely frozen. The molds can then be stored at -80 &#176;C for later use by wrapping them in aluminum foil or cut immediately. 
	NOTE: Avoid open exposure to the air at -20 &#176;C and -80 &#176;C between the two freezing stages. This can cause moisture to develop between the layers.</li>
</ol>
6. Cryosectioning of molds
NOTE: It is generally advisable to prepare and cut a blank mold before cutting experimental group molds. This allows us to ensure the proper functionality of the wheel, blade, and anti-roll glass immediately before sectioning tissue. In addition, allow any molds coming from colder storage to acclimate in the cryostat for 30 min.
<ol>
	<li>Attach the chuck bit to the mold by applying a generous amount of OCT to the block and placing the bit on top, pressing it flat. Allow this to freeze completely in the cryostat, usually within 5 min.</li>
	<li>Release the bit and OCT block from the mold and place it into the chuck, ensuring that the block remains properly oriented from top to bottom. Tighten the chuck key until the bit is secure.</li>
	<li>Align the block with the blade using the adjustment knobs and chuck depth controls. Set the section width to 20 &#181;M or lower. 
	NOTE: Thinner sections are more desirable to increase the number of possible sections collected. The first 2-6 sections of each subject should contain the desired unobstructed portions of the heart which is a function of the orientation and section width. One must try and get the orientation of the chuck right before cutting the important 2-6 sections.</li>
	<li>Cut each slice using slow but consistent motion, allowing the anti-roll glass to capture each slice. Collect sections using warmed slides. Allow slides to dry at room temperature for at least 30 min but no more than 1 hour before the first round of washes.</li>
</ol>
7. Fluorescence staining
<ol>
	<li>Immediately following the drying period, remove excess OCT surrounding the sections using a razor blade. Be careful to avoid encountering the tissue at this stage. If the slide is intended to be stained at a later time point, store them at -80 &#176;C.</li>
	<li>Outline this border on each slide using a hydrophobic marker. This border will serve to keep wash and stain solutions on the tissue. 
	NOTE: This protocol outlines how to perform staining and washing using a pipette method, slide buckets and racks may also be used. Simply substitute the pipetting of solutions on top of the slides with the submerging of slides into each of the fresh solutions. As we have done recently for brain imaging17, an immunostaining procedure can be applied to cardiac staining as well.</li>
	<li>Wash all slides 3x for 5 min each with PBS by pipetting gently on top of the slide, avoiding pipetting directly on top of tissue when possible. Perform this using the 1000 &#181;L pipette and associated tips. Try not to rapidly jet the solution as this can cause tissues to release from the surface of the slide.</li>
	<li>Discard the final wash and pipette the fluorescence stain solution (1:250, 1x Phalloidin) onto the tissues. Allow this to incubate for 1 h at room temperature in the dark.</li>
	<li>After this point subjects are light sensitive. Perform all subsequent steps in the dark or by covering the slides. Discard the fluorescence stain, and, using the 1000 &#181;L pipette, wash 3x for 5 min with PBS.</li>
</ol>
8. Mounting and preparation for imaging
<ol>
	<li>Discard the final wash, leaving a small amount of PBS on the slide. Pipette 3-5 drops of mounting media onto the slide using the 1000 &#181;L pipette, avoiding direct deployment on tissue.</li>
	<li>Place the coverslip on the slide such that the production of air bubbles is minimized. For ease, use forceps to place the coverslip at one edge and gradually lower the other side until it is flush with the slide. Handle the freshly mounted slides with care and store them flat for at least 24 h &#160;at 4 &#176;C.</li>
	<li>Seal slides using nail polish, or if the mounting media is hardening, avoid this. Nail polish containing neon or glitter should not be used. Depending on the fluorescent stain, perform imaging immediately or store slides can be stored for later capture.</li>
</ol>
9. Imaging
NOTE: An Olympus BX 63 microscope with 10x, 40x, and 60x lenses was used for image capture. The appropriate filters for DAPI, Lipid Spot 488, and Phalloidin 594 were used as well.
<ol>
	<li>Before imaging, inspect the slides and clean them using a tissue wipe and 70% ethanol in water solution. If slides are imaged immediately following mounting, take extra care when cleaning to avoid disturbing the coverslip.</li>
	<li>Select proper exposure times for each channel based on the brightest fluorescing subjects within the overall experiment. Avoid oversaturation while generating as bright an image as possible and reducing the background. Quickly survey each experimental group with a live view of each channel to ensure that exposures are set appropriately.</li>
	<li>Focus on each subject, using the same channel across all subjects for consistency. Then, capture each subject. Capture all experimental groups used for comparison on the same day to avoid differences in fluorescent decay between compared groups.</li>
	<li>After imaging, store slides in the dark at 4 &#176;C for later image recapture.</li>
</ol>

<ol>
	<li>Baker, K. D., Thummel, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetic+larvae+and+obese+flies-emerging+studies+of+metabolism+in+Drosophila.">Diabetic larvae and obese flies-emerging studies of metabolism in Drosophila.</a> Cell Metab. 6 (4), 257-266 (2007).</li><li>Bhide, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increasing+autophagy+and+blocking+Nrf2+suppress+laminopathy-induced+age-dependent+cardiac+dysfunction+and+shortened+lifespan.">Increasing autophagy and blocking Nrf2 suppress laminopathy-induced age-dependent cardiac dysfunction and shortened lifespan.</a> Aging Cell. 17 (3), e12747 (2018).</li><li>Bier, E., Bodmer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila,+an+emerging+model+for+cardiac+disease.">Drosophila, an emerging model for cardiac disease.</a> Gene. 342 (1), 1-11 (2004).</li><li>Bodmer, R., Venkatesh, T. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heart+development+in+Drosophila+and+vertebrates:+conservation+of+molecular+mechanisms.">Heart development in Drosophila and vertebrates: conservation of molecular mechanisms.</a> Dev Genet. 22 (3), 181-186 (1998).</li><li>Gill, S., Le, H. D., Melkani, G. C., Panda, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-restricted+feeding+attenuates+age-related+cardiac+decline+in+Drosophila.">Time-restricted feeding attenuates age-related cardiac decline in Drosophila.</a> Science. 347 (6227), 1265-1269 (2015).</li><li>Melkani, Y., Pant, A., Guo, Y., Melkani, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+assessment+of+cardiac+dynamics+in+aging+and+dilated+cardiomyopathy+Drosophila+models+using+machine+learning.">Automated assessment of cardiac dynamics in aging and dilated cardiomyopathy Drosophila models using machine learning.</a> Commun Biol. 7 (1), 702 (2024).</li><li>Ocorr, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=KCNQ+potassium+channel+mutations+cause+cardiac+arrhythmias+in+Drosophila+that+mimic+the+effects+of+aging.">KCNQ potassium channel mutations cause cardiac arrhythmias in Drosophila that mimic the effects of aging.</a> Proc Natl Acad Sci U S A. 104 (10), 3943-3948 (2007).</li><li>Piazza, N., Wessells, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+models+of+cardiac+disease.">Drosophila models of cardiac disease.</a> Prog Mol Biol Transl Sci. 100, 155-210 (2011).</li><li>Wolf, M. J., Rockman, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila,+genetic+screens,+and+cardiac+function.">Drosophila, genetic screens, and cardiac function.</a> Circ Res. 109 (7), 794-806 (2011).</li><li>Lee, J. H., Bassel-Duby, R., Olson, E. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heart-and+muscle-derived+signaling+system+dependent+on+MED13+and+Wingless+controls+obesity+in+Drosophila.">Heart-and muscle-derived signaling system dependent on MED13 and Wingless controls obesity in Drosophila.</a> Proc Natl Acad Sci. 111 (26), 9491-9496 (2014).</li><li>Adams, M. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+genome+sequence+of+Drosophila+melanogaster.">The genome sequence of Drosophila melanogaster.</a> Science. 287 (5461), 2185-2195 (2000).</li><li>Rotstein, B., Paululat, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=On+the+morphology+of+the+Drosophila+heart.">On the morphology of the Drosophila heart.</a> J Cardiovasc Dev Dis. 3 (2), 15 (2016).</li><li>Alayari, N. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+labeling+of+Drosophila+heart+structures.">Fluorescent labeling of Drosophila heart structures.</a> J Vis Exp. (32), e1423 (2009).</li><li>Guo, Y., Abou Daya, F., Le, H. D., Panda, S., Melkani, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diurnal+expression+of+Dgat2+induced+by+time&#8208;restricted+feeding+maintains+cardiac+health+in+the+Drosophila+model+of+circadian+disruption.">Diurnal expression of Dgat2 induced by time&#8208;restricted feeding maintains cardiac health in the Drosophila model of circadian disruption.</a> Aging Cell. 23 (7), e14169 (2024).</li><li>Melkani, G. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Huntington's+disease-induced+cardiac+amyloidosis+is+reversed+by+modulating+protein+folding+and+oxidative+stress+pathways+in+the+Drosophila+heart.">Huntington's disease-induced cardiac amyloidosis is reversed by modulating protein folding and oxidative stress pathways in the Drosophila heart.</a> PLoS Genet. 9 (12), e1004024 (2013).</li><li>Hartley, P. S., Coward, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+podocyte+biology+using+Drosophila+nephrocytes.">Modeling podocyte biology using Drosophila nephrocytes.</a> Methods Mol Biol. 2067, 11-24 (2020).</li><li>Watson, J. R., Roth, J., Melkani, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+cryosectioning+of+Drosophila+heads+for+enhanced+Brain+Fluorescence+Staining+and+Immunostaining.">Direct cryosectioning of Drosophila heads for enhanced Brain Fluorescence Staining and Immunostaining.</a> J. Vis. Exp. , e67791 (2025).</li><li>Kelso, R. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flytrap,+a+database+documenting+a+GFP+protein&#8208;trap+insertion+screen+in+Drosophila+melanogaster.">Flytrap, a database documenting a GFP protein&#8208;trap insertion screen in Drosophila melanogaster.</a> Nuc Acids Res. 32 (suppl_1), D418-D420 (2004).</li><li>Dietzl, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+genome-wide+transgenic+RNAi+library+for+conditional+gene+inactivation+in+Drosophila.">A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila.</a> Nature. 448 (7150), 151-156 (2007).</li><li>Mery, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+muscle+LIM+protein,+Mlp84B,+is+essential+for+cardiac+function.">The Drosophila muscle LIM protein, Mlp84B, is essential for cardiac function.</a> J Exp Biol. 211 (1), 15-23 (2008).</li></ol>

The authors have nothing to disclose.

We have developed an efficient protocol for preparing a Drosophila cardiac tube for visualization using fluorescent or confocal imaging. This is preceded by a discussion of a commonly used yet time- and labor-intensive method for accessing and monitoring the cytological integrity of Drosophila heart. Our innovative and efficient method offers a concise and efficient alternative to traditional approaches by utilizing direct cryo-embedding, which preserves the structural integrity of the Drosophila</e...

Cardiovascular disease (CVD) is the leading cause of death globally, responsible for approximately 17.9 million deaths each year, accounting for nearly 1/3rd of all global deaths. Drosophila melanogaster (commonly known as fruit fly) has been widely used as a model organism for studying the genetic, cellular, and molecular basis of cardiac development, physiology, metabolism, aging, and cardiomyopathies1,2,3,4,5,6,7,8,9. Drosophila models have also been used to study the role of cardiac muscle in the systemic regulation of obesity10, a major risk factor for cardiovascular morbidity and mortality. Drosophila genome sequencing studies11 have revealed significant conservation of genes in humans, including those associated with developing various organs, including the heart. Among these highly conserved genes, some are involved in cardiac dysfunction, such as cardiomyopathies or channelopathies3. The recent development of effective techniques to study cardiac performance has expanded the model&#39;s applications to explore long-term changes in adult cardiac physiology due to factors such as exercise, diet, and aging8. However, technical and logistic challenges often hinder the use of this model system. One challenge with using Drosophila in cardiac studies is a precise dissection of the heart in a manner that preserves the cytoarchitecture and myocardial elements.
The Drosophila heart or the dorsal vessel consists of a tube-like structure made up of a single layer of cardiomyocytes, pericardial cells positioned along the heart wall, supported by alary muscles, and in adults, accompanied by a layer of ventral longitudinal muscle cell12. Accurate dissection to access these delicate structures is a time- and labor-intensive process. The current standard involves technically challenging dissection and capillary vacuum suction, requiring advanced training and motor skills13,14,15,16. Typically, the dissection starts by&#160;incising the ventral body wall, and the challenges present&#160;themselves quickly with the minute anatomy of the heart, its&#160;fragile structure, and difficult-to-access dorsal location.&#160;This combined with traditional dissection techniques, allows for precise analysis of heart structure and function, providing an improved tool for studying cardiovascular diseases&#160;in Drosophila13. For example, using this, Alayari et al. provided a protocol for fluorescently labeling Drosophila heart structures, facilitating the visualization of cardiac morphology and structure. Despite these efforts, traditional heart dissection and staining face several challenges, including the difficulty of maintaining tissue integrity and the specialized training required for effective heart staining.
The method offers an innovative solution to this problem by replacing the whole procedure with a simpler protocol that utilizes cryoembedding of Drosophila thorax and abdomen followed by immunostaining and fluorescence imaging. This easy-to-learn approach ensures faster and more&#160;straightforward visualization of cardiac structures with greater reproducibility. Additionally, we describe a simple method involving dry ice that ensures uniform alignment of the Drosophila abdominal cuticle on the same z-plane streamlining the cryosectioning step downstream. We demonstrate the effectiveness of this protocol in detecting important cardiac markers, heart morphology, and cellular organization with immunofluorescent as well as confocal microscopy. The ease and high efficacy of this approach is particularly helpful for high-throughput Drosophila-based cardiac studies.

<table><tbody><tr><td>1000 &amp;micro;L Pipette</td><td>Eppendorf</td><td>3123000063</td><td></td></tr><tr><td>1000 &amp;micro;L Pipette Tips</td><td>Olympus Plastics</td><td>23-165R</td><td></td></tr><tr><td>10X Phosphate Buffered Saline (PBS)</td><td>Fisher</td><td>J62036.K7</td><td>ph=7.4</td></tr><tr><td>200 Proof Ethanol</td><td>Decon Laboratories</td><td>64-17-5</td><td></td></tr><tr><td>20X Tris Buffered Saline</td><td>Thermo Scientific</td><td>J60877.K2</td><td>pH=7.4</td></tr><tr><td>Anti-Roll Glass</td><td>IMEB</td><td>AR-14047742497</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Fisher</td><td>9048-46-8</td><td></td></tr><tr><td>Centrifuge Tubes 1.5 mL</td><td>Fisher</td><td>05-408-129</td><td></td></tr><tr><td>Charged Slides</td><td>Globe Scientific</td><td>1415-15</td><td></td></tr><tr><td>Cryosectioning Molds</td><td>Fisher</td><td>2363553</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM 3050 S</td><td></td></tr><tr><td>Cryostat Blades</td><td>C.L. Sturkey</td><td>DT554N50</td><td></td></tr><tr><td>Dry Ice</td><td></td><td></td><td></td></tr><tr><td>Fine Forceps</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>Fly Pad</td><td>Tritech Research</td><td>MINJ-DROS-FP</td><td></td></tr><tr><td>Hardening mounting Media with Dapi</td><td>Vectashield</td><td>H-1800</td><td></td></tr><tr><td>Kimwipes</td><td>Kimtech</td><td>34120</td><td></td></tr><tr><td>Microscope</td><td>Olympus</td><td>SZ61</td><td></td></tr><tr><td>Optimal Cutting Temperature Compound</td><td>Fisher</td><td>4585</td><td></td></tr><tr><td>Paraformaldehyde 20%</td><td>Electron Microscopy Sciences</td><td>15713</td><td></td></tr><tr><td>Phalloidin 594</td><td>Abnova</td><td>U0292</td><td></td></tr><tr><td>Razor Blades</td><td>Gravey</td><td>#40475</td><td></td></tr><tr><td>Spring Scissors</td><td>Fine Science Tools</td><td>15000-10</td><td></td></tr><tr><td>Sucrose</td><td>Fisher</td><td>S5-500</td><td></td></tr></tbody></table>

novel and efficient method for drosophila heart fluorescence staining with cryosectioning

Drosophila heart models are widely employed in studying cardiac aging and modeling&#160;human cardiac diseases. However, the dissection of Drosophila hearts before imaging is a meticulous time-intensive process that requires advanced training and motor skills. To address these challenges, we present an innovative protocol that utilizes cryosectioning for the fluorescence imaging of Drosophila heart tissue. The protocol has been demonstrated in imaging the adult Drosophila heart but could be adapted for developmental stages. The method enhances both the efficiency and accessibility of fluorescence staining while preserving the integrity of the tissue. This protocol simplifies the process without compromising the quality of imaging, thereby reducing the dependency on technicians with highly developed training and motor skills. Specifically, we replace complex techniques, such as capillary vacuum suction, with more straightforward methods like tissue embedding. This approach allows for the visualization of cardiac structures with greater ease and reproducibility. We demonstrate the utility of this protocol by effectively detecting key cardiac markers and&#160;achieving high-resolution fluorescence and immunostaining imaging that unveils intricate details of heart morphology and cellular organization. This method provides a robust and accessible tool for researchers exploring Drosophila cardiac biology, facilitating detailed analyses of heart development, function, and disease models.

The method described above facilitates the study of the Drosophila heart using fluorescence imaging without tedious dissection. This is the main benefit of this method, as the conventional method of heart dissection requires the development of complex motor skills. Illustrated in Figure 1, the method is more approachable than heart dissection for new researchers and allows for experiment flexibility. Alternatively, using -80 &#176;C storage during the OCT mold stage, specimens can b...

The Drosophila heart sectioning and fluorescence imaging protocol simplifies studying heart structure and pathologies. This approach involves straightforward sectioning, staining, and imaging, bypassing the technical expertise needed for traditional dissection. It enhances accessibility, making Drosophila a more widely usable model for cardiac-related research within the broader scientific community.

Novel And Efficient Method for Drosophila Heart Fluorescence Staining with Cryosectioning

Drosophila heart models are widely employed in studying cardiac aging and modeling&#160;human cardiac diseases. However, the dissection of Drosophila ...

novel-efficient-method-for-drosophila-heart-fluorescence-staining

Universidad de Cartagena UDEC

Research

JoVE Journal

Genetics

180 Views.  The University of Alabama at Birmingham. The Drosophila heart sectioning and fluorescence imaging protocol simplifies studying heart structure and pathologies. This approach involves straightforward sectioning, staining, and imaging, bypassing the technical expertise needed for traditional dissection. It enhances accessibility, making Drosophila a more widely usable model for cardiac-related research within the broader scientific community.

Video: Novel And Efficient Method for Drosophila Heart Fluorescence Staining with Cryosectioning

Application of MultiColor FlpOut Technique to Study High Resolution Single Cell Morphologies and Cell Interactions of Glia in Drosophila

Cells display different morphologies and complex anatomical relationships. How do cells interact with their neighbors? Do the interactions differ between cell types or even within a given type? What kinds of spatial rules do they follow? The answers to such fundamental questions in vivo have been hampered so far by a lack of tools for high resolution single cell labeling. Here, a detailed protocol to target single cells with a MultiColor FlpOut (MCFO) technique is provided. This method relies on three differently tagged reporters (HA, FLAG and V5) under UAS control that are kept silent by a transcriptional terminator flanked by two FRT sites (FRT-stop-FRT). A heat shock pulse induces the expression of a heat shock-induced Flp recombinase, which randomly removes the FRT-stop-FRT cassettes in individual cells: expression occurs only in cells that also express a GAL4 driver. This leads to an array of differently colored cells of a given cell type that allows the visualization of individual cell morphologies at high resolution. As an example, the MCFO technique can be combined with specific glial GAL4 drivers to visualize the morphologies of the different glial subtypes in the adult Drosophila brain.

Application of MultiColor FlpOut Technique to Study High Resolution Single Cell Morphologies and Cell Interactions of Glia in Drosophila

Cells display different morphologies and establish a variety of interactions with their neighbors. This protocol describes how to reveal the morphology of single cells and to investigate cell-cell interaction by using the well-established Gal4/UAS expression system.

Cells display different morphologies and complex anatomical relationships. How do cells interact with their neighbors? Do the interactions differ between ...

Behavior

Fluorescent Labeling of Drosophila Heart Structures

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

Fluorescent Labeling of Drosophila Heart Structures

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

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

Biology

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and apoptosis, development and ageing, to neuronal synaptic plasticity and regeneration have been associated with Ryanodine receptors (RyRs). Despite the importance of calcium signaling, the exact mechanism of its function in early development is unclear. As an organism with a short gestational period, the embryos of Drosophila melanogaster are prime study subjects for investigating the distribution and localization of CICR associated proteins and their regulators during development. However, because of their lipid-rich embryos and chitin-rich chorion, their utility is limited by the difficulty of mounting embryos on glass surfaces. In this work, we introduce a practical protocol that significantly enhances the attachment of Drosophila embryo onto slides and detail methods for successful histochemistry, immunohistochemistry, and in-situ hybridization. The chrome alum gelatin slide-coating method and embryo pre-embedding method dramatically increases the yield in studying Drosophila embryo protein and RNA expression. To demonstrate this approach, we studied DmFKBP12/Calstabin, a well-known regulator of RyR during early embryonic development of Drosophila melanogaster. We identified DmFKBP12 in as early as the syncytial blastoderm stage and report the dynamic expression pattern of DmFKBP12 during development: initially as an evenly distributed protein in the syncytial blastoderm, then preliminarily localizing to the basement layer of the cortex during cellular blastoderm, before distributing in the primitive neuronal and digestion architecture during the three-gem layer phase in early gastrulation. This distribution may explain the critical role RyR plays in the vital organ systems that originate in from these layers: the suboesophageal and supraesophageal ganglion, ventral nervous system, and musculoskeletal system.

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Here, we describe a protocol for detection and localization of Drosophila embryo protein and RNA from collection to pre-embedding and embedding, immunostaining, and mRNA in situ hybridization.

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and ...

Developmental Biology

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 ...

Neuroscience

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

DNA in situ hybridization (DNA ISH) is a commonly used method for mapping sequences to specific chromosome regions. This approach is particularly effective at mapping highly repetitive sequences to heterochromatic regions, where computational approaches face prohibitive challenges. Here we describe a streamlined protocol for DNA ISH that circumvents formamide washes that are standard steps in other DNA ISH protocols. Our protocol is optimized for hybridization with short single strand DNA probes that carry fluorescent dyes, which effectively mark repetitive DNA sequences within heterochromatic chromosomal regions across a number of different insect tissue types. However, applications may be extended to use with larger probes and visualization of single copy (non-repetitive) DNA sequences. We demonstrate this method by mapping several different repetitive sequences to squashed chromosomes from Drosophila melanogaster neural cells and Nasonia vitripennis spermatocytes. We show hybridization patterns for both small, commercially synthesized probes and for a larger probe for comparison. This procedure uses simple laboratory supplies and reagents, and is ideal for investigators who have little experience with performing DNA ISH.

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

Here, we present a simple method for performing fluorescence DNA in situ hybridization (DNA ISH) to visualize repetitive heterochromatic sequences on slide-mounted chromosomes. The method requires minimal reagents and it is versatile for use with short or long probes, different tissues, and detection with fluorescence or non-fluorescence-based signals.

DNA in situ hybridization (DNA ISH) is a commonly used method for mapping sequences to specific chromosome regions. This approach is particularly ...

Direct Cryosectioning of Drosophila Heads for Enhanced Brain Fluorescence Staining and Immunostaining

Immunostaining Drosophila melanogaster brains is essential for exploring the mechanisms behind complex behaviors, neural circuits, and protein expression patterns. Traditional methods often involve challenges such as performing complex dissection, maintaining tissue integrity, and visualizing specific expression patterns during high-resolution imaging. We present an optimized protocol that combines cryosectioning with fluorescence staining and immunostaining. This method improves tissue preservation and signal clarity and reduces the need for laborious dissection for Drosophila brain imaging. The method entails rapid dissection, optimal fixation, cryoprotection, and cryosectioning, followed by fluorescent staining and immunostaining. The protocol significantly reduces tissue damage, enhances antibody penetration, and yields sharp, well-defined images. We demonstrate the effectiveness of this approach by visualizing specific neural populations and synaptic proteins with high fidelity. This versatile method allows for the analysis of various protein markers in the adult brain across multiple z-planes and can be adapted for other tissues and model organisms. The protocol provides a reliable and efficient tool for researchers conducting high-quality immunohistochemistry in Drosophila neurobiology studies. This method's detailed visualization facilitates comprehensive analysis of neuroanatomy, pathology, and protein localization, making it particularly valuable for neuroscience research.

This study presents a simplified protocol for tissue processing involving decapitation, fixation, cryosectioning, fluorescence staining, immunostaining, and imaging, which can be extended to confocal and multiphoton imaging. The method maintains efficacy comparable to complex dissections, bypassing the need for advanced motor skills. Quantitative image analysis provides extensive investigative potential.

Immunostaining Drosophila melanogaster brains is essential for exploring the mechanisms behind complex behaviors, neural circuits, and protein expression ...

Preparation of Drosophila Heart: A Technique to Study Heart Physiology in the Adult Fly

Source: Vogler, G., Ocorr, K. <a href="https://jove.unicartagenaproxy.elogim.com/video/1425/" target="_blank">Visualizing the Beating Heart in Drosophila</a>. J. Vis. Exp. (2009).
This video describes the fly&#39;s adult heart, or otherwise called the Drosophila dorsal vessel. The featured protocol demonstrates a dissection procedure for live heart preparations.

Novel And Efficient Method for Drosophila Heart Fluorescence Staining with Cryosectioning

Summary

Explore More Videos

Application of MultiColor FlpOut Technique to Study High Resolution Single Cell Morphologies and Cell Interactions of Glia in Drosophila

Fluorescent Labeling of Drosophila Heart Structures

Visualizing the Beating Heart in Drosophila

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

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

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

Direct Cryosectioning of Drosophila Heads for Enhanced Brain Fluorescence Staining and Immunostaining

Preparation of Drosophila Heart: A Technique to Study Heart Physiology in the Adult Fly