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

Research

JoVE Journal

Genetics

290 Views.  The University of Alabama at Birmingham. Het Drosophila-protocol voor hartdoorsnede en fluorescentiebeeldvorming vereenvoudigt het bestuderen van de hartstructuur en pathologieën. Deze aanpak omvat eenvoudige secties, kleuringen en beeldvorming, waarbij de technische expertise die nodig is voor traditionele dissectie wordt omzeild. Het verbetert de toegankelijkheid, waardoor Drosophila een breder bruikbaar model wordt voor hartgerelateerd onderzoek binnen de b...

Video: Nieuwe en efficiënte methode voor Drosophila-hartfluorescentiekleuring met cryosectie 

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

The mitochondrial genome is inherited exclusively through the maternal line. Understanding of how the mitochondrion and its genome are proliferated and transmitted from one generation to the next through the female oocyte is of fundamental importance. Because of the genetic tractability, and the elegant, ordered simplicity by which oocyte development proceeds, Drosophila oogenesis has become an invaluable system for mitochondrial study. An EdU (5-ethynyl-2&#180;-deoxyuridine) labeling method was utilized to detect mitochondrial DNA (mtDNA) replication in Drosophila ovaries. This method is superior to the BrdU (5-bromo-2'-deoxyuridine) labeling method in that it allows for good structural preservation and efficient fluorescent dye penetration of whole-mount tissues.
Here we describe a detailed protocol for labeling replicating mitochondrial DNA in Drosophila adult ovaries with EdU. Some technical solutions are offered to improve the viability of the ovaries, maintain their health during preparation, and ensure high-quality imaging. Visualization of newly synthesized mtDNA in the ovaries not only reveals the striking temporal and spatial pattern of mtDNA replication through oogenesis, but also allows for simple quantification of mtDNA replication under various genetic and pharmacological perturbations.

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

Drosophila oogenesis continues to be exceptionally useful in the study of mitochondrial proliferation and inheritance. This manuscript describes a detailed protocol used to label the replicating mitochondrial DNA (mtDNA) in Drosophila adult ovaries with 5-ethynyl-2&#180;-deoxyuridine (EdU), which facilitates uncovering mechanisms associated with mitochondrial inheritance that were previously debatable.

The mitochondrial genome is inherited exclusively through the maternal line. Understanding of how the mitochondrion and its genome are proliferated and ...

A Fast Silver Staining Protocol Enabling Simple and Efficient Detection of SSR Markers using a Non-denaturing Polyacrylamide Gel

Simple Sequence Repeat (SSR) is one of the most effective markers used in plant and animal genetic research and molecular breeding programs. Silver staining is a widely used method for the detection of SSR markers in a polyacrylamide gel. However, conventional protocols for silver staining are technically demanding and time-consuming. Like many other biological laboratory techniques, silver staining protocols have been steadily optimized to improve detection efficiency. Here, we report a simplified silver staining method that significantly reduces reagent costs and enhances the detection resolution and picture clarity. The new method requires two major steps (impregnation and development) and three reagents (silver nitrate, sodium hydroxide, and formaldehyde), and only 7 min of processing for a non-denaturing polyacrylamide gel. Compared to previously reported protocols, this new method is easier, quicker and uses fewer chemical reagents for SSR detection. Therefore, this simple, low-cost, and effective silver staining protocol will benefit genetic mapping and marker-assisted breeding by a quick generation of SSR marker data.

Here, we report a simple and low-cost silver staining protocol which requires only three reagents and 7 min of processing, and is suitable for fast generation of high-quality SSR data in the genetic analysis.

Simple Sequence Repeat (SSR) is one of the most effective markers used in plant and animal genetic research and molecular breeding programs. Silver ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

Lentiviral Mediated Production of Transgenic Mice: A Simple and Highly Efficient Method for Direct Study of Founders

For almost 40 years, pronuclear DNA injection represents the standard method to generate transgenic mice with random integration of transgenes. Such a routine procedure is widely utilized throughout the world and its main limitation resides in the poor efficacy of transgene integration, resulting in a low yield of founder animals. Only few percent of animals born after implantation of injected fertilized oocytes have integrated the transgene. In contrast, lentiviral vectors are powerful tools for integrative gene transfer and their use to transduce fertilized oocytes allows highly efficient production of founder transgenic mice with an average yield above 70%. Furthermore, any mouse strain can be used to produce transgenic animal and the penetrance of transgene expression is extremely high, above 80% with lentiviral mediated transgenesis compared to DNA microinjection. The size of the DNA fragment that can be cargo by the lentiviral vector is restricted to 10 kb and represents the major limitation of this method. Using a simple and easy to perform injection procedure beneath the zona pellucida of fertilized oocytes, more than 50 founder animals can be produced in a single session of microinjection. Such a method is highly adapted to perform, directly in founder animals, rapid gain and loss of function studies or to screen genomic DNA regions for their ability to control and regulate gene expression in vivo.

Here, we present a protocol to promote transgene integration and production of founder transgenic mice with high efficacy by a simple injection of a lentiviral vector in the perivitelline space of a fertilized oocyte.

For almost 40 years, pronuclear DNA injection represents the standard method to generate transgenic mice with random integration of transgenes. Such a ...

A Fluorescence-based Method to Study Bacterial Gene Regulation in Infected Tissues

Bacterial virulence genes are often regulated at the transcriptional level by multiple factors that respond to different environmental signals. Some factors act directly on virulence genes; others control pathogenesis by adjusting the expression of downstream regulators or the accumulation of signals that affect regulator activity. While regulation has been studied extensively during in vitro growth, relatively little is known about how gene expression is adjusted during infection. Such information is important when a particular gene product is a candidate for therapeutic intervention. Transcriptional approaches like quantitative, real-time RT-PCR and RNA-Seq are powerful ways to examine gene expression on a global level but suffer from many technical challenges including low abundance of bacterial RNA compared to host RNA, and sample degradation by RNases. Evaluating regulation using fluorescent reporters is relatively easy and can be multiplexed with fluorescent proteins with unique spectral properties. The method allows for single-cell, spatiotemporal analysis of gene expression in tissues that exhibit complex three-dimensional architecture and physiochemical gradients that affect bacterial regulatory networks. Such information is lost when data are averaged over the bulk population. Herein, we describe a method for quantifying gene expression in bacterial pathogens in situ. The method is based on simple tissue processing and direct observation of fluorescence from reporter proteins. We demonstrate the utility of this system by examining the expression of Staphylococcus aureus thermonuclease (nuc), whose gene product is required for immune evasion and full virulence ex vivo and in vivo. We show that nuc-gfp is strongly expressed in renal abscesses and reveal heterogeneous gene expression due in part to apparent spatial regulation of nuc promoter activity in abscesses fully engaged with the immune response. The method can be applied to any bacterium with a manipulatable genetic system and any infection model, providing valuable information for preclinical studies and drug development.

Described here is a method for analyzing bacterial gene expression in animal tissues at a cellular level. This method provides a resource for studying the phenotypic diversity occurring within a bacterial population in response to the tissue environment during an infection.

Bacterial virulence genes are often regulated at the transcriptional level by multiple factors that respond to different environmental signals. Some ...

Sequencing of mRNA from Whole Blood using Nanopore Sequencing

Sequencing in remote locations and resource-poor settings presents unique challenges. Nanopore sequencing can be successfully used under such conditions, and was deployed to West Africa during the recent Ebola virus epidemic, highlighting this possibility. In addition to its practical advantages (low cost, ease of equipment transport and use), this technology also provides fundamental advantages over second-generation sequencing approaches, particularly the very long read length, ability to directly sequence RNA, and real-time availability of data. Raw read accuracy is lower than with other sequencing platforms, which represents the main limitation of this technology; however, this can be partially mitigated by the high read depth generated. Here, we present a field-compatible protocol for sequencing of the mRNAs encoding for Niemann-Pick C1, which is the cellular receptor for ebolaviruses. This protocol encompasses extraction of RNA from animal blood samples, followed by RT-PCR for target enrichment, barcoding, library preparation, and the sequencing run itself, and can be easily adapted for use with other DNA or RNA targets.

Nanopore sequencing is a novel technology that allows cost-effective sequencing in remote locations and resource-poor settings. Here, we present a protocol for sequencing of mRNAs from whole blood that is compatible with such conditions.

Sequencing in remote locations and resource-poor settings presents unique challenges. Nanopore sequencing can be successfully used under such conditions, ...

In Vivo Forward Genetic Screen to Identify Novel Neuroprotective Genes in Drosophila melanogaster

There is much to understand about the onset and progression of neurodegenerative diseases, including the underlying genes responsible. Forward genetic screening using chemical mutagens is a useful strategy for mapping mutant phenotypes to genes among Drosophila and other model organisms that share conserved cellular pathways with humans. If the mutated gene of interest is not lethal in early developmental stages of flies, a climbing assay can be conducted to screen for phenotypic indicators of decreased brain functioning, such as low climbing rates. Subsequently, secondary histological analysis of brain tissue can be performed in order to verify the neuroprotective function of the gene by scoring neurodegeneration phenotypes. Gene mapping strategies include meiotic and deficiency mapping that rely on these same assays can be followed by DNA sequencing to identify possible nucleotide changes in the gene of interest.

In Vivo Forward Genetic Screen to Identify Novel Neuroprotective Genes in Drosophila melanogaster

We present a protocol using a forward genetic approach to screen for mutants exhibiting neurodegeneration in Drosophila melanogaster. It incorporates a climbing assay, histology analysis, gene mapping and DNA sequencing to ultimately identify novel genes related to the process of neuroprotection.

There is much to understand about the onset and progression of neurodegenerative diseases, including the underlying genes responsible. Forward genetic ...

Universal and Efficient Electroporation Protocol for Genetic Engineering of Gastrointestinal Organoids

Electroporation is a common method for transfection with different kinds of molecules by electrical permeabilization of the plasma membrane. With the increasing use of organoids as a culturing method for primary patient material in the last years, efficient transfer methods of components for genetic engineering in this 3D culture system are in need. Especially for organoids, the efficiency of genetic manipulations depends on a successful transfection. Thus, this protocol was developed to facilitate the electroporation of organoids and to prove its universal functionality in different entities. Human colorectal, pancreatic, hepatic and gastric cancer organoids were successfully electroporated with small and large plasmids in comparison. Based on GFP encoding vectors, the transfection efficiency was determined by FACS. No extensive preparation of the cells or special, cost-intensive electroporation buffers are necessary, and the protocol can be performed within one day.

This protocol describes an efficient electroporation method for the transfection of four different gastrointestinal organoid entities with larger plasmids (to the extent of 10 kB). It can be performed within one day and does not need extensive preparation or special, cost-intensive electroporation buffers.

Electroporation is a common method for transfection with different kinds of molecules by electrical permeabilization of the plasma membrane. With the ...

A Screening Method for Identification of Heterochromatin-Promoting Drugs Using Drosophila

Drosophila is an excellent model organism that can be used to screen compounds that might be useful for cancer therapy. The method described here is a cost-effective in vivo method to identify heterochromatin-promoting compounds by using Drosophila. The Drosophila&#39;s DX1 strain, having a variegated eye color phenotype that reflects the extents of heterochromatin formation, thereby providing a tool for a heterochromatin-promoting drug screen. In this screening method, eye variegation is quantified based on the surface area of red pigmentation occupying parts of the eye and is scored on a scale from 1 to 5. The screening method is straightforward and sensitive and allows for testing compounds in vivo. Drug screening using this method provides a fast and inexpensive way for identifying heterochromatin-promoting drugs that could have beneficial effects in cancer therapeutics. Identifying compounds that promote the formation of heterochromatin could also lead to the discovery of epigenetic mechanisms of cancer development.

A Screening Method for Identification of Heterochromatin-Promoting Drugs Using Drosophila

Drosophila is a widely used experimental model suitable for screening drugs with potential applications for cancer therapy. Here, we describe the use of Drosophila variegated eye color phenotypes as a method for screening small-molecule compounds that promote heterochromatin formation.

Drosophila is an excellent model organism that can be used to screen compounds that might be useful for cancer therapy. The method described here is a ...

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, Eastern equine encephalitis, lymphatic filariasis, and Saint Louis encephalitis. Notably, avian malaria has played a major role in the extinction of numerous endemic island bird species, while WNV has become an important vector-borne disease in the United States. To gain further insight into C. quinquefasciatus biology and expand their genetic control toolbox, we need to develop more efficient and affordable methods for genome engineering in this species. However, some biological traits unique to Culex mosquitoes, particularly their egg rafts, have made it difficult to perform microinjection procedures required for genome engineering. To address these challenges, we have developed an optimized embryo microinjection protocol that focuses on mitigating the technical obstacles associated with the unique characteristics of Culex mosquitoes. These procedures demonstrate optimized methods for egg collection, egg raft separation and other handling procedures essential for successful microinjection in C. quinquefasciatus. When coupled with the CRISPR/Cas9 genome editing technology, these procedures allow us to achieve site-specific, efficient and heritable germline mutations, which are required to perform advanced genome engineering and develop genetic control technologies in this important, but currently understudied, disease vector.

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

This protocol describes microinjection procedures for Culex quinquefasciatus embryos that are optimized to work with CRISPR/Cas9 gene editing tools. This technique can efficiently generate site-specific, heritable, germline mutations that can be used for building genetic technologies in this understudied disease vector.

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, ...

Novel And Efficient Method for Drosophila Heart Fluorescence Staining with Cryosectioning

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