Novel And Efficient Method for Drosophila Heart Fluorescence Staining with Cryosectioning

Summary

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.

Abstract

Drosophila heart models are widely employed in studying cardiac aging and modeling 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 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.

Introduction

Cardiovascular disease (CVD) is the leading cause of death globally, responsible for approximately 17.9 million deaths each year, accounting for nearly 1/3^rd 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 cardiomyopathies^{1,2,3,4,5,6,7,8,9}. 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'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 skills^13,14,15,16. Typically, the dissection starts by incising the ventral body wall, and the challenges present themselves quickly with the minute anatomy of the heart, its fragile structure, and difficult-to-access dorsal location. This combined with traditional dissection techniques, allows for precise analysis of heart structure and function, providing an improved tool for studying cardiovascular diseases 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 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.

Protocol

1. Preparation of equipment

1. Ensure that the cryostat is powered on and set to -20 °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.
2. Power on the slide warmer or incubator, ensuring it is set to 37 °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.
3. Ensure that dry ice is ready for use. Ice can be removed from storage at – 80 °C once step 5 has been reached.

2. Preparation of solutions

1. 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 µL times the number of experimental groups.
2. Prepare a cleaning solution of 70% ethanol in water in a spray bottle. Prepare all fluorescent staining solutions.
   NOTE: This protocol will detail how to perform simple staining using simple fluorescent stains or dyes. In this case, we will describe the use of 1x Phalloidin 594 at a 1:250 ratio in PBS. Fluorescent stain concentrations will vary depending on the reagent and experimental goals. Preliminary testing should be done to find the appropriate concentration. Antibody staining will not be detailed here but should be applicable using conventional staining methods.

3. Collection of tissue

1. 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.
2. 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.
3. Using a brush, gently place the bodies into labeled 1.5 mL tubes on ice until all experimental groups have been collected.

4. Fixation of whole tissue

1. After removing the tubes from the ice, add 100 µL 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. Table-top centrifuge can be used to facilitate the bodies to submerge.
2. Remove and discard the solution in the tube. Pipette 100 µL of PBS in each tube to wash tissue for 10 min, placing on the orbital shaker. Remove and discard after the time has elapsed.
3. Pipette 100 µL of 4% Paraformaldehyde in PBS solution into each tube. Ensure that all bodies are submerged in the solution.
4. 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.
5. 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.
6. Discarding the previous solution each time, wash the tissue with PBS 2x for 10 min each, for a total of 3 washes.
7. 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.

5. Mold preparation

1. 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.
2. 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.
3. 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.
4. 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.
5. There may be a curvature of the abdomen, particularly in the males; flatten this out to the best of the technician'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.
6. 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.
7. Once this stage is reached, transfer the mold to -20 °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 °C for later use by wrapping them in aluminum foil or cut immediately.
   NOTE: Avoid open exposure to the air at -20 °C and -80 °C between the two freezing stages. This can cause moisture to develop between the layers.

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.

1. 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.
2. 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.
3. Align the block with the blade using the adjustment knobs and chuck depth controls. Set the section width to 20 µM 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.
4. 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.

7. Fluorescence staining

1. 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 °C.
2. 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.
3. 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 µL pipette and associated tips. Try not to rapidly jet the solution as this can cause tissues to release from the surface of the slide.
4. 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.
5. 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 µL pipette, wash 3x for 5 min with PBS.

8. Mounting and preparation for imaging

1. 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 µL pipette, avoiding direct deployment on tissue.
2. 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  at 4 °C.
3. 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.

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.

1. 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.
2. 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.
3. 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.
4. After imaging, store slides in the dark at 4 °C for later image recapture.

Results

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 °C storage during the OCT mold stage, specimens can b...

Discussion

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

Materials

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