Name: Visualizing the Beating Heart in Drosophila (Video) | JoVE Journal
Uploaded: 2009-09-28T21:00:00.000+00:00
Duration: 10 min 15 s
Description: Learn techniques for visualizing the beating heart in larval and adult Drosophila using dissection and immobilization methods. These protocols enable cardiac physiology studies in developmental biology.

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5 minutes

KO is supported by a grant from the American Heart Association.

Before you start
Adult hearts
<ol>
 <li>Freshly prepare artificial hemolymph (AH) solution containing 108mM Na+, 5mM K+, 2mM Ca2+, 8mM MgCl2, 1mM NaH2PO4, 4mM NaHCO3, 10mM sucrose, 5mM trehalose, 5mM HEPES (pH 7.1, all reagents from Sigma Chemicals). The sucrose and trehalose should be added to the AH from refrigerated stock solutions just prior to use in order to prevent bacterial contamination. </li>
 <li>Bring AH to room temperature and oxygenate the solution by air-bubbling for at least 15 min.</li>
 <li>Pull several fine capillaries (e.g. Glass Capillaries, 100ul, VWR) required for removal of fat.</li>
</ol>
Larval hearts
<ol>
 <li>Sylgard-coated coverslips: using a toothpick, transfer about 50-70&mu;l of Sylgard solution onto a 22x22mm coverslip. Let the Sylgard harden at 65&deg;C over night. </li>
 <li>Prepare the Broadie and Bate&#x2019;s buffer (135mM NaCl, 5mM KCl, 4mM MgCl2, 2mM CaCl2, 5mM TES, 36mM Sucrose; pH 7.15. See Broadie and Bate, 1993).</li>
 <li>Prepare the tools required to apply the histoacryl glue. First, pull several fine capillaries (e.g. Science Products GB100T8P). Connect a 1.5ml pipette tip with a small plastic tubing (Tygon, 1/32&#x201D;) by inserting the smaller end into the tube. Insert a capillary with its wide opening to the other end. Load the capillary with Histoacryl glue by dipping the capillary into a drop of glue and gentle suction into the plastic tip. When working with the glue one should have a second glass capillary at hand to remove excessive glue from the tip of the glue capillary. The glue is liquid until it comes into contact with buffered saline where it will harden within a couple of seconds. However, it usually remains plastic long enough to be removed from the glue capillary tip or to while applying it to the specimen.</li>
</ol>
Semi-intact Drosophila Heart from the Adult Fly
<ol>
 <li>Adult Drosophila flies are anesthetized with Fly Nap (Carolina Biological Supply Co.) for 2-5 minutes. Care must be taken not to leave the flies in the Fly Nap for longer exposures. Short term exposure to cold can also be used to anesthetize flies but CO2 is not recommended as it has longer lasting effects on heart rate and rhythmicity.</li>
 <li>Anesthetized flies are placed, dorsal side down, into a Petri dish coated with a thin layer of petroleum jelly. The hydrophobic cuticle in the wings and body reversibly &#x201C;stick&#x201D; to the jelly but the fly can be repositioned if needed. This becomes especially important during subsequent manipulations.</li>
 <li>An initial cut is made using a curved pair of spring scissors (Roboz #5611). The scissor blades are placed under the legs and angled down toward the dorsal surface of the thorax near the neck. The head, ventral nerve cord, and legs are removed with a single cut. The preparation is then submerged in an oxygenated, artificial hemolymph (AH) solution.</li>
 <li>Using spring scissors the posterior tip of the abdomen (comprised of abdominal segments 7 and 8) is removed with a single cut. This provides access for making lateral cuts along both edges of the abdomen and serves to sever the connection between the posterior gut and the abdominal cuticle. The freed flap of ventral abdominal cuticle is then removed.</li>
 <li>In steps 3 and 4 the anterior and posterior connections of the gut are severed so the abdominal organs are now held in place only by tracheols. The gut and other abdominal organs can usually be removed as a single mass using jeweler&#x2019;s forceps (Roboz, #55) and gentle tugging. Removing the internal organs reveals the beating heart tube still attached to the dorsal cuticle surrounded by fat bodies.</li>
 <li>The fat bodies are quite opaque in the light microscope consequently it is often necessary to remove some of this fat in order to see the heart tube clearly. This is accomplished by a liposuction technique using finely drawn glass capillaries. Capillaries are made using a standard pipette puller (E.g. Sutter P-97 Pipette Puller) and are then inserted into plastic tubing (Tygon, 1/16&#x201D;) attached to a vacuum source. This system is used to suction off excess fat from around the heart tube. Suction force is proportional to tip diameter so pipettes with tips larger than 40 microns in diameter should not be used. Extreme care must be taken to avoid touching the heart itself and because the posterior portion of the heart is very fragile that region should be avoided altogether. </li>
 <li>The adult Drosophila heart tube is now exposed and should be beating. If necessary the cuticle and attached heart can be repositioned by gently lifting the cuticle out of the petroleum jelly and carefully tamping it back down, again avoiding any contact with the heart tissue. At this point the solution bathing the preparation should be replaced with fresh AHL. This preparation should be allowed to equilibrate for 20-30 minutes with oxygenation prior to any manipulations. If the preparation is to be maintained for longer than 60 minutes it should be perfused with fresh AHL.</li>
</ol>
Immobilizing Drosophila larvae for optical recording
<ol>
 <li>Place a drop of Histoacryl glue close to each corner of the Sylgard coverslips. Add 50&mu;l of B&amp;B buffer to the glue drops which will harden instantly. Fill the space between the glued spots with B&amp;B &#x2013; the glue will keep the buffer on the slide.</li>
 <li>Take a wandering L3 larva and place it on a piece of wet tissue paper on ice for 2 min. This will slow down its movement. Then transfer the larva into the buffer and orient it with the dorsal side up (this is the side of the larva that lacks the denticle belts). Quickly apply some glue between the head region and the Sylgard. The glue will harden very quickly and prevent the head of the larva from moving. </li>
 <li>Using forceps carefully grab the animal by the posterior spiracles and gently stretch it. Fix the larva in this position by applying the glue along both sides of the larva at the posterior half. To reduce further movement of the cuticle by the body wall muscles, add more glue to the flanks of the animal. The larva is now fixed in this position. The coverslip can be placed into a small Petri dish filled with AHL or PBS. Heart beats can now be recorded without interference from body wall movements.</li>
</ol>
Representative Results:
The semi-intact adult Drosophila heart will beat rhythmically for hours following dissection when maintained in fresh, oxygenated AH (see Supplemental Data, Ocorr, et al., 2007). A representative example is shown in Movie 1. Hearts from 3rd instar larva and young flies (1 &#x2013; 3 weeks post eclosion) generally exhibit a regular heart beat at rates between 1 &#x2013; 3 Hz. Flies older than 3 weeks generally are more arrhythmic; nevertheless, these hearts are still able to beat spontaneously for hours in oxygenated AHL. Hearts in adult flies that are damaged as a result of the dissection procedure typically show localized regions of extreme constriction that are unable to relax. Heartbeat rates slower than 1 Hz are rarely observed in flies younger than 3 weeks of age; consequently such preparations are considered to be damaged and are discarded.
<a href="https://www.jove.com/files/ftp_upload/1425/MOVIE1.avi">Click here to download Movie 1.</a>- A one week old adult wildtype fly (w1118 laboratory strain) showing the exposed abdominal heart tube (anterior to the left). Note regular, rhythmic contractions. Opaque regions in the upper right hand corner are the remaining abdominal fat bodies; round cells on either side of the heart are pericardial cells.

<ol><li>Ocorr, K., Reeves, N., Wessells, R. J., Fink, M., Chen, H. -S. V., Akasaka, T., Yasuda, S., Metzger, J., Giles, W., Posakony, J. W., Bodmer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((KCNQ+potassium+channel+mutations+cause+cardiac+arrhythmias+in+Drosophila+that+mimic+the+effects+of+aging%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">KCNQ potassium channel mutations cause cardiac arrhythmias in Drosophila that mimic the effects of aging.</a> Proc Natl Acad Sci U S A. 104, 3943-3948 (2007).</li>
<li>Broadie, K. S., Bate, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Development+of+the+embryonic+neuromuscular+synapse+of+Drosophila+melanogaster%5BTitle%5D)+AND+%22J+Neurosci%22%5BJournal%5D)">Development of the embryonic neuromuscular synapse of Drosophila melanogaster.</a> J Neurosci. 13, 144-166 (1993).</li>
</ol>

The Drosophila model has proven to be a powerful genetic tool that has been used to address a variety of scientific questions ranging from embryological development to learning and memory. Recently this versatile model organism has been used to investigate the genetics of heart function in the fly. A number of attempts to quantify heart physiology in adult Drosophila have relied on observations made in intact flies through the abdominal cuticle. Most of these approaches have relied on visual observation or recordings of ...

<table><tbody><tr><td>Micro Dissecting Spring Scissors (curved)</td><td>Roboz Surgical Instruments Co.</td><td>RS-5611</td><td>Good for gross cuts (Step 3)</td></tr><tr><td>Micro Dissecting Spring Scissors (straight)</td><td>Roboz Surgical Instruments Co.</td><td>RS-5620</td><td></td></tr><tr><td>Dumont #55 forceps</td><td>Fine Science Tools</td><td>11295-51</td><td></td></tr><tr><td>Dumont #5 forceps</td><td>Fine Science Tools</td><td>11295-10</td><td></td></tr><tr><td>Glass Capillaries, 100ul</td><td>VWR international</td><td>53432-921</td><td></td></tr><tr><td>Glass Capillaries, fine</td><td>Science Products</td><td>GB100T8P</td><td></td></tr><tr><td>Pipette Puller</td><td>Sutter Equipment</td><td>P-97</td><td>Both horizontal and vertical pipette pullers will work</td></tr><tr><td>Plastic tubing</td><td>Tygon</td><td>1/16&amp;#8221; inner diameter</td><td>for 100&amp;#x03BC;l capillaries</td></tr><tr><td>Plastic tubing</td><td>Tygon</td><td>1/32&amp;#8221; inner diameter</td><td>for small capillaries</td></tr><tr><td>Histoacryl&amp;#174; tissue adhesive</td><td>B. Braun Medical</td><td></td><td></td></tr></tbody></table>

visualizing the beating heart in drosophila

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.

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 ...

Two techniques will be presented here for the visualization of a beating heart in Drosophila for the dissection of the adult heart. First, the head and thorax are removed by a single cut along the indicated line. Next, the genitalia are removed and the abdominal body cavity is opened using forceps the gut, and in the case of a female fly, the ovaries are removed.The abdominal fat body in abdominal segments three and four is carefully sucked away to expose the adult heart for the immobilization of the larval heart. A single larva placed in buffer solution is fixed by applying histo acral glue first around the larva head region. The larvae is gently stretched and more glue is applied to the posterior end, and then on the flanks of the larva body wall.Hi, I'm GI Fler. Hi, I'm Karen er. We're both members of the Development and Aging program at the Burnham Institute for Medical Research.Today we will show you the procedures that we've developed to visualize the beating heart of Laval and adult drosophila. We use both of these preparations as a way to visualize, examine, and quantify heartbeat parameters in the fruit fly.Drosophila. Before getting started, we first prepare an artificial hemolymph solution.This solution contains sucrose entry, helo, which should be added to the artificial hemolymph from refrigerated stock solutions just prior to use. In order to prevent bacterial contamination, bring the artificial hemolymph to room temperature and oxygenate the solution by air bubbling for at least 15 minutes. Next, pull several fine capillaries using a standard pipette puller, which will be needed for removal of fat bodies in the fly.Once everything has been prepared, adult flies must be anesthetized. This can be done using brief exposure to fly nap for two to five minutes, but be careful not to leave the flies in the fly nap longer than five minutes. Short term exposure to cold can also be used to anesthetize flies, but carbon dioxide is not recommended since it has longer lasting effects on heart rate and ity.Next, the anesthetized flies are placed dorsal side down into a Petri dish coated with a thin layer of petroleum jelly. The hydrophobic cuticle in the wings and body should reversibly stick to the jelly, but the fly can be repositioned if needed. An initial cut is made using a curved pair of spring scissors.This is done by placing the scissor blades under the legs of the fly and angled down toward the dorsal surface of the near the neck. The head ventral nerve cord and legs are removed with a single cut once cut. The preparation is then submerged in the oxygenated artificial hemo lymph solution using spring scissors.The posterior tip of the abdomen, which is comprised of abdominal segment seven and eight, is removed with a single cut. This provides access for making lateral cuts along both edges of the abdomen and serves to sever the connection between the posterior gut and the abdominal cuticle. Insert the blades of the scissors into the small opening in the tip of the abdomen and cut the cuticle laterally.This frees up one edge of the ventral cuticle. Grasp the flap with forceps and make a similar cut along the opposite lateral edge of the abdomen. The freed flap of ventral abdominal cuticle can now be removed.The gut and other abdominal organs can usually be removed as a single mass using jeweler's forceps and gentle tugging. Removing the internal organs reveals the beating heart tube, which is still attached to the dorsal cuticle by small aery muscles and is surrounded by fat bodies. The pulled capillaries are inserted into plastic, one 16th inch tubing attached to a vacuum source.The suction force is proportional to tip diameter, so pipettes with tips larger than 40 microns in diameter should not be used once set up. This suction system is used to suck off excess fat from around the heart tube. It can also be used to remove air bubbles that accumulate around the cuticle and any debris.Extreme care must be taken to avoid touching the heart itself, and because the posterior portion of the heart is very fragile, that region should be avoided together. The adult drosophila heart tube is now exposed and should be beating if necessary. The cuticle and attached heart can be repositioned by gently lifting the cuticle out of the petroleum jelly and carefully tamping it back down again.Avoiding any contact with the heart tissue. At this point, the solution bathing the preparation should be replaced with fresh artificial hemolymph. This preparation should be allowed to equilibrate for 20 to 30 minutes with oxygenation prior to any manipulations.If the preparation is to be maintained for longer than 60 minutes, it should be reperfused with fresh artificial hemolymph. This preparation is now ready to be used for optical recording and can also be used for electrophysiological recording or for histological manipulations. A different technique is required for visualizing the beating heart intra larvae.First, prepare cover slips the day before the procedure. This is done by using a toothpick to transfer about 50 to 70 microliters of Sard solution onto a 22 by 22 millimeter cover slip. Let the syl guard harden at 65 degrees celsius overnight.Next, make the Brody and Bates buffer once prepared, aliquots can be stored at minus 20 degrees Celsius and thawed when needed. Finally, prepare the tools required to apply the histo acral glue. First, pull several fine capillaries.Next, connect a 1.5 milliliter pipette tip with a small plastic tubing about one 32nd inches in diameter. By inserting the smaller end into the tube, insert a capillary with its wide opening to the other end. Once set up, load the capillary with histo glue by dipping the capillary into a drop of glue and gentle suction into the plastic tip.When working with the glue, one should have a second glass capillary at hand to remove excessive glue from the tip of the glue capillary. The glue is liquid until it comes into contact with buffered saline where it will harden within a couple of seconds. However, it usually remains plastic long enough to be removed from the glue capillary tip or while applying it to the specimen.Now to begin immobilization of the larvae, place a drop of histo acral glue close to each corner of the syl guard. Cover slips add 50 microliters of B and B buffer to the glue drops, which will harden instantly fill the space between the glued spots with b and b. The glue will keep the buffer on the slide.Take a wandering L three larvae and place it on a piece of wet tissue paper on ice for two minutes. This will slow down its movement. Next, transfer the larvae into the buffer and orient it with the dorsal side up.This is the side of the larvae that lacks the identical belts. Quickly apply some glue between the head region and the sill guard. The glue will harden very quickly and prevent the head of the larvae from moving.Using forceps carefully grab the animal by the posterior sparkles and gently stretch it. Fix the larvae in this position by applying the glue along both sides of the larvae at the posterior half. To reduce further movement of the cuticle by the body wall muscles add more glue to the flanks of the animal.The larva is now fixed in this position. The cover slip can be placed into a small Petri dish filled with artificial hemolymph or PBS heartbeats can now be recorded without interference from body wall movements. The semi intact adult drosophila heart will beat rhythmically for hours following dissection when maintained in fresh oxygenated artificial hemolymph.A representative example is shown here and in supplemental movie one hearts from third in star larvae shown here, and young flies those one to three weeks post elo generally exhibit a regular heartbeat at rates between one to three hertz flies older than three weeks generally are more arrhythmic. Nevertheless, these hearts are still able to beat spontaneously for hours in oxygenated artificial hemolymph hearts and adult flies that are damaged as a result of the dissection procedure typically show localized regions of extreme constriction that are unable to relax and often be uncontrollably. Heartbeat rates slower than one Hertz are rarely observed in flies younger than three weeks of age.Consequently, such preparations are considered to be damaged and are discarded. We've just demonstrated how to prepare level and adult fruit fly hearts for optical imaging. Remember, when making the semi intact preparation never touch the heart itself.So that's it. Thank you for watching and good luck with your experiments.

visualizing-the-beating-heart-in-drosophila

Research

JoVE Journal

Biology

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.

Fluorescent Labeling of Drosophila Heart Structures

We have developed a Semi-automated Optical Heartbeat Analysis method (SOHA) for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts. We demonstrate the application of our methodology to the analysis of heart function in fruit fly and embryonic mouse hearts.

Semi-automated Optical Heartbeat Analysis of Small Hearts

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

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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.

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

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

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This video describes the fly's adult heart, or otherwise called the Drosophila dorsal vessel. The featured protocol demonstrates a dissection procedure for live heart preparations.

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

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

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

We describe optimized tools to study the zebrafish heart in vivo with light sheet fluorescence microscopy. Specifically, we suggest bright cardiac transgenic lines and present new gentle embedding and immobilization techniques that avoid developmental and heart defects. A possible data acquisition and analysis pipeline adapted to cardiac imaging is also provided.

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

The present protocol describes the generation of Drosophila melanogaster expressing eNpHR2.0 or ReaChR opsins in the heart for OCT imaging and optogenetic heart pacing. Detailed instructions for Drosophila OCT imaging and heart beating modulation, including the simulation of restorable heart arrest, bradycardia, and tachycardia in live animals at different developmental stages, are reported.

Developing Drosophila melanogaster Models for Imaging and Optogenetic Control of Cardiac Function

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

Learn techniques for visualizing the beating heart in larval and adult Drosophila using dissection and immobilization methods. These protocols enable cardiac physiology studies in developmental biology.

Visualizing the Beating Heart in Drosophila

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