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.remotexs.ntu.edu.sg/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/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, 3943-3948 (2007).</li><li>Broadie, K. S., Bate, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+embryonic+neuromuscular+synapse+of+Drosophila+melanogaster.">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 changes in light intensity transmitter through the abdomen to quantify a single parameter, heart rate. These methods have several limitations: typically only the anterior end of the heart can be observed, what is being observed is the movement of fat bodies secondary to the heart contractions, and the nervous input to the heart is intact. The adult semi-intact preparation provides a more detailed view of a large portion of the functioning heart permitting quantification of a number of parameters in addition to heart rate. It can be used for electrophysiological or optical (see JoVE article, 1435) recording procedures. In addition, this preparation is suitable for histological manipulations (see JoVE article, 1435). It can also be extracted as a relatively clean heart tissue sample for PCR, Western blotting, microarray analysis, etc.
The adult heart in Drosophila is, for the purposes of this dissection, conveniently fixed to the dorsal cuticle of the abdomen by a network of alary muscles. Thus it is possible to dissect away the head and ventral lying nerve cord, and then to remove the internal organs without damaging the heart. However, extreme care should be taken in step 3 to cut only the very tip of the abdomen as the heart extends into abdominal segment 5/6 and a pacemaker cell or cells is undoubtedly present in this region.
Care must also be taken to avoid touching the heart during the dissection. If contact with the heart is suspected the preparation should be discarded. Moderate suction usually suffices to remove some of the fat and tracheols that surround the heart for better visualization of the heart tube but excessive suction can also damage the heart. The suction force is controlled primarily by the size of the pipette tip used for the liposuction; tips larger than approximately 40 microns should be avoided.
Unlike in the adult, the heart tube in Drosophila larva is relatively free to move around the body cavity. Thus a semi-intact larval heart preparation presents difficulties for optical recording techniques. Due to the translucency of the integument it is possible to visualize the heart in the intact larva using brightfield microscopy. The glue technique described here immobilizes larva of any developmental stage and keeps the heart in a fixed position. This permits recording of a relatively stationary heart tube. Heart contractions recorded by such immobilized larva can be subsequently analyzed by the optical recording technique described in JoVE 1435.

<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

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.

Watch this Scientific Journal Video about Visualizing the Beating Heart in Drosophila at JoVE.com

Visualizing the Beating Heart in Drosophila

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

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 ...

visualizing-the-beating-heart-in-drosophila

Nanyang Technological University

Research

JoVE Journal

Biology

18.6K Views.  The Sanford Burnham Institute for Medical Research. Techniek die nodig is voor het visualiseren van het kloppend hart in de larven en volwassen Drosophila Worden gepresenteerd. Elke levensfase vraagt ​​om een ​​andere methodologie.

Video: Visualizing the Beating Heart in Drosophila

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Fluorescent Labeling of Drosophila Heart Structures

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

Fluorescent Labeling of Drosophila Heart Structures

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

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

Semi-automated Optical Heartbeat Analysis of Small Hearts

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our Semi-automatic Optical Heartbeat Analysis (SOHA) uses a novel movement detection algorithm that is able to detect cardiac movements associated with individual contractile and relaxation events. The program provides a host of physiologically relevant readouts including systolic and diastolic intervals, heart rate, as well as qualitative and quantitative measures of heartbeat arrhythmicity. The program also calculates heart diameter measurements during both diastole and systole from which fractional shortening and fractional area changes are calculated. Output is provided as a digital file compatible with most spreadsheet programs. Measurements are made for every heartbeat in a record increasing the statistical power of the output. We demonstrate each of the steps where user input is required and show the application of our methodology to the analysis of heart function in all three genetically tractable heart models.

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.

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

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.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

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

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

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

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

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

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 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.

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

Genetics

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

Using Drosophila melanogaster (fruit fly) as a model organism has ensured significant progress in many areas of biological science, from cellular organization and genomic investigations to behavioral studies. Due to the accumulated scientific knowledge, in recent years, Drosophila was brought to the field of modeling human diseases, including heart disorders. The presented work describes the experimental system for monitoring and manipulating the heart function in the context of a whole live organism using red light (617 nm) and without invasive procedures. Control over the heart was achieved using optogenetic tools. Optogenetics combines the expression of light-sensitive transgenic opsins and their optical activation to regulate the biological tissue of interest. In this work, a custom integrated optical coherence tomography (OCT) imaging and optogenetic stimulation system was used to visualize and modulate the functioning D. melanogaster heart at the 3rd instar larval and early pupal developmental stages. The UAS/GAL4 dual genetic system was employed to express halorhodopsin (eNpHR2.0) and red-shifted channelrhodopsin (ReaChR), specifically in the fly heart. Details on preparing D. melanogaster for live OCT imaging and optogenetic pacing are provided. A lab-developed integration software processed the imaging data to create visual presentations and quantitative characteristics of Drosophila heart function. The results demonstrate the feasibility of initiating cardiac arrest and bradycardia caused by eNpHR2.0 activation and performing heart pacing upon ReaChR activation.

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

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.

Using Drosophila melanogaster (fruit fly) as a model organism has ensured significant progress in many areas of biological science, from cellular ...

Bioengineering

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

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

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

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

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

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid external development, tractable genetics, and translucency of the zebrafish, Danio rerio, LSFM has delivered insights into cardiac form and function at high spatial and temporal resolution without significant phototoxicity or photobleaching. Imaging of beating hearts challenges existing sample preparation and microscopy techniques. One needs to maintain a healthy sample in a constricted field of view and acquire ultrafast images to resolve the heartbeat. Here we describe optimized tools and solutions to study the zebrafish heart in vivo. We demonstrate the applications of bright transgenic lines for labeling the cardiac constituents and present novel gentle embedding and immobilization techniques that avoid developmental defects and changes in heart rate. We also propose a data acquisition and analysis pipeline adapted to cardiac imaging. The entire workflow presented here focuses on zebrafish embryonic heart imaging but can also be applied to various other samples and experiments.

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.

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid ...

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

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

Visualizing the Beating Heart in Drosophila

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