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/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 ...

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Micro Dissecting Spring Scissors (curved)</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5611</td>
<td>Good for gross cuts (Step 3)</td>
</tr>
<tr>
<td>Micro Dissecting Spring Scissors (straight)</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5620</td>
<td>&nbsp;</td>
<tr>
<td>Dumont #55 forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11295-51</td>
<td>&nbsp;</td>
<tr>
<td>Dumont #5 forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11295-10</td>
<td>&nbsp;</td>
<tr>
<td>Glass Capillaries, 100ul</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>53432-921</td>
<td>&nbsp;</td>
<tr>
<td>Glass Capillaries, fine</td>
<td>&nbsp;</td>
<td>Science Products</td>
<td>GB100T8P</td>
<td>&nbsp;</td>
<tr>
<td>Pipette Puller</td>
<td>&nbsp;</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>&nbsp;</td>
<td>Tygon R-3603</td>
<td>1/16&#x201D; inner diameter</td>
<td>for 100&mu;l capillaries</td>
</tr>
<tr>
<td>Plastic tubing</td>
<td>&nbsp;</td>
<td>Tygon R-3603</td>
<td>1/32&#x201D; inner diameter</td>
<td>for small capillaries</td>
</tr>
<tr>
<td>Histoacryl&reg; tissue adhesive</td>
<td>&nbsp;</td>
<td>B.Braun, Melsungen</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

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

Research

JoVE Journal

Biology

18.6K Views.  The Sanford Burnham Institute for Medical Research. Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

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 ...

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 ...

Quantification of &gamma;H2AX Foci in Response to Ionising Radiation

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA lesions with respect to survival and preservation of genomic integrity. An early response to the induction of DSBs is phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX1. Following induction of DSBs, H2AX is rapidly phosphorylated by the phosphatidyl-inosito 3-kinase (PIKK) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Typically, only a few base-pairs (bp) are implicated in a DSB, however, there is significant signal amplification, given the importance of chromatin modifications in DNA damage signalling and repair. Phosphorylation of H2AX mediated predominantly by ATM spreads to adjacent areas of chromatin, affecting approximately 0.03% of total cellular H2AX per DSB2,3. This corresponds to phosphorylation of approximately 2000 H2AX molecules spanning ~2 Mbp regions of chromatin surrounding the site of the DSB and results in the formation of discrete &gamma;H2AX foci which can be easily visualized and quantitated by immunofluorescence microscopy2. The loss of &gamma;H2AX at DSB reflects repair, however, there is some controversy as to what defines complete repair of DSBs; it has been proposed that rejoining of both strands of DNA is adequate however, it has also been suggested that re-instatement of the original chromatin state of compaction is necessary4-8. The disappearence of &gamma;H2AX involves at least in part, dephosphorylation by phosphatases, phosphatase 2A and phosphatase 4C5,6. Further, removal of &gamma;H2AX by redistribution involving histone exchange with H2A.Z has been implicated7,8. Importantly, the quantitative analysis of &gamma;H2AX foci has led to a wide range of applications in medical and nuclear research. Here, we demonstrate the most commonly used immunofluorescence method for evaluation of initial DNA damage by detection and quantitation of &gamma;H2AX foci in &gamma;-irradiated adherent human keratinocytes9.

Quantification of &amp;#947;H2AX Foci in Response to Ionising Radiation

Quantification of DNA double-strand streaks using &gamma;H2AX formation as a molecular marker has become an invaluable tool in radiation biology. Here we demonstrate the use of an immunofluorescence assay for quantification of &gamma;H2AX foci after exposure of cells to radiation.

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA ...

Quantitation of &gamma;H2AX Foci in Tissue Samples

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) processes or by exogenous factors, particularly ionizing radiation and radiomimetic compounds. Phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX, is an early response to DNA double-strand breaks1. This phosphorylation event is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Overall, DSB induction results in the formation of discrete nuclear &gamma;H2AX foci which can be easily detected and quantitated by immunofluorescence microscopy2. Given the unique specificity and sensitivity of this marker, analysis of &gamma;H2AX foci has led to a wide range of applications in biomedical research, particularly in radiation biology and nuclear medicine. The quantitation of &gamma;H2AX foci has been most widely investigated in cell culture systems in the context of ionizing radiation-induced DSBs. Apart from cellular radiosensitivity, immunofluorescence based assays have also been used to evaluate the efficacy of radiation-modifying compounds. In addition, &gamma;H2AX has been used as a molecular marker to examine the efficacy of various DSB-inducing compounds and is recently being heralded as important marker of ageing and disease, particularly cancer3. Further, immunofluorescence-based methods have been adapted to suit detection and quantitation of &gamma;H2AX foci ex vivo and in vivo4,5. Here, we demonstrate a typical immunofluorescence method for detection and quantitation of &gamma;H2AX foci in mouse tissues.

Quantitation of &amp;#947;H2AX Foci in Tissue Samples

Quantitation of DNA double-strand breaks on the basis of &gamma;H2AX foci has become an invaluable tool, particularly in radiation biology, for the evaluation of tissue radiosensitivity and effects of radiation modifying compounds. Here we demonstrate the use of an immunofluorescence assay for quantitation of &gamma;H2AX foci in tissue samples.

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) ...

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &amp;#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

Clonogenic Assay: Adherent Cells

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. Apart from documenting the method, the initial landmark study generated the first radiation-dose response curve for X-ray irradiated mammalian (HeLa) cells in culture1. Basically, the clonogenic assay enables an assessment of the differences in reproductive viability (capacity of cells to produce progeny; i.e. a single cell to form a colony of 50 or more cells) between control untreated cells and cells that have undergone various treatments such as exposure to ionising radiation, various chemical compounds (e.g. cytotoxic agents) or in other cases genetic manipulation. The assay has become the most widely accepted technique in radiation biology and has been widely used for evaluating the radiation sensitivity of different cell lines. Further, the clonogenic assay is commonly used for monitoring the efficacy of radiation modifying compounds and for determining the effects of cytotoxic agents and other anti-cancer therapeutics on colony forming ability, in different cell lines. A typical clonogenic survival experiment using adherent cells lines involves three distinct components, 1) treatment of the cell monolayer in tissue culture flasks, 2) preparation of single cell suspensions and plating an appropriate number of cells in petri dishes and 3) fixing and staining colonies following a relevant incubation period, which could range from 1-3 weeks, depending on the cell line. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cell lines with the use of an immortalized human keratinocyte cell line (FEP-1811)2. Also, our aims are to describe common features of clonogenic assays including calculation of the plating efficiency and survival fractions after exposure of cells to radiation, and to exemplify modification of radiation-response with the use of a natural antioxidant formulation.

The applicability of the clonogenic assay for evaluating reproductive viability has been established for more than 50 years. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cells.

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...