KO est soutenu par une subvention de l&#39;American Heart Association.

Avant de commencer Coeurs adultes <ol><li> Fraîchement préparer hémolymphe artificielle (AH) solution contenant 108mm Na +, 5 mM de K +, 2 mM de Ca 2 +, 8mm MgCl2, 1 mM NaH 2 PO 4, 4mm NaHCO 3, saccharose 10 mM, tréhalose 5mM, HEPES 5mm (pH 7,1, tous les réactifs de Sigma Chemicals). Le saccharose et le tréhalose devrait être ajouté à l&#39;AH à partir de solutions mères réfrigéré juste avant l&#39;utilisation afin de prévenir la contamination bactérienne. </li><li> Apportez AH à la température ambiante et oxygéner la solution par air-bullage pendant au moins 15 min. </li><li> Tirez plusieurs fins capillaires (par exemple capillaires de verre, 100 ul, VWR) requis pour l&#39;élimination des graisses. </li></ol> Cœur des larves <ol><li> Sylgard enduit de lamelles: en utilisant un cure-dent, le transfert d&#39;environ 50-70μl de solution Sylgard sur une lamelle 22x22mm. Laissez durcir le Sylgard à 65 ° C pendant la nuit. </li><li> Préparer le Broadie et le tampon de Bate (135mm NaCl, KCl 5 mM, MgCl 2 4 mM, 2 mM CaCl 2, 5 mM TES, saccharose 36mm;. PH 7,15 Voir Broadie et Bate, 1993). </li><li> Préparer les outils nécessaires pour appliquer la colle Histoacryl. Tout d&#39;abord, tirez plusieurs fins capillaires (par exemple sciences Produits GB100T8P). Connecter une extrémité de la pipette 1,5 ml avec un tuyau en plastique de petite taille (Tygon, 1 / 32 &quot;) en insérant l&#39;extrémité la plus petite dans le tube. Insérez un capillaire avec sa large ouverture à l&#39;autre extrémité. Chargez le capillaire avec de la colle Histoacryl en plongeant le capillaire dans une goutte de colle et une aspiration douce dans l&#39;embout en plastique. Lorsque vous travaillez avec de la colle on devrait avoir un deuxième verre capillaire à la main pour enlever la colle excessive de la pointe de la colle capillaire. La colle est liquide jusqu&#39;à ce qu&#39;il entre en contact avec une solution saline tamponnée où il va durcir en quelques secondes. Cependant, il reste généralement en plastique assez longtemps pour être retiré de la pointe de la colle capillaire ou à tout en l&#39;appliquant à l&#39;échantillon. </li></ol> Semi-Cœur intacte drosophile de la mouche adulte <ol><li> Adulte mouches drosophiles sont anesthésiés avec Nap Fly (Caroline biologique Supply Co.) pendant 2-5 minutes. Des précautions doivent être prises pour ne pas laisser les mouches dans le Nap Fly pour de plus longues expositions. L&#39;exposition à court terme au froid peut également être utilisé pour anesthésier les mouches mais le CO 2 n&#39;est pas recommandée car elle a des effets plus durables sur le rythme cardiaque et de la rythmicité. </li><li> Mouches anesthésiés sont placés, face dorsale vers le bas, dans une boîte de Petri recouverte d&#39;une fine couche de gelée de pétrole. La cuticule hydrophobe dans les ailes et le corps de manière réversible «coller» à la gelée, mais la mouche peut être repositionné si nécessaire. Cela devient particulièrement important lors des manipulations ultérieures. </li><li> Une première coupe est faite en utilisant une paire de ciseaux courbes printemps (Roboz # 5611). Les lames des ciseaux sont placés sous les pattes et inclinées vers le bas vers la surface dorsale du thorax près du cou. La tête, le cordon nerveux ventral, et les jambes sont enlevés avec une seule coupe. La préparation est ensuite immergé dans une oxygénée, hémolymphe artificielle (AH) solution. </li><li> Avec des ciseaux à ressort de la pointe postérieure de l&#39;abdomen (composé de segments abdominaux 7 et 8) est enlevé avec une seule coupe. Cela fournit un accès pour faire des coupes latérales le long des deux bords de l&#39;abdomen et sert à couper les liens entre l&#39;intestin postérieur et la cuticule abdominale. Le volet libéré de la cuticule abdominale ventrale est ensuite retiré. </li><li> Dans les étapes 3 et 4, les liaisons antérieures et postérieures de l&#39;intestin sont coupés de sorte les organes abdominaux sont désormais tenus en place que par tracheols. L&#39;intestin et d&#39;autres organes abdominaux peuvent habituellement être enlevé comme une masse unique à l&#39;aide des pinces de bijoutier (Roboz, # 55) et doux tiraillement. Retrait des organes internes révèle le tube coeur battant encore attaché à la cuticule dorsale entourée de corps gras. </li><li> Les corps gras sont assez opaques au microscope optique, par conséquent, il est souvent nécessaire d&#39;enlever une partie de cette graisse, afin de voir le tube cardiaque clairement. Ceci est accompli en utilisant une technique de liposuccion finement dessinés capillaires en verre. Les capillaires sont faites en utilisant une norme pipette extracteur (Ex. Sutter P-97 Extracteur pipette) et sont ensuite insérées dans un tube en plastique (Tygon, 1 / 16 &quot;) attaché à une source de vide. Ce système est utilisé à l&#39;aspiration hors excès de graisse autour du tube cardiaque. Force d&#39;aspiration est proportionnelle au diamètre pointe afin Pipettes avec pointes de plus de 40 microns de diamètre ne doit pas être utilisé. Un soin extrême doit être pris pour éviter de toucher le cœur lui-même et parce que la partie postérieure du cœur est très fragile de cette région devrait être évitée. </li><li> Le tube cardiaque adulte drosophile est maintenant exposée et devrait être battu. Si nécessaire, le coeur cuticule et attachés peuvent être repositionnées en soulevant doucement la cuticule de la gelée de pétrole et soigneusement bourrage vers le bas, encore une fois en évitant tout contact avec le tissu cardiaque. A tson point la solution de bain de la préparation doit être remplacé par AHL frais. Cette préparation devrait être autorisé à s&#39;équilibrer pendant 20-30 minutes avec l&#39;oxygénation avant toute manipulation. Si la préparation doit être maintenu pendant plus de 60 minutes, il devrait être perfusé avec AHL frais. </li></ol> Immobilisation larves de drosophile pour l&#39;enregistrement optique <ol><li> Déposer une goutte de colle Histoacryl près de chaque coin du Sylgard lamelles. Ajouter 50 pl de B &amp; B tampon pour les gouttes de colle qui durcit instantanément. Remplir l&#39;espace entre les taches collées avec B &amp; B - la colle va garder le tampon sur la diapositive. </li><li> Prenez une larve L3 errance et le placer sur un morceau de papier tissu humide sur la glace pendant 2 min. Cela va ralentir son mouvement. Puis transfert de la larve dans le tampon et l&#39;orienter à la face dorsale vers le haut (c&#39;est le côté de la larve qui n&#39;a pas la ceinture de denticules). Appliquez rapidement un peu de colle entre la région de la tête et le Sylgard. La colle durcit très rapidement et d&#39;empêcher la tête de la larve de se déplacer. </li><li> En utilisant des pinces attentivement attraper l&#39;animal par les stigmates postérieurs et doucement l&#39;étirer. Fixer la larve dans cette position par l&#39;application de la colle le long des deux côtés de la larve à la moitié postérieure. Afin de réduire poursuite du mouvement de la cuticule par des muscles de la paroi du corps, ajouter plus de colle sur les flancs de l&#39;animal. La larve est maintenant fixé dans cette position. La lamelle peut être placé dans une petite boîte de Petri remplie de AHL ou PBS. Battements du cœur peuvent désormais être enregistrées sans interférence de mouvements paroi du corps. </li></ol> Les résultats représentatifs: Le semi-intactes pour adultes Drosophila coeur battra rythmiquement pendant des heures suivantes de dissection lors maintenu en eau douce, AH oxygénée (voir données supplémentaires, Ocorr, et al., 2007). Un exemple représentatif est montré dans le film 1. Coeurs de 3 ème stade larvaire et les mouches jeunes (1 - 3 semaines après l&#39;éclosion) présentent généralement un rythme cardiaque régulier à des taux entre 1 - 3 Hz. Les mouches plus de 3 semaines sont généralement plus arythmique, néanmoins, ces coeurs sont encore en mesure de battre spontanément pendant des heures dans l&#39;AHL oxygénée. Coeurs perdus en mouches adultes qui sont endommagés à la suite de la procédure de dissection montrent typiquement des régions localisées de la constriction extrêmes qui sont incapables de se détendre. Heartbeat taux inférieure à 1 Hz sont rarement observés chez les mouches plus jeunes que 3 semaines d&#39;âge, par conséquent de telles préparations sont considérés comme étant endommagés et sont éliminés. <a href="http://www.jove.com/files/ftp_upload/1425/MOVIE1.avi">Cliquez ici pour télécharger la vidéo 1.</a> - Une volée un semaine anciennes de type sauvage adulte (souche de laboratoire w 1118) montrant le tube cardiaque exposés abdominale (antérieure à la gauche). Remarque réguliers, des contractions rythmiques. Zones opaques dans le coin supérieur droit sont les autres corps gras abdominaux; cellules rondes de chaque côté du cœur sont les cellules péricardiques.

<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>

Le modèle de drosophile s&#39;est avérée être un puissant outil génétique qui a été utilisée pour traiter une variété de questions scientifiques allant du développement embryologique de l&#39;apprentissage et la mémoire. Récemment cet organisme modèle polyvalent a été utilisé pour étudier la génétique de la fonction cardiaque chez la mouche. Un certain nombre de tentatives pour quantifier la physiologie cardiaque chez les adultes de drosophile sont appuyés sur les observations faites chez des mouch...

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<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>
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<td>Micro Dissecting Spring Scissors (straight)</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5620</td>
<td>&nbsp;</td>
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<td>Dumont #55 forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11295-51</td>
<td>&nbsp;</td>
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<td>Dumont #5 forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
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<td>Glass Capillaries, 100ul</td>
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<td>&nbsp;</td>
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<td>Both horizontal and vertical pipette pullers will work</td>
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<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>
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<td>Plastic tubing</td>
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<td>Tygon R-3603</td>
<td>1/32&#x201D; inner diameter</td>
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<td>Histoacryl&reg; tissue adhesive</td>
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visualizing the beating heart in drosophila

L&#39;<em&gt; Drosophile</em&gt; Coeur a récemment émergé comme un système bon modèle pour étudier les mécanismes génétiques, cellulaires et moléculaires sous-jacents de fonction dans les coeurs myogéniques. Une étape clé dans l&#39;examen de la fonction cardiaque chez la mouche est de trouver un moyen d&#39;accéder au cœur d&#39;une manière qui préserve sa fonction myogénique tout en permettant toujours l&#39;organe cœur battant pour être observées et enregistrées. Deux méthodes différentes pour observer et enregistrer le cœur battant à la fois dans la larve et l&#39;adulte<em&gt; Drosophile</em&gt; Sont décrites ici. Notre semi-intactes préparation en utilisant les mouches adultes permet une visualisation claire du cœur abdominale sans ingérence de la cuticule recouvrant pigmentées et corps gras. Pour enregistrer le cœur bat des larves, il est nécessaire d&#39;immobiliser la larve, ce qui minimise les mouvements paroi du corps réduisant ainsi les mouvements du cœur qui ne sont pas associés avec des contractions du myocarde. Nos méthodologies de produire adulte stable et préparations coeur des larves qui peut battre pendant des heures à des taux de Hz 1-3.

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

Visualizing the Beating Heart in Drosophila

Technique requise pour visualiser le cœur battant dans la larve et l&#39;adulte<em&gt; Drosophile</em&gt; Sont présentés. Chaque étape de la vie nécessite une méthodologie différente.

Visualiser le cœur battant Drosophile</em

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

Visualizing the Beating Heart in Drosophila

18.6K vues.  The Sanford Burnham Institute for Medical Research. Technique requise pour visualiser le cœur battant dans la larve et l'adulte Drosophile Sont présentés. Chaque étape de la vie nécessite une méthodologie différente.

Vidéo: 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.

Visualiser le Live Drosophile Gliales-jonction neuromusculaire avec des colorants fluorescents

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

Recherche

Biologie

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.

Marquage fluorescent Drosophile</emOuvrages d&#39;art> Coeur

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.

Surveillance de la fonction cardiaque chez les larves Drosophila melanogaster Pour les études physiologiques

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.

Préparation d&#39;embryons pour la microscopie électronique des Drosophile Tube cardiaque embryonnaire

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.

Quantification des γH2AX Foci en réponse aux radiations ionisantes

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.

La quantification de γH2AX Foci dans les échantillons de tissus

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.

Évaluation de la distribution spatiale des rayonnements ionisants suivantes γH2AX

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.

Test clonogénique: cellules adhérentes

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.

Vivre imagerie de dédouanement cellulaire par apoptose au cours<em&gt; Drosophila</em&gt; Embryogenèse

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.

Évaluation de calcium Sparks dans intactes les fibres musculaires squelettiques

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