KO é apoiado por uma bolsa da American Heart Association.

Antes de começar a Coração do adulto <ol><li> Recém preparar hemolinfa artificial (AH) solução contendo 108mm Na +, 5mM K +, 2mM de Ca 2 + 8mm, MgCl 2, 1 mM NaH 2 PO 4, 4 mM NaHCO 3, sacarose 10mM, trealose 5mM, HEPES 5mM (pH 7.1, todos os reagentes de Chemicals Sigma). A sacarose e trealose deve ser adicionado ao AH a partir de soluções estoque refrigerado imediatamente antes do uso, a fim de evitar a contaminação bacteriana. </li><li> AH trazer à temperatura ambiente e oxigenar a solução por via aérea borbulhando por pelo menos 15 min. </li><li> Puxe vários capilares finos (por exemplo, capilares de vidro, 100ul, VWR) necessário para a remoção de gorduras. </li></ol> Corações larval <ol><li> Sylgard revestido lamínulas: usando um palito de transferência, de cerca de 50-70μl de solução Sylgard em uma lamela 22x22mm. Deixe o endurecem Sylgard a 65 ° C durante a noite. </li><li> Prepare o Broadie e buffer de Bate (135mm NaCl, KCl 5mM, 4mm MgCl 2, CaCl 2 2mM, TES 5mM, Sacarose 36mm;. PH 7,15 Veja Broadie e Bate, 1993). </li><li> Prepare as ferramentas necessárias para aplicar a cola Histoacryl. Primeiro, puxe vários capilares finos (por exemplo, Ciência Produtos GB100T8P). Conecte uma ponta de pipeta de 1,5 ml com um pequeno tubo de plástico (Tygon, 1 / 32 &quot;), inserindo a extremidade menor para dentro do tubo. Inserir um capilar com a sua ampla abertura para o outro lado. Carregar o capilar com cola Histoacryl por imersão do capilar em uma gota de cola e uma sucção suave na ponta de plástico. Ao trabalhar com a cola deve ter um segundo copo capilar em mãos para remover a cola excessiva da ponta da cola capilar. A cola é líquido até que ele entra em contato com solução salina, onde ele irá endurecer em poucos segundos. No entanto, ele geralmente permanece plástica por tempo suficiente para ser removido da ponta cola capilar ou, aplicando-o ao modelo. </li></ol> Semi-intactas Coração Drosophila da mosca adulta <ol><li> Moscas Drosophila são anestesiados com Nap Fly (Carolina Biological Supply Co.) por 2-5 minutos. É preciso ter cuidado para não deixar as moscas no Nap Fly para exposições mais longas. Exposição a curto prazo ao frio também pode ser usado para anestesiar as moscas, mas CO 2 não é recomendado, pois tem mais efeitos duradouros sobre a freqüência cardíaca e ritmo. </li><li> Anestesiados moscas são colocadas, lado dorsal para baixo, em uma placa de Petri revestida com uma fina camada de vaselina. A cutícula hidrofóbica nas asas e no corpo reversivelmente &quot;pau&quot; para a geléia, mas a mosca pode ser reposicionada, se necessário. Isso se torna especialmente importante durante as manipulações subsequentes. </li><li> Um corte inicial é feita usando um par de tesouras curvas mola (Roboz # 5611). As lâminas de tesoura são colocados sob as pernas e anguladas para baixo em direção a superfície dorsal do tórax perto do pescoço. A cabeça, cordão nervoso ventral, e as pernas são removidos com um único corte. A preparação é então imerso em um hemolinfa, oxigenado artificial (AH) solução. </li><li> Usando a tesoura primavera a ponta posterior do abdômen (composto por segmentos abdominais 7 e 8) é removido com um único corte. Isso fornece acesso para fazer cortes laterais ao longo de ambas bordas do abdome e serve para cortar a ligação entre o intestino posterior e da cutícula abdominal. O retalho livre de cutícula abdominal ventral é então removido. </li><li> Nos passos 3 e 4 as conexões anterior e posterior do intestino são cortados para os órgãos abdominais são agora mantidos no lugar apenas por tracheols. O intestino e outros órgãos abdominais geralmente pode ser removida como uma única massa com a pinça joalheiro (Roboz, # 55) e suave puxando. Remover os órgãos internos revela o tubo coração batendo ainda ligado à cutícula dorsal cercado por corpos de gordura. </li><li> Os corpos de gordura são bastante opacos no microscópio de luz, consequentemente, muitas vezes é necessário remover algumas dessa gordura, a fim de ver o tubo cardíaco claramente. Isto é realizado por uma técnica de lipoaspiração usando finamente desenhadas capilares de vidro. Capilares são feitas usando um padrão pipeta extrator (Eg Sutter P-97 Pipetar Puller) e depois são inseridos em tubos de plástico (Tygon, 1 / 16 &quot;) ligado a uma fonte de vácuo. Este sistema é usado para sugar o excesso de gordura ao redor do tubo de coração. Força de sucção é proporcional ao diâmetro da ponta para pipetas com pontas maiores que 40 mícrons de diâmetro não deve ser usado. Extremo cuidado deve ser tomado para evitar tocar o próprio coração e porque a parte posterior do coração é muito frágil naquela região deve ser evitada. </li><li> O tubo de coração adulto Drosophila está agora exposta e deve ser de bater. Se necessário, o coração cutícula e anexado pode ser reposicionada, delicadamente levantar a cutícula fora da vaselina e cuidadosamente tamping-lo de volta para baixo, novamente evitando qualquer contato com o tecido do coração. Em tseu ponto de vista a solução que banha a preparação deve ser substituído por AHL fresco. Esta preparação deve ser autorizada a estabilizar durante 20-30 minutos com oxigenação antes de qualquer manipulação. Se a preparação deve ser mantida por mais de 60 minutos deve ser perfundido com AHL fresco. </li></ol> Imobilizando larvas Drosophila para gravação óptica <ol><li> Coloque uma gota de cola Histoacryl perto de cada canto do lamínulas Sylgard. Adicionar 50μl de tampão B &amp; B para as gotas de cola que vai endurecer instantaneamente. Preencher o espaço entre os pontos colados com B &amp; B - a cola vai manter o buffer no slide. </li><li> Dê uma vagando larva L3 e colocá-lo em um pedaço de lenço de papel molhado em gelo por 2 min. Isto irá desacelerar seu movimento. Em seguida, transferir a larva no buffer e orientá-la com o lado dorsal para cima (este é o lado da larva que não tem os cintos dentículo). Aplicar rapidamente um pouco de cola entre a região da cabeça e do Sylgard. A cola vai endurecer muito rapidamente e impedir que a cabeça da larva de se mover. </li><li> Utilizando uma pinça cuidadosamente agarrar o animal pela espiráculos posterior e gentilmente esticá-lo. Fixar a larva nesta posição, aplicando a cola ao longo de ambos os lados da larva na metade posterior. Para reduzir ainda mais o movimento da cutícula pelos músculos da parede do corpo, adicione mais cola para os flancos do animal. A larva é agora fixa nesta posição. A lamela pode ser colocado em uma placa de Petri pequena cheia de AHL ou PBS. Batimentos cardíacos podem ser gravados sem a interferência de movimentos parede do corpo. </li></ol> Resultados representativos: A semi-intactas adulto coração vai bater ritmicamente Drosophila por horas após a dissecação, quando mantidos em fresco, AH oxigenado (ver Dados Suplementar, Ocorr, et al., 2007). Um exemplo representativo é mostrado no filme 1. Corações de 3 º instar larva e moscas jovens (1-3 semanas após a eclosão) geralmente apresentam um batimento cardíaco regular em taxas entre 1-3 Hz. Voa com mais de três semanas, geralmente são mais arrítmicos, no entanto, esses corações ainda são capazes de bater espontaneamente por horas em AHL oxigenado. Corações em moscas adultas que estão danificados, como resultado do procedimento de dissecção normalmente apresentam regiões localizadas de constrição extremas que são incapazes de relaxar. Taxas de batimento cardíaco mais lento do que 1 Hz são raramente observadas em moscas com menos de 3 semanas de idade, conseqüentemente tais preparações são consideradas danificadas e são descartados. <a href="http://www.jove.com/files/ftp_upload/1425/MOVIE1.avi">Clique aqui para baixar Filme 1.</a> - A uma semana fly tipo selvagem de idade adulta (w cepa de laboratório 1118) mostrando o tubo de coração exposto abdominal (anterior à esquerda). Nota regular, contrações rítmicas. Regiões opacas no canto superior direito estão os restantes órgãos de gordura abdominal; células redondas de cada lado do coração são as células do pericárdio.

<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>

O modelo de Drosophila provou ser uma poderosa ferramenta genética que tem sido utilizado para tratar uma variedade de questões científicas que vão desde desenvolvimento embrionário para a aprendizagem e memória. Recentemente, este organismo modelo versátil, tem sido usado para investigar a genética da função cardíaca na mosca. Uma série de tentativas para quantificar a fisiologia do coração em adultos Drosophila têm contado com observações feitas em moscas intacta através da cutícula abdominal. A mai...

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		<th>Type</th>
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		<th>Catalogue Number</th>
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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>
</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>
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<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>
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<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>
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<tr>
<td>Histoacryl&reg; tissue adhesive</td>
<td>&nbsp;</td>
<td>B.Braun, Melsungen</td>
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visualizing the beating heart in drosophila

O<em&gt; Drosophila</emCardíaca&gt; surgiu recentemente como um sistema bom modelo para examinar os mecanismos genéticos, celulares e moleculares subjacentes à função em corações miogênica. Um passo fundamental na análise da função cardíaca na mosca é encontrar uma maneira de acessar o coração de uma forma que preserva a sua função miogênica enquanto ainda permitindo que o órgão coração batendo a serem observados e registrados. Dois métodos diferentes para observar e registrar o coração batendo em ambos larva e adulto<em&gt; Drosophila</em&gt; São descritos aqui. Nossa preparação semi-intactas usando moscas adultas permite a visualização clara do coração abdominal, sem interferência da cutícula pigmentadas e sobrejacente corpos gordos. Para gravar coração bate larval é necessário imobilizar a larva, o que minimiza os movimentos do corpo, reduzindo assim os movimentos da parede do coração que não estão associados com as contrações do miocárdio. Nossas metodologias produzir adulto estável e preparações de coração larval que pode bater por horas a taxas 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

Técnica necessária para visualizar o coração batendo em larvas e adultos<em&gt; Drosophila</em&gt; São apresentados. Cada fase da vida exige uma metodologia diferente.

Visualizar o coração batendo no Drosophila</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 visualizações.  The Sanford Burnham Institute for Medical Research. Técnica necessária para visualizar o coração batendo em larvas e adultos Drosophila São apresentados. Cada fase da vida exige uma metodologia diferente.

Vídeo: 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.

Visualizando o Live Drosophila Junção neuromuscular-Glial com corantes fluorescentes

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

Biologia

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.

Labeling fluorescentes de Drosophila Estruturas Coração

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.

Função de monitorização cardíaca em larval Drosophila melanogaster Para estudos fisiológicos

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.

Preparação de embriões de Microscopia Eletrônica do Drosophila Tubo cardíaco embrionário

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.

Quantificação de γH2AX Focos em resposta à radiação ionizante

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.

Quantificação de γH2AX Focos em amostras de tecido

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.

Avaliação da Distribuição Espacial da γH2AX seguintes Radiação Ionizante

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.

Clonogênica Ensaio: células aderentes

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.

Imagens ao vivo de Apuramento celular por apoptose durante<em&gt; Drosophila</em&gt; Embriogênese

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.

Avaliação do cálcio Faíscas intactas fibras musculares esqueléticas

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