Este trabalho foi financiado por subvenções GM096076 do programa ÁREA NIH.

1. Preparando Moscas para Dissecção <ol><li> Anestesiar moscas saudáveis ​​com o genótipo desejado (por exemplo, expressando uma proteína marcada com GFP) que estão a menos de uma semana de idade. Selecione 10-15 fêmeas grandes com arredondadas de cor creme abdomens. </li><li> Transferir as moscas seleccionados e alguns (3-5) do sexo masculino para um frasco contendo alimentos ligeiramente fermentado novo e permitir que as moscas para engordar durante dois dias a 25 ° C. Referem-se a Weil et al. 6 para detalhes de preparação Drosophila. Engorda as moscas garante que uma variedade de fases, especialmente fases de 8-12, estará disponível para a imagem latente. Mal engordados ou unfattened voa geralmente contêm apenas estágios iniciais (1-6) e de maturação (estádio 14) ovócitos. </li></ol> 2. Preparação de oócitos for Imaging Usando um microscópio composto Vertical <ol><li> Prepare uma lâmina de vidro padrão pela aposição duas lamínulas para o slide, cerca de 1 cm de distância, usando greas siliconee. Só use graxa suficiente para garantir uma boa vedação entre a lamela e slides. Pressionando firmemente em cada lamínula a vai se espalhar a massa fina e uniforme. Adicionar algum óleo halocarbono 27 (Sigma) para a junção entre as lamelas e corrediça. Eles servirão como espaçadores para evitar ferir ovócitos isolados em etapas subseqüentes. Não. 1,5 lamínulas trabalhar bem, com uma espessura média (0,16-,18 mm) que coincide com a de oócitos no estágio final. </li><li> Colocar 2-3 gotas de óleo de halocarbono 27 sobre a superfície de uma lamela de 22 mm 2. Para a obtenção dos oócitos saudáveis ​​possíveis, uma fêmea individual é removida do frasco através de pipetagem alimento boca e suavemente depositada no óleo hidrocarboneto halogenado sem anestesiante primeiro. </li><li> Usando afiadas pinças de dissecação (como Dumont n º 5), pino a voar contra a lamínula e retirar a ponta do abdômen. Remover os ovários, arrastando-os ao longo da superfície da lamela, ao abrigo do óleo. Isto promoverá adhesion dos oócitos para a lamela. </li><li> Suavemente separar os ovários, procurando oócitos que são pelo menos 10B fase de oogênese. Oócitos, nesta fase pode ser identificada por câmaras de ovo em que o oócito ocupa, pelo menos, 50-60% do volume total da câmara do ovo. Neste ponto da oogênese, as células foliculares que recobrem o oócito se tornaram colunar, e rodeiam o oócito em todos os lados com excepção de uma pequena região onde as ligações para as células permanecem enfermeira. Use a pinça para remover oócitos individuais da bainha ovaríolo. </li><li> Prossiga, usando fêmeas 1-3, 5-8 até oócitos não danificados foram isolados na lamela. Remova qualquer tecido inutilizável. </li><li> Cuidadosamente inverter a lamela contendo os oócitos na lâmina pré-preparada de modo a que os oócitos estão posicionadas entre os dois espaçadores. Não pressione a lamela invertido como que vai danificar os ovócitos. Se for necessário, adicionar uma pequena quantidade de óleo de halocarbono para tele bordas do &quot;sanduíche&quot; para garantir o espaço é preenchido. (Opcional:. Deixe resto corrediça durante vários minutos para permitir a sedimentação de lamela) A lâmina está agora pronto para a imagiologia. </li></ol> 3. Preparação de oócitos for Imaging Usando um microscópio composto invertido <ol><li> Os oócitos são dissecados, essencialmente como descrito na secção 2, exceto que as pequenas placas de Petri com um inserto de fundo de vidro (MatTek, 35 mm) são usados ​​em vez de lamelas. Não é necessário para cobrir a amostra, e após o esvaziamento, o tecido pode ser trabalhada diretamente nos pratos, mas os cuidados devem ser tomados para garantir os oócitos permanecem cobertas com óleo e não são autorizados a secar. </li></ol> 4. Imagens ao vivo <ol><li> Traga um ovócito em foco usando DIC ou óptica de contraste de fase e examinar o oócito por quaisquer sinais de danos, tais como vazamento de citoplasma, ou abaulamento de células do folículo. Oócitos imagem apenas de aparência saudável e sem danos. </li><li> Streaming pode ser monitorada directamente, sem a necessidade de rotulagem GFP, uma vez que as vesículas de gema no citoplasma são autofluorescente 7. Este sinal pode ser captado na faixa FITC do escopo, mas provavelmente vai exigir ajustes de ganho mais elevado do que os utilizados para proteínas GFP-rotulados. </li><li> As melhores imagens do fluxo são obtidas a partir de secções ópticas que estão logo abaixo da superfície do oócito que está encostada ao vidro de vidro lamela / Petri. Aqui, o oócito é ligeiramente achatada, para fazer um melhor plano de foco. As imagens podem ser capturadas a 200X - 1000 X. Usando a disposição aqui descrita, os objectivos de ar deve ser utilizado, uma vez que a lamela forma uma barreira entre o oócito e objectiva. Isto é óptimo porque minimiza bastante deriva e movimento da amostra. </li><li> Uma vez que uma região de interesse está em foco, desligue a ótica brighfield e mudar para imagem confocal. Usando a função de digitalização / visualização rápida do software, ajustar o foco e ganharna medida do necessário. </li><li> Definições para captura irão variar de microscópio para microscópio, mas parâmetros gerais são as seguintes: Utilizar um comprimento de onda de excitação de ~ 488 nm e um comprimento de onda de emissão de ~ 515 nm. Defina-se a captura do eixo Z de modo que o eixo Z permanece constante, com um atraso de 10 segundos entre os quadros. Capturar uma sequência de teste de quadros 5-10 para assegurar o ovócito é estacionário e o plano de foco é apropriado. </li><li> Uma sequência típica de lapso de tempo irá capturar entre 20 - 50 quadros, e podem ser analisados ​​imediatamente após a conclusão de avaliar a qualidade de vídeo. O processo pode ser repetido para outras células do slide, ou fêmeas adicionais podem ser dissecados conforme necessário. </li></ol> 5. Solução de problemas e comum <ol><li> Deriva - Ao utilizar um microscópio vertical, as células podem, por vezes, deriva devido à propagação de óleo. Isto pode ser minimizado utilizando apenas óleo suficiente para cobrir a área sob a lamela. Se o problema persistir, use g menos siliconerease a espaçadores colar no slide. </li></ol> Nota: Alguns pacotes de software permitem confocal para rastreamento de células, mas em geral, é melhor abandonar imagem de oócitos à deriva e, em vez usar um outro espécime. <ol start="2"><li> Sem sinais de transmissão de streaming - Se não é evidente, é provável devido a uma de duas causas: ou o plano focal para a captura de imagem não está correto (geralmente muito profundamente no interior do ovócito) ou o oócito está danificado. O tempo ea prática pode minimizar os erros. </li></ol>

<ol>
	<li>Becalska, A. N., Gavis, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lighting+up+mRNA+localization+in+Drosophila+oogenesis.">Lighting up mRNA localization in Drosophila oogenesis.</a> Dev. 136, 2493-2503 (2009).</li><li>He, L., Wang, X., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shining+light+on+Drosophila+oogenesis:+Live+imaging+of+egg+development.">Shining light on Drosophila oogenesis: Live imaging of egg development.</a> Curr. Opin. Gen. Dev. 21, 612-619 (2011).</li><li>King, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Ovarian development Drosophila melanogaster. , (1970).</li><li>Spradling, A. C., Bate, M., Martinez-Arias, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+genetics+of+oogenesis.">Developmental genetics of oogenesis.</a> The Development of Drosophila melanogaster. 1, 1-70 (1993).</li><li>Serbus, L. R., Cha, B. -. J., Theurkauf, W. E., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynein+and+the+actin+cytoskeleton+control+kinesin-driven+cytoplasmic+streaming+in+Drosophila+oocytes.">Dynein and the actin cytoskeleton control kinesin-driven cytoplasmic streaming in Drosophila oocytes.</a> Dev. 132, 3743-3752 (2005).</li><li>Weil, T. T., Parton, R. M., Davis, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparing+Individual+Drosophila+Egg+Chambers+for+Live+Imaging.">Preparing Individual Drosophila Egg Chambers for Live Imaging.</a> J. Vis. Exp. (60), e3679 (2012).</li><li>Theurkauf, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Premature+microtubule-dependent+cytoplasmic+streaming+in+cappuccino+and+spire+mutant+oocytes.">Premature microtubule-dependent cytoplasmic streaming in cappuccino and spire mutant oocytes.</a> Science. 265, 2093-2096 (1998).</li><li>Morin, X., Daneman, R., Zavortink, M., Chia, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protein+trap+strategy+to+detect+GFP-tagged+proteins+expressed+from+their+endogenous+loci+in+Drosophila.">A protein trap strategy to detect GFP-tagged proteins expressed from their endogenous loci in Drosophila.</a> PNAS. 98, 15050-15055 (2001).</li><li>R&#246;per, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rtnl1+is+enriched+in+a+specialized+germline+ER+that+associates+with+ribonucleoprotein+granule+components.">Rtnl1 is enriched in a specialized germline ER that associates with ribonucleoprotein granule components.</a> J. Cell Sci. 120, 1081-1092 (2007).</li><li>Pokrywka, N. J., Payne-Tobin, A., Raley-Susman, K. M., Swartzman, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubules,+the+ER+and+Exu:+New+associations+revealed+by+analysis+of+mini+spindles+mutations.">Microtubules, the ER and Exu: New associations revealed by analysis of mini spindles mutations.</a> Mech. Dev. 126, 289-300 (2009).</li><li>Petri, W. H., Mindrinos, M. N., Lombard, M. F., Margaritis, L. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+development+of+the+Drosophila+chorion.ih+a+chemically+defined+organ+culture+medium.">In vitro development of the Drosophila chorion.ih a chemically defined organ culture medium.</a> Wilhelm Roux's Arch., Devl. Biol. 186, 351-362 (1979).</li></ol>

Eu não tenho nada para divulgar.

Oócitos Drosophila são um sistema modelo versátil para abordar uma série de questões relacionadas com o funcionamento e organização das células e componentes subcelulares. Uma compreensão completa da função da proteína requer a monitorização de ambos os aspectos espacial e temporal do comportamento de proteínas. Avanços em ambos os sistemas de imagiologia e as ferramentas genéticas disponíveis para Drosophilists tornar possível, mesmo para pequenos laboratórios para executar as experiências de imagiologia de alta qualidade em oócitos de vida. Aqui I delinear um protocolo para analisar o comportamento das proteínas GFP marcadas com ou autofluorescente e estruturas durante a oogénese mid-late-que pode ser utilizado em conjunto com uma variedade de origens genéticas para sondar a função da proteína. Neste protocolo I descrever o processo pelo qual os oócitos são preparadas para a imagem latente e as configurações básicas confocal que permitirão aquisição de vídeo de lapso de tempo de proteínas fluorescentes e estruturas. Detai adicionalls de preparação de um oócito são descritos por Weil et al. 6 Uma grande variedade de linhagens de moscas que expressam proteínas que foram marcadas endogenamente via exon-trapping estão disponíveis 8, e suas variantes proteicas infinitamente mais podem ser manipuladas e reintroduzidos moscas. Ao trabalhar com o tecido vivo, a saúde dos espécimes é de importância primordial. Câmaras de ovos de 10B estágio pode completar oogênese de uma forma essencialmente autônomos 11, tornando-os ideais para estudos de curto prazo de imagem. Cuidados devem ser tomados em vários passos-chave para obter dados de alta qualidade. Durante a dissecção, é importante para manipular ovários e oócitos tão pouco quanto possível, e para minimizar a quantidade de tempo entre a dissecção e a conclusão de imagiologia. Idealmente, uma estação de dissecção pequena deve estar localizado na mesma sala que abriga o escopo confocal, e apenas a poucas câmaras de ovos de uma vez deve ser trabalhada. O número recomendado aqui (5-8) Garante que o tempo de dissecção é minimizada, enquanto maximiza a probabilidade de obtenção de imagens de boa qualidade. Eu recomendo capturar uma pequena série de imagens para avaliar a qualidade de imagem e confirmar que as configurações de captura são otimizados, antes de capturar uma sequência mais lapso de tempo. A maioria dos softwares permite quadros individuais a ser salva, e essas imagens podem então ser analisados ​​numa variedade de maneiras, usando ImageJ 12 ou outro software.

<table border="1"><tbody><tr><td> Nome </td><td> Companhia </td><td> Número de catálogo </td><td> Comentários </td></tr><tr><td> Halocarbon óleo 27 </td><td> Sigma-Aldrich </td><td> H8773-100ML </td><td></td></tr><tr><td> # 5SF Fórceps </td><td> Belas Science Tools </td><td> 11252-00 </td><td></td></tr><tr><td> Graxa de silicone </td><td> Sigma Aldrich </td><td> Z273554-1EA </td><td></td></tr><tr><td> Com fundo de vidro placas de Petri </td><td> MatTek </td><td> P35G-0-20-C </td><td></td></tr></tbody></table>

live imaging of gfp-labeled proteins in drosophila oocytes

O oócito Drosophila tem sido estabelecido como um sistema versátil para a investigação de questões fundamentais, como função do citoesqueleto, a organização das células e organelas estrutura e função. A disponibilidade de várias proteínas GFP-tagged significa que muitos processos celulares podem ser controlados em células vivas ao longo de alguns minutos ou horas, e usando esta técnica, os processos tais como o transporte de RNP, morfogénese epitelial, e remodelação de tecidos têm sido descritos em grande detalhe em Drosophila 1,2 oócitos. A capacidade de realizar imagens de vídeo combinado com um rico repertório de mutantes permite uma enorme variedade de genes e processos a serem examinados em detalhe incrível. Um exemplo disso é o processo de fluxo contínuo, que inicia ooplasmic em meados de oogénese 3,4. Este movimento vigoroso de vesículas citoplasmáticas é microtúbulos e kinesin-dependente 5 e fornece um sis úteisma de investigação da função do citoesqueleto desses estágios. Aqui eu apresento um protocolo para imagens de lapso de tempo de oócitos de vida de praticamente qualquer configuração de microscopia confocal.

Imagiologia de proteínas GFP marcadas com variará dependendo de quão fortemente a proteína marcada é expresso. A fase mid-oócito expressando Reticulon-like 1 (Rtnl-1), marcado com GFP exon 8,9 produz um sinal forte. Rtnl-1 parece co-localizar com os microtúbulos longas presentes durante o fluxo ooplasmic 10 (Figura 1A). Rtnl-1 é um bom marcador para a observação de transmissão, e é também um indicador de organização de microtúbulos em oócitos de fase tardia. Streaming de Ooplasmic numa câmara de ovo tipo selvagem é evidente que o movimento perceptível de vesículas gema autofluorescente ao usar uma taxa de captura de um quadro a cada 10 segundos (Figura 1B). Uma projecção máxima de cinco quadros podem ser gerados para marcar o caminho de vesículas em movimento. <img alt="Figura 1" src="/files/ftp_upload/50044/50044fig1.jpg" /> Figura 1. Vivo imaging de ambos autofluorescent e GFP-estruturas. marcado A) Uma imagem lapso de tempo único de um oócito fase 11 Rtnl1 expressando-GFP em 400X. Rtnl1-GFP fibras mover com um movimento ondulatório em oócitos streaming. B) A projeção máxima de cinco quadros de uma câmara de ovo fase 10B em que as vesículas autofluorescente gema foram fotografadas a cada 10 segundos para 50 total seg, com ampliação de 400X. Barra de escala = 30 mm

Watch this Scientific Journal Video about Live Imaging of GFP-labeled Proteins in Drosophila Oocytes at JoVE.com

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Um protocolo para geração de imagens ao vivo das proteínas GFP-tagged ou estruturas autofluorescente no indivíduo<em&gt; Drosophila</em&gt; Oócitos é descrito.

Imagens ao vivo de Proteínas GFP-rotulados em<em&gt; Drosophila</em&gt; Oócitos

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

live-imaging-of-gfp-labeled-proteins-in-drosophila-oocytes

Research

JoVE Journal

Biology

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

11.3K visualizações.  Vassar College. Um protocolo para geração de imagens ao vivo das proteínas GFP-tagged ou estruturas autofluorescente no indivíduo Drosophila Oócitos é descrito.

Vídeo: Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Imagens ao vivo da migração das células da glia no Disco Drosophila Eye Imaginal

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

Biologia

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Elaboração de Drosophila Os embriões para Live-Imaging Usando o protocolo gota suspensa

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

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

Imagens ao vivo de Drosophila melanogaster Embrionárias Migrações hemócitos

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

Labeling fluorescentes de COS-7 expressando as proteínas SNAP tag-Fusion para imagens de células vivas

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

Preparação de Desenvolvimento e Adultos Drosophila Brains e retina de imagens ao vivo

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Imagem vertical de Drosophila Os embriões

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Imagem Alterar Forma Celular em Viver Drosophila Os embriões

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Preparação Individual Drosophila Ovos Câmaras de imagens ao vivo

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Duas e três dimensões imagens de células vivas de Proteínas de DNA resposta a danos

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.