Agradecemos p. Verrijzer, J. Johansen H., Okhura por anticorpos e A. Debec, os centros de ações bloomington e viena para moscas. Obrigado a C. Jackson por sugestões e discussões estimulantes.

1. Coleção de testículos de Drosophila
NOTA: Machos jovens (0-15 h pós-eclosão) são necessários para examinar células germinativas diploides (ou seja, espermatogonia e espermatocitocitos).
<ol><li>Selecione moscas jovens o máximo possível para minimizar a interferência do desenvolvimento de espermatóides.</li>
	<li>Anesthetize voa por breve exposição ao dióxido de carbono e transferência para uma plataformade moscas 14,15.</li>
	<li>Sob um microscópio dissecando (ampliação de 10x), use fórceps para remover a cabeça.</li>
	<li>Transfira 5 a 10 moscas decapitadas para 100 μL do buffer de Ringer (183 mM KCI, 47 mM NaCl, 10 mM Tris-HCI, 1 mM EDTA, inibidor completo de proteína, pH 6.8) em uma lâmina de vidro revestida de silicone em um suporte de fundo preto.</li>
	<li>Com microscópio dissecado em uma ampliação de 40x, use um par de fórceps número 5 para segurar uma mosca entre o tórax e o abdômen, e, com outros fórceps, puxe o abdômen para longe do tórax. Isole rapidamente os testículos e tecidos adjacentes da carcaça15 e coloque em uma nova gota do tampão de Ringer no mesmo slide.</li>
	<li>Quando todos os testículos estiverem prontos, limpe os testículos dos tecidos restantes (glândula acessório e genitália externa).</li>
	<li>Remova o excesso de tampão da gota contendo os testículos dissecados. Normalmente, use uma pipeta p200 com uma ponta de 10 μL instalada em cima de uma ponta de 200 μL. Isso permite a remoção do excesso de líquido através de uma pequena abertura. Coloque uma queda de 4% de paraformaldeído fixador em cima da gota contendo os testículos dissecados e, em seguida, transfira rapidamente os testículos para um tubo contendo solução fixativa fresca.</li>
</ol>2. Fixação de paraformaldeído
<ol><li>Prepare 2 mL de solução fixa pouco antes do uso: 600 μL de 4% de paraformaldeído diluído em soro fisiológico tamponado de fosfato (PBS) com 30 μL de NP40 20% mais 1200 μL de heptano em um tubo de microcentrifuge siliconado de 2 mL. Use 4% de paraformaldeído em PBS preparado em alíquotas de 600 μL e armazenado a -20°C. Atenção: Heptano e paraformaldeído são tóxicos e devem ser manuseados em um capô de fumaça. Elimine-os de acordo com as normas do instituto anfitrião.</li>
	<li>Transfira os testículos usando uma micropipette p200 com uma ampla ponta de abertura para a solução PFA/NP40/heptane. Aperte o tubo para cima e para baixo por 30 s à mão. Continue consertando os testículos em uma batedeira rotativa à temperatura ambiente por 30 minutos.</li>
	<li>Pare a batedeira rotativa e coloque o tubo em um rack de tubo para permitir a separação de fase e para que os testículos se acomodem na parte inferior do tubo.</li>
	<li>Remova todo o heptano e a solução fixa e enxágue os testículos rapidamente três vezes com 1x PBS.</li>
	<li>Transfira para um tubo individual limpo de 200 μL. Realizar todas as incubações nestes pequenos tubos para minimizar as quantidades de anticorpos empregados.</li>
	<li>Remova o excesso de tampão usando uma micropipette de 200 μL com uma p200 e uma ponta P10 anexada. Tome cuidado para não aspirar testículos.</li>
	<li>Mantenha os testículos em 1x PBS no gelo até que todas as amostras estejam prontas.</li>
</ol>3. Coloração de anticorpos em solução
<ol><li>Descarte o PBS e deixe testículos em 0,3% PBS-T por 30 minutos à temperatura ambiente para permeabilizar as membranas celulares.</li>
	<li>Pré-incubação no mesmo tampão mais 5% de soro de cabra normal (NGS) por 1 h.</li>
	<li>Adicionar 200 μL do anticorpo primário diluído em 0,3% PBS-T mais 5% NGS(Tabela de Materiais).</li>
	<li>Incubar o anticorpo primário durante a noite a 4 °C (ou temperatura ambiente) com agitação suave em um roqueiro.</li>
	<li>Lave 3 vezes em 0,3% PBS-T por 15 minutos em um roqueiro à temperatura ambiente. Deixe os testículos se acomodarem na parte inferior do tubo pela gravidade.</li>
	<li>Adicione 200 μL de anticorpo secundário diluído 1:500 da série conjugada DyLight.</li>
	<li>Incubar o anticorpo secundário em uma câmara escura por 4h à temperatura ambiente.</li>
	<li>Lave em 0,3% PBS-T por 15 minutos em um roqueiro à temperatura ambiente. Em seguida, repita com 0,2% PBS-T e 0,1% PBS-T, cada vez por 30 min em um roqueiro à temperatura ambiente. Faça uma lavagem final em 1x PBS por 30 minutos para remover o detergente.</li>
	<li>Adicione 180 μL de solução DAPI a 1 μg/mL em 1x PBS por 15 min. A solução de estoque é de 10 mg/mL em H2O. Evite fazer solução de estoque em PBS, o que pode reduzir o sinal de coloração da DAPI em cromatina difusa em espermatocitocitos maiores.</li>
	<li>Lave 3x por 5 min cada.</li>
	<li>Use uma caneta de barreira hidrofóbica para desenhar um círculo em um escorregador limpo.</li>
	<li>Aspirar testículos e colocá-los no slide. Pré-enxaguar a ponta de abertura larga com 0,1% pbs-t impede que os testículos grudem na ponta. Remova o excesso de PBS.</li>
	<li>Adicione uma gota (aproximadamente 100 μL) do Citifluor AF1.</li>
	<li>Coloque delicadamente uma tampa de vidro sobre o tecido, tomando cuidado para evitar a captura de bolhas de ar.</li>
	<li>Sele a mancha de cobertura no slide usando esmalte transparente.</li>
	<li>Observe slides em um microscópio.</li>
</ol>4. Análises de colocalização
<ol><li>Adquira imagens confocal. Utilizamos um microscópio confocal Zeiss LSM 710 (óleo apocromático plano 63x) com uma resolução z de 0,5 ou 1 μm.</li>
	<li>Selecione diferentes células de testículos independentes.</li>
	<li>Importar imagens para software de processamento (por exemplo, Imaris).</li>
	<li>Defina o núcleo por segmentação manual, a fim de excluir os núcleos vizinhos. Use o software Coloc; ele coleta os coeficientes de correlação (PCCs) do Pearson dentro de todo o volume entre 2 fluoroforos.</li>
</ol>

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Os autores não têm nada a revelar.

Este protocolo fornece um método para o bem sucedido imunolabeling de testículos de Drosophila adultos de montagem inteira. Conforme relatado em outros procedimentos, devem ser utilizados os machos jovens (até encurtamos a idade para menos de 20 horas após a eclosão) para minimizar a presença de espermatozoides em desenvolvimento, o que corre o risco de obstruir os espermatocitos. A dissecção deve ser rápida e a diluição do anticorpo deve ser ajustada para otimizar a relação sinal-ruído21,...

Os testículos de drosophila são um modelo valioso para o estudo da arquitetura nuclear. Em células espermatogoniais mitóticas, o volume nuclear é em grande parte ocupado por cromatina. Essas células passam por quatro rodadas de divisão, produzindo um cisto de 16 espermatos primários. Nos dias seguintes, os espermatocitos passam por 6 estágios de desenvolvimento (S1-S6), aumentando consideravelmente em volume, aproximadamente 20vezes 1,2. Eles também mudam sua morfologia nuclear. O contorno do núcleo, revelado pelo lamin Dm0, torna-se irregular e mostra profundas invaginações1,3. Isso é acompanhado por extensas alterações de expressão genética e modificações na organização cromatina1,4,5,6,7. Os cromossomos interfásicos são bem definidos no estágio S4-S5, formando três massas distintas correspondentes aos 2 principais autossomos bivalentes e o par X-Y ancorado com o nucleolus.
Mad1 foi originalmente isolado como um componente-chave do ponto de verificação mitótico (revisado em8). Na interfase, é descrito como localizado principalmente no envelope nuclear, mas também no nucleoplasma9,10,11. Em Drosophila testis, relatamos que Mad1 é proeminente no nucleoplasma, intimamente associado com as duas principais massas de cromatina autossômica e em menor grau com a cromatina XY. Definimos esta estrutura associada à cromatina como MINT (Território IntraNuclear contendo Mad1)12. Quatro outras proteínas foram associadas com MINTs em espermatotos: o nucleoporina Mtor/Tpr, o SUMO peptidase Ulp1, a proteína de checkpoint mitotístico Mad2 e uma subunidade de um complexo de Policomb Raf212.
Para completar a descrição dos MINTs, precisávamos usar anticorpos e fomos confrontados com os problemas técnicos geralmente encontrados com imunostendências em testículas: uma má permeabilidade resultando em uma significativa não uniformidade da coloração na profundidade dos testítes, particularmente nos grandes espermatocitos. Aspreparações anuladas aumentaram o acesso a anticorpos, mas levaram a imagens compactadas, restringindo as análises 3D e impedindo a medição da fluorescência quantitativa, incluindo a colocalização. O problema da penetração de anticorpos também poderia ser resolvido por agentes permeabilizadores como 0,3% Na desoxicolato13,mas este procedimento não resultou em nossas mãos resultados reprodutíveis. Como realizamos muitos experimentos de co-rotulagem, procuramos melhorar o protocolo de permeabilização. A combinação de 1% NP40 mais heptano resultou em mais reprodutibilidade, particularmente no estudo dos grandes espermatos.
Este protocolo realiza a fixação e a imunostenção de testículos na suspensão, melhorando a qualidade da amostra, bem como melhorando a reprodutibilidade, e tornando-a adequada para microscopia confocal. Usamos o protocolo para definir melhor as relações espaciais de certas proteínas de envelope nuclear com as estruturas nucleoplasmáticas de grandes espermatotos. Esse método permitirá melhores investigações das mudanças que acompanham o desenvolvimento de espermatotocitos, como por exemplo as dinâmicas mudanças estruturais do núcleo, incluindo cromatina, nucleoplasma e os poros nucleares.

<table><tbody><tr><td>&lt;em&gt;&lt;strong&gt;Drosophila srains&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>ketelGFP y; ketelGFP/y+CyO</td><td></td><td></td><td>Gift from A. Debec (Institut Jacques Monod, UMR 7592, Paris, France)</td></tr><tr><td>Mad1-GFP</td><td></td><td></td><td>J Cell Sci. 2011 May 15;124(Pt 10):1664-71.</td></tr><tr><td>Mtor RNAi&amp;nbsp;</td><td>VDRC</td><td>ID 110218</td><td>Vienna Drosophila RNAi Center</td></tr><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DAPI&amp;nbsp;</td><td>Thermo</td><td>D1306</td><td>Stock solution must be prepared in H2O</td></tr><tr><td>Dumont # 5 Fine Forseps&amp;nbsp;</td><td>Fine Science Tools</td><td></td><td></td></tr><tr><td>MBP Wide Bore Pipette Tips</td><td>Fisherbrand</td><td>11917744</td><td>Wide aperture tip</td></tr><tr><td>Thermo Scientific&amp;nbsp;0.2 mL Individual Tube</td><td>Thermo</td><td>AB-0620</td><td></td></tr><tr><td>2 ml micro centrifuge tubes&amp;nbsp;</td><td>Sigma</td><td>T3531-200EA&amp;nbsp;</td><td>Siliconized 2ml tube</td></tr><tr><td>Citifluor AF1</td><td>Citifluor</td><td>AF1</td><td></td></tr><tr><td>Dakopen&amp;nbsp;</td><td>Sigma</td><td>Z377821-1EA&amp;nbsp;</td><td></td></tr><tr><td>Normal Goat Serum (NGS)</td><td>Sigma</td><td>NS02L-1ML</td><td></td></tr><tr><td>Paraformaldehyde 4%</td><td>Sigma</td><td>P6148-500G&amp;nbsp;</td><td>Paraformaldehyde (4%) dissolved in PBS 1X&amp;nbsp;; aliquot by 600 ml&amp;nbsp;</td></tr><tr><td>PBS 1X</td><td>Thermo</td><td>14190094</td><td>DPBS, no calcium, no magnesium</td></tr><tr><td>0.1% (0.2 or 0.3%) PBS-T&amp;nbsp;</td><td></td><td></td><td>PBS 1X&amp;nbsp; containing 0.1% (0.2 % or 0.3%) Triton X-100</td></tr><tr><td>Triton X-100, for molecular biology,&amp;nbsp;</td><td>Sigma</td><td>T8787</td><td></td></tr><tr><td>&lt;strong&gt;Antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>GFP booster</td><td>Chromotek</td><td>gba488</td><td>1/200</td></tr><tr><td>Mouse anti-Mtor</td><td></td><td></td><td>1/40 gift from J. Johansen, Iowa State University</td></tr><tr><td>Rat anti-Nup62</td><td></td><td></td><td>1/200 gift from H. Ohkura (University of Edinburgh, UK</td></tr><tr><td>Guinea-pig anti-Ulp1</td><td></td><td></td><td>1/200 gift from C. P. Verrijzer, Erasmus University, Rotterdam</td></tr><tr><td>Rabbit anti-Raf2</td><td></td><td></td><td>1/200 gift from C. P. Verrijzer, Erasmus University, Rotterdam</td></tr><tr><td>DyLight conjugated series&amp;nbsp;</td><td>Thermo</td><td></td><td>1/500 Donkey anti-(guinea-pig, mouse,&amp;nbsp; rabbit or rat) as second antibodies</td></tr></tbody></table>

immunostaining of whole-mount drosophila testes for 3d confocal analysis of large spermatocytes

Os testículos de drosophila são um poderoso sistema modelo para estudar processos biológicos, incluindo biologia de células-tronco, arquitetura nuclear, meiose e desenvolvimento de espermatozoides. No entanto, a imunolabelação de todo o teste de Drosophila é frequentemente associada com significativa não uniformidade da coloração devido à penetração de anticorpos. As preparações esmagadas só superam parcialmente o problema, pois diminui a qualidade 3D das análises. Aqui, descrevemos um protocolo de montagem inteira usando NP40 e heptano durante a fixação, juntamente com a imunolabelagem em mídia líquida. Preserva o volume adequado para microscopia confocal juntamente com rotulagem reprodutível e confiável. Mostramos diferentes exemplos de reconstituição 3D de núcleos espermatocitocitos de seções confocal. A reprodutibilidade intra e inter-testes permite quantificação 3D e comparação da fluorescência entre células únicas de diferentes genótipos. Utilizamos diferentes componentes da estrutura de MINT intranuclear (Território Intra Nuclear Contendo Mad1), bem como dois componentes associados ao complexo de poros nucleares para ilustrar este protocolo e suas aplicações nas maiores células dos testis, os espermatozoides S4-S5.

O principal obstáculo para a obtenção de uma coloração adequada dos testículos de Drosophila é a penetrabilidade limitada dos anticorpos, que foi parcialmente resolvida por meio de preparações esmagadas, mas ao custo de induzir uma restrição das análises 3D (por exemplo, representação 3D ou medida de colocalização). O procedimento permite rotulagem uniforme e preserva o volume de todas as células do testis. Aqui focamos a ilustração do método na representação 3D dos espermatos S4-S5, pois são as ma...

Watch this Scientific Journal Video about Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes at JoVE.com

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Apresentamos aqui um protocolo aprimorado para a preparação e imunossagem completa de testículos Drosophila, adequados para microscopia confocal e também permitindo rotulagem reprodutível e confiável. Ilustramos este protocolo por representação 3D e quantificação de experimentos de colocalização nos grandes espermatos S4-S5.

Imunostaining de Testes Drosophila de Monte Inteiro para Análise Confocal 3D de Grandes Espermatocitos

Drosophila testes are a powerful model system for studying biological processes including stem cell biology, nuclear architecture, meiosis and sperm ...

immunostaining-whole-mount-drosophila-testes-for-3d-confocal-analysis

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JoVE Journal

Biology

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

4.1K visualizações.  Université de Paris, F 75006, Paris, France. Apresentamos aqui um protocolo aprimorado para a preparação e imunossagem completa de testículos Drosophila, adequados para microscopia confocal e também permitindo rotulagem reprodutível e confiável. Ilustramos este protocolo por representação 3D e quantificação de experimentos de colocalização nos grandes espermatos S4-S5.

Vídeo: Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Imagem Vertical de<em&gt; Drosophila</em&gt; Câmaras de ovo

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Biologia do Desenvolvimento

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Preparação de Drosophila Larval e Pupal Testes para Análise de Divisão Celular em Tecido Vivo e Intacto

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells entering meiotic prophase I are eliminated during Fetal Oocyte Attrition (FOA) and the first week of postnatal life. Three major mechanisms regulate the number of oocytes that survive during development and establish the ovarian reserve in females entering puberty. In the first wave of oocyte loss, 30-50% of the oocytes are eliminated during early FOA, a phenomenon that is attributed to high Long interspersed nuclear element-1 (LINE-1) expression. The second wave of oocyte loss is the elimination of oocytes with meiotic defects by a meiotic quality checkpoint. The third wave of oocyte loss occurs perinatally during primordial follicle formation when some oocytes fail to form follicles. It remains unclear what regulates each of these three waves of oocyte loss and how they shape the ovarian reserve in either mice or humans.
Immunofluorescence and 3D visualization have opened a new avenue to image and analyze oocyte development in the context of the whole ovary rather than in less informative 2D sections. This article provides a comprehensive protocol for whole ovary immunostaining and optical clearing, yielding preparations for imaging using multiphoton microscopy and 3D modeling using commercially available software. It shows how this method can be used to show the dynamics of oocyte attrition during ovarian development in C57BL/6J mice and quantify oocyte loss during the three waves of oocyte elimination. This protocol can be applied to prenatal and early postnatal ovaries for oocyte visualization and quantification, as well as other quantitative approaches. Importantly, the protocol was strategically developed to accommodate high-throughput, reliable, and repeatable processing that can meet the needs in toxicology, clinical diagnostics, and genomic assays of ovarian function.

Here, we present an optimized protocol for imaging entire ovaries for quantitative and qualitative analyses using whole-mount immunostaining, multiphoton microscopy, and 3D visualization and analysis. This protocol accommodates high-throughput, reliable, and repeatable processing that is applicable for toxicology, clinical diagnostics, and genomic assays of ovarian function.

Imunofluorescência total do ovário, compensação e microscopia multifotínica para análise quantitativa 3D da reserva ovariana em desenvolvimento em mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Dissecção e coloração de Drosophila Larval Ovários

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Biologia

Drosophila Pupal Abdomen Immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic morphologies 1-3. During pupation, which spans approximately 96 hours (at 25 &deg;C), proliferating populations of imaginal cells replace the larval epidermis to generate the adult abdominal segments. These imaginal cells, born during embryogenesis, exist as lateral pairs of histoblast nests in each abdominal segment of the larvae. Four pairs of histoblast nests give rise to the adult dorsal cuticle (anterior and posterior dorsal nests), the ventral cuticle (ventral nests) and the spiracles associated with each segment (spiracle nests) 4. Upon puparation, these diploid cells (distinguishable by size from the larger polyploid larval epidermal cells- LECs) begin a stereotypical process of proliferation, migration and replacement of the LECs. Various molecular and genetic tools can be employed to investigate the contributions of genetic pathways involved in morphogenesis of the adult abdomen. Ultimate adult phenotypes are typically analyzed following dissection of adult abdominal cuticles. However, investigation of the underlying molecular processes requires immunohistochemical analyses of the pupal epithelium, which present unique challenges. Temporally dynamic morphogenesis and the interactions of two distinct epithelial populations (larval and imaginal) generate a fragile tissue prone to excessive cell loss during dissection and subsequent processing. We have developed methods of dissection, fixation, mounting and imaging of the Drosophila pupal abdominem epithelium for immunohistochemical studies that generate consistent high quality samples suitable for confocal or standard fluorescent microscopy.

Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal abdomen.

Drosophila Imunohistoquímica Abdomen Pupal

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic ...

Imaging Centrosomes in Fly Testes

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell differentiation. Centrosomes are essential to determine the cell division axis during mitosis and to nucleate cilia during interphase. The identity of the proteins that mediate these dynamic changes remains only partially known, and the function of many of the proteins that have been implicated in these processes is still rudimentary. Recent work has shown that Drosophila spermatogenesis provides a powerful system to identify new proteins critical for centrosome function and formation as well as to gain insight into the particular function of known players in centrosome-related processes. Drosophila is an established genetic model organism where mutants in centrosomal genes can be readily obtained and easily analyzed. Furthermore, recent advances in the sensitivity and resolution of light microscopy and the development of robust genetically tagged centrosomal markers have transformed the ability to use Drosophila testes as a simple and accessible model system to study centrosomes. This paper describes the use of genetically-tagged centrosomal markers to perform genetic screens for new centrosomal mutants and to gain insight into the specific function of newly identified genes.

Imaging of centrosomal proteins during Drosophila spermatogenesis is a powerful method to identify new proteins critical for centrosome biology as well as to elucidate the particular function of known players in this process.

Centrossomas de imagem na mosca Testículos

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Citológica Análise da espermatogênese: Preparados ao vivo e fixo<em&gt; Drosophila</em&gt; Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and organogenesis. Immunostaining allows for the visualization of developing tissues and organs. However, a protective cuticle that forms at the end of embryogenesis prevents permeation of antibodies into late-stage embryos and larvae. While dissection prior to immunostaining is regularly used to analyze Drosophila larval tissues, it proves inefficient for some analyses because small tissues may be difficult to locate and isolate. Sonication provides an alternative to dissection in larval Drosophila immunostaining protocols. It allows for quick, simultaneous processing of large numbers of late-stage embryos and larvae and maintains in situ morphology. After fixation in formaldehyde, a sample is sonicated. Sample is then subjected to immunostaining with antigen-specific primary antibodies and fluorescently labeled secondary antibodies to visualize target cell types and specific proteins via fluorescence microscopy. During the process of sonication, proper placement of a sonicating probe above the sample, as well as the duration and intensity of sonication, is critical. Additonal minor modifications to standard immunostaining protocols may be required for high quality stains. For antibodies with low signal to noise ratio, longer incubation times are typically necessary. As a proof of concept for this sonication-facilitated protocol, we show immunostains of three tissue types (testes, ovaries, and neural tissues) at a range of developmental stages.

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Facilitou-Sonication Imunofluorescência Coloração de Late-estágio embrionário e larval<em&gt; Drosophila</em&gt; Tecidos<em&gt; In Situ</em

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and ...

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes differentiation from a stem cell population, mitotic amplification, and meiosis. In addition, post-meiotic germ cells undergo a dramatic morphological reshaping process as well as a global epigenetic reconfiguration of the germ line chromatin&#8212;the histone-to-protamine switch.
Studying the role of a protein in post-meiotic spermatogenesis using mutagenesis or other genetic tools is often impeded by essential embryonic, pre-meiotic, or meiotic functions of the protein under investigation. The post-meiotic phenotype of a mutant of such a protein could be obscured through an earlier developmental block, or the interpretation of the phenotype could be complicated. The model organism Drosophila melanogaster offers a bypass to this problem: intact testes and even cysts of germ cells dissected from early pupae are able to develop ex vivo in culture medium. Making use of such cultures allows microscopic imaging of living germ cells in testes and of germ-line cysts. Importantly, the cultivated testes and germ cells also become accessible to pharmacological inhibitors, thereby permitting manipulation of enzymatic functions during spermatogenesis, including post-meiotic stages.
The protocol presented describes how to dissect and cultivate pupal testes and germ-line cysts. Information on the development of pupal testes and culture conditions are provided alongside microscope imaging data of live testes and germ-line cysts in culture. We also describe a pharmacological assay to study post-meiotic spermatogenesis, exemplified by an assay targeting the histone-to-protamine switch using the histone acetyltransferase inhibitor anacardic acid. In principle, this cultivation method could be adapted to address many other research questions in pre- and post-meiotic spermatogenesis.

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

This protocol describes the dissection and cultivation of intact testes and germ-line cysts from Drosophila melanogaster pupae. This method allows microscopic observation of spermatogenesis ex vivo. Furthermore, we describe a pharmacological assay of the effect of inhibitors on specific stages of germ-cell development in pupal testes.

Ex vivo Cultura de Drosophila Pupal Testículo e único macho da linha germinal Cistos: Dissection, Imaging, eo tratamento farmacológico

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

Imagem celular ao vivo super-resolução de estruturas subcelulares

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

Imunostaining de Testes Drosophila de Monte Inteiro para Análise Confocal 3D de Grandes Espermatocitos

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