We thank Stephan Uphoff for discussions. 
R.C. was supported by Linacre College, Oxford University. A.P. was supported by the German Academic Exchange Service (DAAD), the German National Academic Foundation and EPSRC. M.S. was supported by the Wellcome Trust. A.N.K. was supported by a UK BBSRC grant (BB/H01795X/1), and a European Research Council Starter grant (261227).

<ol>
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The authors have nothing to disclose.

Muitos parâmetros podem variar durante a electroporação de células e de aquisição de dados, dependendo do sistema biológico de interesse e a natureza exacta da experiência (de nível celular ou análise de uma única molécula). Por exemplo, quando electroporating ADN em bactérias de 0,25 a 5 pmol de fragmentos de dsDNA marcados conduz a uma baixa eficiência de internalização, permitindo que uma única molécula de detecção directa (isto é., Sem a necessidade de fotobranqueamento previamente). Acima de 5 pmol dsDNA, as células tendem a ser muito carregado, um regime mais adequado para a análise de uma única célula. Todos os DNAs marcados devem também ser previamente, a fim de eliminar qualquer vestígio de corante livre purificado em gel (fluoróforo não reagiu) a partir da solução de DNA. Além disso, possíveis problemas com a degradação do DNA, especialmente para experimentos smFRET, podem ser tratadas usando DNAs com ácidos nucleicos não naturais, ou motivos que protegem terminais exonuclease acessível como loops hairpin. Outra adjustable em parâmetro de electroporação é a força do campo aplicado durante electroporação. Força Low campo (~ 1 kV / cm) vai levar a uma baixa eficiência de carga apropriado para estudos de molécula única. Maior força de campo (até 1,8 kV / cm) irá aumentar a eficiência de carregamento; No entanto, existe uma correlação inversa entre a intensidade do campo e a viabilidade celular após a electroporação (ver Figura 4). Para referência, uma intensidade de campo normal utilizado para bactérias e leveduras electroporação é ~ 1,5 kV / cm. A constante de tempo, que representa o comprimento deste deterioração, é um parâmetro conveniente para seguir, uma vez que as gotas de constantes de tempo, assim que qualquer formação de arco fenómeno ocorre na cuvete. Em configurações normais, a constante de tempo deve ser superior a 4 ms; valores mais baixos levará a baixa eficiência de carga ou células danificadas, mesmo não-carregados. A maioria dos outros electroporators oferecer graus de liberdade (tais como &quot;truncagem de impulso&quot; ou &quot;forma de pulso&quot;) que pode ser modificado para ajustar tantocarregamento e viabilidade celular. Aplicou-se este método para ambas as bactérias e leveduras, contudo procedimentos semelhantes também deve permitir a internalização de biomoléculas marcadas em células de mamíferos usando o electroporador configurações adequadas desde a sua membrana é, na verdade, menos complexo (única bicamada lipídica), e uma vez que já electroporação foi usado com tais células 21. Quando internalizar proteínas marcadas, todos corante livre precisa ser removido a partir da solução de proteína marcada de electroporação anteriores. As moléculas livres de corantes, devido ao seu menor tamanho, podem ser internalizadas preferencialmente sobre as proteínas de interesse, e são difíceis de discriminar durante a análise de dados (não obstante o seu esperado difusão mais rápida). Como um guia, para uma amostra de proteína marcada orgânica como sendo adequados para a electroporação, a quantidade de corante remanescente livre deve ser inferior a 2% (detectado utilizando um varrimento fluorescente de um SDS-PAGE) 22. Este processo é particularmente importante,como algumas moléculas pode aderir às membranas externas de bactérias ou leveduras a electroporação. A este respeito, a amostra de controlo negativo devem exibir a intensidade de fluorescência por célula claramente mais baixo do que as células electroporadas, idealmente tão baixa quanto o nível de autofluorescência das células vazias (células que não foram incubadas com quaisquer biomoléculas marcadas com fluorescência ou a electroporação, Figura 2). Tal como acontece com dsDNA, a eficiência de internalização proteínas marcadas está relacionada com o valor de biomoléculas adicionados às células antes da electroporação. No entanto, outros parâmetros, tais como o tamanho e carga, desempenhar um papel na internalização. As pequenas proteínas apresentam alta eficiência de internalização, enquanto que as proteínas maiores (até 98 kDa) podem ser internalizados com êxito, mas com uma eficiência mais baixa (Figura 5) 26. O ponto isoelétrico da proteína, potenciais interacções com a membrana celular e os outros parâmetros físico-químicos tambémcélula de carregamento influência durante a electroporação. Como resultado, os usuários precisam otimizar experiências para o seu próprio sistema, sabendo que a concentração inicial de alta proteína marcada (&gt; 50 M) vai lhe dar a melhor chance para o carregamento de sucesso. A electroporação também oferece uma nova ferramenta para perturbar e analisar a função celular mediante a introdução de proteínas e outras biomoléculas para as células (ou marcados ou não marcados). As experiências de ARN polimerase de T7 (Figura 5C) apresentam um exemplo de uma experiência em que se pode introduzir uma biomolécula que pode alterar a expressão de genes in vivo utilizando electroporação. Ao realizar experiências de fluorescência de molécula única, iluminação TIR é geralmente favorecido sobre os outros modos de iluminação, uma vez que oferece a melhor relação sinal-para-ruído de apenas fluoróforos excitantes dentro de uma secção fina sobre a superfície lamela (~ 100 nm). No entanto imagem rotulada biomoléculas difundem dentro microorganismos vivos pode require iluminação mais profundas (até 0,8 mm para E. coli). Iluminação mais profundo é alcançado no modo de Hilo, preservando ao mesmo tempo uma elevada relação de sinal-para-ruído. Por outro lado, a imagem de campo largo é particularmente importante para a análise de fotobranqueamento passo a passo, em que o utilizador está a estimativa do número de moléculas internalizadas por fotobranqueamento toda uma célula carregada com elevado poder de laser e dividindo a intensidade de fluorescência celular inicial pela intensidade unitária produzida por uma única molécula (fotobranqueamento passo único, Figura 3). Widefield imagiologia também é necessário para a molécula de seguimento de longo prazo, a fim de localizar as moléculas de difusão de interesse mesmo que as suas trajectórias cobrir o volume da célula inteira. Neste protocolo, apresentamos como eletroporação, uma técnica padrão para biólogos e bioquímicos para a entrega de ácidos nucleicos em células, constitui um método simples para a entrega de biomoléculas fluorescentes em vários tipos de células. Thé nova, a técnica de alto rendimento oferece uma ferramenta única para observar moléculas marcadas em seu ambiente nativo. Em adição biomoléculas marcadas com fluoróforos que abrange uma ampla gama de comprimentos de onda, a electroporação pode entregar moléculas modificadas com vários grupos químicos, tais como nucleótidos e aminoácidos não naturais, quelantes de metais, agentes de reticulação, e grupos obtido confinando. Se o sistema biológico de interesse não é essencial para o desenvolvimento celular, o gene que codifica para a proteína alvo pode também ser eliminado (ou batido para baixo), assegurando que as proteínas observadas após a internalização representam todos (ou mais) do banco de proteína intracelular . Em essência, eletroporação pode &quot;transplante&quot; a flexibilidade do in vitro bioconjugates em células vivas e, portanto, beneficiar esforços em biologia sintética, biologia de sistemas, e in vivo de detecção de uma única molécula.

A maioria dos estudos de fluorescência dentro de células vivas depender de fusões de proteínas com proteínas fluorescentes (PQ), tais como 1 GFP. Estes marcadores fluorescentes permitir estudos do número de cópias, padrão de difusão ou localização de proteínas envolvidas em processos, tais como a expressão de genes ou de transporte de membrana 2-7. PQ oferecem alta especificidade rotulagem, de fácil aplicação, e estão disponíveis em um grande estoque de variantes com vários photophysical e propriedades químicas 1. No entanto, fluorophores orgânicos continuam a ser a principal escolha para experiências in vitro, devido à sua maior fotoestabilidade (até 100 vezes mais estável que o PQ) 8,9, tamanho pequeno (até 100 vezes menor do que o volume PQ) e facilidade de rotulagem intramolecular (principalmente através do uso de resíduos de cisteína). Todos estes factores são particularmente importantes para uma única molécula de fluorescência e FRET estuda 10. Vários métodos de internalização combining as vantagens da rotulagem orgânica e na detecção vivo foram introduzidas ao longo da última década; No entanto, tais métodos ou empregar polipéptidos relativamente grandes etiquetas (tags por exemplo, TMP, halo, ou 20 kDa SNAP) 11-14, requerem a utilização de aminoácidos não naturais 15, ou estão limitadas a células eucarióticas, grandes único à membrana (por ex. , raspe o carregamento, seringa carregamento, microinjeção) 16-19. Este protocolo descreve um novo, simples e de alto rendimento método internalização que acopla as vantagens de fluoróforos orgânicos com observação in vivo. Para o desenvolvimento desta técnica, foi adaptado do procedimento de electroporação comumente usado para transformar células com o plasmídeo de ADN 20,21 a fim de carregar os microrganismos, tais como E. coli ou S. cerevisiae com biomoléculas rotulados como orgânicos. O protocolo consiste em 4 passos simples: a incubação de células com biomoléculas marcadas,electroporação, a recuperação de células, e lavagem das células para remover biomoléculas não internalizado. Aqui, apresentamos este protocolo eletroporação, bem como os processos de imagem de células e de análise de dados para o estudo de fluorescência baseada em células e única molécula e traste sinais. A electroporação depende de descarregar um campo eléctrico de alta tensão através de uma suspensão de células de força iónica baixa para formar poros da membrana através da qual transientes biomoléculas podem entrar nas células (Figura 1) 20,21. Assim como com a transformação de bactérias ou de leveduras com DNA de plasmídeo, as células têm que ser preparada antes da electroporação a garantir a sua electrocompetency. Este procedimento, que consiste em vários passos de lavagem com água, aumenta a permeabilidade da membrana e diminui a força iónica da solução de células para evitar a formação de arco eléctrico na cuvete de electroporação. Neste protocolo, as células podem ser preparadas como descrito abaixo (Veja PROTOCOLO: 1.1) ou comprados a partir de provedor comercials. <img alt="Figura 1" src="/files/ftp_upload/52208/52208fig1.jpg" /> Figura 1: Representação esquemática do protocolo internalização Da esquerda para a direita:. Adicionar alguns microlitros de biomoléculas marcadas para a alíquota de células electrocompetentes (fragmentos de DNA duplamente marcada e bactérias, neste exemplo); incubar 1 a 10 min em gelo e transferir para uma cuvete de electroporação de pré-arrefecida; electroporate e depois adicionar 0,5-1 ml meio rico para as células imediatamente após; incubar a 37 ° C (ou a temperatura requerida pelo organismo, por exemplo, 29 ° C durante levedura) para permitir que as células recuperar; executar 5 passos de lavagem para remover qualquer excesso de moléculas não-internalizados marcados; ressuspender o sedimento final em 100-200 ul de tampão PBS e pipeta de 10 uL sobre uma almofada de agarose; cobrir o teclado com uma lamela e imagem limpa em um microscópio de fluorescência (no modo de campo largo ou modo HILO). <p class = &quot;jove_content&quot;&gt; células electrocompetentes são incubadas com as biomoléculas marcadas pouco antes eletroporação, que podem ser executadas usando electroporators padrão encontrados na maioria dos laboratórios de bioquímica. Imediatamente após a electroporação, as células são incubadas em meio enriquecido que permita a sua recuperação antes da lavagem (Figura 1). O excesso de biomoléculas marcadas não-internalizados é removido em primeiro lugar por lavagem com um tampão contendo uma concentração razoavelmente elevada de sal e um pouco de detergente (Ver PROTOCOLO: 3,3). A presença de sal perturba interacções electrostáticas não-específicos formados por não internalizados biomoléculas marcadas que de outra forma podem colar na membrana exterior. De igual modo, a presença de detergente no tampão de lavagem rompe as interacções hidrófobas não específicas. Enquanto internalização DNA é simples (Figura 2), as precauções devem ser tomadas quando internalizar proteínas marcadas utilizando eletroporação. Em primeiro lugar, a amostra estoque de proteína organicamente marcado ainda pode conter uma pequena percentagem de corante livre. As moléculas livres de corantes são muito menores do que as proteínas e pode, portanto, ser internalizada preferencialmente. Para assegurar que a vasta maioria das moléculas fluorescentes internalizadas observados correspondem à proteína de interesse, a amostra de proteína inicial deve conter menos de ~ 2% de corante livre (Figura 5) 22. O excesso de proteínas marcadas não-internalizadas também pode furar a membrana exterior da célula após a electroporação; este fenómeno é específico para a proteína e deve ser verificada para cada nova proteína. Propomos várias opções que permitem a remoção de proteínas não-internalizado a partir da amostra celular carregado (Ver PROTOCOLO: 3.3.3). Finalmente, as células são ressuspensas num pequeno volume de tampão de fosfato e pipetou-se para uma almofada de agarose, permitindo a sua imagem através de um microscópio de fluorescência. A imobilização sobre almofadas de agarose é um simple e forma eficiente de imagiologia células em uma lamela sem prejudicar sua integridade. A almofada deve conter um meio de cultura de baixa fluorescência. Imagens de células pode ser realizada tanto em microscopia widefield, o total de fluorescência reflexão interna (TIRF) ou utilizando HILO (altamente inclinadas e laminado folha óptica). Na configuração Hilo, o feixe de laser penetra mais profundamente do que a amostra em TIRF, ainda não ilumina toda a amostra como para campo amplo, permitindo uma maior razão sinal-para-ruído de 23. Dependendo da resolução de energia laser e tempo de utilização, biomoléculas internalizados podem ser contadas (utilizando a análise de fotobranqueamento-gradual, Figura 3), localizada, ou rastreados 24-28. Internalização de construções duplamente marcados com um par de fluoróforos FRET permite a quantificação de FRET tanto a uma única célula ou níveis de molécula única (Figura 6). Diferentes parâmetros podem ser variadosdependendo do resultado desejado e o sistema biológico estudado. Em primeiro lugar, a quantidade de material internalizados por células pode ser ajustado através da alteração da concentração de biomoléculas marcadas adicionado à electroporação as células antes (Figura 2). A intensidade do campo electroporação também irá influenciar tanto a eficiência do carregamento e a viabilidade celular; como esperado, enquanto que a eficiência aumenta com o aumento da carga de intensidade de campo, a viabilidade das células electroporadas diminui (Figura 4A). Ambos os parâmetros podem ser quantificados por registo da percentagem de células em divisão e carregado após a electroporação. Este ensaio de viabilidade juntamente com imagens de fluorescência também verifica a observação de biomoléculas internalizados em células vivas e permite a observação contínua ao longo de várias gerações (Figura 4B). Em resumo, este protocolo permite que a internalização de DNA e marcado por fluorescência em moléculas proteicasE. coli ou S. cerevisiae 26. Moléculas individuais marcadas com fluoróforos orgânicos podem ser rastreados com alta resolução espaço-temporal para escalas de tempo uma ordem de magnitude maior do que PQ. Finalmente, este método é compatível com widefield, TIRF e detecção confocal, bem como regimes de excitação por impulsos, tais como Alex (alternando excitação laser 28,29).

<table border="0" cellpadding="0" cellspacing="0" width="834">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Name of Material/ Equipment</td>
			<td style="width:227px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:321px;">ElectroMax DH5-alpha Comptent cells</td>
			<td style="width:227px;">Invitrogen</td>
			<td style="width:119px;">11319-019</td>
			<td>or any other commercial or lab-mage electrocompetant bacteria or yeast.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">EZ Rich Defined Madia</td>
			<td style="width:227px;">Teknova</td>
			<td style="width:119px;">M2105</td>
			<td>low fluorescence rich media</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MicroPulser Electroporation Apparatus&#160;</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">165-2100</td>
			<td>or any classical electroporator for microorganism transformation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Certified Molecular Biology agarose</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">161-3100</td>
			<td>low fluorescence agarose for agarose pad</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:321px;">Microscope coverslips No 1.5 thickness</td>
			<td style="width:227px;">Menzel</td>
			<td style="width:119px;">BB024060SC</td>
			<td>remove background particles by heating slides in furnace at 500 &#176;C for 1h</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Single-molecule fluorescence microscope</td>
			<td style="width:227px;">Home-built</td>
			<td style="width:119px;"></td>
			<td>described in REFs</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Localization software</td>
			<td style="width:227px;">Custom-written, available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB and C++ software package that can be adapted for localization analysis.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Tracking software</td>
			<td style="width:227px;">Available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB implementation by Blair and Dufresne.&#160;</td>
		</tr>
	</tbody>
</table>

internalization and observation of fluorescent biomolecules in living microorganisms via electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Watch this Scientific Journal Video about Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation at JoVE.com

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

Internalização e Observação de Fluorescent Biomoléculas em microorganismos vivos via Eletroporação

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

internalization-observation-fluorescent-biomolecules-living

Research

JoVE Journal

Biology

16.7K visualizações.  University of Oxford. Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

Vídeo: Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

Eletroporação de Micobactérias

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

Biologia

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

Imagens de células vivas de F-actina Dynamics via Microscopia fluorescente Speckle (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

Cultura primária e Eletroporação Plasmid do Órgão de Corti murino.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

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 ...

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron (dgn)-destabilized Green Fluorescent Protein (GFP)-based Reporter Protein

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. Maintenance of proteasome activity was implicated in many key cellular processes, like cell's stress response 3, cell cycle regulation and cellular differentiation 4 or in immune system response 5. The dysfunction of the ubiquitin-proteasome system has been related to the development of tumors and neurodegenerative diseases 4, 6. Additionally, a decrease in proteasome activity was found as a feature of cellular senescence and organismal aging 7, 8, 9, 10. Here, we present a method to measure ubiquitin-proteasome activity in living cells using a GFP-dgn fusion protein. To be able to monitor ubiquitin-proteasome activity in living primary cells, complementary DNA constructs coding for a green fluorescent protein (GFP)&#x2013;dgn fusion protein (GFP&#x2013;dgn, unstable) and a variant carrying a frameshift mutation (GFP&#x2013;dgnFS, stable 11) are inserted in lentiviral expression vectors. We prefer this technique over traditional transfection techniques because it guarantees a very high transfection efficiency independent of the cell type or the age of the donor. The difference between fluorescence displayed by the GFP&#x2013;dgnFS (stable) protein and the destabilized protein (GFP-dgn) in the absence or presence of proteasome inhibitor can be used to estimate ubiquitin-proteasome activity in each particular cell strain. These differences can be monitored by epifluorescence microscopy or can be measured by flow cytometry.

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron dgn-destabilized Green Fluorescent Protein GFP-based Reporter Protein

A method to monitor ubiquitin-proteasome activity in living cells is described. A degron-destabilized GFP- (GFP-dgn) and a stable GFP-dgnFS fusion protein are generated and transduced into the cell using a lentiviral expression vector. This technique allows to generate a stable GFP-dgn/GFP-dgnFS expressing cell line in which ubiquitin-proteasome activity can be easily assessed using epifluorescence or flow cytometry.

Monitoramento da ubiquitina-proteassoma Atividade em células vivas Usando um Degron (dgn) desestabilizado Green Fluorescent Protein Protein Repórter (GFP) com base em

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. ...

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific properties, such as binding affinity, agonist or antagonist behavior, or stability, relative to the native sequence. Protein design lies at the center of current advances drug design and discovery. Not only does protein design provide predictions for potentially useful drug targets, but it also enhances our understanding of the protein folding process and protein-protein interactions. Experimental methods such as directed evolution have shown success in protein design. However, such methods are restricted by the limited sequence space that can be searched tractably. In contrast, computational design strategies allow for the screening of a much larger set of sequences covering a wide variety of properties and functionality. We have developed a range of computational de novo protein design methods capable of tackling several important areas of protein design. These include the design of monomeric proteins for increased stability and complexes for increased binding affinity.
To disseminate these methods for broader use we present Protein WISDOM (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>), a tool that provides automated methods for a variety of protein design problems. Structural templates are submitted to initialize the design process. The first stage of design is an optimization sequence selection stage that aims at improving stability through minimization of potential energy in the sequence space. Selected sequences are then run through a fold specificity stage and a binding affinity stage. A rank-ordered list of the sequences for each step of the process, along with relevant designed structures, provides the user with a comprehensive quantitative assessment of the design. Here we provide the details of each design method, as well as several notable experimental successes attained through the use of the methods.

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

We developed computational de novo protein design methods capable of tackling several important areas of protein design. To disseminate these methods we present Protein WISDOM, an online tool for protein design (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>). Starting from a structural template, design of monomeric proteins for increased stability and complexes for increased binding affinity can be performed.

SABEDORIA Protein: A Workbench para<em&gt; In silico</em&gt;<em&gt; De novo</em&gt; Projeto de biomoléculas

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

In Vivo microinjeção e Eletroporação de testículos de camundongo

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

Tracking Drug-induced Changes in Receptor Post-internalization Trafficking by Colocalizational Analysis

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and overall cell responsiveness to ligands and is, itself, influenced by intra- and extracellular conditions, including ligand-induced signaling. Optimized for use with monolayer-plated cultured cells, but extendable to free-floating tissue slices, this protocol uses immunolabelling and colocalizational analysis to track changes in intracellular receptor trafficking following both chronic/prolonged and acute interventions, including exogenous drug treatment. After drug treatment, cells are double-immunolabelled for the receptor and for markers for the intracellular compartments of interest. Sequential confocal microscopy is then used to capture two-channel photomicrographs of individual cells, which are subjected to computerized colocalizational analysis to yield quantitative colocalization scores. These scores are normalized to permit pooling of independent replicates prior to statistical analysis. Representative photomicrographs may also be processed to generate illustrative figures. Here, we describe a powerful and flexible technique for quantitatively assessing induced receptor trafficking.

Receptor trafficking modulates signaling and cell responsiveness to ligands and is, itself, responsive to cell conditions, including ligand-induced signaling. Here, we describe a powerful and flexible technique for quantitatively assessing drug-induced receptor trafficking using immunolabeling and colocalizational analysis.

Rastreando alterações induzidas pela droga em Receptor de Pós-internalização Tráfico de Análise Colocalizational

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

De alta resolução de imagem Time-lapse e análise automatizada de microtúbulos Dynamics em Viver umbilical humano Células Endoteliais da Veia

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. To improve the mating efficiency of B. Xylophilus and to increase the chances of mating observation, virgin adults were cultured and used for the investigation. Eggs were obtained by keeping the nematodes in water and allowing the females to lay eggs for 10 min. The second-stage juveniles (J2) were synchronized by incubating the eggs for 24 h at 25 &#176;C in the dark, and the early J4 were obtained by culturing the J2 with grey mold, Botrytis cinerea, for another 52 h. At this time point, most J4 nematodes could be clearly distinguished as being male or female using their genital morphology. The male and female J4 were collected and cultured separately in two different Petri dishes for 24 h to get virgin adult nematodes. A virgin male and a virgin female were paired in a drop of water in the well of a concave slide. The mating behavior was filmed with a video recorder under a stereomicroscope. The whole period of the mating process was 82.8 &#177;3.91 min (mean &#177;SE) and could be divided into 4 different phases: searching, contacting, copulating, and lingering. The mean minutes of duration were 21.8 &#177; 2.0, 28.0 &#177; 1.9, 25.8 &#177; 0.7 and 7.2 &#177; 0.5, respectively. Eleven sub-behaviors were described: cruising, approaching, encountering, touching, hooping, locating, attaching, ejaculating, separating, quiescence, and roaming. Interestingly, obvious intra-sexual competition was observed when one female was grouped with 3 males or one male with 3 females. This protocol is useful and valuable, not only in investigating the mating behavior of B. xylophilus, but also in acting as a reference for ethological studies of other nematodes.

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A protocol for investigating the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, is presented. Behavioral features of B. xylophilus are described in the mating process.

Observação e quantificação do comportamento de acasalamento no nemátodo do pinheiro,<em&gt; Bursaphelenchus xylophilus</em

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. ...