Reconhecemos as doações tipo de cepas de bactérias do grupo do Prof Judith Armitage (Universidade de Oxford, UK) e Conrad Prof Mullineaux (Queen Mary University of London, UK). IMD é financiado conjuntamente pelo Departamento de Bioquímica (Universidade de Oxford) e OCISB; AR é financiado por uma Engenharia e Ciências Físicas Research Council (EPSRC) studentship DTC; ND é financiado pelo Biotecnologia e Ciências Biológicas Research Council (BBSRC); MCL é financiada por uma Universidade Royal Society Fellowship Research.

<ol><li> Para iniciar este procedimento, 50 ul de estoques congelados de proteína fluorescente expressar Escherichia coli células bacterianas são os primeiros descongelados e cresceu aerobicamente com agitação em 5 ml de mídia LB crescimento durante a noite a 37 graus C. Pela manhã, 50 ul desta cultura saturada é extraído e sub-cultivadas em um mínimo M63 media glucose cultura, incubando a 30 graus C para 4 a 6 horas. Aqui demonstramos usando duas linhagens celulares diferentes, um dos quais exprime um transporte de elétrons do citocromo fundido a GFP, o outro que expressa uma proteína envolvida no motor flagelar bacteriano fundido a GFP. </li><li> Células podem ou colhidas diretamente do crescente sub-cultura se vê-las como amostras de imobilizado, ou podem ser cortados para truncar flagelos bacterianos se ver em &quot;amarrados&quot; condições. </li><li> Shearing tipicamente envolve a colocação de 1-5ml da sub-cultura em um dispositivo composto por duas seringas estéreis unidas por tubos estéreis. O corte é feito pela alternância empurrando em cada bomba de seringa para empurrar a cultura através do tubo estreito. Isto é feito 50-100 vezes, dependendo da extensão do corte necessário. A cultura é então centrifugado para agregar as células, que é ressuspenso em meio mínimo para remover fragmentos de flagelos. </li><li> Nós, então, preparar limpos BK7 lamínulas de vidro por imersão em uma solução saturada de KOH em etanol por 20 min, em seguida, enxaguar bem em água desionizada e etanol e deixar secar ao ar por pelo menos 1 h. </li><li> Nós construímos uma simples célula de fluxo para a casa das células ao microscópio. Isto envolve o desenho de linhas de graxa de parafina em uma lâmina de microscópio BK7 vidro e em seguida, criar um túnel sanduíche colocando uma das coverlips limpo em cima e pressionando delicadamente com uma pinça, dando um volume de célula de fluxo de 50-10 mL . </li><li> Para observar células imobilizadas enchemos o fluxo de células por injeção com uma solução de 0,1% w / v de poli-L-lisina, e permitir que ele deixe em temperatura ambiente por pelo menos 1 min. Em seguida, lave com o uso mínimo de 100 l de mídia, injetando os meios de comunicação de uma extremidade da célula de fluxo e absorção com papel de tecido da outra. </li><li> Em seguida, através de pavio 20 l de uma diluição de 1:500 de 200 nm de diâmetro microesferas de látex (Polysciences) em meio mínimo para marcar a superfície lamela. A célula de fluxo é invertida de modo que a lamela está voltada para baixo, colocado sobre uma plataforma clara da superfície em uma câmara de umidade simples, e incubadas em temperatura ambiente por 5 min. Contas não ligados são então lavados por absorção através de 100 l media mínima. </li><li> Se quisermos observar as células amarrados omitimos o passo poli-L-lisina e, em vez encher a célula de fluxo com 5 mg / ml de anticorpos anti-flagelina. A célula de fluxo é lugar na câmara úmida por 10 min e depois é lavado por meio de absorção. </li><li> 20 l da cultura de células é então mau através da célula de fluxo, utilizando a amostra cisalhada se observar células de presos ou a amostra unsheared em ver células imobilizadas. </li><li> A célula de fluxo é invertido e colocado na câmara úmida por 20 min. Células não ligados são então lavados por absorção através de 100 l media mínima. </li><li> Uma gota de óleo de imersão é colocada no centro da superfície superior da lamela eo fluxo de células é então colocada gentilmente sobre o porta-amostras do microscópio construído sob encomenda de fluorescência, fazendo contato óptico com a objectiva de alta abertura numérica. </li><li> O microscópio de elétron-multiplicando câmera é ligada e definida para a câmera ser resfriado a -70 graus C, o software está definido para adquirir imagens a uma taxa de quadros típicos de 25 Hz no quadro de transferência de modo, inicialmente disenabling o controle de ganho na câmera. </li><li> A iluminação de campo claro é ligado ea imagem é colocada em foco, selecionando uma célula adequada ou grupo de células a ser trabalhada com base em seu ser preso com o seu longo eixo paralelo à superfície da lamínula. O foco é ajustado com precisão para garantir que as 200 contas nm latex preso à superfície lamela são apenas em foco. </li><li> Uma seqüência de imagens é adquirido em campo claro para gravar o contorno do corpo celular. A iluminação de campo claro está desligado eo ganho câmera é ativada ao máximo. </li><li> Para uma aquisição padrão usando fluorescência de reflexão interna total (ou TIRF) um feixe de laser sobre comprimento de onda apropriado (aqui, para a excitação da proteína verde fluorescente, usamos um laser de 473 nm) é pré-fixado de modo a ser focado no plano focal posterior da a lente objetiva, mas deslocadas lateralmente do eixo óptico para gerar um campo evanescente de excitação de fluorescência na amostra de células. </li><li> A aquisição da câmera é iniciado eo obturador aberto a laser para excitar as proteínas fluorescentes no interior da bactéria. Os parâmetros para a intensidade do laser ea velocidade de aquisição precisa ser otimizado para o sistema particular sob investigação biológica através da experimentação de diferentes valores, mas uma gama típica relevant a estudar complexos de proteína celular na membrana celular são a potência do laser 10-10 mW sobre uma área circular de excitação de diâmetro ~ 30 mm, com um tempo de exposição por quadro de 50-40 ms. Amostras são iluminadas até Foto-descoloração, tipicamente para ~ 10 s. </li><li> Este protocolo pode ser utilizado tanto para as células amarrados em que o corpo da célula está girando sobre um ponto de ligação entre as lamelas e um esboço flagelar, e por células imobilizadas que são rigidamente fixados à superfície lamela. </li><li> Algumas cepas de células que têm um número elevado de cópias de complexos beneficiar de uma lixívia de laser inicial de difração limitada focada em um pólo da célula antes TIRF imagem. Esta lixívia é equivalente ao utilizado na recuperação de fluorescência após a fotodegradação (ou FRAP). Usando nosso microscópio personalizado um pouco da luz do laser de excitação pode ser alimentado fora em um segundo caminho independente usado para FRAP tipo de clareamento. Normalmente 1-10mW de potência do laser é usado, com um tempo de lixívia típico na faixa de 10-300 ms. Isso resulta em contraste de imagem muito maior na zona branqueada da célula permitindo complexos individuais, que posteriormente difusa em que a área a ser mais facilmente visualizadas. </li><li> Para visualizar complexos no citoplasma da célula, que difundem muito mais rápido do que os da membrana da célula um modo de iluminação diferente chamado &quot;slimfield&quot; é empregado. Aqui, um laser focalizado é expandido lateralmente para apenas abranger uma única célula. Isso produz um campo muito intenso permitindo exposições muito mais rápido do normalmente um milésimo de segundo a tomar. </li><li> Após a aquisição de dados, as imagens são alimentados em software escrito personalizado (codificados em LabVIEW 8.5). Este detecta automaticamente as posições dos pontos fluorescentes nas células para uma precisão de um nanômetros geralmente poucos e extratos de seu tamanho e brilho. O brightnessof o traço fotobranqueamento com respeito ao tempo de um complexo molecular rastreado é então usada para estimar a estequiometria, ou seja, quantos individuais proteínas fluorescentes compõem um único complexo molecular. </li></ol> Resultados representativos: Quando o protocolo é feito corretamente as imagens das células vista em campo claro é muito distintos, com os perímetros dos corpos cela escura contra um corpo celular branco / cinza (Figura 1a). Na fluorescência usando células imobilizadas, podemos ver a intensidade pontos distintos, tipicamente de 250-300 nm de largura (Figura 1b). Saudável, as células amarrados será visto para girar em torno do ponto de fixação tether em imagens de campo claro. Sob excitação de fluorescência alguns complexos moleculares no nosso caso também pode ser visto no ponto de ligação, indicando uma localização da proteína marcada com o motor flagelar. Essas manchas são individuais complexos moleculares e do número deles visto vai depender do modo de iluminação usados ​​e como muitos dos complexos estão realmente presentes na célula de cada vez. A mobilidade dos pontos depende do sistema biológico específico em estudo. Se a densidade de pontos é inicialmente muito alta, como é o caso com os citocromos rotulados usado aqui, então executar uma lixívia FRAP inicial pode melhorar o contraste de imagem. <img src="/files/ftp_upload/1508/1508Figure1tmb.jpg" border="0" alt="Figura 1" /> Figura 1. (A) Brightfield e (B) imagem TIRF (cor falsa) de uma célula imobilizada Escherichia coli expressando uma proteína fundida a proteína verde fluorescente (GFP), que é conhecida por estar envolvida nos motores flagelares de bactérias. Por favor, <a href="https://www.jove.com/files/ftp_upload/1508/1508Figure1.jpg">clique aqui</a> para ver uma versão ampliada da figura 1.

<ol>
	<li>Leake, M. C., Chandler, J. H., Wadhams, G. H., Bai, F., Berry, R. M., Armitage, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stoichiometry+and+turnover+in+single,+functioning+membrane+protein+complexes.">Stoichiometry and turnover in single, functioning membrane protein complexes.</a> Nature. 443, 355-358 (2006).</li><li>Leake, M. C., Greene, N. P., Godun, R. M., Granjon, T., Buchanan, G., Chen, S., Berry, R. M., Palmer, T., Berks, B. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variable+stoichiometry+of+the+TatA+component+of+the+twin-arginine+protein+transport+system+observed+by+in+vivo+single-molecule+imaging.">Variable stoichiometry of the TatA component of the twin-arginine protein transport system observed by in vivo single-molecule imaging.</a> Proc Natl Acad Sci U S A. 105, 15376-15381 (2008).</li><li>Leake, M. C., Wilson, D., Gautel, M., Simmons, R. M., M, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+elasticity+of+single+titin+molecules+using+a+two-bead+optical+tweezers+assay.">The elasticity of single titin molecules using a two-bead optical tweezers assay.</a> Biophys. J. 87, 1112-1135 (2004).</li><li>Leake, M. C., Wilson, D., Bullard, B., Simmons, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+elasticity+of+single+kettin+molecules+using+a+two-bead+laser-tweezers+assay.">The elasticity of single kettin molecules using a two-bead laser-tweezers assay.</a> FEBS Lett. , 535-555 (2003).</li><li>Lenn, T., Leake, M. C., Mullineaux, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+clustering+and+dynamics+of+cytochrome+bd+complexes+in+the+Escherichia+coli+plasma+membrane.">In vivo clustering and dynamics of cytochrome bd complexes in the Escherichia coli plasma membrane.</a> Mol. Microbiol. 70, 1397-1407 (2008).</li><li>Lenn, T., Leake, M. C., Mullineaux, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Are+Escherichia+coli+OXPHOS+complexes+concentrated+in+specialised+zones+within+the+plasma+membrane.">Are Escherichia coli OXPHOS complexes concentrated in specialised zones within the plasma membrane.</a> Biochem. Soc. Trans. 36, 1032-1036 (2008).</li><li>Lo, C. J., Leake, M. C., Pilizota, T., Berry, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+measurements+of+Membrane+Potential,+Sodium-Motive+Force+and+Flagellar+Motor+Speed+in+Escherichia+coli.">Single-cell measurements of Membrane Potential, Sodium-Motive Force and Flagellar Motor Speed in Escherichia coli.</a> Biophys. 93, 294-302 (2007).</li><li>Lo, C. J., Leake, M. C., Berry, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+measurement+of+intracellular+sodium+concentration+in+single+Escherichia+coli+cells.">Fluorescence measurement of intracellular sodium concentration in single Escherichia coli cells.</a> Biophys. J. 90, 357-3565 (2006).</li></ol>

The authors have nothing to disclose.

É preciso ter cuidado para não &quot;sobre cisalhamento&quot; célula para olhar as bactérias amarrados, pois isso pode prejudicar a funcionalidade dos motores flagelares. É importante o uso de células por muito mais tempo do que uma hora uma vez na lâmina de microscópio, uma vez que pode tornar-se falta de oxigênio. Otimização considerável pode ser necessária para encontrar as melhores condições de imagem do microscópio servidos ao seu sistema biológico específico sob investigação. Pode ser sábio pa...

visualizing single molecular complexes in vivo using advanced fluorescence microscopy

Pleno conhecimento sobre os mecanismos de células vivas só pode ser alcançada por investigar os processos-chave que provocam direta e eventos em um nível celular. Até o momento a complexidade de cisalhamento de sistemas biológicos tem causado experimentação molécula única precisa ser demasiado exigentes, em vez enfocando estudos de sistemas simples utilizando granel relativamente rudimentar ensemble da média das medições. No entanto, muitos processos importantes ocorrem na célula viva no nível de apenas uma ou algumas poucas moléculas; medições ensemble geralmente mascaram a natureza estocástica e heterogénea desses eventos. Aqui, utilizando microscopia óptica avançada e ferramentas de análise de análise de imagens que demonstram como monitorar proteínas dentro de uma única célula viva bacteriana com uma precisão de moléculas individuais e como podemos observar a dinâmica dentro de complexos moleculares em funcionamento máquinas biológicas. As técnicas são diretamente relevantes fisiologicamente. Eles são minimamente perturbativos e não-invasivo para a amostra biológica em estudo e estão totalmente sintonizadas para as investigações na vida material, recursos não disponíveis para outros uma única molécula de abordagens de biofísica. Além disso, as amostras biológicas estudadas todas as produzem a proteína fluorescente marcadas em níveis que são quase idênticas às cepas de células não modificadas (&quot;codificação genômica&quot;), ao contrário da abordagem mais comum, mas menos ideal para a geração de proteínas significativamente mais do que ocorreria naturalmente (&#39;expressão plasmídeo). Assim, as amostras biológicas reais que serão investigadas são significativamente mais dos organismos naturais, e, portanto, as observações mais relevantes para os reais processos fisiológicos.

Watch this Scientific Journal Video about Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy at JoVE.com

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Aqui demonstramos os protocolos para a realização de uma única molécula de microscopia de fluorescência em células vivas de bactérias para habilitar funcionais complexos moleculares a serem detectados, rastreados e quantificados.

Visualizando Único complexos moleculares In Vivo Usando microscopia de fluorescência avançada

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

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Research

JoVE Journal

Biology

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

9.2K visualizações.  University of Oxford. Aqui demonstramos os protocolos para a realização de uma única molécula de microscopia de fluorescência em células vivas de bactérias para habilitar funcionais complexos moleculares a serem detectados, rastreados e quantificados.

Vídeo: Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

Replicação de DNA visualizar Único molécula com microscopia de fluorescência

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Biologia

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Lapso de tempo de imagens de Mitose Depois de Transfection siRNA

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

Visualizando bactérias em nematóides por microscopia de fluorescência

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Quantitative Analysis of Autophagy using Advanced 3D Fluorescence Microscopy

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, traditional chemotherapeutic approaches have been largely unsuccessful. Hormone therapy is effective at early stage, but often fails with the eventual development of hormone-refractory tumors. We have been interested in developing therapeutics targeting specific metabolic deficiency of tumor cells. We recently showed that prostate tumor cells specifically lack an enzyme (argininosuccinate synthase, or ASS) involved in the synthesis of the amino acid arginine1. This condition causes the tumor cells to become dependent on exogenous arginine, and they undergo metabolic stress when free arginine is depleted by arginine deiminase (ADI)1,10. Indeed, we have shown that human prostate cancer cells CWR22Rv1 are effectively killed by ADI with caspase-independent apoptosis and aggressive autophagy (or macroautophagy)1,2,3. Autophagy is an evolutionarily-conserved process that allows cells to metabolize unwanted proteins by lysosomal breakdown during nutritional starvation4,5. Although the essential components of this pathway are well-characterized6,7,8,9, many aspects of the molecular mechanism are still unclear - in particular, what is the role of autophagy in the death-response of prostate cancer cells after ADI treatment? In order to address this question, we required an experimental method to measure the level and extent of autophagic response in cells - and since there are no known molecular markers that can accurately track this process, we chose to develop an imaging-based approach, using quantitative 3D fluorescence microscopy11,12.
Using CWR22Rv1 cells specifically-labeled with fluorescent probes for autophagosomes and lysosomes, we show that 3D image stacks acquired with either widefield deconvolution microscopy (and later, with super-resolution, structured-illumination microscopy) can clearly capture the early stages of autophagy induction. With commercially available digital image analysis applications, we can readily obtain statistical information about autophagosome and lysosome number, size, distribution, and degree of colocalization from any imaged cell. This information allows us to precisely track the progress of autophagy in living cells and enables our continued investigation into the role of autophagy in cancer chemotherapy.

Autophagy is a ubiquitous process that enables cells to degrade and recycle proteins and organelles. We apply advanced fluorescence microscopy to visualize and quantify the small, but essential, physical changes associated with the induction of autophagy, including the formation and distribution of autophagosomes and lysosomes, and their fusion into autolysosomes.

Análise Quantitativa de Autophagy usando microscopia de fluorescência avançada 3D

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, ...

Visualizing Cytoplasmic Flow During Single-cell Wound Healing in Stentor coeruleus

Although wound-healing is often addressed at the level of whole tissues, in many cases individual cells are able to heal wounds within themselves, repairing broken cell membrane before the cellular contents leak out. The giant unicellular organism Stentor coeruleus, in which cells can be more than one millimeter in size, have been a classical model organism for studying wound healing in single cells. Stentor cells can be cut in half without loss of viability, and can even be cut and grafted together. But this high tolerance to cutting raises the question of why the cytoplasm does not simply flow out from the size of the cut. Here we present a method for cutting Stentor cells while simultaneously imaging the movement of cytoplasm in the vicinity of the cut at high spatial and temporal resolution. The key to our method is to use a &#34;double decker&#34; microscope configuration in which the surgery is performed under a dissecting microscope focused on a chamber that is simultaneously viewed from below at high resolution using an inverted microscope with a high NA lens. This setup allows a high level of control over the surgical procedure while still permitting high resolution tracking of cytoplasm.

Visualizing Cytoplasmic Flow During Single-cell Wound Healing in Stentor coeruleus

The giant ciliate Stentor coeruleus is a classical system for studying regeneration and wound healing in single cells.&#160;By imaging Stentor cells simultaneously at low and high magnification it is possible to measure cytoplasmic flows before, during, and after wounding.

Although wound-healing is often addressed at the level of whole tissues, in many cases individual cells are able to heal wounds within themselves, ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Análise rápida e Exploração de microscopia de fluorescência Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Visualizando Frequency Stromule com microscopia de fluorescência

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

Functional Imaging of Viral Transcription Factories Using 3D Fluorescence Microscopy

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to understand where and when transcription is taking place within nuclear space and to visualize the relationship between episomes infected within the same cell's nucleus. Here, both immunofluorescence (IFA) and RNA-FISH have been combinedto identify actively transcribing Kaposi's sarcoma-associated herpesvirus (KSHV) episomes. By staining KSHV latency-associated nuclear antigen (LANA), it is possible to locate where viral episomes exist within the nucleus. In addition, by designing RNA-FISH probes to target the intron region of a viral gene, which is expressed only during productive infection, nascent RNA transcripts can be located. Using this combination of molecular probes, it is possible to visualize the assembly of large viral transcription factories and analyze the spatial regulation of viral gene expression during KSHV reactivation. By including anti-RNA polymerase II antibody staining, one can also visualize the association between RNA polymerase II (RNAPII) aggregation and KSHV transcription during reactivation.

Viral transcriptional factories are discrete structures that are enriched with cellular RNA polymerase II to increase viral gene transcription during reactivation. Here, a method to locate sites of actively transcribing viral chromatin in 3D nuclear space by a combination of immunofluorescence staining and in situ RNA hybridization is described.

Imagem latente funcional das fábricas de transcrição Viral usando microscopia de fluorescência 3D

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to ...

Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are strategically interposed in the path of the lymphatic vessels, allowing intimate contact of tissue antigens with all resident immune cells in the LN. Thus, understanding the cellular composition, distribution, location and interaction using ex vivo whole LN imaging will add to the knowledge on how the body coordinates local and systemic immune responses. This protocol shows an ex vivo imaging strategy following an in vivo administration of fluorescent-labeled antibodies that allows a very reproducible and easy-to-perform methodology by using conventional confocal microscopes and stock reagents. Through subcutaneous injection of antibodies, it is possible to label different cell populations in draining LNs without affecting tissue structures that can be potentially damaged by a conventional immunofluorescence microscopy technique.

This protocol describes a technique to image different cell populations in draining lymph nodes without alterations in the organ structure.

Visualizando estrutura de linfonodos e localização celular usando microscopia confocal ex-vivo

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are ...

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Examinação da dinâmica nuclear do fermento mitotic e Meiotic da fissão pela microscopia viva-pilha da fluorescência

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...