我们感谢实验室的Gelfand谁促成了这些协议的开发成员。研究报告本出版物中由美国国立卫生研究院国家普通医学科学根据奖号R01GM052111国家学院VIG和教育巴斯克政府部门，大学和研究下奖号BFI-2011-295 UDC得到了支持。

1， 果蝇 S2细胞<ol><li>刀豆蛋白A包被盖玻片的制备<ol><li>放置盖玻片在陶瓷机架和用铬酸浸渍冲洗1小时。 [注意：铬酸具有腐蚀性，可引起眼睛发炎，鼻子，喉咙和皮肤。 </li><li>与不断运行DH 2 O 30分钟，直至酸被完全冲洗掉彻底冲洗盖玻片。 </li><li>让盖玻片空气干燥，被毛他们的ConA（0.5毫克/毫升的溶液在DH 2 O）30分钟。 </li><li>冲洗盖玻片与DH 2 O 15分钟，让他们风干。刀豆蛋白A包被盖玻片可以存储长达1个月。 </li></ol></li><li>电镀细胞<ol><li>允许S2细胞在T25或T75厘米2烧瓶成倍增长取决于实验所需的细胞数。 </li><li>使用血细胞计数器计数细胞密度。加入1 ml增长米edium（例如昆虫随心）到35毫米组织培养皿用ConA-涂覆的盖玻片，并轻轻〜1×10 5个细胞转移到培养皿中。此细胞密度是最佳的成像，因为从不同的细胞过程中不重叠。 </li><li>为了诱导形成的过程中，电镀加入1ml培养基用5μMCytoD到培养皿（以2.5μMCytoD的终浓度）后立即使用。由成像前25ºC培养细胞至少2小时让流程全面发展。 </li></ol></li></ol> 2， 果蝇初级神经元培养<ol><li>神经元培养制备<ol><li>制备果蝇神经细胞生长培养基（Schneider的培养基补充有20％胎牛血清，热失活，在55℃，30分钟，5微克/毫升的胰岛素，100微克/毫升青霉素，100微克/毫升链霉素，10微克/毫升四环素） 。这SUP施耐德执行完成的介质可以储存在4℃6个月。 </li><li>制备苹果汁琼脂小瓶中，收集果蝇胚胎（22.5克琼脂，12.0克蔗糖和750毫升水2 O中的2升烧瓶中，高压釜为20-25分钟;添加250毫升苹果汁和倒〜10毫升到每个果蝇小瓶;凝固后，插上小瓶用棉球和小瓶包装用塑料食品包装，避免了对粮食的干燥）。在琼脂瓶苹果汁可以储存在4℃在密封良好的塑料包装袋，共4周。 </li><li>外套酸洗25毫米圈盖玻片经ConA（见议定书果蝇 S2细胞中的细节）和消毒的紫外光下10-15分钟。这些盖玻片可存放一个月。 </li></ol></li><li> 果蝇胚胎神经元培养<ol><li>保持年轻的成年苍蝇在收集胚胎f前辅以干酵母1-2天苹果汁琼脂小瓶或神经元的准备。 </li><li>在收集日，以同步采集胚胎，转成蝇的新鲜苹果汁琼脂小瓶中光1小时precollection并丢弃这些胚胎。 </li><li>转让苍蝇到一个新的苹果汁琼脂小瓶中，收集的胚胎为1小时，并在果蝇转移到另一个小瓶中一小时，无论是在黑暗中以提高胚胎产量。 </li><li>从苹果汁琼脂小瓶中取出成蝇，让胚胎发育另4小时到9-11级在室温下（4-6小时大的胚胎阶段是9-11之间）。 </li><li>通过水喷到小瓶中，并从松动的苹果汁琼脂用小画笔的胚胎（需要至少10胚胎一个良好的神经元准备）将胚胎。 </li><li>胚胎转移到一个100微米的尼龙网细胞滤网用移液管排出的水（预先清洗吸在果蝇胚胎的洗涤缓冲液，0.4％氯化钠，0.03％的Triton移液管X-100）。 </li><li>从使用小画笔过滤收集胚胎，并把它们转移至1.5 ml微量离心管装入1ml 果蝇胚胎洗涤（注：使用画笔或移液管尖端清洁任何琼脂或棉纤维）。在这个协议中，1.5mL微管带有一个匹配的一次性颗粒杵。 </li><li>在果蝇胚胎洗3次洗胚胎。 </li><li> Dechorionate胚胎在一个新的1点01分解决方案的商业漂白剂和95％乙醇10分钟[注意：漂白剂，会刺激眼睛，鼻子，喉咙和皮肤。 </li><li>移动到组织培养罩，并在补充施耐德的媒体洗胚胎2倍。 </li><li>离开200-300微升培养基在管中的胚胎，并使用已灭菌的70％乙醇（碾10-15X）金布尔大通弃沉淀杵轻轻机械解离dechorionated胚胎。 </li><li>在20×g离心离心匀浆混合物，持续2分钟，和TRAnsfer上清至新管。 [这一步是可选的，它有助于消除胚胎碎片] </li><li>离心的上清液在550×g离心2分钟以沉淀细胞。 </li><li>重悬浮颗粒在500微升，在550×g离心2分钟再补充施耐德的媒体和自旋向下的细胞，重复的再悬浮，动感单车一次。 </li><li>最后悬浮在〜100微升补充施耐德的介质中的颗粒和板上的细胞到25毫米的无菌35毫米培养皿刀豆蛋白A包被的盖玻片。孵育10分钟以使细胞附着。 </li><li>加入2ml补充施耐德的介质，具有5微米CytoD淹没盖玻片与贴壁细胞。 </li><li>成像前孵育菜在湿润的25℃培养箱中过夜。 </li></ol></li></ol>

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作者宣称，他们有没有竞争的财务权益。

在这里，我们提出了用于研究微管相关货物运输在果蝇 S2细胞和原代神经元培养的详细协议。既S2流程和突起结构是由充满了作为细胞器运输轨道捆绑微管阵列。 两种策略通常用于标签的细胞器：载体与细胞器靶向序列或染色用直接添加到培养物的荧光染料编码荧光蛋白的转染。的货物标签，常见的例子列于表1中 。 果蝇 S2细胞中瞬时转染或RNAi治疗...

果蝇 S2细胞中都获得了学习许多细胞过程日益普及。这些细胞最初是从OregonR 果蝇的胰酶消化后期胚胎获得和维持巨噬细胞样特性1。 果蝇 S2细胞中提供了许多优于其他细胞系。它们被培养于25℃无CO 2，并且可以成像在室温下几个小时，无需加热或气体交换的需要。更重要的是， 果蝇 S2细胞对RNAi的治疗，允许感兴趣的蛋白质的高效率击倒非常敏感。 S2细胞的高效转染和稳定的细胞系可以在不到四周的时间进行选择，以便感兴趣的蛋白的稳定表达。 S2细胞正常生长，无论是松散连接，但不能涂在烧瓶或悬浮状态。它们可以诱导铺展在盖玻片预涂用植物凝集素孔卡navalin A（刀豆）。扩散细胞的周围是特别薄的，因此是理想的活细胞显微镜检查。为S2细胞培养，RNA干扰治疗，现场光成像和免疫组化的详细方案可以在文献2,3中找到。我们的实验室已采用S2细胞作为模型系统来研究微管为基础的货物运输。与任何药物解聚或切断的F-肌动蛋白，如细 ​​胞松弛素D（CytoD）处理过的果蝇 S2细胞，长长的过程充满了均匀极化微管4 -10，这使它成为理想的系统来研究货物运输沿微管阵列。交通工具在这些过程中不同的参数（ 即轨迹长度，速度，方向性等 ）可以使用自动化粒子跟踪软件，DiaTrack（Semasopht很容易地测量：http://www.microscopy.info/Organization/Details/4423;博士。帕斯卡尔·瓦洛：http://www.csiro.au/en/Organisation-Structure/Divisions/Mathematics-Informatics-and-Statistics/PascalVallotton.aspx＃A4）。这些特性使得S2细胞一个有吸引力的制度研究货物运输以及在组织培养细胞中的微管。 在S2细胞试验实验可以扩展到在果蝇原代神经元的货物运输研究的一个重要的生理模型。类似于S2细胞，神经元的解离胚胎培养是可渗透的小分子物质，例如染料和抑制剂，并且可以在神经元中，在室温下进行的细胞器运输的高分辨率实时成像。利用果蝇与不同的遗传背景，我们可以通过基因敲除（丧失功能的基因突变）研究基因功能的初级神经元，击倒（双链RNA注射或表达式）或异位表达（转基因果蝇）。具体来说，使用CytoD治疗肌动蛋白丝的碎片不会阻止轴突生长，相反，他们成长得更快，因此，我们可以研究微管depende在CytoD处理的神经元新台币细胞器运输无肌动蛋白的影响丝11。因此，初级神经元培养结合组织培养细胞的优点和飞遗传学，这使它成为一个伟大的系统来研究货物运输在生理相关的系统10,12。因为几个家族性神经变性疾病可能由以下原因引起或与货物运输的缺陷（ 例如，帕金森氏病13相关联，亨廷顿氏病14，阿尔茨海默氏病15,16，肌萎缩性侧索硬化症/ ALS 17，HMN7B和佩里综合征18,19，和脊髓小脑性共济失调型5/SCA5 20），原代神经元培养物可以是在特定突变对微管依赖性的传输造成这些疾病的影响的分析是有用的。 最后，微管依赖性运输的重要性能哺乳动物和果蝇之间非常保守，但<em&gt;果蝇基因和细胞培养物是不太昂贵和费时的，正当的使用培养的果蝇细胞在正常和病理条件下研究细胞器运输的基本方面。

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			<td height="61" style="height:61px;width:124px;">Powder Insect cell medium (Schneider&#8217;s)</td>
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			<td style="width:179px;">S9895</td>
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			<td height="61" style="height:61px;width:124px;">Penicillin</td>
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			<td height="21" style="height:21px;width:124px;">Cytochalasin D</td>
			<td style="width:133px;">VWR</td>
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			<td style="width:179px;">L7528</td>
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			<td height="41" style="height:41px;width:124px;">100 &#181;m Cell strainer</td>
			<td style="width:133px;">Fisher Scientific</td>
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			<td height="41" style="height:41px;width:124px;">25 mm Circle cover glass</td>
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			<td style="width:179px;">48380 080</td>
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			<td height="41" style="height:41px;width:124px;">Drosophila Vials</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">89092-730</td>
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			<td height="81" style="height:81px;width:124px;">Disposable pellet pestles with matching microtubes</td>
			<td style="width:133px;">Kimble Chase</td>
			<td style="width:179px;">749520-0000</td>
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organelle transport in cultured drosophila cells: s2 cell line and primary neurons.

果蝇 S2细胞中填充有一致极化微管任何肌动蛋白解聚药物形式长直链的过程的存在下接种在盖玻片上。沿着这些过程通过微管马达的细胞器移动。易维护，高灵敏度RNAi介导的蛋白敲除和有效的程序来创建稳定的细胞系使果蝇 S2细胞中一种理想的模型系统，通过实时成像研究货物运输。与S2细胞中获得的结果可被进一步应用到多种生理上相关的系统：在从分离的果蝇胚胎培养的原代神经元的轴突运输。培养神经元长出长突起充满微管捆绑在一起，非常类似于S2的过程。像S2细胞，在培养的神经元细胞器可以通过两种细胞器特异性的荧光染料，或通过使用由DNA编码的荧光标记的细胞器可视化注入到早期胚胎或表达transgenIC苍蝇。因此，细胞器运输可以很容易地记录在神经元上使用生活成像盖玻片培养。在这里，我们描述了在果蝇 S2细胞和原代神经元培养和可视化货物运输手续。我们相信，这些协议使两个系统的实验室研究货物运输方便。

果蝇 S2细胞附着在刀豆蛋白A包被的盖玻片蔓延成扁圆形“煎饼”形（ 图1A和1C）。然而，当镀上盖玻片S2细胞中CytoD的存在下，并使其分散为2-3小时，便形成细长的进程填充平行微管（ 图1B和1D）。这些微管具有统一的极性与加结束了7。的果蝇 S2稳定表达的mCherry-微管蛋白的时间推移荧光成像表明，过程是非常动态的，特别?...

Watch this Scientific Journal Video about Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons. at JoVE.com

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

果蝇 S2细胞和培养的神经元是用于体内电机驱动的细胞器运输的成像大的系统。在这里，我们将详细介绍协议进行培养两种细胞类型，它们的成像和运输分析。

细胞器运输的培养<em&gt;果蝇</em&gt;细胞：S2细胞系和原代神经元。

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

organelle-transport-cultured-drosophila-cells-s2-cell-line-primary

Research

JoVE Journal

Biology

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

13.6K 次观看.  Feinberg School of Medicine, Northwestern University.  果蝇 S2细胞和培养的神经元是用于体内电机驱动的细胞器运输的成像大的系统。在这里，我们将详细介绍协议进行培养两种细胞类型，它们的成像和运输分析。 

视频: Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

从晚期果蝇蛹的大脑原发性神经文化

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

科研

生物学

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

在原代培养海马神经元的密集核心囊泡的实时成像

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

制备果蝇 S2细胞，光学显微镜

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Adenovirus-mediated Genetic Removal of Signaling Molecules in Cultured Primary Mouse Embryonic Fibroblasts

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction analysis. One particular method to achieve this conditional deletion is via the use of the Cre-loxP system. This method involves flanking the gene of interest with loxP sites, which are specific recognition sequences for the Cre recombinase protein. Exposure of the so-called floxed (flanked by loxP site) DNA to this enzyme results in a Cre-mediated recombination event at the loxP sites, and subsequent excision of the intervening gene3. Several different methods exist to administer Cre recombinase to the site of interest. In this video, we demonstrate the use of an adenovirus containing the Cre recombinase gene to infect primary mouse embryonic fibroblasts (MEFs) obtained from embryos containing a floxed Rac1 allele1. Our rationale for selecting Rac1 MEFs for our experiments is that clear morphological changes can be seen upon deletion of Rac1, due to alterations in the actin cytoskeleton2,5. 72 hours following viral transduction and Cre expression, cells were stained using the actin dye phalloidin and imaged using confocal laser scanning microscopy. It was observed that MEFs which had been exposed to the adeno-Cre virus appeared contracted and elongated in morphology compared to uninfected cells, consistent with previous reports2,5. The adenovirus method of Cre recombinase delivery is advantageous as the adeno-Cre virus is easily available, and gene deletion via Cre in nearly 100% of the cells can be achieved with optimized adenoviral infection.

In this video we use an adenovirus carrying the Cre recombinase gene to infect primary mouse embryonic fibroblasts carrying a floxed Rac1 allele.

腺病毒介导的基因去除信号分子在原代培养的小鼠胚胎成纤维

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

原代细胞培养从果蝇原肠胚

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...

Isolation and Primary Culture of Rat Hepatic Cells

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology and other related subjects. The method based on two-step collagenase perfusion for isolation of intact hepatocytes was first introduced by Berry and Friend in 1969 1 and, since then, has undergone many modifications. The most commonly used technique was described by Seglenin 1976 2. Essentially, hepatocytes are dissociated from anesthetized adult rats by a non-recirculating collagenase perfusion through the portal vein. The isolated cells are then filtered through a 100 &mu;m pore size mesh nylon filter, and cultured onto plates. After 4-hour culture, the medium is replaced with serum-containing or serum-free medium, e.g. HepatoZYME-SFM, for additional time to culture. These procedures require surgical and sterile culture steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol, which allow consistently in the generation of viable hepatocytes in large numbers.

Primary hepatocytes provide a valuable tool to evaluate biochemical, molecular, and metabolic functions in a physiologically relevant experimental system. We describe a reliable protocol for rat in situ liver perfusion, which consistently generates viable hepatocytes up to 1.0 &times; 108 cells per preparation with cell viability between 88 ~ 96%.

大鼠肝细胞的分离和小学文化

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

成像本地CA<sup&gt; 2+</sup&gt;在培养的哺乳动物细胞信号

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

Generating a &#34;Humanized&#34; Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper distribution of the genetic material requires that sister kinetochores on every chromosome become bioriented by attaching to microtubules from opposite spindle poles before progressing into anaphase. However, erroneous, non-bioriented attachment states are common and cellular pathways exist to both detect and correct such attachments during cell division. The process by which improper kinetochore-microtubule interactions are destabilized is referred to as error correction. To study error correction in living cells, incorrect attachments are purposely generated via chemical inhibition of kinesin-5 motor, which leads to monopolar spindle assembly, and the transition from mal-orientation to biorientation is observed following drug washout. The large number of chromosomes in many model tissue culture cell types poses a challenge in observing individual error correction events. Drosophila S2 cells are better subjects for such studies as they possess as few as 4 pairs of chromosomes. However, small molecule kinesin-5 inhibitors are ineffective against Drosophila kinesin-5 (Klp61F). Here we describe how to build a Drosophila cell line that effectively replaces Klp61F with human kinesin-5, which renders the cells sensitive to pharmacological inhibition of the motor and suitable for use in the cell-based error correction assay.

Generating a &quot;Humanized&quot; Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

This protocol describes how to generate a Drosophila S2 cell line that is sensitive to small molecule inhibitors of kinesin-5. The use of these cells in a cell-based error correction assay is also outlined.

生成一个“人性化”<em&gt;果蝇</em&gt; S2细胞敏感的驱动蛋白-5的药理抑制

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper ...

Glucose Uptake Measurement and Response to Insulin Stimulation in In Vitro Cultured Human Primary Myotubes

Skeletal muscle is the largest glucose deposit in mammals and largely contributes to glucose homeostasis. Assessment of insulin sensitivity of muscle cells is of major relevance for all studies dedicated to exploring muscle glucose metabolism and characterizing metabolic alterations. In muscle cells, glucose transporter type 4 (GLUT4) proteins translocate to the plasma membrane in response to insulin, thus allowing massive entry of glucose into the cell. The ability of muscle cells to respond to insulin by increasing the rate of glucose uptake is one of the standard readouts to quantify muscle cell sensitivity to insulin. Human primary myotubes are a suitable in vitro model, as the cells maintain many features of the donor phenotype, including insulin sensitivity. This in vitro model is also suitable for the test of any compounds that could impact insulin responsiveness. Measurements of the glucose uptake rate in differentiated myotubes reflect insulin sensitivity.
In this method, human primary muscle cells are cultured in vitro to obtain differentiated myotubes, and glucose uptake rates with and without insulin stimulation are measured. We provide a detailed protocol to quantify passive and active glucose transport rates using radiolabeled [3H] 2-deoxy-D-Glucose ([3H]2dG). Calculation methods are provided to quantify active basal and insulin-stimulated rates, as well as stimulation fold.

Glucose Uptake Measurement and Response to Insulin Stimulation in In Vitro Cultured Human Primary Myotubes

In this method, human primary muscle cells are cultured in vitro to obtain differentiated myotubes and glucose uptake rates are measured. We provide a detailed protocol to quantify rates in basal and insulin-stimulated states using radiolabeled [3H] 2-deoxy-D-Glucose.

葡萄糖摄取测量和对胰岛素刺激的反应<em&gt;体外</em&gt;培养的人类初级肌管

Skeletal muscle is the largest glucose deposit in mammals and largely contributes to glucose homeostasis. Assessment of insulin sensitivity of muscle ...

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory cells, EC cells sense luminal stimuli and convert them into serotonin (5-hyroxytryptamine, 5-HT) release events. However, the electrophysiology of these cells is poorly understood because they are difficult to culture and to identify. The method presented in this paper outlines primary EC cell cultures optimized for single cell electrophysiology. This protocol utilizes a transgenic cyan fluorescent protein (CFP) reporter to identify mouse EC cells in mixed primary cultures, advancing the approach to obtaining high-quality recordings of whole cell electrophysiology in voltage- and current-clamp modes.

Enterochromaffin (EC) cells comprise a small subset of gastrointestinal epithelial cells. EC cells are electrically excitable and release serotonin, yet difficulties in culturing and identifying EC cells have limited physiological studies. The method presented here establishes a primary culture model amenable to examination of single EC cells by electrophysiology.

原代培养小鼠肠嗜铬细胞的全细胞电生理

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory ...