我们要感谢Claycomb博士提供HL-1细胞和详细的维护协议。我们还要感谢埃琳娜·卡里略女士和赛斯Robia博士为他们与产生细胞电影和皮特卡隆先生的协助做一些配件为双通道光学标测系统的援助。 这项工作是由美国心脏协会（10GRNT3770030＆12GRNT12050478为XA），国立卫生研究院（HL113640到XA），和Loyola大学的研究发展基金（以XA）的支持。

1.溶液制备<ol><li>去甲肾上腺素（NP）溶液<ol><li>溶解40毫克的NP的成将25ml 30mM的抗坏血酸（0.1475克抗坏血酸在25ml纯化的Milli-Q（MQ）水。该溶液在制备过程中避免直接光照，因为去甲肾上腺素（NP）是光敏感的。 </li><li>过滤用0.22μm注射器滤器对NP溶液。分装，在-20℃下将溶液在1ml无菌微量管和存储。一旦冻结，NP为稳定一个月。 </li></ol></li><li>辅以洗净，最后Claycomb媒体<ol><li>先从43.5毫升Claycomb培养基和补充用5毫升胎牛血清（FBS），0.5 ml的青霉素/链霉素，0.5毫升的NP和1：100稀释的L-谷氨酰胺，使50最后补充Claycomb培养基中。这种溶液的制备过程中应避免直射光照射，因为Claycomb介质对光敏感。 </li><li>使用相同的配方为SUP准备洗Claycomb媒体除了执行完成Claycomb中添加5％FBS和省略NP和L-谷氨酰胺的组件。 注：存放于4℃无直射光照射时，最后的补充和冲洗Claycomb媒体是很好的两周。 </li></ol></li><li>大豆胰蛋白酶抑制解决方案<ol><li>溶解的大豆胰蛋白酶抑制剂25毫克到100ml无钙和镁的Dulbecco磷酸盐缓冲盐水（PBS）中。 </li><li>过滤用0.22微米注射器过滤该溶液。 注：储存在4℃下时，灭菌溶液是稳定一个月。 </li></ol></li><li>明胶/盘纤连蛋白涂液<ol><li>将0.1g明胶到500ml MQ水中，使0.02％的明胶。 </li><li>反应釜，然后稀释1毫升纤维连接蛋白进入199毫升0.02％的明胶，轻轻混匀。 </li><li>等分6毫升的明胶溶液成单独的管中并储存在-20℃。 </li></ol></li><li>汉克斯&#39;盐溶液<ol><li>使用搅拌板，溶解在MQ水中（毫摩尔/升）以下的盐类，氯化钠（136.9），氯化钾（5.366），KH 2 PO 4（0.4409），将NaH 2 PO 4（0.4000），硫酸镁 （0.8142），D-葡萄糖（5.051）， 碳酸氢钠 （4.162），HEPES（5.042）， 氯化钙 （1.261）。 </li></ol></li><li>钙敏感染料（Rhod2）解决方案<ol><li>稀释的1mg Rhod2干粉在400微升20％的Pluronic F-127的DMSO作Rhod2染料原液并储存在-20℃。 </li><li>这种稀释原液1：1000用Hanks&#39;盐溶液在实验开始前，作最后的工作方案。 </li></ol></li><li>电压敏感染料（Rh237）解决方案<ol><li>稀释5毫克Rh237的用2毫升DMSO中，使储液并储存在-20℃。稀释原液1：1000用Hanks&#39;盐溶液在实验开始前，作最后的工作方案。 </li></ol></li></ol> ＆＃160;注：1-4的解决方案是基于Claycomb HL-1细胞培养协议轻微修改。 2.传代培养和HL-1心房肌细胞到盖玻片注：HL-1细胞可以从Claycomb博士（路易斯安娜州立大学）获得。 <ol><li>生长的HL-1细胞在T25烧瓶和饲料用5ml补充Claycomb中等每日。通道中的细胞在T25根据Claycomb HL-1细胞培养物的协议（以下描述）10烧瓶每五天。 </li><li>地方圆形盖玻片（20毫米直径）到12孔培养板中之前24小时，将细胞在T25烧瓶中完全汇合，并准备进行传代，然后盖上明胶/纤连蛋白过夜，在37℃的盖玻片。 </li><li>第二天，吸出并丢弃从培养板中的明胶/纤连蛋白和替换用1.5ml在每个补充最终Claycomb介质的井。 </li><li>从T25文化性拆分单元格Ë烧瓶中，在5天达到完全汇合后，弃从含有汇合培养的T25烧瓶中的培养基和冲洗培养简要地用PBS在37℃。 </li><li>抽吸PBS并且加入3毫升0.05％胰蛋白酶/ EDTA，在37℃吹打的胰蛋白酶/ EDTA到T25烧瓶的底部，避免与细胞直接接触。孵育在37℃下1分钟。 </li><li>通过抽吸除去胰蛋白酶/ EDTA，并添加每个T25烧瓶另一个1.3毫升胰蛋白酶/ EDTA中。在室温下孵育1分钟。检查在显微镜下，如果细胞仍然附着在烧瓶中，点击要充分打跑剩余的细胞。 </li><li>添加1.3毫升大豆胰蛋白酶抑制剂，从烧瓶内的细胞和细胞转移到15ml离心管中。冲洗用5毫升洗净Claycomb介质的空烧瓶中。漂洗添加到细胞中，在15ml离心管中。离心，在500×g离心5分钟。 </li><li>离心，吸出上清液，并轻轻后重新悬浮细胞沉淀6毫升补充决赛Claycomb介质。 </li><li>种子在12孔平板的每个孔用0.5ml从细胞传代细胞悬浮液。每天喂细胞新鲜辅以最终Claycomb媒介。 注：密切监测细胞的生长，通常细胞单层达到汇合状态在第4天，并自发活动保持可见。然后，将培养的单层将准备在4-5天的光学测绘的。 </li></ol> 3.装载钙和电压敏感染料的细胞<ol><li>用于光学作图研究，轻轻地从12孔细胞培养板除去盖玻片和冲洗用Hanks&#39;盐溶液。 </li><li>染色的细胞用钙敏感染料Rhod2，孵育在盖玻片的细胞单层用3ml的充氧，在37℃的水浴中30分钟Rhod2工作溶液（用5％的CO 2/95％O 2气鼓泡） 。 </li><li>染色的细胞在V 米敏感的染料Rh237，浸入Rhod2加载细胞单层分为3毫升含氧Rh237工作溶液，在37℃的水浴中另外15分钟。 </li></ol> 4.双通道光映射<ol><li>将单层细胞，这是双沾满V M和钙染料，插入一个倒置荧光显微镜循环室。使用加热的循环体系，以保持细胞室的温度在35±1℃。小心通过调节循环流的速度通过​​腔室，以避免超过0.3℃，在整个腔室的梯度控制所述灌流液的温度。 <ol><li>记录V M /钙信号之前，用新鲜的Hanks液洗细胞单层约5分钟，以从先前的染色步骤除去剩余的染料。 </li></ol></li><li>使用双极电极的玻璃吸移管填充用Hanks&#39;盐溶液和SILV组成步伐健康细胞单层呃丝枪头周围卷绕如前所述12,17。 </li><li>吴佩慈健康的细胞单层在200-500毫秒，脉冲2毫秒宽度的周期长，期待起搏阈值约为1毫伏。 注：对于每一批培养单层的，至少有两个非处理单层应当用于测试细胞批次的状态。健康对照单层应当能够被电起搏在200-500毫秒周期长度捕获。如果控制单层不能被电节奏，这表明，整个细胞批次可能不适合用于光学测绘实验。 </li><li>激励两个V M和钙敏感性染料，可使用的LED光源（520纳米波长）加上一个激励滤光器（510纳米/ 40）和一个二向色镜（561纳米）。 <ol><li>收集通过40X物镜和分割用分色镜（594纳米）成V 米组分和Ca成分由细胞发出的荧光信号。过滤在V 米通道的发射光具有长通滤波器（700纳米），发射的光中的钙通道有一个带通滤波器（四十分之五百八十五纳米）。 </li><li>收集用两台CCD照相机（80×80像素，1000 fps的;红衫成像）将过滤的荧光。 </li><li>为了避免染料光漂白，使录音为小于2秒上的每个视场限制光曝光的时间。这里，典型的图像的每个单层四个不同的区域。 </li></ol></li></ol>的V M或Ca录音5.数据处理<ol><li>为V M和Ca信号（80×80像素），平均的25（5×5）的强度值的相邻像素成在不同的记录时间点的一个强度值以减少背景噪声。 注：该数据平均后，个人动作电位和钙瞬变每次录制时间点产生含有16×16的平均亮度值的数据矩阵（平均数据信道）为50微米的空间分辨率。 </li><li>定义的记录，V M或Ca信号激活时间（AT）。 <ol><li>以限定的单个动作电位或钙瞬变的激活时间（AT），确定的在V m或者使用一个基于MATLAB定做算法钙信号的峰值幅度的50％的时间，如前所述。12 </li><li>手动校对所分析的AT来消除程序分析期间可能发生的任何错误。 </li><li>存储所述所定义的AT矩阵（16×16平均数据信道）分别记录动作电位和钙瞬变，在所使用的软件。 </li></ol></li><li>计算上使用矢量场分析方法如先前稍作修改12,18中描述的AT矩阵动作电位或钙瞬变传导速度（CV）的载体。 <ol><li>计算数据信道相对于其周围的AT的数据点的CV载体（5×; 5相邻平均数据信道），使用中的MATLAB程序12,18最小二乘算法，模拟的激活时间（AT表面光滑多项式表面）。 </li><li>使用以下公式计算传导矢量 式1： <img alt="式（1）" fo:content-width="1.5in" src="/files/ftp_upload/52542/52542eq1.jpg" /> 配方2： θ=反正切[（DY / DT）/（DX / DT）] 注：Ve为在x轴和y轴投射在简历矢量; DX / DT是在X轴上的CV矢量的投影; DY / DT是在y轴的CV向量的投影。 T是所述AT表面上;的x，y指的是2-D坐标为的T x =∂T/狓是T的关于x的偏导数; ŤY =∂T/狔是T的相对于y的偏导数; θ是一个简历矢量的最终方向。 </li><li>所述AT矩阵的组中的所有矢量成36通路，其每一个覆盖指南针上的10度扇区。&lt;/ li&gt; <li>选择包含​​36传导通路之间的CV向量的人口最多的途径。计算这个途径的平均CV代表单层的CV。 </li></ol></li></ol> 6.可视化的波前传播<ol><li>正常化的在相对于利用MATLAB软件其峰值不同的记录时间点的单个动作电位或Ca瞬态信号的强度。 </li><li>每记录的时间点的归一化强度数据暂时存储到在所使用的MATLAB软件一个16×16的强度矩阵。 </li><li>构造从使用surf.m函数的每个记录时间点的存储的强度矩阵的彩色编码强度地图。 注：在强度图，红色代表的V 米或钙信号强度高，蓝色体现的荧光信号的强度低。 </li><li>创建使用avifile.m功能的.avi文件。 </li><li>保存所有的I在动作电位或Ca瞬态不同的记录时间点使用addframe.m函数ntensity地图。 注意：在这一点上，将产生的V M和Ca传播两个独立的AVI文件。 </li></ol>

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No conflicts of interest declared.

本文介绍了培养HL-1心房肌细胞单层染色钙和电压敏感的荧光染料的光映射的关键方面。它包括培养的最佳的HL-1细胞单层，设置的映射设备，映射培养单层，和数据分析。 要成功映射培养细胞，关键是要准备一个均匀分布的单层细胞。当播种到盖玻片细胞，一定要均匀地分散细胞。始终映射完全长大，完​​全融合细胞单层，因为这些将最好的电起搏回应。同样重要的是，保...

独特的钙（Ca）和电压（V M）双通道光映射技术1-5正在成为一种有效的工具，同时记录V M和钙信号在两个完整的心和培养细胞单层。该技术使得有可能获得关于钙瞬变和动作电位之间的关系，以便更好地理解心律失常的底层电机制有力的信息。 培养的细胞单层已被证明是研究心脏电生理和心律失常的基本机制的有用的细胞模型。4,6-8的HL-1细胞是充分表征的心房肌细胞培养线。 HL-1细胞也保持着独特的差异化基因型和表现型，包括形态，电生理和成人心房肌细胞的药理特性。这些细胞表达心肌基因和蛋白质，包括性重要nt的心脏离子通道（ 即 ，L和T型钙通道）9和肌节收缩蛋白的成熟异构体通常在成人心房肌细胞作为其他人发现，我们以前曾报道10-13此外，HL-1细胞可以是培养，以形成一个2维（2-D）的肌细胞单层。因此，使用培养的HL-1单层的优点包括：1）较低的成本，更容易保持一个心肌细胞培养系不是分离和培养原代新生心肌细胞; 2）细胞的汇合单层减少而导致的心脏的3-D结构的结构的复杂性; 3）细胞单层可以消除间质纤维化的发生在完整心脏的干扰。这可以用来解剖一组肌细胞的特定电功能，而不成纤维细胞和间质基质的干扰; 4）药物或基因操纵的小路评估功能的后果捕获的原始图像的细胞单层，可以有效地实现。因此，HL-1细胞已经成为研究肌细胞生理学的各个方面以及起搏诱导的异常电活动的一个广泛使用的细胞模型13-16然而，特殊的处理和护理需要培养健康细胞单层，为了响应外部电起搏光学定位研究。此外，双荧光染料染色过程可能会很容易损坏的培养融合细胞单层的完整性。因此，在执行V 米/ CA双通道培养HL-1单层光学标测受到了极大的挑战。 该方法的目标是提供的关键步骤为在培养的HL-1单层成功执行双通道光学测绘的。在这里，我们提供了大量具体的细节上的优化协议HL-1细胞单层的准备，培养的细胞单层双通道光学测绘，测绘数据处理。

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voltage and calcium dual channel optical mapping of cultured hl-1 atrial myocyte monolayer

光学映射已被证明是有价值的技术来检测心电活动上都完好体外心和在培养的心肌细胞单层。的HL-1细胞已被广泛地用作用于研究心脏生理学的各个方面的2维细胞模型。但是，它已经受到了极大的挑战光学地图钙（Ca）瞬态和动作电位同时从相同的视野中培养HL-1心房细胞单层。这是因为特殊的处理和护理，需要准备一个可以电捕获并光学映射健康细胞。因此，我们已开发了一种最佳的工作协议进行双通道光学测绘的。在这个手稿中，我们已经详细介绍了如何进行双通道光的映射实验。这个协议是一个有用的工具，以提高动作电位传播和Ca动力学心律失常发展的理解。

有文化的融合单层呈现有规律的内在节奏表现在电影1。然后，我们执行的V 米/钙双通道光映射在一个完全融合的HL-1的单层。 图1A显示了V M和钙信号的例子痕迹从记录单击败。均匀地传播V M和使用该双通道光学测绘系统的Ca信号代表等时的地图显示在图1B和1C。均匀地传播动作电位和钙瞬变代表时间顺序的图像是从同一的HL-...

Watch this Scientific Journal Video about Voltage and Calcium Dual Channel Optical Mapping of Cultured HL-1 Atrial Myocyte Monolayer at JoVE.com

Voltage and Calcium Dual Channel Optical Mapping of Cultured HL-1 Atrial Myocyte Monolayer

This article describes the technique used to perform dual channel optical mapping in cultured HL-1 atrial cell monolayers. This unique protocol allows the simultaneous visualization of both calcium (Ca) and voltage (Vm) activity in the same area for the detailed detection and analysis of electrophysiological properties of culture monolayers.

培养HL-1心房肌细胞单层的电压和钙双通道光映射

Optical mapping has proven to be a valuable technique to detect cardiac electrical activity on both intact ex vivo hearts and in cultured myocyte ...

voltage-calcium-dual-channel-optical-mapping-cultured-hl-1-atrial

Research

JoVE Journal

Biology

10.1K 次观看.  Loyola University Chicago. This article describes the technique used to perform dual channel optical mapping in cultured HL-1 atrial cell monolayers. This unique protocol allows the simultaneous visualization of both calcium (Ca) and voltage (Vm) activity in the same area for the detailed detection and analysis of electrophysiological properties of culture monolayers.

视频: Voltage and Calcium Dual Channel Optical Mapping of Cultured HL-1 Atrial Myocyte Monolayer

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

适当的照顾和清洁的显微镜

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

科研

生物学

Phase Contrast and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by Zernike, in 1942, who received the Nobel prize for his achievement. DIC microscopy, introduced in the late 1960s, has been popular in biomedical research because it highlights edges of specimen structural detail, provides high-resolution optical sections of thick specimens including tissue cells, eggs, and embryos and does not suffer from the  phase halos  typical of phase-contrast images. This protocol highlights the principles and practical applications of these microscopy techniques.

Phase Contrast and Differential Interference Contrast DIC Microscopy

This protocol highlights the principles and practical applications of Phase and Differential Interference Contrast (DIC) Microscopy

相衬，微分干涉对比（DIC）显微镜

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by ...

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

光学测绘的Langendorff灌注大鼠心脏

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

Optical Recording of Electrical Activity in Guinea-pig Enteric Networks using Voltage-sensitive Dyes

The enteric nervous system (ENS) is a self-contained network with identified functions, capable of performing complex behaviors in isolation. Its neurons (10 to 25 &mu;m in diameter) are arranged in plexuses that are confined to distinct planes of the gut wall 1; the myenteric plexus can be found between the longitudinal and circular muscle layers, and the submucous plexus between the circular muscle layer and the mucosa. Since the effector systems for these plexuses (transporting epithelium, endocrine cells, immune elements, blood vessels and smooth muscle) are also contained within the gut wall, semi-intact preparations can be dissected that preserve individual components of different reflex pathways. The behavior of the effector systems is controlled by the submucous and myenteric plexuses acting in concert. Therefore, detailed knowledge of synaptic interactions within and between ganglia, and of communication between the plexuses, is essential for understanding normal gastrointestinal function. The ENS, as an intact nervous system, is a unique experimental model in which one can correlate molecular and cellular events with the electrical behavior of the neuronal network and its physiological outputs. Because of the quasi-two-dimensional organization of its plexuses, the ENS is particularly well suited for the study of neural networks using multiple site optical recording techniques that employ voltage-sensitive dyes 2,7,8,9. We will illustrate here the use of a relatively new naphthylstyryl-pyridinium dye (di-4-ANEPPDHQ) 3 that offers multiple advantages over its predecessors, including very low phototoxicity, slow rate of internalization, and remarkable chemical stability. When used in conjunction with a camera that permits sub-millisecond time resolution, this dye allows us to monitor the electrical activity of all the neurons in the field of view with a maximal spatial resolution of ~ 2.5 &mu;m at 100X magnification. At lower magnification (10X or 20X), the sacrifice of single-cell resolution is compensated by a gain in perspective, revealing the intricacies of the inter-ganglionic circuitry.

This protocol illustrates how voltage-sensitive dyes enable optical recording of electrical activity from intact neural networks such as the plexuses of the guinea-pig enteric nervous system, with an adjustable resolution that ranges from single-cells to multi-ganglionic circuitry.

在豚鼠肠道网络，利用电压敏感染料光学记录的电活动

The enteric nervous system (ENS) is a self-contained network with identified functions, capable of performing complex behaviors in isolation. Its neurons ...

Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of fluorescence can be changed by exposure to light of a specific wavelength. Optical highlighting allows noninvasive marking of a subpopulation of fluorescent molecules, and is therefore ideal for tracking single cells or organelles.
Critical parameters for efficient photoconversion are the intensity and the exposure time of the photoconversion light. If the intensity is too low, photoconversion will be slow or not occur at all. On the other hand, too much intensity or too long exposure can photobleach the protein and thereby reduce the efficiency of photoconversion.
This protocol describes a general approach how to set up a confocal laser scanning microscope for pc-FP photoconversion applications. First, we describe a procedure for preparing purified protein droplet samples. This sample format is very convenient for studying the photophysical behavior of fluorescent proteins under the microscope. Second, we will use the protein droplet sample to show how to configure the microscope for photoconversion. And finally, we will show how to perform optical highlighting in live cells, including dual-probe optical highlighting with mOrange2 and Dronpa.

This protocol describes a general approach to perform photoconversion of fluorescent proteins on a confocal laser scanning microscope. We describe procedures for the photoconversion of puried protein samples, as well as for dual-probe optical highlighting in live cells with mOrange2 and Dronpa.

在活细胞荧光蛋白的双探头光学突出纯净的光转化的

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

基于荧光测定活细胞中的钙进入商店经营：从培养的肿瘤细胞，骨骼肌纤维

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Isolation and Kv Channel Recordings in Murine Atrial and Ventricular Cardiomyocytes

KCNE genes encode for a small family of Kv channel ancillary subunits that form heteromeric complexes with Kv channel alpha subunits to modify their functional properties. Mutations in KCNE genes have been found in patients with cardiac arrhythmias such as the long QT syndrome and/or atrial fibrillation. However, the precise molecular pathophysiology that leads to these diseases remains elusive. In previous studies the electrophysiological properties of the disease causing mutations in these genes have mostly been studied in heterologous expression systems and we cannot be sure if the reported effects can directly be translated into native cardiomyocytes. In our laboratory we therefore use a different approach. We directly study the effects of KCNE gene deletion in isolated cardiomyocytes from knockout mice by cellular electrophysiology - a unique technique that we describe in this issue of the Journal of Visualized Experiments. The hearts from genetically engineered KCNE mice are rapidly excised and mounted onto a Langendorff apparatus by aortic cannulation. Free Ca2+ in the myocardium is bound by EGTA, and dissociation of cardiac myocytes is then achieved by retrograde perfusion of the coronary arteries with a specialized low Ca2+ buffer containing collagenase. Atria, free right ventricular wall and the left ventricle can then be separated by microsurgical techniques. Calcium is then slowly added back to isolated cardiomyocytes in a multiple step comprising washing procedure. Atrial and ventricular cardiomyocytes of healthy appearance with no spontaneous contractions are then immediately subjected to electrophysiological analyses by patch clamp technique or other biochemical analyses within the first 6 hours following isolation.

Kv channel dysfunction is associated with cardiac arrhythmias. In order to study the molecular mechanisms that lead to such arrhythmias we utilize a systematic protocol for isolation of atrial and ventricular cardiomyocytes from Kv channel ancillary subunit knockout mice. Isolated cardiomyocytes can then immediately be used for cellular electrophysiological studies, biochemical or immunofluorescence (IF) assays.

在小鼠的心房和心室肌细胞的分离及钾离子通道的录音

KCNE genes encode for a small family of Kv channel ancillary subunits that form heteromeric complexes with Kv channel alpha subunits to modify their ...

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

跨膜蛋白的重组，电压门控离子通道，KvAP，进入巨人单层囊的显微镜和膜片钳研究

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Single-channel Analysis and Calcium Imaging in the Podocytes of the Freshly Isolated Glomeruli

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells in the pathogenesis of various renal diseases has been established since podocyte injury leads to proteinuria and foot process effacement. It was previously reported that various endogenous agents may cause a dramatic overload in intracellular Ca2+ concentration in podocytes, presumably leading to albuminuria, and this likely occurs via calcium-conducting ion channels. Therefore, it appeared important to study calcium handling in the podocytes both under normal conditions and in various pathological states. However, available experimental approaches have remained somewhat limited to cultured and transfected cells. Although they represent a good basic model for such studies, they are essentially extracted from the native environment of the glomerulus. Here we describe the methodology of studying podocytes as a part of the freshly isolated whole glomerulus. This preparation retains the functional potential of the podocytes, which are still attached to the capillaries; therefore, podocytes remain in the environment that conserves the major parts of the glomeruli filtration apparatus. The present manuscript elaborates on two experimental approaches that allow 1) real-time detection of calcium concentration changes with the help of ratiometric confocal fluorescence microscopy, and 2) the recording of the single ion channels activity in the podocytes of the freshly isolated glomeruli. These methodologies utilize the advantages of the native environment of the glomerulus that enable researchers to resolve acute changes in the intracellular calcium handling in response to applications of various agents, measure basal concentration of calcium within the cells (for instance, to evaluate disease progression), and assess and manipulate calcium conductance at the level of single ion channels.

Changes in the intracellular calcium levels in the podocytes are one of the most important means to control the filtration function of glomeruli. Here we explain a high-throughput approach that allows detection of real-time calcium handling and single ion channels activity in the podocytes of the freshly isolated glomeruli.

单通道分析和钙成像在新鲜分离的肾小球足细胞

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells ...

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a significant role in certain types of arrhythmia. Because arrhythmias are spatially distinct, emergent phenomena, they must be investigated at the tissue level. However, methods for directly probing SR Ca2+ in the intact heart remain limited. This article describes the protocol for dual optical mapping of transmembrane potential (Vm) and free intra-SR [Ca2+] ([Ca2+]SR) in the Langendorff-perfused rabbit heart. This approach takes advantage of the low-affinity Ca2+ indicator Fluo-5N, which has minimal fluorescence in the cytosol where intracellular [Ca2+] ([Ca2+]i) is relatively low but exhibits significant fluorescence in the SR lumen where [Ca2+]SR is in the millimolar range. In addition to revealing SR Ca2+ characteristics spatially across the epicardial surface of the heart, this approach has the distinct advantage of simultaneous monitoring of Vm, allowing for investigations into the bidirectional relationship between Vm and SR Ca2+ and the role of SR Ca2+ in arrhythmogenic phenomena.

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

This article describes the detailed protocol and equipment necessary for dual optical mapping of transmembrane potential (Vm) and free intra-sarcoplasmic reticulum (SR) Ca2+ in the Langendorff-perfused rabbit heart. This method allows for direct observation and quantification of Vm and SR Ca2+ dynamics in the intact heart.

的内肌浆网钙光学标测<sup&gt; 2+</sup&gt;和的Langendorff灌注兔心脏跨膜电位

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a ...