Name: flash-and-freeze: a novel technique to capture membrane dynamics with electron microscopy
Uploaded: 2017-05-01T12:00:00.000+00:00
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Description: Watch this Scientific Journal Video about Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy at JoVE.com

这项工作是由来自约翰霍普金斯大学（SW）的资金支持。我们感谢医学显微镜基金的约翰霍普金斯大学为他们提供技术支持。我们感谢埃里克·乔琴森和克里斯蒂安·罗斯蒙及其实验室成员，该技术的发展。我们感谢M.韦恩·戴维斯初始设备的设计。我们感谢保罗·齐杰，克韦塔·托莫瓦和Delgermaa Luvsanjav为他们提供技术援助。我们也感谢纳塔利·R·哈密尔顿和格兰特·F·库西克对稿件他们的批判性阅读。

所有的实验都按照规则和动物使用的法规由美国国立卫生研究院进行的。该协议被批准的动物护理和使用委员会（IACUC）医学院的约翰霍普金斯大学。 1.隔离老鼠的海马神经元的文化和<ol><li>从0日龄解剖皮质-天2（P0 - 2）小鼠脑18。隔离从皮质的星形胶质细胞。 注:鼠标脑斩首后分离。星形胶质细胞作为海马神经元饲养层。 </li><li>治疗用0.05％胰蛋白酶-EDTA的800μL的皮质15分钟，在37℃以解离星形胶质细胞。 </li><li>培养在T-75烧瓶中的星形胶质细胞用13毫升含有10％胎牛血清（FBS）和0.2％青霉素-链霉素的一周，在37℃和5％CO 2的DMEM。 </li><li>放置一个酸洗涤和灭菌18毫米玻璃盖玻片PE一个12孔板的孔 - [R。 </li><li>简言之洗26毫米碳涂覆的蓝宝石盘在70％乙醇，并放置在每个玻璃盖玻片的顶部。 </li><li>通过混合3毫升的17 nM的乙酸用1mL鼠尾胶原和1mL PDL（1毫克/毫升）制备聚-D-赖氨酸（PDL）溶液。适用的PDL溶液200μL的蓝宝石盘和玻璃盖玻片上，在室温5分钟。 </li><li>卸下PDL溶液并风干。在使用之前，如步骤1.4制得消毒的板 - 1.6在紫外光下30分钟。 </li><li>种子来自步骤1.3在2mL DMEM的星形胶质细胞以5×10 4个细胞/孔的密度在板中的步骤制备1.4 - 1.6。它们生长于37℃和5％CO 2的一个星期。 </li><li>培养神经元前向每个孔至少几个H:加入20μL氟脱氧尿苷（80μM终浓度）。 注:氟脱氧尿苷星形胶质细胞停止分裂。 </li><li>介质改变为1.5毫升的神经元的BAS含有1％L-丙氨酰-L-谷氨酰胺（参见材料的表 ），对神经元细胞的2％无血清补充物（参见材料的表 ）和0.2％青霉素-链霉素人培养基（参见材料的表 ） 。 </li><li>通过加入20个单位的木瓜蛋白酶的5毫升的酶溶液（1.65毫半胱氨酸，1mM的CaCl 2的，和0.5mM EDTA在DMEM）制备木瓜蛋白酶溶液。酸化通过使CO 2气体20分钟将溶液。用0.22μm过滤器过滤灭菌。 </li><li>收获来自P0脑- 2鼠标18。立即大脑转移到冰冷的Hanks&#39;，平衡盐溶液（HBSS）。解剖海马体视显微镜下，保持在HBSS淹没的组织。 </li><li>将两个海马在1mL在步骤1.11中制备的木瓜蛋白酶溶液。在37℃和750rpm下孵育上的热混合器1个小时。 </li><li>用1mL灭活含有溶液代替木瓜蛋白酶2.5毫克胰蛋白酶抑制剂和0.5毫克每DMEM毫升白蛋白。在37℃下孵育5分钟。吸掉失活的解决方案。 </li><li>添加的神经元基础培养基（参见材料的表 ），以分离海马200μL。磨碎用200μl移液管尖端来离解细胞。等到未解细胞沉淀在底部。小心地从顶部清除细胞培养基。 </li><li>重复步骤3倍1.15。池中的所有解离的细胞的一个新的1.5毫升离心管中。 </li><li>计数使用血球细胞的数量。在6.5×10 4个细胞的密度板神经元/孔的在步骤1.1中制备的星形胶质细胞层的顶部- 1.10。 </li><li>感染慢病毒在DIV 3（图3d 体外 ）表达18的通道视紫红神经元。 </li><li>上DIV 14执行快速和冻结，如在步骤2.1中所述 - 2.5，下方。 </li></ol> 2.闪存的D-冻结<ol><li> 固色剂的制备 <ol><li>下化学通风橱中，下列物质添加至锥形管中，以制备固定剂:2.5毫升戊二醛（10％丙酮的库存）0.25g的四氧化锇，0.25毫升的水，和22.25毫升无水丙酮中。 注:各成分的最终浓度应为1％在丙酮中。戊二醛加入冷冻置换过程中保持该蛋白质的结构。注意:四氧化锇的急性毒性高。接触蒸汽可能会损坏眼睛的角膜。它应该只在经过认证的化学罩处理。 </li><li>分装1毫升固定剂的成编号的低温小瓶（2mL）中。保持冷冻在液氮中直至使用的固定液。 注:将四氧化锇和戊二醛交叉反应并沉淀;因此，一旦混合，等分试样，立即盖上管，并淹没冻存管在液氮中冻结固定剂。用铅笔编号的低温核不小瓶，因为丙酮可以洗掉标记。 </li></ol></li><li> 生理盐水准备 <ol><li>通过混合HEPES（10毫米，pH值7.5），NaCl的（140毫摩尔），氯化钾（2.4毫摩尔），和葡萄糖（10毫摩尔）使生理盐水溶液中。 注:这些值是终浓度。低温防护剂不用于单层培养。然而，需要对试样大于5μm厚的适当冷冻保护。通常建议使用20％BSA的。酵母膏和大肠杆菌OP50也可以用作冷冻保护剂对蝇幼虫或线虫 。 </li><li>在4 mM和MgCl 2的添加氯化钙2以1mM的终浓度。 注: 氯化钙和氯化镁的浓度取决于实验。为了确保胞吐中间体的捕获，4mM的钙用于这些特定实验，以增加囊泡18的释放概率。 </li><li>检查摩尔渗透压浓度使用渗压计。确保它是300±5毫渗量。 </li><li>到3μm和GABA受体拮抗剂（荷包牡丹碱）的终浓度添加AMPA受体拮抗剂（NBQX），以30μM的终浓度。 注:加入的神经递质受体拮抗剂，以避免以下的神经元的刺激18经常性网络活动。 </li><li>预热生理盐水至37℃使用。 </li></ol></li><li> 专门高压冷冻和一个自动冷冻置换单元的制备 <ol><li>之前的高压冷冻，通过填充用液氮罐冷却的自动冷冻置换单元至-90℃。 </li><li>通过将其放置在试样室内部中一个小杯冷却丙酮至-90℃。 </li><li>填充高压冷冻的液体硝基液氮杜瓦瓶和存储杜瓦（参见材料的表 ）根。 </li><li>设置使用触摸屏显示器的光刺激方案。 <ol><li>名称点击"编辑"旁边的"程序名称"光刺激窗上的程序。另一个窗口会弹出。 </li><li>通过键入"在"期间"15000毫秒"，"黑暗阶段"，"100毫秒"，"10毫秒"设立程序"脉冲，"和"1"，在为10的单个刺激"周期数"女士。冷冻所述细胞90毫秒后（ 图1C）。 注意:"黑暗阶段"允许细胞样品加载期间从曝光恢复。 "周期"定义刺激频率。例如，如果刺激应以20Hz被应用，则此栏应在50毫秒设置。 "脉冲"定义了光刺激的持续时间。最后，"周期数"定义施加刺激的总数。为了建立一个高频率的刺激，请参阅<strong&gt;图1E。 </li></ol></li><li>对于"无刺激"控制，在光类型"15秒"，"黑暗阶段"，"0毫秒"，在"期限"，"1毫秒"，在"脉冲，"和"0"，在"周期数"刺激设置窗口，在步骤2.3.4所描述的。 注:默认情况下，"脉搏"必须至少为1毫秒。 </li><li>在主屏幕上，确保了对光刺激的复选框被选中。 </li><li>通过点击"样本存放"在主屏幕上设置的存储协议。点击"编辑"。在下面的窗口中，使用"+"或" - "，以选择"2"存储在每个通道2盘（3个通道在总）。选中"存储LN2启用。" </li></ol></li><li> 样和在高压冷冻冷冻 注:所有样品装置和装载步骤与7.5-60X放大范围为立体显微镜下完成的。一个tweezER在步骤2.4.1使用 - 2.4.4操作标本。实验必须在生理温度下进行。 <ol><li>放置一个蓝宝石盘，电池侧朝上，在黑色，中间板（ 图1B）的孔中。 </li><li>放置在所述蓝宝石磁盘（ 图1B）100μm的间隔环。 </li><li>盘的一侧从步骤2.2浸渍在预热的盐溶液后放置在间隔环（ 图1B）空白蓝宝石磁盘。确保没有气泡被困在两个蓝宝石圆盘之间。 </li><li>放置另一个为100μm间隔环和一个400μm的间隔环（ 图1B）。取出用滤纸多余的液体。 </li><li>放置组件从步骤2.4.3两个透明半圆柱体（ 图1A）之间。关上红色封面的启动冻结过程。 注意:一旦盖被关闭预设协议自动运行。将样品保持在在冷冻室中的相同的方向，和光的闪光从样品组件的顶部上。红色封面的弹出自动备份，一旦冷冻过程完成。 </li><li>存储在存储杜瓦瓶样品。 注:冷冻后，试样被自动滴入充满液氮的存储杜瓦并存储在那里，直到进一步处理。存储杜瓦具有三个腔室，并且每个腔室可以最多容纳3个样品。典型地，两个样品在相同的刺激条件下冻结，并且高压冷冻被编程为既存储在相同的腔室。 </li><li>重复步骤2.4.1 - 2.4.6每个样品。 </li><li>继续执行步骤2.5一旦所有的室都满了。 注:设备设定为存储多达6个样品在存储杜瓦在一个时间。因此，每6 个样本之后，步骤2.5必须执行。一旦卸载，重复步骤2.4.1 - 2.4.6。 </li></ol></li><li> 样品采集和传送到自动冷冻置换部件 <ol><li>开门到存储杜瓦，它位于所述高压冷冻机的表下。除去存储杜瓦并将其放在工作台上。 </li><li>使用手，从杜瓦瓶取出样品室和在充满液氮的专门样品盘传送它。解锁按钮释放样品杯中。 </li><li>除去从试样杯使用一对镊子的后预冷的镊子的顶端用液氮透明半圆柱体（〜-196℃）。小心地将中间，黑盘转移到含有液氮的小杯。 注意:镊子的尖端必须预冷却至液氮的温度。样品必须在液氮下在任何时候都保持在防止冰晶形成。 </li></ol></li></ol> 3.冷冻置换在金tomated冷冻置换部<ol><li>使用预冷镊子（提示在〜-196℃），迅速地从步骤2.5.3转移中间板到预冷的丙酮（-90℃）。 </li><li>通过轻轻晃动或拍打独立于中板蓝宝石盘。 注意:有时，可能难以将蓝宝石圆盘从中间分隔板。在这种情况下，留在预冷丙酮中间板（-90℃）几分钟。用镊子轻轻拍打也有助于从中间板分离的蓝宝石。 </li><li>将包含置换单元的样品室内部的固定剂（步骤2.1）的低温小瓶中。蓝宝石磁盘传输到低温冻存管，并放置在瓶帽。 </li><li>设置冷冻置换程序如下:（ⅰ）-90℃下进行5 - 30小时，（ⅱ）-90 - -20℃下在14个小时（5℃/小时），（ⅲ）-20℃ 12个小时，和（iv）-20℃ - 20℃下，在4小时（10℃/小时）。 注:杜在-90℃下的第一步骤的定量可以进行改变。冷冻置换的总持续时间在该程序在早晨（〜为1.5d postexperiment）上午08点周围，以便能够在白天期间执行后续步骤结束这样的方式设置。 </li></ol> 4.渗透和塑料包埋用环氧树脂<ol><li>一旦程序结束，用戴手套的手到包含蓝宝石磁盘低温小瓶从替代单元的试样腔室转移到化学通风橱。 </li><li>使用移液管，添加丙酮（室温）到每个小瓶中低温和洗涤各蓝宝石盘4 - 6×1 - 2小时。 </li><li>任选地，如果需要额外的对比度孵育在0.1％乙酸双氧铀的试样1个小时。洗4 - 2小时 - 用丙酮超过1 6倍。 注:注意:没有与乙酸双氧铀，这将导致上呼吸道的刺激的吸入以下内部曝光相关的风险。高曝光可以damagË血细胞。醋酸铀工作应在通风排气设备来完成。建议防护服。 </li><li>通过称量6.2克甘油多缩水甘油醚的，4.4克双酚A型环氧树脂，以及12.2克十二碳烯基琥珀酸酐（DDSA）的制备液体环氧树脂平台。调匀并添加苄基二甲胺（BDMA）的800μL，同时混合。除气10分钟。 </li><li>从在步骤4.4中制备的100％环氧树脂在丙酮中制备30，70，和90％的环氧树脂。 </li><li> 30％的环氧树脂加入到含有蓝宝石磁盘低温小瓶中，并孵育2 - 以120rpm在轨道摇床在室温3小时。 </li><li>通过移液代替70％的环氧树脂的30％环氧树脂和孵育3 - 在定轨摇床中在室温下4小时。 </li><li>使用镊子，转移的每个蓝宝石磁盘，细胞朝上，给嵌入胶囊的帽。 90％的环氧树脂加入到帽。孵育O / N在4℃下。 </li><li>第二天，使新鲜环氧树脂的介质，因为在第4步0.4。 </li><li>转移的每个蓝宝石盘，电池侧朝上，给嵌入胶囊的帽。填充用新鲜制备的100％环氧树脂的帽。改变为新鲜100％环氧树脂，每2小时，重复三次。 </li><li>放置样品在60℃烘箱中48小时以聚合。 </li></ol> 5.安装样品<ol><li>放置上，使得蓝宝石磁盘位于所述树脂块的顶部立体显微镜样品颠倒。通过用刀片刮擦它除去从顶部环氧树脂的薄层。 </li><li>使用刀片，切割沿蓝宝石盘的边缘的浅线;这条线有助于将蓝宝石盘从环氧树脂在步骤5.3中分离出来。 </li><li>通过将其浸入液氮中约10秒分开环氧树脂蓝宝石磁盘。 注:细胞将留在环氧树脂。 </li><li>去除蓝宝石磁盘后，使用解剖范围找到与细胞的区域（4-10×放大倍数）。使用刀片的感兴趣区域（〜2×2mm）的周围切割（双刃）。保持试样在适当位置，而将试样贴在显微镜的表面执行此步骤。 </li><li>安装使用含有由环氧树脂制成的圆柱形虚设块上乙基氰基丙烯酸酯胶含有细胞的小的塑料片（〜2×2×5毫米）。孵育在60℃下的块1个小时。 </li></ol> 6切片<ol><li>使用超薄切片机到部分样品。 <ol><li>修剪塑料包埋标本用玻璃刀的表面以3mm / s的速度和厚度为200nm /截面。切4 - 从其中星形胶质细胞位于表面5部分。 </li><li>切换到金刚石刀和部分以0.8mm / s的速度和厚度为40纳米/截面。切20 - 25节。 </li></ol></li><li>收集覆盖有0.7％聚乙酸乙烯酯在单时隙铜格栅部分的带（见材料的表 ）。 </li></ol> 7.成像用透射电子显微镜（TEM） <ol><li>之前TEM成像，染色在甲醇中，用2.5％乙酸双氧铀的部分5分钟。 </li><li>洗，在50％甲醇中的网格15x连接。然后，洗净在超纯水中15倍，与每次洗涤持续30秒。 </li><li>简言之空气干燥部和网格放置到TEM的试样保持器。 </li><li>图片在93,000X的放大倍率。 </li><li>采集图像。 注:成像通常做盲目的时间点或基因型，通常〜200图像采集/时间点。 </li></ol> 8.图像分析<ol><li>分析使用软件（ 例如，ImageJ的）用自定义宏（渡边，戴维斯和约根森，未发表）的图像。 注:X / Y坐标是从囊泡记录，质膜，有源区的膜，和在突触所有其它膜结合的细胞器。文本FIL包含该信息的ES被导出到另一软件（ 例如，MATLAB）。数据使用自定义程序的进一步分析。 </li></ol>

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"闪蒸和冻结"的方法通过诱导与光遗传学特定细胞事件，并通过刺激19后冻结确定的时间点的细胞可视化膜动力学。在此演示中，我们使用通道视紫红，光敏阳离子通道，刺激神经元和捕获在突触末端融合和突触小泡的回收。近年来，许多光遗传学工具已经开发22，23，所有这些都与闪存和冻结兼容。例如，细胞器贩卖可使用隐和CIB1 24的光诱导的异二聚化来诱导。类似地，质膜的脂质组合物可以通过磷酸肌醇磷酸酶的光诱导易位到质膜25来改变。此外，像偶氮苯CH小，光敏感的化合物安格构象取决于照明波长。这种构象变化可以被用来激活配体门控通道或改变脂质组合物在膜26，27。笼化合物也可用于诱导细胞活性。然而，在目前的安装中使用的LED可能不产生用于解笼锁足够的能量;因此，该系统的进一步优化是可能必要的。然而，这些光可激活工具的应用是灵活的，很多细胞事件可以由闪光引起。 "闪存和冻结"可以捕获产生的膜动力学。 有以"闪存和冻结"法两个主要局限。首先，它捕获来自不同小区的特定事件的"快照"。换句话说，它是不可能在一段时间跟随膜动力学在一个小区中。因此，对于任何蜂窝的重建，即使吨，必须获得和分析大量的来自每个样品和在每个时间点的图像。此外，在神经元中，图像的更大的数目是必要的，因为突触小泡的融合只需要地点20 -突触的30％小鼠海马神经元18,28。如此大的数据集的分析需要时间的巨大数额。在未来，图像采集和分析需要实现自动化，使办法更有效29,30。 第二个限制是由高压冷冻技术的性质强加的。当细胞冷冻，细胞水重排形成冰晶如果冷冻速率是低于100 K / S 21。这些冰晶可以渗透膜或浓缩溶质，以改变局部的渗透压，从而导致膜的破裂。为了避免冰晶，HIGh压力（〜2000大气压）施加到标本。由于过冷却效果，100 K / s的冷冻速率是足够的，以防止水从在此压力下21形成冰晶。从理论上讲，如500微米可以在没有冰晶被冻结，但大约为200μm是容易的实际限制，因为冰的立方形形式倾向于形成在厚组织，从而损害形态标本一样厚。当处理标本厚于5μm，则使用适当的冷冻保护的，如BSA，是必要的。然而，BSA将改变溶液的渗透压，并可能影响细胞的生理响应。因此，需要大量的对照实验来验证尤其系统中使用的BSA。冰晶也可以高压冷冻后，如果标本无意中从液氮浴中取出形成。因此，关键是要在保持液氮将样品在任何时候，使用预冷却镊子操作它们。 在规划的实验，以下三点应予以考虑。首先，光的最大强度（在460nm的线）是5.5 - 8.0毫瓦/毫米2。是否该强度足以诱导活动都必须与上之前闪存和冷冻实验的荧光显微镜的活细胞成像进行验证。其次，实验必须在生理温度下进行。高压冷冻机的阶段升温至37℃与小鼠海马神经元31中的实验。最后，时间点必须仔细选择捕捉膜动力学。初步研究表明，细胞内吞作用是后100毫秒刺激的完整。因此，三个附加的时间点（15，30，和50毫秒）还进行了检查以跟随膜动力学17,18。这些时间点是必要的，可视化的膜贩运事件突触传递期间秒。然而，对于时间点的数目的要求在每个细胞事件不同。因此，有几个时间点应该启动大数据集收集之前进行采样。

一个小区内的可视化膜和蛋白质动力学是理解特定过程的细胞生物学中的关键步骤。动态广告投放管理事件可以使用光或荧光显微镜来捕获。然而，亚细胞状况很大程度上在这样的图像丢失，因为亚细胞结构不能完全的"绘制"由染料或荧光探针和空间分辨的光谱和1，2。在另一方面，当电子显微镜可以以精致的细节描绘亚细胞结构，它不能捕获细胞动力学，因为标本必须成像前固定。因此，它通常是不够的仅使用一个成像模态完全理解细胞动力学。 为了克服光学和电子显微镜的局限，相关的显微镜技术已经开发出来。相关光学和电子显微镜（CLEM）可视化使用光镜和电镜底层的亚细胞结构细胞内动力学。在CLEM，从事各种过程，例如胞质和胞吞作用3，4，5，6个细胞，是活成像，然后用于电子显微镜进行处理。虽然CLEM捕捉细胞内动态的某些方面，有四个因素限制了这种方法的实用性。分由于缓慢扩散和固定剂7的反应-首先，时间分辨率被细胞能多快被固定，后者通常需要限量。其次，亚细胞结构中观察到事后 8;因此，动态的形态变化，不能使用这种方法捕获。第三，荧光和电子显微照片不能精确由于组织SH对准用于电子显微镜9，10中的样品制备过程中通过脱水rinkage引起的。第四，像细胞分裂和内吞作用的事件不采取在同一时间发生在每一个细胞5，11，因此，这是从事活动的特定细胞必须从一个人口众多的细胞进行鉴定。这个过程通常是费力。因此，一种新的方法是必要的，以诱导在每个细胞中的特定事件，并在确定的时间点，以捕获由细胞的快速固定化所得到的细胞动力学。 最近，一些工具被开发出来，以引起利用光（光遗传学）特定细胞动力学。紫红质通道是从莱茵衣藻 12,13分离的光敏感，非选择性阳离子通道。当紫红质通道中neurona表示升膜，光的简要闪光诱导钠离子的流入神经元和触发动作电位14，15。然后将动作电位传播到突触终端，其中，突触小泡融合毫秒16,17内，18因此，通道视紫红诱导神经元活性。跟随在突触末端膜动力学，神经元必须在限定的时间点与MS精度刺激后固定。 为了诱导神经元活性后捕获膜动力学，我们耦合光刺激用高压冷冻17，18，19。高压冷冻允许具有降低的冰晶形成20个细胞的近瞬时固定。冰晶CAÑ破裂膜和破坏的亚细胞结构21。通过改变刺激和冷冻之间的时间间隔，突触末端内的膜运输被捕获的动作电位的感应以下。 在这里，我们证明了用商业化的高压冷冻，夫妻光刺激的高压冻结一毫秒的时间控制实验程序。与需要的外部设备来控制光刺激和冷冻其它仪器，光刺激完全集成在该系统中，并且可以与毫秒精度19来施加。这个过程涉及多个步骤。 1）小鼠海马神经元是在蓝宝石上的磁盘培养并慢病毒感染携带表达载体紫红质通道18。 2）神经元刺激和刺激后在确定的时间点冷冻。 3）玻璃化水替代通过固定剂20d与有机溶剂，而脂质和蛋白质是交联的以保持细胞内结构。 4）将样品渗透并包埋在环氧树脂中。 5）超薄切片使用超微切片收集。 6）薄切片进行成像上的透射型电子显微镜。 7）图像采集和分析相对于时间点或基因型盲目地进行的。细胞动力学可以通过时间分辨的图像17，18的重建来确定。样品制备（步骤2 - 5以上）需要一个星期，但随后的图像分析需要半年到一年。

<table><tbody><tr><td>Freeze-substitution and Low-temperature Embedding System&amp;nbsp; for Light and Electron Microscopy -AFS II</td><td>Leica&amp;nbsp;</td><td></td><td></td></tr><tr><td>High Pressure Freezer - EM-ICE</td><td>Leica&amp;nbsp;</td><td></td><td>EM ICE is a specialized high pressure freezer that allows precise control of light stimulation and freezing.</td></tr><tr><td>Osmometer</td><td>Gonotec</td><td></td><td></td></tr><tr><td>Ultramicrotome UC7</td><td>Leica&amp;nbsp;</td><td></td><td></td></tr><tr><td>Oven</td><td>Blue M</td><td></td><td></td></tr><tr><td>Razor blade</td><td>Personna</td><td></td><td></td></tr><tr><td>Glutaraldehyde</td><td>EMS</td><td>16530</td><td></td></tr><tr><td>Osmium tetroxide</td><td>EMS</td><td>RT19132</td><td>Toxic, open only under certified chemical hood</td></tr><tr><td>Acetone</td><td>EMS</td><td>RT10016</td><td></td></tr><tr><td>HEPES</td><td>Emdmillipore</td><td>391340-250GM</td><td></td></tr><tr><td>Glucose</td><td>Sigma</td><td>49159-1KG</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>746436-1KG</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653-1KG</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>21115-250ML</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>J.T.Baker</td><td>2444-01</td><td></td></tr><tr><td>Liquid epoxy resin Eponate 12</td><td>Ted Pella</td><td>18028</td><td></td></tr><tr><td>Bisphenol A epoxy resin Araldite 502</td><td>Ted Pella</td><td>18028</td><td></td></tr><tr><td>Dodecenyl succinic anhydride (DDSA)</td><td>Ted Pella</td><td>18028</td><td></td></tr><tr><td>Benzyl dimethyl amine (BDMA)</td><td>Ted Pella</td><td>18241</td><td></td></tr><tr><td>Special embedding (BEEM) capsule</td><td>EMS</td><td>70021</td><td></td></tr><tr><td>Copper Grid</td><td>Ted Pella</td><td>1GC12H</td><td></td></tr><tr><td>Polyvinyl acetate (Pioloform F)</td><td>Ted Pella</td><td>19244</td><td></td></tr><tr><td>Uranyl acetate</td><td>Polysciences</td><td>21447-25</td><td></td></tr><tr><td>Ethyl cyanoacrylate (Super glue)</td><td>Scotch</td><td>170497</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Themo scientific</td><td>25300-120</td><td></td></tr><tr><td>DMEM (Dulbecco&#039;s Modified Eagle&#039;s Medium)</td><td>Thermo scientific</td><td>10569-044</td><td>Should be warmed to 37 &amp;deg;C before use</td></tr><tr><td>FBS (Fetal Bovine Serum)</td><td>Thermo scientific</td><td>26140-079</td><td></td></tr><tr><td>Pen-Strep</td><td>Thermo scientific</td><td>15140-122</td><td></td></tr><tr><td>Fluoro-deoxyuridine (FUDR)</td><td>Sigma</td><td>F0503</td><td></td></tr><tr><td>Glass cover slip</td><td>Fisher</td><td>S175223</td><td>Should be acid-washed</td></tr><tr><td>Sapphire disc</td><td>Technotrade</td><td>616-100</td><td></td></tr><tr><td>Acetic acid</td><td>Emdmillipore</td><td>1000631011</td><td></td></tr><tr><td>Poly-D-Lysine (PDL)</td><td>Sigma</td><td>P6407</td><td></td></tr><tr><td>Rat tail collagen</td><td>Thermo scientific</td><td>A10483-01</td><td></td></tr><tr><td>Neurobasal A</td><td>Fisher</td><td>10888022</td><td>Should be warmed to 37 &amp;deg;C before use</td></tr><tr><td>L-alanyl-L-glutamine (Glutamax)</td><td>Fisher</td><td>35050-079</td><td></td></tr><tr><td>Seraum free supplement&amp;nbsp; (B-27)&amp;nbsp;</td><td>Fisher</td><td>17504044</td><td></td></tr><tr><td>Hanks&#039;-balanced Salt Solution (HBSS)</td><td>Thermo scientific</td><td>24020117</td><td></td></tr><tr><td>Papain</td><td>Worthington</td><td>LS003126</td><td>Active Unit should be calculated for each batch</td></tr><tr><td>Thermomixer</td><td>Eppendorf&amp;nbsp;</td><td>Thermomixer C</td><td></td></tr><tr><td>Trypsin inhibitor</td><td>Sigma</td><td>T9253</td><td></td></tr><tr><td>NBQX</td><td>Tocris</td><td>03-731-0</td><td></td></tr><tr><td>Bicculine</td><td>Tocris</td><td>01-091-0</td><td></td></tr><tr><td>Whatman I filter paper</td><td>GE</td><td></td><td></td></tr><tr><td>Transmission Electron Microscope</td><td>Philips CM20</td><td></td><td></td></tr></tbody></table>

flash-and-freeze: a novel technique to capture membrane dynamics with electron microscopy

细胞不断地改变它们的膜结构和蛋白质的分布，但它是非常困难的，在MS和纳米量级的时间和空间分辨率分别可视化这些事件。我们已经开发出一种时间分辨电子显微技术，"闪光和冻结，"诱导细胞事件与光遗传学和刺激后在规定的时间点冷冻细胞可视化所产生的膜动力学。为了证明这种技术，我们表达通道视紫红，光敏阳离子通道，在小鼠海马神经元。光的闪光刺激神经元活动，并通过突触小泡的融合诱导从突触末端释放神经递质。神经元的光遗传学刺激加上高压冷冻跟随突触传递中的形态学变化。使用商业工具，我们捕捉到突触小泡的融合和SYNA恢复普天泡膜。为了显现事件序列，产生和分析盲目的，因为形态学变化在不同的细胞随着时间的推移，接着大的数据集。尽管如此，闪存和冻结允许膜动力学的电子显微照片与毫秒时间分辨率的可视化。

使用上文描述的协议中，我们执行在小鼠海马神经元"闪光和冻结"实验表达紫红质通道。这些神经元被冻结或者15毫秒或光后发病100毫秒。我们先前已经证明，突触小泡的胞吐作用和胞吞作用发生在在15毫秒分别为18突触终端和100个毫秒的时间点，。这些事件在适当的时间（ 图2）被成功地捕获，这表明快速和冻结实验可以在所选择的专门的高压冷冻成功地进行（参见材料的表 ）。 <img alt="图1" src="/files/ftp_upload/55664/55664fig1.jpg" /> 图1.样和编程的高压冷冻。 A）高-p的样本载物台ressure冰柜。中间板，在插图中用于结构详细地示出，被放置在上样一个克莱姆·霍尔德。一项所述的半圆柱体的被放置在样品载物台的底部部分，和另一个连接用夹子到顶盖。一旦样品被加载，中间板被向前推到下半圆筒和盖是关闭的，以启动冻结。 B）试样组件。含神经元的蓝宝石盘放置在中间板的孔，与所述电池侧朝上。 A 100微米环直接井内蓝宝石盘上方放置。然后，在生理食盐水中浸渍空蓝宝石盘放置与溶液的一面朝下。气泡必须避免。最后，一个100μm的环和一个400μm的环紧贴地放在上面。任何额外的液体用滤纸除去。 C）在环氧树脂浸没蓝宝石磁盘包埋胶囊的横截面。该SAPP雇用盘被放置在胶囊的底部，与细胞面朝上并覆盖有环氧树脂用于渗透和嵌入。 D）编程为一个单一的，10毫秒刺激高压冷冻机中。该标本的光脉冲后冻结90毫秒。 E）编程为10个刺激高压冷冻在20赫兹。该标本的最后一个光脉冲后冻结5毫秒。 <a href="https://ecsource-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55664/55664fig1large.jpg" target="_blank">请点击此处查看该图的放大版本。</a> <img alt="图2" src="/files/ftp_upload/55664/55664fig2.jpg" /> 图2.可视化胞吐和胞吞作用在小鼠海马神经元。海马神经元受到刺激一次，并在指定的时间冻结。电子显微照片显示突触小泡A的胞外分泌）和超快的内吞作用<s仲&gt; B）。 PSD，突触后密度。 <a href="https://ecsource-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55664/55664fig2large.jpg" target="_blank">请点击此处查看该图的放大版本。</a>

Watch this Scientific Journal Video about Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy at JoVE.com

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

我们开发的电子显微镜，"闪光和冻结，"一种新的技术，使膜动力学与毫秒的时间分辨率的可视化。该技术结合了高压冷冻神经元的光遗传学刺激。这里，我们证明的程序和详细描述了协议。

闪存和冻结:一种新的技术来捕获膜动力学与电子显微镜

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

flash-freeze-novel-technique-to-capture-membrane-dynamics-with

Universidad San Sebastian

Research

JoVE Journal

Biology

13.9K 次观看.  Johns Hopkins School of Medicine. 我们开发的电子显微镜，"闪光和冻结，"一种新的技术，使膜动力学与毫秒的时间分辨率的可视化。该技术结合了高压冷冻神经元的光遗传学刺激。这里，我们证明的程序和详细描述了协议。 

视频: Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

方法可视化和用透射电子显微镜分析膜蛋白的相互作用

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

科研

生物化学

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, direct visualization of individual viral complexes in their host cellular environment with cryoET is challenging2, due to the infrequent and dynamic nature of viral entry, particularly in the case of HIV-1. While time-lapse live-cell imaging has yielded a great deal of information about many aspects of the life cycle of HIV-13-7, the resolution afforded by live-cell microscopy is limited (~ 200 nm). Our work was aimed at developing a correlation method that permits direct visualization of early events of HIV-1 infection by combining live-cell fluorescent light microscopy, cryo-fluorescent microscopy, and cryoET. In this manner, live-cell and cryo-fluorescent signals can be used to accurately guide the sampling in cryoET. Furthermore, structural information obtained from cryoET can be complemented with the dynamic functional data gained through live-cell imaging of fluorescent labeled target.
In this video article, we provide detailed methods and protocols for structural investigation of HIV-1 and host-cell interactions using 3D correlative high-speed live-cell imaging and high-resolution cryoET structural analysis. HeLa cells infected with HIV-1 particles were characterized first by confocal live-cell microscopy, and the region containing the same viral particle was then analyzed by cryo-electron tomography for 3D structural details. The correlation between two sets of imaging data, optical imaging and electron imaging, was achieved using a home-built cryo-fluorescence light microscopy stage. The approach detailed here will be valuable, not only for study of virus-host cell interactions, but also for broader applications in cell biology, such as cell signaling, membrane receptor trafficking, and many other dynamic cellular processes.

We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, ...

生物工程

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

从快速荧光成像在分子扩散法对活细胞的膜在商业显微镜

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular communication. EVs consist of three populations of vesicles, namely microvesicles (MVs), exosomes, and apoptotic bodies. The limiting membrane of EVs is crucially involved in the interactions with the recipient cells, which could lead to the transfer of biologically active molecules to the recipient cells and, consequently, affect their behavior. The freeze-fracture electron microscopy technique is used to study the internal organization of biological membranes. Here, we present a protocol for MV isolation from cultured cancerous urothelial cells and the freeze-fracture of MVs in the steps of rapid freezing, fracturing, making and cleaning the replicas, and analyzing them with transmission electron microscopy. The results show that the protocol for isolation yields a homogenous population of EVs, which correspond to the shape and size of MVs. Intramembrane particles are found mainly in the protoplasmic face of the limiting membrane. Hence, freeze-fracture is the technique of choice to characterize the MVs' diameter, shape, and distribution of membrane proteins. The presented protocol is applicable to other populations of EVs.

We present a protocol for the isolation and freeze-fracturing of extracellular vesicles (EVs) originating from cancerous urothelial cells. The freeze-fracture technique revealed the EVs' diameter and shape and-as a unique feature-the internal organization of the EV membranes. These are of immense importance in understanding how EVs interact with the recipient membranes.

用于细胞外囊泡分析的冻裂-断裂电子显微镜

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular ...

癌症研究

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

同步干涉反射和全内反射荧光显微镜，用于动态微管和相关蛋白质的成像

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

荧光寿命成像显微镜观察细胞内圈套的贩运

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

免疫与感染

Fundamental Technical Elements of Freeze-fracture/Freeze-etch in Biological Electron Microscopy

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated to generate a carbon/platinum &#8220;cast&#8221; intended for examination by transmission electron microscopy. Specimens are subjected to ultrarapid freezing rates, often in the presence of cryoprotective agents to limit ice crystal formation, with subsequent fracturing of the specimen at liquid nitrogen cooled temperatures under high vacuum. The resultant fractured surface is replicated and stabilized by evaporation of carbon and platinum from an angle that confers surface three-dimensional detail to the cast. This technique has proved particularly enlightening for the investigation of cell membranes and their specializations and has contributed considerably to the understanding of cellular form to related cell function. In this report, we survey the instrument requirements and technical protocol for performing freeze-fracture, the associated nomenclature and characteristics of fracture planes, variations on the conventional procedure, and criteria for interpretation of freeze-fracture images. This technique has been widely used for ultrastructural investigation in many areas of cell biology and holds promise as an emerging imaging technique for molecular, nanotechnology, and materials science studies.

Basic techniques and refinements of freeze-fracture processing of biological specimens and nanomaterials for examination by transmission electron microscopy are described. This technique is a preferred method for revealing ultrastructural features and specializations of biological membranes and for obtaining ultrastructural level dimensional and spatial data in materials sciences and nanotechnology products.

冷冻断裂/冷冻蚀刻的生物电子显微技术的基本要素

Freeze-fracture/freeze-etch describes a process whereby specimens, typically biological or nanomaterial in nature, are frozen, fractured, and replicated ...

生物学

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

通过低温扫描电子显微镜研究光合膜内冻结，割断叶片组织的超分子组织

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

Plunge Freezing: A Tool for the Ultrastructural and Immunolocalization Studies of Suspension Cells in Transmission Electron Microscopy

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize macromolecular complexes at very high resolution. However, to get as close as possible to the native state, perfect sample preservation is required. Conventional electron microscopy (EM) fixation with aldehydes, for instance, does not provide good ultrastructural preservation. The slow penetration of fixatives induces cell reorganization and loss of various cell components. Therefore, conventional EM fixation does not allow for an instantaneous stabilization and preservation of structures and antigenicity. The best choice for examining intracellular events is to use cryofixation followed by the freeze-substitution fixation method that keeps cells in their native state. High-pressure freezing/freeze-substitution, which preserves the integrity of cellular ultrastructure, is the most commonly used method, but requires expensive equipment. Here, an easy-to-use and low-cost freeze fixation method followed by freeze-substitution for suspension cell cultures is presented.

This manuscript describes an easy-to-use and low-cost cryofixation method for visualizing suspension cells by transmission electron microscopy.

暴跌冻结:在透射电子显微镜的超微结构和免疫组化研究工具悬架细胞

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

透射电镜的小型化样品制备方法

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...