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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Freeze Substitution and Low Temperature Embedding System&#160; for Light and Electron Microscopy -AFS II</td>
			<td>Leica&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>High Pressure Freezer - EM-ICE</td>
			<td>Leica&#160;</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&#160;</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>CaCl2</td>
			<td>Sigma</td>
			<td>21115-250ML</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</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</td>
			<td>Thermo scientific</td>
			<td>10569-044</td>
			<td>Should be warmed at 37&#176;C before use</td>
		</tr>
		<tr>
			<td>FBS</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</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 at 37&#176;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&#160; (B-27)&#160;</td>
			<td>Fisher</td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr>
			<td>Hanks 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&#160;</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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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

Nanyang Technological University

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

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

研究膜蛋白的构象动力学原位网站的荧光标记

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

科研

生物学

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

从基因组DNA选择性捕获的5  - 羟甲基

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Capture Compound Mass Spectrometry - A Powerful Tool to Identify Novel c-di-GMP Effector Proteins

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis (diguanylate cyclases) and degradation (phosphodiesterases) of the second messenger c-di-GMP. In contrast, little information is available regarding the molecular mechanisms and cellular components through which this signaling molecule regulates a diverse range of cellular processes. Most of the known effector proteins belong to the PilZ family or are degenerated diguanylate cyclases or phosphodiesterases that have given up on catalysis and have adopted effector function. Thus, to better define the cellular c-di-GMP network in a wide range of bacteria experimental methods are required to identify and validate novel effectors for which reliable in silico predictions fail.
We have recently developed a novel Capture Compound Mass Spectrometry (CCMS) based technology as a powerful tool to biochemically identify and characterize c-di-GMP binding proteins. This technique has previously been reported to be applicable to a wide range of organisms1. Here we give a detailed description of the protocol that we utilize to probe such signaling components. As an example, we use Pseudomonas aeruginosa, an opportunistic pathogen in which c-di-GMP plays a critical role in virulence and biofilm control. CCMS identified 74% (38/51) of the known or predicted components of the c-di-GMP network. This study explains the CCMS procedure in detail, and establishes it as a powerful and versatile tool to identify novel components involved in small molecule signaling.

The ubiquitous second messenger c-di-GMP controls growth and behavior of many bacteria. We have developed a novel Capture Compound Mass Spectrometry based technology to biochemically identify and characterize c-di-GMP binding proteins in virtually any bacterial species.

捕获复合质谱 - 一个强大的工具来确定新型C-二GMP效应蛋白

Considerable progress has been made during the last decade towards the identification and characterization of enzymes involved in the synthesis ...

Correlative Confocal and 3D Electron Microscopy of a Specific Sensory Cell

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused throughout tissues made of diverse cell types, such as enteroendocrine cells of the intestinal epithelium. These gastrointestinal sensors of food and bacteria have been difficult to study because they are dispersed among other epithelial cells at a ratio of 1:1,000. Recently, transgenic reporter mice have been generated to identify enteroendocrine cells by means of fluorescence. One of those is the peptide YY-GFP mouse. Using this mouse, we developed a method to correlate confocal and serial block-face scanning electron microscopy. We named the method cocem3D and applied it to identify a specific enteroendocrine cell in tissue and unveil the cell&#8217;s ultrastructure in 3D. The resolution of cocem3D is sufficient to identify organelles as small as secretory vesicles and to distinguish cell membranes for volume rendering. Cocem3D can be easily adapted to study the 3D ultrastructure of other specific cell types in their native tissue.

Here, we introduce a method, cocem3D, to unveil the ultrastructure of a specific cell in its native tissue by bridging confocal and serial block-face scanning electron microscopy.

一个具体的感觉细胞的共聚焦相关和3D电子显微镜

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

微灌技术在调查微血管渗透性的大鼠肠系膜条例

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

具有可变角度荧光显微镜植物细胞表面动力学的实时成像

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

扫描浸软组织的电子显微镜可视化的细胞外基质

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

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 ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

高压冷冻、微波辅助对比度增强和最小树脂嵌入在体积成像中的生物样品制备

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

超结构保存细胞内膜蛋白定位电子显微镜分析的CryoAPEX方法

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...