作者感谢瑞典研究理事会，斯德哥尔摩郡议会和KI资金的支持。在卡罗林斯卡研究所/ SciLifeLab蛋白质科学研究平台基金（http://PSF.ki.se）进行MSP的表达和纯化。作者也想感谢帕西Purhonen博士和马蒂尔达·舍贝里博士分享他们的技术专长，并为他们及时的援助。

1.纳米圆盘的制备<ol><li>表达和膜支架蛋白8的纯化，35 <ol><li>表达在大肠杆菌 BL21的His-标记MSP1E3D1（DE3）中T1R pRARE2应变在烧瓶中。准备与在37℃下补充有50微克/毫升卡那霉素的LB培养基的50mL过夜起始培养。稀释过夜起始培养在2L补充50微克/毫升卡那霉素了不起的肉汤培养基。 </li><li>生长的细胞在37℃，直至在600nm（OD 600）的光密度在18℃达到约3.诱导用0.5mM异丙基β-D-1-硫代半乳糖苷（IPTG）为3小时的蛋白的表达。 </li><li>制备裂解缓冲液（100mM的Tris-HCl，pH值8.0; 100毫摩尔NaCl; 10％甘油）。临用前加1毫米TCEP。 </li><li>在18℃诱导3小时后，收获，离心所述细胞10分钟，在4500 xg离心和4℃。弃去上清液，称重收获的细胞，并在裂解缓冲液以每克细胞的2裂解毫升缓冲液中的比例重新悬浮它们。 </li><li>裂解的细胞用脉冲超声处理（重复率:4秒开，4秒OFF）为3分钟，在80％的幅度。 </li><li>离心20分钟的裂解物在49000 xg离心和4℃。舍弃这个离心沉淀。 </li><li>注入上清液成5毫升镍+螯合柱连接到自动化液体色谱系统优选保持在4℃。 </li><li>洗脱步骤（步骤1.1.9）之前，洗涤用缓冲液1列 - 3的下方，以除去除了His-标记的MSP的所有蛋白质。监测280nm处跟随纯化过程中的蛋白质含量;在280处的UV记录应该每次洗涤后回到基线。 <ol><li>的40mM的Tris-HCl，300mM NaCl的10 mM咪唑，和1％的Triton，pH值8.0:用洗涤缓冲液1洗涤。 </li><li> 40毫米的Tris-HCl，300 MM:用洗涤液洗2的NaCl，20mM的咪唑，和50mM胆酸钠，pH值8.0。 </li><li>的40mM的Tris-HCl，300mM NaCl的，​​和50mM咪唑，pH 8.0的:用洗涤缓冲液3洗涤。 </li></ol></li><li>用40mM的Tris-HCl，300mM的NaCl和500mM咪唑，pH 8.0的洗脱MSP。 </li><li>改变缓冲器到MSP标准缓冲液（20mM的Tris-HCl，pH值7.5; 100毫摩尔NaCl;和0.5mM EDTA）中通过凝胶过滤8，35;每批产量应该在7毫克L -1。 </li></ol></li><li>磷脂原液的制备<ol><li>添加1-棕榈酰-2-油酰- SN溶于氯仿-glycero -3-磷酸胆碱（POPC）的305微升以25毫克/毫升到一个玻璃烧杯中的浓度，并通过用氮气缓流吹扫它蒸发氯仿。干燥在真空干燥器脂质过夜。注:执行该步骤以从脂质，留下脂质的薄膜在玻璃烧杯底部除去溶剂。</li><li>悬浮在通过涡旋管，直到溶液变得透明含有100mM胆酸钠的MSP标准缓冲器200微升脂质饼;这将提供具有50毫米的POPC浓度的溶液。 注意:一般来说，脂质洗涤剂此时的摩尔比应为1:2（脂:清洁剂）8。此脂质洗涤剂混合物可以储存在-80℃近2个月。 </li></ol></li><li>用于洗涤剂除去疏水珠粒制备<ol><li>代替5-克珠（干重），在50毫升的管中。 </li><li>用30ml 100％的甲醇洗珠，然后用40mL超纯水。 </li><li>与MSP毫升10标准缓冲液洗珠。最后，在4℃下用15ml的MSP标准缓冲的存储珠。 </li></ol></li><li>纳米圆盘重建<ol><li>免除MSP1E3D1（0.124毫米）的190微升到离心管中，加入的磷脂原液（50毫米毫米P 61.5微升OPC），以相同的管中。孵育在湿冰上重构混合物1小时。注意:此等于1的摩尔比:（:POPC MSP1E3D1）130。 </li><li>以0.5克每毫升珠粒重构混合物的浓度添加疏水珠以引发自组装过程。孵育在旋转恒温箱中在4℃下16小时。 </li><li>温育后，离心在13,000rpm XG 10分钟，4℃以除去沉淀物和聚集体。弃沉淀，并保存上清液。 </li><li>平衡的大小排阻层析柱（安装一个自动液相色谱系统上）与MSP标准缓冲液，直到在280nm处的吸光度是稳定的。注入上清液到柱并收集峰级分。 </li><li>测量的纳米圆盘的浓度在280nm处的峰的级分。使用MSP1E3D1（ε=29910厘米-1 M -1）的摩尔消光系数为纳米圆盘浓度的计算。注意:每纳米圆盘脂质的数量可以通过放射性标记的脂质和磷酸分析4的组合或单独磷酸盐分析来测定。 </li></ol></li></ol> 2.制备Monotopic蛋白5-脂氧合酶35 <ol><li>准备过夜起始培养。补充50毫升，100微克/毫升氨苄青霉素的LB培养基。接种用含有5-脂氧化酶基因（ALOX5）的质粒的大肠杆菌 BL21（DE3）中的介质。稀释过夜起始培养在含有42毫摩尔的 Na 2 HPO 4，24毫KH 2 PO 4，9毫摩尔NaCl，19毫摩尔氯化铵 ，1mM的MgSO 4上，0.1毫氯化钙 ，0.2％D-葡萄糖，0.1表达培养基％，5微米的FeSO 4，和100微克/ mL氨苄青霉素。生长细胞直到OD 600为〜0.5，在25℃。在20℃35诱导蛋白表达用0.2mM的IPTG 16小时。 </li><li>收获离心在7000 xg离心和4℃10分钟并弃去上清液。称重在含有裂解缓冲液收获细胞，重悬收集的细胞在管的底部的沉淀含有蛋白酶抑制剂和0.5（100毫摩尔Tris-HCL，pH8.0中;和1mM TCEP 100mM NaCl的; 10％甘油）中毫克/毫升35溶菌酶以2mL每克细胞的裂解缓冲液的比例。 </li><li>裂解在80％振幅的细胞通过超声处理5×15秒。离心10分钟在7000 XG，4℃取出细胞碎片。执行硫酸铵沉淀到30 - 60％的饱和溶液中的35，以沉淀蛋白质。离心16,000 xg离心15分钟，4℃。注:5LO粒料可以快速冷冻并储存在-80℃下长达6个月。 </li><li>悬浮用20ml裂解缓冲液和离心分离的沉淀物为以40,000 xg离心15分钟，4℃。 </li><li>孵育上清<html>在4℃下进行的ATP琼脂糖柱30分钟。用含有0.5M NaCl的裂解缓冲液中的一列体积洗柱一次。洗脱用20mM ATP的5LO在含有10μM的FeSO 4和20微克/毫升过氧化氢酶的裂解缓冲液。执行凝胶过滤层析以除去ATP的35。 注:5LO是不稳定的，应在纯化后立即使用。否则，推荐在硫酸铵沉淀的步骤（见步骤2.1.3后音符）停止。 </li></ol> 3.纳米圆盘蛋白复合物的制备<ol><li>制备100μL的复合物的总体积。拌0.8μMND和0.8μM的5LO与1毫米的Ca 2+存在于MSP标准缓冲液（20mM的Tris-HCl，pH值7.5; 100毫摩尔NaCl; 0.5mM EDTA的;和1.5mM的氯化钙2）和孵育10分钟在冰上。注:样品可以储存在4℃下35至一个月。 </li></ol>样品的ove_title"&gt; 4。分析<ol><li>凝胶电泳<ol><li>非变性电泳<ol><li>混合使用装载缓冲液的5微升（50毫BisTris，6当量盐酸，50 mM氯化钠，10％（重量/体积）甘油，和0.001％丽春红，pH值7.2）49样品15微升并加载在4 - 16％双Tris凝胶电泳执行。 </li><li>填充光阴极缓冲阴极罐（50毫双的Tris; 50mM的曲辛，pH值6.8;以及0.002％考马斯G-250），并与运行缓冲液（50mM二的Tris和50mM麦黄酮，PH值6.8）在阳极罐。在150 V.使用恒定电压开始分离</li><li>停止电泳分离时考马斯前沿到达凝胶的末端。 </li><li>染色用标准考马斯蓝协议的凝胶。 </li></ol></li><li>变性电泳<ol><li>混合样品40微升用含SDS上样缓冲液10微升（0.05％（重量/体积）溴酚蓝;的0.2M的Tris-HCl，pH值6.8; 20％（体积/体积）甘油; 10％（重量/体积）SDS中和10mM 2-巯基乙醇），并加载在4 - 20％的Tris甘氨酸凝胶进行电泳。 </li><li>用运行缓冲液填充在阴极和阳极罐（25毫摩尔Tris盐酸，pH值6.8; 200毫甘氨酸;及0.1％（重量/体积）SDS）。在150 V.使用恒定电压开始分离</li><li>停止电泳分离时装载染料前沿到达凝胶的末端。 </li><li>染色用标准考马斯蓝协议的凝胶。 </li></ol></li></ol></li><li>的NAPT溶液的制备<ol><li>溶解1g盐磷钨酸钠在50毫升水通过搅拌在室温下，得到2％（重量/体积）的酸性溶液。 </li><li>将pH调节至7.4，用1M的NaOH。使用0.22微米的注射器过滤器除去颗粒。储存在室温下或4℃的溶液中。 </li></ol></li><li>样品的TEM分析的制备<ol><li>辉光放电碳包覆铜网格（400目）为20秒，在30 mA到呈现样品的吸附前亲水的网格。放置在网格上的样品（0.8μM相对于所述纳米圆盘）的3.5微升并孵育30秒。注:施加的体积可以变化微升2.5和5之间，这取决于样品浓度。对于纳米圆盘合适的浓度范围为0.5 - 1微米。 </li><li>用滤纸吸干多余的解决方案。 </li><li>立即染色用2％NAPT的30秒的下降电网。吸去多余的溶液，并离开网格风干。 </li><li>评估通过TEM网格。与120-200千电子伏加速电压显微镜足以估计堆积的程度。注:在本协议中，使用了校准的透射电子显微镜，配备了一枚200千电子伏场发射枪。 </li><li>记录TEM图像。对于呈现多头堆栈的图像，不进一步处理;图像显示的纳米圆盘的复合物，与外在蛋白可能包含几短栈，但大多数颗粒是在复杂的。记录多个图像和过程中，这些根据标准方法来获得类的平均值和低分辨率，复杂的三维模型。 注:对于负染色数据的收集，网格被至少在三个不同的日子进行。每一天，涂敷负染色前新鲜样品孵育（如在步骤3.1）中进行。 </li></ol></li></ol>

The authors have nothing to disclose.

该方法可以被分为三个部分:空纳米圆盘的重组，制备蛋白质 - 纳米圆盘复合物，以及用于这些配合物的TEM的阴性染色。每个部分将分别关于该技术，关键步骤，和有用的改进的局限性来解决。 空纳米圆盘重建。关键步骤和限制在生产和使用纳米圆盘。 为空的纳米圆盘的制备中，有必要优化MSP与脂质的比例。对于最常见的MSP和脂质（使用胆作为洗涤剂），?...

在医学研究中，许多注意力集中在膜蛋白，无论是内在的或外在的，参与多种脂质的相互作用。与脂相互作用的蛋白质的工作包括任一选择的替代给脂质，如清洁剂，amphipols 1，或小蛋白质2，或发现的膜替代，保持该蛋白的可溶性和活性。硫辛酸膜替代品包括脂质体和纳米圆盘（ND）3,4。 纳米圆盘是通过改造蛋白的一部分，载脂蛋白A-1，高密度脂蛋白（HDL）在血液天然存在的开发近天然膜的平台。载脂蛋白A-1是243残留长的短的两亲性α螺旋链，并有一个免费的脂溶性构象。 在体外 ，当在脂质的存在下，蛋白质的ApoA-1的两个副本自发重新排列以环绕HYDR脂质双层补丁5 ophobic酰基链部。的ApoA-1的工程化的版本通常被称为膜支架蛋白（MSP），以及越来越多的是可商购的质粒或纯化的蛋白质。在6较长或较短的7膜支架蛋白重复或α螺旋的缺失载脂蛋白A-1的结果。这又使得能够形成大约6毫微米光盘7至17纳米的直径8英寸有不同类型的用于纳米圆盘3,9的应用程序。最常用的应用是提供一种完整的膜蛋白8的稳定的近天然膜的环境，预先3,9审查。 A-探索较少使用是为研究提供一个纳米膜表面外周膜蛋白10，11，12，13，14，15，16，17。下面的协议的部分1可视化制备的磷脂和膜支架蛋白组成的纳米圆盘的过程。 样品制备是在大多数方法的瓶颈。具体方法-样品可以添加特定的信息，但它们也使结果的比较困难。因此，它是简单的，当样品是多峰，并且可以在多种不同的方法可直接使用。与使用的纳米圆盘的一个优点是纳米圆盘的相比，脂质体（ 例如，样品可以直接用于两个TEM和非变性凝胶电泳，如在本协议）的小尺寸。 18。然而，嵌入脂质体膜中的monotopic膜蛋白没有高分辨率三维结构还没有公开发表，据我们所知。金纳米粒子或抗体可用于显现结合的蛋白质用TEM 19的脂质体或囊泡。尽管这些探针是非常具体的，它们可能是由光帷的膜结合位点或通过掩模的与柔性部件感兴趣的区域与膜结合蛋白干扰。金标记或抗体 - 复合的蛋白质也许可以在一个凝胶上分析，但是这将增加实验的成本。 虽然脂质体是一个很好的平台，我们不能肯定，流行ulation具有每脂质体蛋白质的特定的比率，可以通过使用纳米圆盘20的探索的特征。在脂质体，辅因子和底物可被截留在水溶性内部。这是膜水溶性物质将共享相同的命运两种膜的模拟物的。尽管如此，由于该双层区域是在纳米圆盘较小，需要物质的量较小以饱和纳米圆盘膜。 通过原子结构的测定蛋白质的认识功能的研究很多领域是必不可少。蛋白质结构测定方法包括X射线21;核磁共振（NMR）22，23;和透射电子显微镜（TEM）24在低温下，cryoEM。通过cryoEM分辨率近来有了很大的提高，主要是由于直接使用电子脱离的tectors 25,26。的大分子在近天然状态下拍摄的薄，玻璃体冰27。然而，由于生物分子的低对比度，它们变得很难在100的尺寸范围，以检测 - 200 kDa的。为适当大小的样品，数据收集可以制成并且可以应用于单粒子重建的方法，得到结构28。 然而，蛋白质结构的通过TEM测定是一个多步骤的过程。它通常与样品单分散性的使用负染色TEM 29像phosphotungsten（PT）30或铀31重金属盐的评估开始。的负染大分子的低分辨率模型的重构通常制成，并且可以得到在分子结构29的重要信息。在平行下，使用cryoEM可以开始收集数据。应谨慎评估负染色TEM数据时，为了避免假象形成的误解作出。一个特定的人工制品是在PT污点上的磷脂的效果和脂质体32，从而导致长棒类似，从侧面33观察硬币堆叠的形成。这种"ROULEAU"或"堆栈"（以下简称记为"堆"）观察早期对HDL 34，后来还为纳米圆盘中35。 膜的堆叠和重塑可能有很多原因发生。例如，它可以通过共因素，如铜，通过TEM成像钼酸铵染色36所示来诱导。膜脂质的脂质体的部分含有一个亚氨基二乙酸头组用EDTA模仿金属络合，从而在加入铜离子后堆叠脂质体<s补课="外部参照"&gt; 36。堆叠也可能是由于在或在脂质双层通过的蛋白质的蛋白质-蛋白质相互作用（使用的染色未提及）37。观察到初期由PT磷脂的叠层形成;然而，后来的工作重点是拆除或取消此神器形成38。 在这里，我们建议采取NAPT诱导纳米圆盘堆叠的膜结合蛋白的透射电镜研究优势的方法。总之，蛋白的纳米圆盘结合会阻止纳米圆盘从堆叠。虽然为堆叠的原因尚不清楚，有人提议39存在的磷脂和PT的磷酰基之间的静电相互作用，使光盘粘到彼此（ 图1A）。我们的协议背后的假设是，当一个蛋白结合至纳米圆盘，大部分磷脂表面的不availa<html>竹叶提取用于与PT中的相互作用由于由蛋白质空间位阻。这将防止堆的形成（ 图1B）。两个结论可以得出。首先，预防堆叠意味着感兴趣的蛋白质已结合到膜上。其次，蛋白质- ND复合体可以用标准的单粒子加工方法24，40被处理，以获得复合物的粗形态。此外，通过像非变性凝胶电泳或动态光散射方法的分析可以被执行。 为了证明这一假设，我们使用的膜结合蛋白5-脂氧合酶（5LO），其涉及许多炎性疾病41,42。这个78-kDa蛋白需要钙离子结合其膜43。虽然这种膜协会已被广泛研究使用脂质体S ="外部参照"&gt; 44，45,46和膜部分47，这些不能被用于TEM分析和结构确定。 纳米圆盘的制备通过混合脂重新悬浮于胆洗涤剂钠MSP开始。在冰上孵育1小时后，该洗涤剂慢慢地从使用吸附剂树脂重构混合物中除去。这种材料常常由聚苯乙烯形成小珠。他们是相对疏水并有较脂类结合48洗涤剂有强烈的偏好。除去疏水珠和使用离心进行澄清后，将纳米圆盘通过尺寸排阻色谱（SEC）纯化。纯化的纳米圆盘与以等摩尔比率（或几个比率滴定）一个monotopic膜蛋白（和可能的辅因子）混合，并留于rEACT（15分钟）。通过TEM分析是通过将样品的微升-量到辉光放电，碳包被的网格，然后通过与NAPT进行负染色进行。从当等分试样施加到TEM格栅上相同的样品可通过非变性或SDS-PAGE凝胶电泳，以及由各种活动的测量可用于分析，没有大的变化。

<table><tbody><tr><td>Transmission electron microscope: JEOL2100F</td><td>JEOL</td><td></td><td></td></tr><tr><td>CCD camera</td><td>Tiez Video and Imaging Processing System GmbH, Germany</td><td></td><td></td></tr><tr><td>Glow discharger</td><td>Baltec</td><td></td><td></td></tr><tr><td>TEM grid: 400 mesh</td><td>TAAB</td><td>GM016/C</td><td></td></tr><tr><td>Size exclusion chromatography: Agilent SEC-5</td><td>Agilent Technologies</td><td>5190-2526</td><td></td></tr><tr><td>Superdex 200 HR 10/300</td><td>GE Healthcare Life Sciences</td><td>17-5172-01</td><td></td></tr><tr><td>Plasmid: MSP1E3D1</td><td>Addgene</td><td>20066</td><td></td></tr><tr><td>Bacteria: BL21DE3</td><td>NEB</td><td>C2527H</td><td></td></tr><tr><td>Bacteria: BL21 (DE3) T1R pRARE2</td><td>Protein Science Facility, KI, Solna</td><td></td><td></td></tr><tr><td>Purification Matrix: ATP agarose</td><td>Sigma Aldrich</td><td>A2767</td><td></td></tr><tr><td>Purification Matrix: HisTrap HP-5 mL</td><td>GE Healthcare Life Sciences</td><td>17-5247-01</td><td></td></tr><tr><td>Lipid: POPC</td><td>Avanti polar lipids</td><td>850457C</td><td>25 mg/mL in chloroform</td></tr><tr><td>Hydrophobic beads: Bio-Beads, SM-2 Resin</td><td>Bio-Rad</td><td>1523920</td><td></td></tr><tr><td>13 mm syringe filter: 0.2 &amp;mu;m</td><td>Pall life sciences</td><td>PN 4554T</td><td></td></tr><tr><td>Stain: Sodium phosphotungstate tribasic hydrate</td><td>Sigma Aldrich</td><td>31648</td><td></td></tr><tr><td>2-mercaptoethanol</td><td>Sigma Aldrich</td><td>M3148-250ML</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate (SDS)</td><td>Bio-Rad</td><td>161-0301</td><td></td></tr><tr><td>Protease inhibitor cocktail</td><td>Sigma Aldrich</td><td>4693132001</td><td></td></tr><tr><td>TCEP</td><td>Sigma Aldrich</td><td>646547</td><td></td></tr><tr><td>Detergent: Sodium cholate hydrate</td><td>Sigma Aldrich</td><td>C6445-10G</td><td></td></tr><tr><td>Sodium Cholate</td><td></td><td></td><td>500 mM Sodium cholate. Resuspend in miliQ water and store at -20 &amp;deg;C.</td></tr><tr><td>Lipid Stock</td><td></td><td></td><td>50 mM POPC, 100 mM sodium cholate, 20 mM Tris-HCl pH 7.5, 100 mM NaCl. &lt;bt /&gt;Store at 4 &amp;deg;C for a week; or&lt;br/&gt; Store -80 &amp;deg;C for a month, after purging the solution with nitrogen.</td></tr><tr><td>MSP standard buffer</td><td></td><td></td><td>20 mM Tris-HCl, pH 7.5, 100 mM NaCl, 0.5 mM EDTA. &lt;br /&gt;Store at 4 &amp;deg;C.</td></tr><tr><td>Non-Denaturaing Electrophoresis Anode Buffer</td><td>Thermo Fisher Scientific</td><td>BN2001</td><td>50 mM Bis-Tris, 50 mM Tricine, pH 6.8</td></tr><tr><td>Non-Denaturaing Electrophoresis Cathode Buffer</td><td>Thermo Fisher Scientific</td><td>BN2002</td><td>50 mM Bis-Tris, 50 mM Tricine, pH 6.8, 0.002% Coomassie G-250</td></tr><tr><td>Non-Denaturaing Electrophoresis 4x Sample loading Buffer</td><td>Thermo Fisher Scientific</td><td>BN2003</td><td>50 mM Bis-Tris, pH 7.2, 6 N HCl, 50 mM NaCl, 10% (w/v) glycerol, 0.001% Ponceau S</td></tr><tr><td>Denaturaing Electrophoresis Running Buffer</td><td></td><td></td><td>In-house recipe: 25 mM Tris-HCl, pH 6.8, 200 mM Glycine, 0.1% (w/v) SDS</td></tr><tr><td>Denaturaing Electrophoresis 5x Sample loading Buffer</td><td></td><td></td><td>In-house recipe: 0.05% (w/v) Bromophenolblue, 0.2 M Tris-HCl, pH 6.8, 20% (v/v) glycerol, 10% (w/v) SDS, 10 mM 2-mercaptoethanol</td></tr><tr><td>Terrific broth</td><td></td><td></td><td>Tryptone - 12.0 g, Yeast Extract - 24.0 g, 100 mL 0.17 M KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; and 0.72 M K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, Glycerol - 4 mL.&lt;br /&gt;Tryptone, yeast extract and glycerol were prepared to 900 mL and autoclaved seperately. KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; and K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; were prepared and autoclaved separately. Both were mixed before using the medium.</td></tr></tbody></table>

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.

我们提出的方法，取决于制备纳米圆盘提供monotopic膜蛋白结合膜表面。因为有嵌入到纳米圆盘脂质双层无跨膜蛋白，所述纳米圆盘在这里表示为"空纳米圆盘"（ 图2A）。这些具有256 kDa的计算分子量两个MSP1E3D1骨架蛋白和大约260 POPC 8的分子的组合物。使用这种蛋白质:用于重建脂质比，一个主要的峰（ 图2A）的凝胶过滤中洗脱。约9?...

Watch this Scientific Journal Video about Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy at JoVE.com

通过透射电子显微镜可视化和分析膜相互作用蛋白的方法生物化学|视频

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

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 ...

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13.1K 次观看.  Karolinska Institutet. 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.

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

Visualizing Proteins and Macromolecular Complexes by Negative Stain EM: from Grid Preparation to Image Acquisition

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural biology 1. In recent years, this method has achieved great success in studying structures of proteins and macromolecular complexes 2, 3. Compared with electron cryomicroscopy (cryoEM), in which frozen hydrated protein samples are embedded in a thin layer of vitreous ice 4, negative staining is a simpler sample preparation method in which protein samples are embedded in a thin layer of dried heavy metal salt to increase specimen contrast 5. The enhanced contrast of negative stain EM allows examination of relatively small biological samples. In addition to determining three-dimensional (3D) structure of purified proteins or protein complexes 6, this method can be used for much broader purposes. For example, negative stain EM can be easily used to visualize purified protein samples, obtaining information such as homogeneity/heterogeneity of the sample, formation of protein complexes or large assemblies, or simply to evaluate the quality of a protein preparation.
In this video article, we present a complete protocol for using an EM to observe negatively stained protein sample, from preparing carbon coated grids for negative stain EM to acquiring images of negatively stained sample in an electron microscope operated at 120kV accelerating voltage. These protocols have been used in our laboratory routinely and can be easily followed by novice users.

Visualizing protein samples by negative stain electron microscopy (EM) has become a popular structural analysis method. It is useful for quantitative structural analysis, such as calculating a 3D reconstruction of the molecules being studied, and also for qualitative examination of the quality of protein preparations. In this article we present detailed protocols for preparing the EM grids, staining the sample and visualizing the sample in an electron microscope. Novice users can follow these protocols easily and to utilize negative stain EM as a routine assay, in addition to other biochemical assays, for evaluating their protein samples.

负染色的EM可视化蛋白质和大分子复合物：从电网的制备图像采集

Single particle electron microscopy (EM), of both negative stained or frozen hydrated biological samples, has become a versatile tool in structural ...

科研

生物工程

Analyzing Dynamic Protein Complexes Assembled On and Released From Biolayer Interferometry Biosensor Using Mass Spectrometry and Electron Microscopy

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this work, the pre-endosomal anthrax toxin is assembled and transitioned into the endosomal complex. First, the N-terminal domain of a cysteine mutant lethal factor (LFN) is attached to a biolayer interferometry (BLI) biosensor through disulfide coupling in an optimal orientation, allowing protective antigen (PA) prepore to bind (Kd 1 nM). The optimally oriented LFN-PAprepore complex then binds to soluble capillary morphogenic gene-2 (CMG2) cell surface receptor (Kd 170 pM), resulting in a representative anthrax pre-endosomal complex, stable at pH 7.5. This assembled complex is then subjected to acidification (pH 5.0) representative of the late endosome environment to transition the PAprepore into the membrane inserted pore state. This PApore state results in a weakened binding between the CMG2 receptor and the LFN-PApore and a substantial dissociation of CMG2 from the transition pore. The thio-attachment of LFN to the biosensor surface is easily reversed by dithiothreitol. Reduction on the BLI biosensor surface releases the LFN-PAprepore-CMG2 ternary complex or the acid transitioned LFN-PApore complexes into microliter volumes. Released complexes are then visualized and identified using electron microscopy and mass spectrometry. These experiments demonstrate how to monitor the kinetic assembly/disassembly of specific protein complexes using label-free BLI methodologies and evaluate the structure and identity of these BLI assembled complexes by electron microscopy and mass spectrometry, respectively, using easy-to-replicate sequential procedures.

Here we present a protocol to monitor the assembly and disassembly of the anthrax toxin using biolayer interferometry (BLI). Following assembly/disassembly on the biosensor surface, the large protein complexes are released from the surface for visualization and identification of components of the complexes using electron microscopy and mass spectrometry, respectively.

用质谱和电镜分析 Biolayer 干涉传感器上的动态蛋白复合物的组装和释放

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this ...

生物化学

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

线粒体和内质视网膜成像由相关光和体积电子显微镜

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

Native Cell Membrane Nanoparticles System for Membrane Protein-Protein Interaction Analysis

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal transduction; from sensing environmental signals to biological response; from metabolic regulation to developmental control. Accurate structural information of protein-protein interactions is crucial for understanding the molecular mechanisms of membrane protein complexes and for the design of highly specific molecules to modulate these proteins. Many in vivo and in vitro approaches have been developed for the detection and analysis of protein-protein interactions. Among them the structural biology approach is unique in that it can provide direct structural information of protein-protein interactions at the atomic level. However, current membrane protein structural biology is still largely limited to detergent-based methods. The major drawback of detergent-based methods is that they often dissociate or denature membrane protein complexes once their native lipid bilayer environment is removed by detergent molecules. We have been developing a&#160;native cell membrane nanoparticle system for membrane protein structural biology. Here, we demonstrate the use of this system in the analysis of protein-protein interactions on the cell membrane with a case study of the oligomeric state of AcrB.

Presented here is a protocol for the determination of oligomeric state of membrane proteins that utilizes a native cell membrane nanoparticle system in conjunction with electron microscopy.

用于膜蛋白-蛋白质相互作用分析的原生细胞膜纳米粒子系统

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal ...

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

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient association of pore-forming and auxiliary subunits, lateral diffusion, and clustering with other proteins. However, the relationship between lateral diffusion and function is poorly understood. To approach this problem, we describe how lateral mobility and activity of individual channels in supported lipid membranes can be monitored and correlated using total internal reflection fluorescence (TIRF) microscopy. Membranes are fabricated on an ultrathin hydrogel substrate using the droplet interface bilayer (DIB) technique. Compared to other types of model membranes, these membranes have the advantage of being mechanically robust and suitable for highly sensitive analytical techniques. This protocol measures Ca2+ ion flux through single channels by observing the fluorescence emission of a Ca2+-sensitive dye in close proximity to the membrane. In contrast to classical single-molecule tracking approaches, no fluorescent fusion proteins or labels, which can interfere with lateral movement and function in the membrane, are required. Possible changes in ion flux associated with conformational changes of the protein are only due to protein lateral motion in the membrane. Representative results are shown using the mitochondrial protein translocation channel TOM-CC and the bacterial channel OmpF. In contrast to OmpF, the gating of TOM-CC is very sensitive to molecular confinement and the nature of lateral diffusion. Hence, supported droplet-interface bilayers are a powerful tool to characterize the link between lateral diffusion and the function of ion channels.

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

This protocol describes how to use TIRF microscopy to track individual ion channels and determine their activity in supported lipid membranes, thereby defining the interplay between lateral membrane movement and channel function. It describes how to prepare the membranes, record the data, and analyze the results.

使用全内反射荧光（TIRF）显微镜对脂质双层中的横向迁移率和离子通道活性进行单分子成像

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient ...

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 ...

生物学

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as cells, organelles, membrane vesicles, or viruses at molecular detail. To achieve this, the aqueous sample is rapidly vitrified in liquid ethane, which preserves it in a close-to-native, frozen-hydrated state. In the electron microscope, tilt series are recorded at liquid nitrogen temperature, from which 3D tomograms are reconstructed. The signal-to-noise ratio of the tomographic volume is inherently low. Recognizable, recurring features are enhanced by subtomogram averaging, by which individual subvolumes are cut out, aligned and averaged to reduce noise. In this way, 3D maps with a resolution of 2 nm or better can be obtained. A fit of available high-resolution structures to the 3D volume then produces atomic models of protein complexes in their native environment. Here we show how we use electron cryo-tomography to study the in situ organization of large membrane protein complexes in mitochondria. We find that ATP synthases are organized in rows of dimers along highly curved apices of the inner membrane cristae, whereas complex I is randomly distributed in the membrane regions on either side of the rows. By subtomogram averaging we obtained a structure of the mitochondrial ATP synthase dimer within the cristae membrane.

We present a protocol of how to collect and process electron cryo-tomograms of whole mitochondria. The technique provides detailed insights into the structure, function, and organization of large membrane protein complexes in native biological membranes.

ATP合酶二聚体线粒体通过冷冻电子断层扫描可视化

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as ...

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

确定膜蛋白拓扑使用荧光蛋白酶保护（FPP）

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

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 ...