我们感谢埃文斯实验室的成员对这份手稿的深思熟虑的反馈。这项工作得到了杜克怀特黑德学者，杜克科学和技术学者和霍华德休斯医学研究所（HHMI）汉娜格雷奖学金的支持。 图1A 是使用 BioRender.com 制作的。

1. 生物样品的制备
注意：在生物安全柜中使用无菌技术执行以下步骤。用70%乙醇喷洒机柜表面和所有材料。
<ol><li>HeLa细胞培养和转染<ol><li>在Dulbecco的改良鹰培养基（DMEM）中培养30，000个HeLa细胞，含有4.5g / L葡萄糖，补充有10%胎牛血清和1%L-谷氨酰胺溶液（HeLa培养基;参见 材料表）。将细胞铺在含有 2 mL 预热 HeLa 培养基的 35 mm 玻璃底成像皿（参见材料 T 上）。将HeLa培养物保持在5%CO2 和37°C26，27。</li>
		<li>电镀后的第二天，使用光学显微镜检查35毫米成像皿，以确保细胞~50%-60%汇合。汇合后，用空的黄色荧光蛋白（YFP）载体，YFP-ParkinWT或YFP-ParkinT240R 转染HeLa细胞（图1A）。</li>
		<li>每次转染时，在无菌微量离心管中制备两种单独的混合物。通过敲击试管或轻轻上下移液进行混合。<ol><li>在管 1 中，将 200 μL 还原血清培养基（参见 材料表）与 2 μg 质粒 DNA 混合。</li>
			<li>在试管 2 中，将 200 μL 还原血清培养基与 6 μL 转染试剂混合（参见 材料表）28。</li>
		</ol></li>
		<li>将试管在室温（RT）下孵育5分钟。将管2加入管1中，通过上下移液混合，并在室温下孵育20分钟。</li>
		<li>将转染复合物滴加到含有HeLa细胞培养物的现有成像培养皿中，确保在整个培养皿中均匀分布。将成像培养皿置于5%CO2 和37°C培养箱中过夜。 注意：用适当的转染质粒标记每个培养皿。对于每个实验，标记三个YFP-ParkinWT 培养皿（二甲基亚砜[DMSO;见 材料表]，5μM CCCP和20μM CCCP）。用三个YFP-ParkinT240R 培养皿和三个空的YFP载体培养皿重复标签。</li>
	</ol></li>
	<li>CCCP和荧光染料的制备 注意：荧光染料通常对光敏感。使用铝箔或棕色离心管将染料存放在黑暗中，以减少暴露在光线下。<ol><li>通过将 5 mg CCCP 溶解在 4.89 mL DMSO 中，制备 5 mM 的 CCCP 工作储备液（参见 材料表）。通过将 5 mg CCCP 溶解在 1.22 mL DMSO 中，制备 20 mM 的 CCCP 工作储备液。</li>
		<li>在 390 μL DMSO 中稀释 10 μL 1 mM MitoTracker 深红色（参见 材料表）储备溶液，制成 25 μM 工作储备液。</li>
		<li>将 50 μg MitoSOX 红25 （参见 材料表）溶解在 13 μL DMSO 中，以创建 5 mM 工作储备液。</li>
		<li>将 10 mg TMRE24 （参见 材料表）溶解在 19.4 mL DMSO 中，制成 1 mM 储备溶液。通过将 10 μL 1 mM TMRE 加入 990 μL DMSO 中来稀释 TMRE，制成 10 μM 工作储备液。</li>
	</ol></li>
</ol>2. HeLa细胞中荧光探针的CCCP处理和线粒体标记
注意：快速执行以下步骤，以确保HeLa细胞不会长时间离开培养箱。
<ol><li>转染后第二天，向每个实验板中加入 2 μL 的 5 或 20 mM CCCP（终浓度：分别为 5 μM 和 20 μM）。对于对照板，加入 2 μL DMSO。将板返回到5%CO2 和37°C培养箱中1.5小时（图1A）。 注意：CCCP治疗总共2小时。然而，在CCCP治疗的最后30分钟内用荧光探针标记线粒体。</li>
	<li>1.5小时后，从培养箱中取出板并将荧光探针添加到成像培养皿中。将板返回到5%CO2 和37°C培养箱中30分钟（图1A）。<ol><li>TMRE 实验：加入 2 μL 25 μM MitoTracker（终浓度：25 nM）和 2 μL 10 μM TMRE（最终浓度：10 nM）。</li>
		<li>MitoSOX 实验：加入 2 μL 25 μM MitoTracker（终浓度：25 nM）和 1 μL 5 mM MitoSOX（最终浓度：2.5 μM）。</li>
	</ol></li>
	<li>在2小时CCCP处理后，准备HeLa细胞进行成像（图1A，B）。<ol><li>TMRE实验：HeLa细胞立即准备好成像;确保TMRE保留在HeLa细胞培养基中。</li>
		<li>MitoSOX 实验：用 2 mL 预热的 HeLa 培养基洗涤细胞 3 倍，以去除游离的 MitoSOX 染料。加入 2 mL 预热培养基。HeLa细胞现在已准备好成像。 注意：用新鲜的预热HeLa培养基洗涤HeLa细胞，该培养基含有与实验条件相同的DMSO或CCCP浓度;不包括 MitoSOX 或 MitoTracker。</li>
	</ol></li>
</ol>3. 共聚焦显微镜图像采集设置
注意：使用配备63x/1.40数值孔径（NA）油浸物镜和环境室（见材料表）的共聚焦显微镜（见材料表）对HeLa细胞进行成像。
<ol><li>在成像前1小时，通过打开罐阀打开CO2。按开按钮打开显微镜的环境控制器。使用触摸板上的向上和向下箭头将温度调节到 37 °C，将 CO 2 调节到 5%。完成后按设置。 注意：确保环境室的门已关闭，并等待条件稳定下来。</li>
	<li>激光设置<ol><li>打开 白光激光器 （WLL）并将 激光 功率设置为 85%， 将 激发控制 设置为 最大功率。单击“ 获取 ”选项卡，选择 “添加激光器”，然后在出现的对话框中，将 WLL 切换为 “开”。单击 激光功率 按钮并输入 85%。单击 励磁控制 按钮，然后从下拉菜单中选择 最大功率 （图2A）。</li>
		<li>TMRE实验： 设置YFP，MitoTracker和TMRE的激发和发射光谱。<ol><li>对于YFP，将激发激光设置为514 nm，将发射光谱窗口设置为524-545 nm。 在“获取”选项卡中，单击“添加新设置”按钮;然后，单击添加激光并将其拖到设置 1 中。双击激发线并在对话框中输入 514 作为波长。双击相应的检测器，输入 524 作为起始波长，输入 545 作为结束波长。</li>
			<li>对于MitoTracker深红，将激发激光设置为641，将发射光谱窗口设置为650-750 nm。在“获取”选项卡中，单击“添加激光”按钮并将其拖到“设置1”中。双击激发线并在对话框中输入 641 作为波长。双击相应的检测器，输入 650 作为开始波长，输入 750 作为结束波长。</li>
			<li>对于TMRE，将激发激光设置为555 nm，将发射光谱窗口设置为557-643 nm。在“获取”选项卡中，单击“添加新设置”按钮;然后，点击 添加激光 按钮并将其拖到设置 2.双击激发线，然后在对话框中输入 555 作为波长。双击相应的检测器，输入 557 作为起始波长，输入 643 作为结束波长。</li>
		</ol></li>
		<li>MitoSOX 实验： 设置YFP，MitoTracker和MitoSOX的激发和发射光谱（图2B）。<ol><li>对于YFP，将激发激光设置为507 nm，将发射光谱窗口设置为517-540 nm。在“获取”选项卡中，单击“添加新设置”按钮;然后，单击 添加激光 按钮并将其拖到设置 1。双击激发线并在对话框中输入 507 作为波长。双击相应的检测器，输入 517 作为起始波长，输入 540 作为结束波长。</li>
			<li>对于 MitoSOX，将 激发 激光设置为 547 nm， 将发射 光谱窗口设置为 564-636 nm。在“ 获取 ”选项卡中，单击“ 添加新设置 ”按钮;然后，点击 添加激光 按钮并将其拖到 设置 2.双击 激发线并在对话框中输入 547 作为波长。双击相应的 检测器 ，输入 564 作为起始波长，输入 636 作为结束波长。</li>
			<li>对于MitoTracker深红，将激发激光设置为641 nm，将发射光谱窗口设置为652-742 nm。 在“获取”选项卡中，单击“添加新设置”按钮;点击 添加激光 按钮并将其拖到设置 3。双击激发线并在对话框中输入 641 作为波长。双击相应的检测器，输入 652 作为起始波长，输入 742 作为结束波长。</li>
		</ol></li>
	</ol></li>
	<li>图像采集设置（图2C）<ol><li>将 格式 设置为 1，024 x 1，024， 将扫描速度 设置为 600 Hz，将 线平均值 设置为 3。<ol><li>选择“ 采集”选项卡，然后单击“ 格式 ”按钮。从下拉菜单中，选择 1，024 x 1，024。单击 速度 按钮，然后从下拉菜单中选择 600 。然后，单击 线平均值 按钮，然后从下拉菜单中选择 3。</li>
			<li>打开 双向扫描 并将 相位 和 变焦系数 分别设置为 22.61 和 1.50。<ol><li>在“ 采集”选项卡中，将“ 双向 ”按钮切换为 “开”。单击 阶段 X 设置并将其设置为 22.61。单击 缩放系数 设置并将其设置为 1.50。</li>
			</ol></li>
		</ol></li>
	</ol></li>
</ol>4. 图像采集
注意：实验者必须根据YFP荧光信号做出视觉判断以选择细胞。避免像素过饱和，因为它们会显着影响荧光强度定量。使用指示像素饱和度的过/欠查找表，以避免获取饱和图像。
<ol><li>单击感兴趣的 设置 ，然后按快速实时以提供荧光图像的 实时预览。</li>
	<li>通过双击相应的检测器来调整增益和强度。在出现的对话框中，使用滑块调整增益。要更改强度，请双击激发线并使用弹出窗口中的向上和向下箭头更改强度。单击“停止”结束预览。<ol><li>TMRE实验：首先对DMSO控制板进行成像。调整TMRE信号的增益和强度（设置2），使线粒体网络强度略低于饱和度; 保持实验的增益和强度恒定。调整丝线虫追踪器和YFP的增益和强度（设置1），使线粒体网络可见但暗淡。</li>
		<li>MitoSOX 实验：首先对 DMSO 控制板进行成像。调整 MitoSOX（设置 2）信号的增益和强度，使荧光可见但暗淡;保持实验的增益和强度恒定。调整YFP（设置1）和MitoTracker（设置3）的增益和强度，使线粒体网络可见但暗淡。 注意：记录TMRE和MitoSOX的增益和强度，因为这些值在整个实验过程中必须保持不变。YFP和MitoTracker的荧光信号在该协议中未量化。因此，可以针对每个图像调整增益和强度。最有效的是具有增益和强度设置，其中细胞 易于看到但仍很暗，以确保细胞不会暴露在导致细胞损伤或光漂白的过度激光强度下。</li>
	</ol></li>
	<li>增益和强度设置完成后，单击 “开始 ”以获取图像。每个实验条件获取 20 个细胞的图像（例如，五个图像，每个图像四个细胞; 图 2D）。</li>
</ol>5. 使用 ImageJ 进行荧光强度定量
注意：图像文件保存为“.lif”，并与 ImageJ29 兼容。与 ImageJ 兼容的文件类型在其网站上指定。如果文件类型不兼容，则可能需要转换文件类型。
<ol><li>创建并保存感兴趣区域 （ROI）。<ol><li>在 ImageJ 中，单击“ 文件”|”新品 |图像。 在弹出的对话框中，单击“ 确定”。</li>
		<li>单击矩形工具按钮并绘制一个 6 微米 x 6 微米的框。通过单击“分析”|”工具 |ROI 管理器，然后等待出现带有 ROI 管理器的对话框（图 3A）。通过单击管理器对话框中的添加将矩形 ROI 添加到管理器。通过选择“更多”来保存投资回报率，然后单击“保存”按钮和“确定”。</li>
	</ol></li>
	<li>测量荧光强度。<ol><li>在图像 J 中，单击 文件 并选择 打开。在对话框中，选择实验的映像文件，然后单击“ 打开”。</li>
		<li>等待 “生物格式导入选项” 窗口出现（图3B）。选择 拆分通道 ，然后单击 打开。 注意：拆分通道功能允许将图像中的每个通道作为单独的窗口打开。通道顺序对应于获取顺序。<ol><li>TMRE 实验： YFP （设置 1）是第一个通道 （c = 0）; MitoTracker （设置 1）是第二个通道 （c = 1）;TMRE（设置 2）是第三个通道 （c = 2）。</li>
			<li>MitoSOX 实验： YFP （设置 1）是第一个通道 （c = 0）; MitoSOX （设置 2）是第二个通道 （c = 1）; MitoTracker （设置3）是第三个通道（c = 2）。</li>
		</ol></li>
		<li>通过选择 “图像 |调整 |亮度 （图3C）。 注：请勿按“ 设置”或 “应用”以确保图像亮度不会永久更改。有时，荧光强度在TMRE或MitoSOX通道中不可见。调整亮度以简化可视化。改变亮度不会影响原始荧光强度值。</li>
		<li>单击 分析 ，然后选择 设置测量。选中 面积 和 平均灰度值 框，然后单击 确定 （图 3D）。 注意：平均灰度值是荧光强度。</li>
		<li>打开 ROI 管理器 ，然后通过单击 “分析”|”工具 |投资回报率经理。在 ROI 管理器中，单击 更多， 然后从显示的列表中单击 打开，然后选择保存的 ROI。</li>
		<li>通过从ROI管理器中选择保存的ROI来测量单个细胞中五个随机区域的荧光强度，并将ROI移动到细胞内的随机位置（图3E）。按 M测量荧光强度。对另外四个不重叠区域重复此步骤。当出现包含面积和平均灰度值的对话框（图 3F）时，将这些值复制并粘贴到电子表格中进行分析。<ol><li>TMRE 实验：选择图像（c = 2）。</li>
			<li>MitoSOX 实验：选择图像 （c = 2）。</li>
		</ol></li>
		<li>获得平均荧光强度值。使用电子表格软件（参见 材料表），平均五个平均灰度值。对每个单元格重复此过程。</li>
		<li>使用统计软件程序分析和比较实验条件下的TMRE或MitoSOX平均灰度值（参见 材料表;图 4 和 图 5）。将数据显示为条形图或小提琴图。</li>
	</ol></li>
</ol>

线粒体膜电位和超氧化物水平的定量

作者没有竞争利益要声明。

此处概述的工作流程可用于使用基于荧光的成像稳定且可重复地量化线粒体膜电位和超氧化物水平30。在设计这些实验时，需要考虑重要的技术限制。用空的YFP载体YFP-ParkinWT或YFP-ParkinT240R转染HeLa细胞。空的YFP载体被用作对照，以确认实验结果是帕金特有的。对于瞬时转染，通过实验确定DNA与转染试剂的质量比为1：3，以产生最高的转染率，其中每个构建体使用2 μ...

线粒体网络是一系列相互连接的细胞器，在能量产生1，先天免疫2，3和细胞信号传导4，5中起着至关重要的作用。线粒体失调与帕金森病（PD）等神经退行性疾病有关6，7。PD是一种进行性神经退行性疾病，影响黑质的多巴胺能神经元，影响全球近1000万人8。PD与线粒体自噬有遗传联系，线粒体自噬是维持细胞稳态所必需的线粒体质量控制途径，选择性地去除受损的线粒体9，10。研究已经确定了多种独立的线粒体自噬途径，包括含有 1 （FUNDC1） 介导的线粒体自噬的 FUN14 结构域、Bcl-2 相互作用蛋白 3 （BNIP3） 促进的线粒体自噬、NIX 依赖性线粒体自噬以及特征明确的 PTEN 诱导激酶 1 （PINK1）/帕金调节线粒体自噬10，11。PINK1（一种假定的激酶）和帕金（一种E3泛素连接酶）与磷酸泛素受损线粒体协同工作，后者驱动自噬体的形成，吞噬受损细胞器并与溶酶体融合以启动降解12，13，14，15，16。帕金的突变与PD相关的表型有关，例如通过多巴胺能神经元丢失引起的神经变性17，18。
在这里，描述了一个协议，其中HeLa细胞，常规使用的源自宫颈癌的永生化细胞，用于研究Parkin在维持线粒体网络健康中的作用。HeLa细胞表达的内源性帕金水平可以忽略不计，因此需要外源性帕金表达19。为了研究Parkin在线粒体网络健康中的作用，用野生型Parkin（ParkinWT），Parkin突变体（ParkinT240R）或空对照载体转染HeLa细胞。ParkinT240R是一种常染色体隐性幼年帕金森综合征突变，影响Parkin E3连接酶活性，显着降低线粒体自噬途径的效率20。HeLa细胞暴露于轻度（5μM）或严重（20μM）浓度的羰基氰间氯苯基腙（CCCP），一种线粒体解偶联剂。用高浓度的CCCP处理通常用于在各种细胞系中诱导帕金介导的线粒体自噬，例如HeLa和COS-7细胞21，22，23。
治疗后，该方案使用两种当前可用的线粒体靶向荧光染料对线粒体网络进行实时成像。四甲基罗丹明、乙酯、高氯酸盐 （TMRE） 是一种阳离子染料，其荧光基于线粒体膜电位24，而 MitoSOX 是一种线粒体超氧化物指示剂，其中荧光强度是超氧化物浓度25 的函数。最后，概述的方案使用基于荧光的定量和简单的工作流程来有效地量化线粒体膜电位和超氧化物水平，而用户偏差的范围最小。尽管该协议旨在研究HeLa细胞中的线粒体功能，但它可以适用于其他细胞系和原代细胞类型，以定量表征线粒体网络健康。

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	<tbody>
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			<td>Chemicals, Peptides, and Recombinant Proteins</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CCCP (carbonyl cyanide m-chlorophenyl hydrazone)&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>C2759</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM (1x) with 4.5 g/L glucose</td>
			<td>Gibco</td>
			<td>11-965-084</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO, Anhydrous</td>
			<td>ThermoFisher Scientific</td>
			<td>D12345</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Hyclone</td>
			<td>SH3007103</td>
			<td></td>
		</tr>
		<tr>
			<td>FuGENE 6 (Tranfection Reagent)</td>
			<td>Promega</td>
			<td>E2691</td>
			<td></td>
		</tr>
		<tr>
			<td>GlutaMAX 100x (L-Glutamine Solution)&#160;</td>
			<td>Gibco&#160;</td>
			<td>35-050-061</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoescht 33342</td>
			<td>ThermoFisher Scientific</td>
			<td>62249</td>
			<td></td>
		</tr>
		<tr>
			<td>MitoSOX&#160; Red&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>M36008</td>
			<td></td>
		</tr>
		<tr>
			<td>MitoTracker Deep Red</td>
			<td>ThermoFisher Scientific</td>
			<td>M7514</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-MEM (Redued Serum media)</td>
			<td>ThermoFisher scientific</td>
			<td>31985070</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetramethylrhodamine, Ethyl Ester, Perchlorate (TMRE)&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>T669</td>
			<td></td>
		</tr>
		<tr>
			<td>Experimental models: Organisms/Strains</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HeLa-M (Homo sapiens)</td>
			<td>A. Peden (Cambridge Institute for Medical Research)</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant DNA</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EYFP Empty Vector</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>YFP-Parkin T240R</td>
			<td>This Paper</td>
			<td>Generated by site-directed mutagenesis from YFP-Parkin</td>
			<td></td>
		</tr>
		<tr>
			<td>YFP-Parkin WT</td>
			<td>Addgene; PMID:19029340</td>
			<td>RRID:Addgene_23955</td>
			<td></td>
		</tr>
		<tr>
			<td>Software and Algorithms</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Adobe Illustrator</td>
			<td>Adobe Inc.</td>
			<td>https://www.adobe.com/products/illustrator</td>
			<td>(Schindelin, 2012)</td>
		</tr>
		<tr>
			<td>Excel (Spreadsheet Software)</td>
			<td>Microsoft Office&#160;</td>
			<td>https://www.microsoft.com/en-us/microsoft-365/excel</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td></td>
			<td>https://imagej.net/software/fiji/</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica Application Suite (LAS X)</td>
			<td>Leica</td>
			<td>https://www.leica-microsystems.com/products/microscope-software/p/leica-las-x-ls/</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsoft Excel</td>
			<td>Microsoft Office</td>
			<td>https://www.microsoft.com/excel</td>
			<td></td>
		</tr>
		<tr>
			<td>Prism9 (Statistical Analysis Software)</td>
			<td>GraphPad Software</td>
			<td>https://www.graphpad.com</td>
			<td></td>
		</tr>
		<tr>
			<td>Other</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Dish, No. 1.5 Coverslip, 20 mm Glass Diameter, Uncoated</td>
			<td>MatTek</td>
			<td>P35G-1.5-20-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Cage Incubator (Environmental Chamber)</td>
			<td>Okolab</td>
			<td>https://www.oko-lab.com/cage-incubator</td>
			<td></td>
		</tr>
		<tr>
			<td>DMiL Inverted Microscope</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>LIGHTNING Deconvolution Software</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>STELLARIS 8 confocal microscope</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

fluorescence-based quantification of mitochondrial membrane potential and superoxide levels using live imaging in hela cells

线粒体是通过ATP合成 控制 能量产生，对代谢稳态至关重要的动态细胞器。为了支持细胞代谢，各种线粒体质量控制机制合作以维持健康的线粒体网络。其中一种途径是线粒体自噬，其中PTEN诱导的激酶1（PINK1）和受损线粒体的Parkin磷酸化泛素化促进自噬体隔离并随后通过溶酶体融合 从 细胞中去除。线粒体自噬对细胞稳态很重要，帕金的突变与帕金森病（PD）有关。由于这些发现，人们非常重视研究线粒体损伤和周转，以了解线粒体质量控制的分子机制和动力学。在这里，活细胞成像用于可视化HeLa细胞的线粒体网络，以量化线粒体解偶联剂羰基氰间氯苯基腙（CCCP）处理后的线粒体膜电位和超氧化物水平。此外，表达抑制帕金依赖性线粒体自噬的帕金PD连锁突变（ParkinT240R）以确定与表达野生型帕金的细胞相比突变如何影响线粒体网络。此处概述的方案描述了一种简单的工作流程，使用基于荧光的方法有效地量化线粒体膜电位和超氧化物水平。

在该协议中，基于荧光的定量用于测量CCCP处理后线粒体网络的膜电位和超氧化物水平（图1）。该工作流程使用了HeLa细胞，这是一种源自宫颈癌的永生化细胞系。HeLa细胞通常用于研究线粒体生物学，并且相对平坦，因此很容易使用显微镜可视化线粒体网络。为了研究Parkin在维持线粒体网络健康中的作用，用空对照载体ParkinWT或ParkinT240R （一种破坏线粒体自噬...

Watch this Scientific Journal Video about Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells at JoVE.com

作者聚焦：在 HeLa 细胞中使用实时成像对线粒体膜电位和超氧化物水平进行基于荧光的定量  

该技术描述了一种有效的工作流程，使用基于荧光的实时成像可视化和定量测量HeLa细胞内的线粒体膜电位和超氧化物水平。

使用HeLa细胞中的实时成像对线粒体膜电位和超氧化物水平进行基于荧光的定量

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, ...

本研究的重点是神经退行性疾病中的线粒体失调。我们相信，这项研究可用于了解疾病发作的潜在原因，这些原因可能导致对抗帕金森病和ALS等神经退行性疾病的治疗方法。STED和SIM等超分辨率显微镜或扩增显微镜等技术提高了准确了解单个细胞器内的蛋白质分布和整个细胞线粒体分布的能力。该技术的结果可以作为研究帕金森病相关突变对线粒体周转影响的起点，并开始了解调节活性氧水平和线粒体膜电位对维持神经元健康的重要性。我们的实验室旨在机械地表征单个线粒体质量控制途径，以了解这些途径之间的相互作用。通过深入了解通路动力学，人们可以了解线粒体如何维持以及线粒体失调如何导致神经退行性疾病的发作。

fluorescence-based-quantification-mitochondrial-membrane-potential

Research

JoVE Journal

Biology

Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells

6.9K 次观看.  Duke University Medical Center. 本研究的重点是神经退行性疾病中的线粒体失调。我们相信，这项研究可用于了解疾病发作的潜在原因，这些原因可能导致对抗帕金森病和ALS等神经退行性疾病的治疗方法。STED和SIM等超分辨率显微镜或扩增显微镜等技术提高了准确了解单个细胞器内的蛋白质分布和整个细胞线粒体分布的能力。该技术的结果可以作为研究帕金森病相关突变对线粒体周转影响的起点，并开...

视频: 作者聚焦：在 HeLa 细胞中使用实时成像对线粒体膜电位和超氧化物水平进行基于荧光的定量  

Monitoring Dynamic Changes In Mitochondrial Calcium Levels During Apoptosis Using A Genetically Encoded Calcium Sensor

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, differentiation, and apoptosis1. During apoptosis, calcium accumulation in mitochondria promotes the release of pro-apoptotic factors from the mitochondria into the cytosol2. It is therefore of interest to directly measure mitochondrial calcium in living cells in situ during apoptosis. High-resolution fluorescent imaging of cells loaded with dual-excitation ratiometric and non-ratiometric synthetic calcium indicator dyes has been proven to be a reliable and versatile tool to study various aspects of intracellular calcium signaling. Measuring cytosolic calcium fluxes using these techniques is relatively straightforward. However, measuring intramitochondrial calcium levels in intact cells using synthetic calcium indicators such as rhod-2 and rhod-FF is more challenging. Synthetic indicators targeted to mitochondria have blunted responses to repetitive increases in mitochondrial calcium, and disrupt mitochondrial morphology3. Additionally, synthetic indicators tend to leak out of mitochondria over several hours which makes them unsuitable for long-term experiments. Thus, genetically encoded calcium indicators based upon green fluorescent protein (GFP)4 or aequorin5 targeted to mitochondria have greatly facilitated measurement of mitochondrial calcium dynamics. Here, we describe a simple method for real-time measurement of mitochondrial calcium fluxes in response to different stimuli. The method is based on fluorescence microscopy of 'ratiometric-pericam' which is selectively targeted to mitochondria. Ratiometric pericam is a calcium indicator based on a fusion of circularly permuted yellow fluorescent protein and calmodulin4. Binding of calcium to ratiometric pericam causes a shift of its excitation peak from 415 nm to 494 nm, while the emission spectrum, which peaks around 515 nm, remains unchanged. Ratiometric pericam binds a single calcium ion with a dissociation constant in vitro of ~1.7 &mu;M4. These properties of ratiometric pericam allow the quantification of rapid and long-term changes in mitochondrial calcium concentration. Furthermore, we describe adaptation of this methodology to a standard wide-field calcium imaging microscope with commonly available filter sets. Using two distinct agonists, the purinergic agonist ATP and apoptosis-inducing drug staurosporine, we demonstrate that this method is appropriate for monitoring changes in mitochondrial calcium concentration with a temporal resolution of seconds to hours. Furthermore, we also demonstrate that ratiometric pericam is also useful for measuring mitochondrial fission/fragmentation during apoptosis. Thus, ratiometric pericam is particularly well suited for continuous long-term measurement of mitochondrial calcium dynamics during apoptosis.

This protocol describes a method for real-time measurement of mitochondrial calcium fluxes by fluorescent imaging. The method takes advantage of a circularly permutated YFP-based dual-excitation ratiometric calcium sensor (ratiometric pericam-mt) selectively expressed in mitochondria.

使用一种基因编码的钙传感器来监测细胞凋亡过程中线粒体钙水平的动态变化

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, ...

科研

生物学

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

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

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

基于荧光测定活细胞中的钙进入商店经营：从培养的肿瘤细胞，骨骼肌纤维

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

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

在活细胞中蛋白质聚集的4D成像

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

活体显微镜成像的亚细胞结构的活体小鼠表达荧光蛋白

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

单线粒体超氧闪烁在完整心脏或共聚焦成像<em&gt;在体内</em

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

线粒体钙和线粒体膜的同时测量通过荧光显微镜活细胞的潜在

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

哺乳动物细胞基态衰竭超分辨成像

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

Improving the Accuracy of Flow Cytometric Assessment of Mitochondrial Membrane Potential in Hematopoietic Stem and Progenitor Cells Through the Inhibition of Efflux Pumps

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic homeostasis have been extensively studied by HSC researchers. Mitochondrial activity levels are reflected in their membrane potentials (&#916;&#936;m), which can be measured by cell-permeant cationic dyes such as TMRM (tetramethylrhodamine, methyl ester). The ability of efflux pumps to extrude these dyes from cells can limit their usefulness, however. The resulting measurement bias is particularly critical when assessing HSCs, as xenobiotic transporters exhibit higher levels of expression and activity in HSCs than in differentiated cells. Here, we describe a protocol utilizing Verapamil, an efflux pump inhibitor, to accurately measure &#916;&#936;m across multiple bone marrow populations. The resulting inhibition of pump activity is shown to increase TMRM intensity in hematopoietic stem and progenitor cells (HSPCs), while leaving it relatively unchanged in mature fractions. This highlights the close attention to dye-efflux activity that is required when &#916;&#936;m-dependent dyes are used, and as written and visualized, this protocol can be used to accurately compare either different populations within the bone marrow, or the same population across different experimental models.

Xenobiotic efflux pumps are highly active in hematopoietic stem and progenitor cells (HSPCs) and cause extrusion of TMRM, a mitochondrial membrane potential fluorescent dye. Here, we present a protocol to accurately measure mitochondrial membrane potential in HSPCs by TMRM in the presence of Verapamil, an efflux pump inhibitor.

通过抑制Efflux泵,提高造血干细胞和造血细胞中线粒体膜电位流细胞测定的准确性

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic ...

Measurement of Mitochondrial Mass and Membrane Potential in Hematopoietic Stem Cells and T-cells by Flow Cytometry

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production of all mature blood cells. In recent years cellular metabolism has emerged as a crucial regulator of HSC function and fate. We have previously demonstrated that modulation of mitochondrial metabolism influences HSC fate. Specifically, by chemically uncoupling the electron transport chain we were able to maintain HSC function in culture conditions that normally induce rapid differentiation. However, limiting HSC numbers often precludes the use of standard assays to measure HSC metabolism and therefore predict their function. Here, we report a simple flow cytometry assay that allows reliable measurement of mitochondrial membrane potential and mitochondrial mass in scarce cells such as HSCs. We discuss the isolation of HSCs from mouse bone marrow and measurement of mitochondrial mass and membrane potential post ex vivo culture. As an example, we show the modulation of these parameters in HSCs via treatment with a metabolic modulator. Moreover, we extend the application of this methodology on human peripheral blood-derived T cells and human tumor infiltrating lymphocytes (TILs), showing dramatic differences in their mitochondrial profiles, possibly reflecting different T cell functionality. We believe this assay can be employed in screenings to identify modulators of mitochondrial metabolism in various cell types in different contexts.

Here we describe a reliable method to measure mitochondrial mass and membrane potential in ex vivo cultured hematopoietic stem cells and T cells.

流细胞测定方法测量造血干细胞和T细胞的线粒体质量及膜电位

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production ...

Isolating and Imaging Live, Intact Pacemaker Regions of Mouse Renal Pelvis by Vibratome Sectioning

The renal pelvis (RP) is a funnel-shaped, smooth muscle structure that facilitates normal urine transport from the kidney to the ureter by regular, propulsive contractions. Regular RP contractions rely on pacemaker activity, which originates from the most proximal region of the RP at the pelvis-kidney junction (PKJ). Due to the difficulty in accessing and preserving intact preparations of the PKJ, most investigations on RP pacemaking have focused on single-cell electrophysiology and Ca2+ imaging experiments. Although important revelations on RP pacemaking have emerged from such work, these experiments have several intrinsic limitations, including the inability to accurately determine cellular identity in mixed suspensions and the lack of in situ imaging of RP pacemaker activity. These factors have resulted in a limited understanding of the mechanisms that underlie normal rhythmic RP contractions. In this paper, a protocol is described to prepare intact segments of mouse PKJ using a vibratome sectioning technique. By combining this approach with mice expressing cell-specific reporters and genetically encoded Ca2+ indicators, investigators may be able to more accurately study the specific cell types and mechanisms responsible for peristaltic RP contractions that are vital for normal urine transport.

The goal of this protocol is to isolate intact pacemaker regions of the mouse renal pelvis using vibratome sectioning. These sections can then be used for in situ Ca2+ imaging to elucidate Ca2+ transient properties of pacemaker cells and other interstitial cells in vibratome slices.

隔离和成像现场， 通过颤音分割老鼠肾佩尔维斯的精致起搏器区域

The renal pelvis (RP) is a funnel-shaped, smooth muscle structure that facilitates normal urine transport from the kidney to the ureter by regular, ...