我们要感谢郭 Chiuan 和 Bharatkumar 博士的宝贵意见, 改进了手稿。

在这些研究中进行的所有实验都是按照《食品和药物管理局 (FDA) 研究机构机构动物保育和使用委员会批准的议定书》进行的。
1. 小鼠胚胎成纤维细胞的分离培养 (MEFs)
<ol><li>通过标准协议7,8,9, 从 PP2A-B56γ鼠株和野生型窝中分离出小鼠胚成纤维细胞 (MEFs)。</li>
	<li>ExpandMEFs 3 通道, 冻结和存储, 直到实验需要8。</li>
</ol>2. 在2井腔盖玻璃中培养 Mefs 的活体成像技术
<ol><li>准备500毫升的 MEF 生长培养基, 其中含有 Dulbecco 修饰的鹰培养基 (DMEM/F12), 10% 胎牛血清 (血清), 1 X 青霉素/链霉素抗生素, l-谷氨酰胺 (200 毫米) 和1X 非必需氨基酸 (NEAA) 在500毫升的培养基瓶中。</li>
	<li>在37摄氏度前预热水浴中, 从野生型和 PP2A-B56γ小鼠处解冻 MEFs 的冷冻瓶。</li>
	<li>转移解冻 MEFs 到15毫升管和慢慢地添加滴状20毫升 DMEM/F12 介质到15毫升管使用10毫升吸管。</li>
	<li>转移 MEFs 与20毫升的 DMEM/F12 生长培养基在 T75 烧瓶, 并扩大它们, 直到细胞70% 汇合。融合估计使用的是4x 或10x 放大的逆显微镜。</li>
	<li>使用附着在罩内真空系统上的玻璃牧场吸管吸入生长培养基, 加入3毫升0.25% 胰蛋白酶/EDTA, 在37摄氏度处孵育5分钟. 添加3毫升的 DMEM/F12 生长培养基以停止反应。</li>
	<li>在室温下将细胞以低速 (300 x g) 的速度离心成5分钟。小心移除上清, 并在水浴中在37摄氏度前预热1毫升的 DMEM/F12 生长培养基中重新悬浮细胞颗粒。</li>
	<li>使用台盼蓝排除方法或其他适当的单元计数方法8枚举单元格。</li>
	<li>种子大约 2万 MEFs/井在2井腔盖玻璃。添加200µL DMEM/F12 生长培养基, 通过在37摄氏度和 5% CO2上过夜, 使细胞通过孵化来附加。</li>
</ol>3. 同步
<ol><li>同步 MEFs 在 G0/G1 阶段通过孵化细胞在 DMEM/F12 生长培养基中含有0.1% 的24小时, 以获得最大数量的细胞在 G0/G1 阶段。 注: 虽然血清饥饿被用作一种首选方法, 但各种其他方法都可以根据需要被逮捕的细胞的有丝分裂阶段10来使用。</li>
</ol>4. 标签
<ol><li>在亚砜中制备1.5 毫克/毫升 nocodazole 的溶液。Nocodazole 通过抑制微管形成10, 在 G2/M 阶段逮捕细胞。</li>
	<li>三天后同步, 添加200µL 生长培养基 (10% FCS), 200 ng/毫升 nocodazole 和 CL-HB (组蛋白 2 b-GFP BacMam 2.0)。</li>
	<li>计算每个单元格 (PPC) 中的 CL-HB 晶粒子, 以如下所示添加到单元格中: <img alt="Equation 1" src="/files/ftp_upload/57389/57389eq1.jpg"> 如果细胞数是标记时的估计总细胞数, PPC 是每个细胞的粒子数, 1 x 108是试剂每毫升的粒子数。例如, 将2万个单元格标记为30的 PPC <img alt="Equation 2" src="/files/ftp_upload/57389/57389eq2.jpg"> 注意: CL-HB 在10和 50 PPC 之间的大部分细胞类型工作良好。然而, 30 PPC 为本研究提供了最好的效果。</li>
	<li>孵化 MEFs 为18小时在37°c 和 5% CO2。 注: 在目前的实验中, 在 nocodazole 细胞周期骤停后, 有丝分裂细胞中脱离纺锤体组装检查点的染色体可视化发生18小时。</li>
</ol>5. 滞后染色体的可视化
<ol><li>用旋转圆盘共聚焦显微镜系统 MEFs 小鼠胚胎成纤维细胞 (), 配备一个环境室和一个油浸泡, 63x 物镜。<ol>
<li>使用 488 nm 和发射波长 450 nm 的激发波长进行 GFP 通道图像采集。</li>
		<li>采用旋转圆盘共焦显微镜系统, 具有机动扫描阶段、Z 压电插入、顶部孵化和光漂后直接荧光恢复等功能。</li>
	</ol>
</li>
	<li>在成像前的前一天, 将环境室的电源打开, 并在一夜之间将整个房间预热37摄氏度。</li>
	<li>打开显微镜架、照相机、旋转圆盘单元、光源、氩激光、计算机和机动舞台的电源。</li>
	<li>让系统预热3分钟;启动氩激光通过打开点火键。将氩激光器的切换开关从 "待机" 切换到 "激光运行"。</li>
	<li>启动数据采集和处理软件。</li>
	<li>为舞台顶部孵化器启动 co2控制器, 并将 co2的浓度设置为5%。这必须在成像开始前完成。</li>
	<li>从孵化器中取出腔盖玻璃, 并将其放置在舞台上进行可视化。通过油浸泡63x 物镜 (NA1.4) 可视化细胞。</li>
	<li>通过眼镜查看, 聚焦图像, 识别核包络击穿 (NEBD) 阶段的细胞。</li>
	<li>启动适当的激光 (488 nm 氩激光可视化 Histone2B-GFP)。</li>
	<li>打开获取控制窗口并设置 GFP 通道的曝光时间。标识处于 NEBD 中的单元格。<ol>
<li>手动确定目标单元格的顶部和底部焦点平面, 然后输入 xyz 光学切片设置。</li>
		<li>观察20分钟;如果单元格不进行单元格划分, 请在20分钟后停止图像获取, 然后移动到 NEBD 中的下一个单元格。</li>
	</ol>
</li>
	<li>在 PP2A-B56γ细胞有丝分裂过程中, 每细胞需要大约1小时来成像滞后染色体。要获取影片的数据, 请每隔3分钟拍照一次。 注: 野生型细胞在 nocodazole 治疗的过程中, 在 NEBD 的情况下不会取得进展。</li>
	<li>以齐文件格式保存图像以进行进一步分析。</li>
</ol>6. 图像处理与分析
注意: 使用任何可用的软件 (如 Axiovision 版本4.8.2 或 Imaris 版本 8.2) 执行三维图像处理和分析。为这项研究使用了 ImageJ 软件。
<ol><li>打开图像序列。如果图像已处于堆栈格式, 请继续执行下一步。如果不是, 将所有相关的图像合并到一个堆栈中, 使用 "图像 > 堆栈 > 图像堆叠" 在菜单栏中。</li>
	<li>根据需要对亮度/对比度和级别执行任何调整。</li>
	<li>要向影片中添加时间戳:<ol>
<li>在菜单栏中转到 "图像 > 堆栈 > 标签..."。</li>
		<li>选择相应的格式、起始时间值和每个图像之间的时间间隔。</li>
		<li>检查 "预览" 框并调整位置和格式设置。按 "确定" 以应用时间戳。</li>
	</ol>
</li>
	<li>要向影片中添加缩放栏:<ol>
<li>在菜单栏中的 "分析 > 设置比例" 下设置图像缩放。</li>
		<li>在 "距离像素" 字段中, 输入距离已知的多个像素, 并在 "已知距离" 字段中输入距离。在µm 中设置距离的正确长度单位,例如. 按 "确定" 将比例应用于堆栈。</li>
		<li>转到 "分析 > 工具 > 缩放栏..." 以添加缩放条。将条形图的大小设置为 "µm 中的宽度", 并根据需要调整其余的格式选项。选中 "标签所有切片" 框, 将缩放条添加到整个堆栈中。</li>
	</ol>
</li>
	<li>通过单击图像窗口左下边缘的三角形播放按钮预览影片。在菜单栏中使用 "图像 > 堆栈 > 动画 > 动画选项" 调整帧速率。</li>
	<li>通过选择 "文件" > "保存为 > AVI..." 来导出影片文件, 并选择帧速率和压缩。</li>
	<li>按 "确定" 选择保存位置和文件名。按 "保存" 保存影片文件。</li>
</ol>

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作者声明他们没有竞争的财政利益。

细胞周期控制检查点, 确保准确的染色体隔离防止体检和细胞转换1,2,3。在本研究中, 我们发现 PP2A-B56γ的失活导致了一个薄弱的主轴装配检查点。活体细胞成像允许我们观察染色体错误隔离在有丝分裂期间 PP2A-B56γ MEFs 治疗 nocodazole 9。
该协议利用 BacMam 2.0 技术生成的 Histone2B-GFP 标签来标记活细胞成像的染色体。标记的关键步骤之一是计算每个细胞 (PPC) 的适量粒子, 以达到足够的标签。通过滴定发现, 10-50 PPC 效果最好。这种试剂可能对细胞有毒;因此, 有必要滴定 PPC 用于实现标签和毒性的标记细胞之间的平衡。此外, 这是一个瞬变转染, 可以检测到只有5天, 因此, 它可能不适合的研究, 将需要更长的观察期。
传统的蛋白质或 DNA 标记可能需要复杂的协议, 但在这里我们使用的试剂, 只需要一夜孵化与期望细胞。此外, 使用旋转椎间盘共焦显微镜进行活细胞成像, 与共焦激光扫描显微镜相比, 在活细胞中速度更快, 减少了毒性和漂白效果11。该协议允许简单, 方便的方法来标记有丝分裂细胞和可视化染色体分离的其他技术, 如鱼类, 免疫组化和流式细胞术。该协议还允许在整个有丝分裂过程中捕获滞后的染色体图像, 为使用传统的免疫细胞化学、鱼类或流式术后获得的信息提供额外的资料, 从而使其能够捕获表型, 如作为滞后的染色体在空间和时间。该协议允许在多种细胞系、主细胞、干细胞、神经元和永生化 T 细胞 ( 12,13,14) 中快速、方便地标记染色体。
目前正在开发各种类型的细胞治疗产品, 希望治疗各种疾病情况15。这些细胞疗法的发展面临的挑战之一是由于可能存在遗传不稳定和肿瘤发生而引起的安全问题。该协议描述了一种方便简便的方法来标记染色体的各种类型的细胞。在今后的研究中, 可以利用该方法对细胞治疗产品的遗传不稳定性和质量进行检测和检测。

在下面的协议中, 我们描述了一个方便的方法来可视化的染色体分离和有丝分裂进展在小鼠胚胎成纤维细胞使用组蛋白 2 b-GFP, BacMam 2.0 标记和活细胞成像。
细胞周期控制检查点监视染色体隔离, 并在维护细胞的基因完整性方面发挥重要作用1,2,3。分离的染色体的积累可能导致体检, 这是大多数实体肿瘤的标志4。因此, 对滞后染色体的检测可以作为一种研究染色体不稳定性的方法。
荧光标记的蛋白质可用于可视化活染色体分离和染色体滞后, 但 mCherry 标记或 H2B-GFP 标记蛋白的产生需要大量的基因传递和分子生物学知识5。在这里, 我们描述使用 CellLight Histone2B-GFP BacMam 2.0 试剂, 此后称为 CL-HB 摄政, 为了简单。这种试剂可以立即使用, 从而消除了对矢量质量和完整性的关注。此外, 该试剂不需要使用潜在的有害治疗或脂质和染料加载化学品。不同于传统的荧光标签, CL-HB 摄政污点独立功能 (i. e, 膜电位)。CL-HB 摄政可以简单地添加到细胞和孵化过夜的蛋白质表达。CL-HB 不复制哺乳动物细胞, 可用于生物安全水平 (BSL) 1 实验室设置。此外, 这种瞬变转染可以检测后, 夜间孵化长达5天, 足够的时间进行最动态的细胞分析。
另外, 可以通过流式细胞术、免疫组化或荧光原位杂交 (鱼) 6等多种技术研究染色体异常。流式细胞术可用于研究体检, 可根据细胞周期中 DNA 含量和细胞相的相位进行测量。虽然流式细胞术可以用来测量体检, 但它并没有提供染色体错误隔离的信息。鱼类和免疫组化技术使用荧光探针绑定到 DNA 或染色体。虽然这些技术提供了一个细胞种群状态的快照, 但它们不允许活细胞成像, 从而错过了任何信息, 通过时空可视化胞质分裂在特定的细胞后一段时间。
本协议用于研究 nocodazole 处理的小鼠胚胎成纤维细胞 (MEFs) 离体 PP2A-B56γ的滞后染色体或染色体错隔离。除上述应用外, 本协议还提供了一种简单的工具, 用于标记和可视化各种细胞类型的染色体分离, 可用于研究肿瘤细胞的细胞周期调节或染色体不稳定性。此外, 它还可用于研究各种药物治疗引起的染色体不稳定性, 或研究基因敲出导致染色体错误隔离的效果。

<table><tbody><tr><td>DMEM/F12</td><td>Gibco</td><td>11320082</td><td></td></tr><tr><td>L-Glutamine</td><td>Gibco</td><td>25030081</td><td></td></tr><tr><td>MEM Non-Essential amino acids solution (100X)</td><td>Gibco</td><td>11140050</td><td></td></tr><tr><td>Penicillin-Streptomycin (10,000 U/mL)</td><td>Gibco</td><td>15140122</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Phosphate Buffered saline</td><td>Gibco</td><td>14190144</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Gibco</td><td>25300054</td><td></td></tr><tr><td>Fetal bovine serum</td><td>Atlanta Biologicals</td><td>S11150</td><td></td></tr><tr><td>DMSO</td><td>EMD Millipore</td><td>MX 1458-6</td><td></td></tr><tr><td>Cryogenic vial storage boxes</td><td>Fisherbrand</td><td>10-500-28</td><td></td></tr><tr><td>Cryogenic vials</td><td>Corning/costar</td><td>431416</td><td></td></tr><tr><td>T75 flasks</td><td>Cellstar</td><td>658175</td><td></td></tr><tr><td>2 well chambered cover glass</td><td>Nunc</td><td>155380PK</td><td></td></tr><tr><td>Cellometer Vision Cell Profiler</td><td>Nexcelom Bioscience LLC</td><td>Cellometer Vision Trio</td><td></td></tr><tr><td>Nocodazole</td><td>Sigma</td><td>M1404</td><td></td></tr><tr><td>CellLight Histone 2B-GFP, BacMam 2.0</td><td>Thermo Fisher Scientific Inc.</td><td>C10594</td><td></td></tr><tr><td>Zeiss Cell Observer Spinning Disk Confocal Microscope system</td><td>Carl Zeiss Microscopy</td><td>Zeiss Cell Observer SD</td><td></td></tr><tr><td>Water Bath</td><td>Thermoscientific</td><td>280 series</td><td></td></tr><tr><td>Incubator</td><td>Sanyo commercial solutions</td><td>MCO-18AIC (UV)</td><td></td></tr><tr><td>Class II Biological safety cabinet</td><td>The Baker Company</td><td>SterilGard</td><td></td></tr><tr><td>Axiovision software</td><td>Zeiss</td><td>Ver.4.8.2</td><td></td></tr><tr><td>ImageJ software</td><td>National Institute of Health</td><td>Ver. 1.51r</td><td></td></tr></tbody></table>

live cell imaging of chromosome segregation during mitosis

染色体在有丝分裂细胞分裂过程中必须可靠地和均匀地隔离成子细胞。染色体隔离的保真度由包括主轴装配检查点 (SAC) 在内的多个机制控制。该囊是一个复杂的反馈系统的一部分, 负责防止细胞的进展, 通过有丝分裂, 除非所有染色体动粒连接到纺锤管。染色体滞后和异常染色体分离是功能障碍细胞周期控制检查点的指示器, 可用于测量细胞的基因组稳定性。解除对囊的放松会导致正常细胞通过染色体分离过程中的错误积累而转化为恶性细胞。该囊的实施和动粒复合体的形成由激酶和磷酸酶 (如蛋白磷酸酶 2A (PP2A)) 之间的相互作用紧密调节。该协议描述了小鼠胚胎成纤维细胞中的滞后染色体的活细胞成像, 这些老鼠的 PP2A-B56γ调节亚基被挖空。这种方法克服了其他细胞周期控制成像技术的缺点, 如流式细胞术或免疫细胞化学, 只提供一个胞质分裂的状态的快照, 而不是一个动态的时空可视化的染色体有丝分裂期间。

MEFs 从野生型和 PP2A-B56γ小鼠种子在2井室盖玻璃, 并允许附加。在2天, MEFs 的同步使用0.1% 的血清24小时。在3天, MEFs 在 200 ng/毫升 nocodazole 和 30 PPC CL regentwere 的介质中, 在 37°c 和 5% CO2上孵化18小时。在4天, 这些细胞是使用旋转圆盘共聚焦显微镜系统 (图 1) 进行成像的。活细胞成像被用来可视化的命运, 个别细胞, 因为他们从 NEBD 的进展, 通过有丝分裂 (图 2)。Nocodazole 在 G2M 阶段通过干扰微管的形成来逮捕野生型细胞。据观察, 后 nocodazole 治疗野生型细胞被逮捕在中期上部面板 (图 2)。我们观察到, 缺乏 PP2A-B56γ削弱了细胞周期控制, 这使得 PP2A-B56γ MEFs 能够克服 nocodazole 9诱导的细胞周期骤停。此外, 在 62% (13 出 21) 的有丝分裂 PP2A-B56γ-MEFs 治疗与 nocodazole (在图 2下面板中的黄色箭头显示的异常染色体隔离)。五的 PP2A-B56γ MEFs 在 NEBD 和3被捕, 经过有丝分裂, 没有明显的缺陷。相比之下, 所有21种被成像的野生类型细胞都没有进行有丝分裂, 被发现被中期 (上部面板图 2) 逮捕。
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57389/57389fig1.jpg"> 图 1.活细胞成像的基本流程图.野生型和 PP2A-B56γ小鼠的 MEFs 在一夜间2井腔盖玻璃中播种。2天, 细胞被同步使用0.1% 血清24小时 3, 细胞被治疗与 200 ng/毫升 nocodazole 在生长媒体和 30 PPC 组蛋白 2 b GFP, BacMam 2.0 试剂 18 h. 天4这些细胞使用旋转圆盘共聚焦显微镜系统进行成像。<a href="//cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57389/57389fig1large.jpg" target="_blank">请单击此处查看此图的较大版本.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57389/57389fig2.jpg"> 图 2。PP2A-B56γ-MEFs 中检测到染色体异常.nocodazole 治疗野生型和 PP2A-B56γ MEFs 的有丝分裂 (0-51 分钟) 不同时间点的时间失效显微图像实时采集。野生型细胞被发现在中期 (上板) 被逮捕, 而 PP2A-B56γ-MEFs 能够通过有丝分裂 (下板)。染色体滞后或错误隔离可以看到在 PP2A-B56γ-MEFs 显示的黄色箭头在下部面板。缩放条 = 5 µm.<a href="//cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57389/57389fig2large.jpg" target="_blank">请单击此处查看此图的较大版本.</a>

Watch this Scientific Journal Video about Live Cell Imaging of Chromosome Segregation During Mitosis at JoVE.com

Live Cell Imaging of Chromosome Segregation During Mitosis

该协议描述了一种简便、方便的方法, 用 Histone2B-GFP BacMam 2.0 标签和旋转椎间盘共焦显微系统对有丝分裂细胞中的活染色体进行标记和可视化。

有丝分裂过程中染色体分离的活体细胞成像

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

live-cell-imaging-of-chromosome-segregation-during-mitosis

Nanyang Technological University

Research

Education

JoVE Journal

JoVE Core

Cell Biology

Developmental Biology

Unit 1

Cell Division

SCIENTISTS IN ACTION

9.2K 次观看.  US Food and Drug Administration. 该协议描述了一种简便、方便的方法, 用 Histone2B-GFP BacMam 2.0 标签和旋转椎间盘共焦显微系统对有丝分裂细胞中的活染色体进行标记和可视化。

视频: Live Cell Imaging of Chromosome Segregation During Mitosis

Long-term Live-cell Imaging to Assess Cell Fate in Response to Paclitaxel

Live-cell imaging is a powerful technique that can be used to directly visualize biological phenomena in single cells over extended periods of time. Over the past decade, new and innovative technologies have greatly enhanced the practicality of live-cell imaging. Cells can now be kept in focus and continuously imaged over several days while maintained under 37 &#176;C and 5% CO2 cell culture conditions. Moreover, multiple fields of view representing different experimental conditions can be acquired simultaneously, thus providing high-throughput experimental data. Live-cell imaging provides a significant advantage over fixed-cell imaging by allowing for the direct visualization and temporal quantitation of dynamic cellular events. Live-cell imaging can also identify variation in the behavior of single cells that would otherwise have been missed using population-based assays. Here, we describe live-cell imaging protocols to assess cell fate decisions following treatment with the anti-mitotic drug paclitaxel. We demonstrate methods to visualize whether mitotically arrested cells die directly from mitosis or slip back into interphase. We also describe how the fluorescent ubiquitination-based cell cycle indicator (FUCCI) system can be used to assess the fraction of interphase cells born from mitotic slippage that are capable of re-entering the cell cycle. Finally, we describe a live-cell imaging method to identify nuclear envelope rupture events.

Live-cell imaging provides a wealth of information on single cells or whole populations that is unattainable by fixed cell imaging alone. Here, live-cell imaging protocols to assess cell fate decisions following treatment with the anti-mitotic drug paclitaxel are described.

长期活细胞成像对紫杉醇反应细胞命运的评估

Live-cell imaging is a powerful technique that can be used to directly visualize biological phenomena in single cells over extended periods of time. Over ...

科研

癌症研究

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

使用果蝇S2细胞进行细胞分裂的活成像

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

遗传学

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

有丝分裂中的不同发育阶段小鼠胚胎皮层实时成像

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

神经科学

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

有丝分裂的siRNA转染后的时间推移成像

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

生物学

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

龙虎斗动粒蛋白的动态行为与染色体运动，在生活学习有丝分裂关卡<em&gt;果蝇</em&gt;合胞胚胎

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Combining Mitotic Cell Synchronization and High Resolution Confocal Microscopy to Study the Role of Multifunctional Cell Cycle Proteins During Mitosis

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The synchronization of cell populations at specific stages of the cell cycle has been found to be very useful in such experimental endeavors. Synchronization of cells by treatment with chemicals that are relatively less toxic can be advantageous over the use of pharmacological inhibitory drugs for the study of consequent cell cycle events and to obtain specific enrichment of selected mitotic stages. Here, we describe the protocol for synchronizing human cells at different stages of the cell cycle, including both in S phase and M phase with a double thymidine block and release procedure for studying the functionality of mitotic proteins in chromosome alignment and segregation. This protocol has been extremely useful for studying the mitotic roles of multifunctional proteins which possess established interphase functions. In our case, the mitotic role of Cdt1, a protein critical for replication origin licensing in G1 phase, can be studied effectively only when G2/M-specific Cdt1 can be depleted. We describe the detailed protocol for depletion of G2/M-specific Cdt1 using double thymidine synchronization. We also explain the protocol of cell fixation, and live cell imaging using high resolution confocal microscopy after thymidine release. The method is also useful for analyzing the function of mitotic proteins under both physiological and perturbed conditions such as for Hec1, a component of the Ndc80 complex, as it enables one to obtain large sample sizes of mitotic cells for fixed and live cell analysis as we show here.&#160;&#160;

We present a protocol for double thymidine synchronization of HeLa cells followed by analysis using high resolution confocal microscopy. This method is key to obtaining large number of cells that proceed synchronously from S phase to mitosis, enabling studies on mitotic roles of multifunctional proteins which also possess interphase functions.

有丝分裂细胞同步与高分辨共聚焦显微镜联合应用研究细胞周期蛋白在有丝分裂过程中的作用

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The ...

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

荧光活细胞显微镜对微孔裂变酵母核动力学的检测

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...

Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, misunderstood, or misinterpreted by the fixed-cell analysis. While the fixed cell imaging and analysis is robust and sufficient to observe cellular steady-state, it can be limited in defining a temporal order of events at the cellular level and is ill-equipped to assess the transient nature of dynamic processes including mitotic progression. In contrast, live cell imaging is an eloquent tool that can be used to observe cellular processes at the single-cell level over time and has the capacity to capture the dynamics of processes that would otherwise be poorly represented in fixed cell imaging. Here we describe an approach to generate cells carrying fluorescently labeled markers of chromatin and microtubules and their use in live cell imaging approaches to monitor metaphase chromosome alignment and mitotic exit. We describe imaging-based techniques to assess the dynamics of spindle formation and mitotic progression, including the identification of cells at various stages in mitosis, identification&#160;and tracking of mitotic defects, and analysis of spindle dynamics and mitotic cell fate following the treatment with mitotic inhibitors.

Here we present a protocol to assess the dynamics of spindle formation and mitotic progression. Our application of time-lapse imaging enables the user to identify cells at various stages of mitosis, track and identify mitotic defects, and analyze spindle dynamics and mitotic cell fate upon exposure to anti-mitotic drugs.

活细胞成像，以评估在斜接主轴扰动后元相定时和细胞命运的动态

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

亚细胞结构的超分辨率活细胞成像

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Time-lapse fluorescence microscopy has revolutionized the understanding of meiotic cell-cycle events by providing temporal and spatial data that is often not seen by imaging fixed cells. Budding yeast has proved to be an important model organism to study meiotic chromosome segregation because many meiotic genes are highly conserved. Time-lapse microscopy of meiosis in budding yeast allows the monitoring of different meiotic mutants to show how the mutation disrupts meiotic processes. However, many proteins function at multiple points in meiosis. The use of loss-of-function or meiotic null mutants can therefore disrupt an early process, blocking or disturbing the later process and making it difficult to determine the phenotypes associated with each individual role. To circumvent this challenge, this protocol describes how the proteins can be conditionally depleted from the nucleus at specific stages of meiosis while monitoring meiotic events using time-lapse microscopy. Specifically, this protocol describes how the cells are synchronized in prophase I, how the anchor away technique is used to deplete proteins from the nucleus at specific meiotic stages, and how time-lapse imaging is used to monitor meiotic chromosome segregation. As an example of the usefulness of the technique, the kinetochore protein Ctf19 was depleted from the nucleus at different time points during meiosis, and the number of chromatin masses was analyzed at the end of meiosis II. Overall, this protocol can be adapted to deplete different nuclear proteins from the nucleus while monitoring the meiotic divisions.

Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Time-lapse microscopy is a valuable tool for studying meiosis in budding yeast. This protocol describes a method that combines cell-cycle synchronization, time-lapse microscopy, and conditional depletion of a target protein to demonstrate how to study the function of a specific protein during meiotic chromosome segregation.

使用延时显微镜和蛋白质的阶段特异性核耗竭来研究酿酒酵母的减数分裂

Time-lapse fluorescence microscopy has revolutionized the understanding of meiotic cell-cycle events by providing temporal and spatial data that is often ...