我们感谢他们仔细阅读这篇稿子哈拉尔胡特尔和海伦娜德克尔。我们也感谢徕卡Microsystems的注册Sidhu，他的技术专长。这项研究是由美国国家科学与工程理事会，加拿大，奖＃327100-06，西蒙弗雷泽大学理学院的支持。

第1部分：用脂质体2000（Invitrogen）的神经元的转设备设置： 培养的大鼠或小鼠海马神经元是根据Kaech和银行家，2006 [4]。典型的转染的细胞密度为25万cells/6-cm菜。所需的试剂和设备，包括50 mM的kynurenic酸，定期台式管支架，台式冷却器（最好保持脂质体转染试剂冷）， 脂质体转染试剂，MEM，离心管，micropipetters和无菌镊子。 程序： <ol><li>对于每一个转标签两个1.5 mL管;含有质粒DNA，含有转染试剂。以下的孵化可以在室温下进行。 <ol><li>结合100μL，纪念在一管1.0微克质粒DNA（不包括任何类型的补充; mem没有需要的pH值平衡的温度）。 </li><li>结合在1.5毫升管100μL的MEM 6.0μL脂质体。 <ol><li>双转染没有调整的需要，即使脂质体：DNA比例将在这种情况下，减少了一半。 ：DNA的脂质体的比例仍然是在制造商的建议。最好是每个质粒测试的最佳表达0.5-1.0微克。 </li><li>为了保持脂质体转染试剂的保质期，它应该被删除从冰箱尽可能短的时间内，台式散热器放置在不使用时。 出冰箱的总时间应保持在最低限度。 </li></ol></li></ol></li><li>为5分钟，在室温下孵育管。 </li><li>在纪念解决方案的DNA转移到脂质管，并用吸管轻轻混匀，然后30分钟，在室温下孵育。 </li><li> 25分钟后，翻转盖玻片前，加入60μL50毫米kynurenic酸，以尽量减少excitoxic损害的培养神经元的菜。 <ol><li>小心翻转盖玻片脚端，用无菌镊子，并安排，使它们不重叠。要小心不要划伤盖玻片的神经元表面涂层或神经胶质涂层表面的菜。 </li></ol></li><li>从6厘米的菜约0.5毫升培养基，并轻轻地用DNA混合管，转移DNA溶液均匀介质表面上的菜淋漓。 </li><li>不要漩涡，但轻轻滑动盘在一个方向来回，然后停止并在垂直方向重复，均匀分布的DNA -脂质体复合物。孵育90分钟，在组织文化的孵化器（37 ° C，5％CO 2）。 </li><li>翻转盖玻片脚面朝下，并安排，使它们不重叠。允许表达，以进行所需的时间，通常是隔夜至48小时。 </li></ol>第2部分：实时成像GFP标记的培养海马神经元的密集核心囊泡设备设置： <ol><li>活细胞成像需要一个装有CCD相机的荧光显微镜，我们使用了Leica DMI 6000B倒置显微镜配备徕卡变量，荧光灯。还需要的是一个温度控制器和客观加热器成像室。我们使用华纳仪器RC - 21BR修改为18毫米的盖玻片，在另外一个平台热水器（货号＃PH2）和温度控制器（货号＃TC324B）。室包含没有开放的端口，在订货时，可包括室修改。客观加热器强烈建议，以帮助保持温度盖玻片，并尽量减少由于温度的波动变化重点。图像采集与使用Metamorph（Molecular Devices公司）的软件，包括“流式”收购模式下拉一个滨松ORCA - ER。注意，我们不使用一个气候室，包围整个显微镜。这除了是昂贵的，而不是必要的，这里所描述的短期成像。 </li><li>其他设备和试剂，包括新鲜配制的实时成像介质（1X汉克斯与Ca 2 +和Mg 2 +，葡萄糖，0.6％和10mm的HEPES），额外的18毫米的玻璃盖玻片，无菌镊子，非无菌镊子，Kimwipes，真空润滑脂，和一个注射器（没有针，或改良大口径针）与申请油脂。 </li></ol>程序： <ol><li>准备鲜活成像介质（1X汉克斯与Ca 2 +和Mg 2 +，葡萄糖，0.6％和10mm的HEPES）。我们通常准备50mls在摄氏4度，在不使用时的时间和存储。最好是准备足够时间不超过一个星期。约5-10毫升，每盘的神经元。 </li><li>启动了显微镜，摄像机，荧光灯，和图像采集软件。还可以打开电源室平台和客观热水器。 </li><li>准备的想象克室。 <ol><li>华纳室包括一个安装的盖玻片聚四氟乙烯插入。为了便于操纵，一个标签的一侧插入“顶”永久性标记。 </li><li>应用的油脂环周围的凹槽孔一侧标有“顶”总商会。避免使用多余的，因为它会进入会议厅，减少观察的区域。 </li><li>使用镊子，附加一个干净的，未使用的18厘米的盖玻片厅一侧的“顶”，适用于温和的压力，在其边缘盖玻片内室的凹沟，以创建一个密封。 </li><li>关闭室和一个类似戒指的油脂，适用于相反的（底部）会议厅一侧的凹槽。 </li><li>广场上5​​0毫升管的盖子，这是一个理想的商会持有的准备室底部。 </li><li>加入750μL的成像介质会议厅。这是一个量，但过多的油脂应防止蔓延的中期和多余的，将有助于防止泡沫细胞覆盖盖玻片应用时被困。 </li><li>维持在组织培养孵化器的准备室，直到需要。长期孵化，〜1小时，应避免成像介质蒸发，改变组成。 </li></ol></li><li>执行最后的商会大会和活细胞成像。 <ol><li>从孵化器的准备室和一碟转盖玻片移动组织培养罩。 </li><li>最终商会大会应快速进行，以确保细胞保持在中期和浸泡在37℃不断。 </li><li>用无菌镊子，小心取出转碟盖玻片，触摸盖玻片边缘Kimwipe绘制了生长介质。 </li><li>将盖玻片神经端上的准备室。触摸盖玻片一侧的油脂和应用的边缘周围的压力，带来不诱捕气泡盖玻片平。如预期超额成像介质将洒出。 </li><li>擦去多余的成像介质，但要小心，不要滑出位置的盖玻片。 </li><li>移动平台加热器预热室，并拧紧到位室。 </li><li>室转移到舞台上。 </li><li>为了减少光漂白和photoxicity，调节灯泡所需的最低数额，发现转染细胞的强度。 </li><li>查找感兴趣的细胞与一个低倍率的油浸物镜（40X）。坚持有计划的搜索模式，以确保最大的盖玻片的覆盖面，以避免收购多余的图像，例如，左上角盖玻片，扫描和工作的权利，这是有益的。 </li><li>一次所需的领域一直位于切换到高放大倍率的石油目标（60 - 100X）。 </li><li>使用“流收购”或相媲美的图像处理软件的功能，记录荧光标记的蛋白质动力学的视频。 </li><li>以一个阶段的形象，为以后的分析和介绍，和任何其他（如可溶性GFP）的陪同图像。 </li><li>进一步探索盖玻片，并记录下视频之前保存的所有图像。 </li><li>通常情况下3-5影片从一个盖玻片被认为是成功的。光毒性和一般细胞的健康，应该牢记，作为运输中受损的细胞减少。可以监测使用相差显微镜观察细胞的健康。通常每盖玻片约30分钟的录音。 </li></ol></li></ol>第3部分：代表性的成果： 活细胞成像观测通常由视频（又称“栈”的图像）显示来看（图1A）领域内的荧光标记的细胞器的运动。伴随着每部影片往往是支持的图像，说明细胞体，蛋白表达和/或细胞的健康水平方面的视野方向。运输的影片，可以定量分析的转变受到kymographs（图1B）。 Kymographs是并列kymograph列在每个时间点代表的扫描线的图像，说明粒子的运动。图2演示了如何在相反的方向移动的两个粒子在kymograph代表。 kymographs进一步分析可以提供关于粒子通量，速度，运行长度，反复，和固定颗粒的信息。 <img src="/files/ftp_upload/1144/fig1.jpg" alt="" /> 图1代表活细胞成像观测（A）选自mCherry的标签组织型纤溶酶原激活物（tPA）交通视频帧。单个粒子移动的顺（绿色箭头）和（二）逆行（红色箭头）的方向指示。Kymographs说明TPA - mCherry在运动轴突。水平线代表在kymographs静止的物体，而对角线代表（正斜率）或向（负斜率）的细胞体，细胞器。长的绿色和红色箭头显示运行相应的粒子在（A）。由于微管解聚剂，噻氨酯哒唑，在运输中的变化也是一个kymograph说明。注意在运动细胞器的对角线的情况下减少。

<ol>
	<li>Kulic, I. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+microtubule+movement+in+bidirectional+organelle+transport.">The role of microtubule movement in bidirectional organelle transport.</a> Proc Natl Acad Sci U S A. 105 (29), 10011-10016 (2008).</li><li>Jacobson, C., Schnapp, B., Banker, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+change+in+the+selective+translocation+of+the+Kinesin-1+motor+domain+marks+the+initial+specification+of+the+axon.">A change in the selective translocation of the Kinesin-1 motor domain marks the initial specification of the axon.</a> Neuron. 49 (6), 797-804 (2006).</li><li>Barkus, R. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+an+Axonal+Kinesin-3+Motor+for+Fast+Anterograde+Vesicle+Transport+that+Facilitates+Retrograde+Transport+of+Neuropeptides.">Identification of an Axonal Kinesin-3 Motor for Fast Anterograde Vesicle Transport that Facilitates Retrograde Transport of Neuropeptides.</a> Mol Biol Cell. 19 (1), 274-283 (2008).</li><li>Kaech, S., Banker, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culturing+hippocampal+neurons.">Culturing hippocampal neurons.</a> Nat Protoc. 1 (5), 2406-2415 (2006).</li></ol>

活细胞成像是一个具有挑战性，但强大的技术，直接观察培养的神经元细胞器运输。困难可能会出现上游与神经元的健康欠佳，由于文化并发症程序。因此，细胞的健康（例子见文献4）应评估转染前后，而在生长介质中使用组织培养，光镜下观察盖玻片。转移过程中的神经元含有盖玻片成像室，可紧张的细胞，因此重要的是要小心操纵的盖玻片。它可以是共同的，健康的细胞没有运输过程中的延...

A correction was made to <a href="http://www.jove.com/details.php?ID=1144">Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons</a>. There was an error in the author's name. The author name was corrected to include a middle initial as follows:

David M. Kwinter, Michael A. Silverman

instead of:

David Kwinter, Michael Silverman.

A correction to author names has been made for article Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons.

Erratum: Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons.

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live imaging of dense-core vesicles in primary cultured hippocampal neurons

动态细胞过程的观测和特征，可以产生对细胞活性，不能从静态图像获得的重要信息。重要的荧光探针，尤其是绿色荧光蛋白（GFP）已经彻底改变了细胞生物学能力标签特定的细胞内的车厢和细胞结构产生。例如，允许实时成像的GFP（和其光谱变种）嵌合体的细胞骨架，细胞器运输，并在众多生物和细胞类型[1-3]膜动态动态分析。虽然实时成像已成为普遍的，这种做法仍然引起了许多技术挑战，特别是在原代培养的神经元，。挑战之一是GFP标记的蛋白质在有丝分裂后神经元的表达，另一种是能够捕捉到的荧光图像，同时最大限度地减少光毒性，漂白，并保持细胞的健康一般。在这里，我们提供了一个协议，它描述了一个脂质体转染法，收益率相对较低的转染率（0.5％），但是是完全极化神经元成像的理想选择。低转染率是至关重要的，使单个轴突和树突细胞体内的方向的特点，以确认的方向性，即运输，顺行诉逆行。我们的方法来成像GFP的表达神经元依赖于一个标准的宽视场荧光显微镜与CCD摄像头，图像采集软件，和热成像室配备。我们有成像的细胞器或结构的种类繁多，例如，没有任何特殊的光学或激发需求，比荧光灯光源的致密核心囊泡，线粒体，生长锥，与肌动蛋白。此外，不同的光谱，荧光标记的蛋白质，例如，绿色荧光蛋白和蛋白DsRed的标签，可附近同时可视化表征的合作运输或其他协调的细胞活动。成像方法，这里描述的是各种成像应用灵活，可以由实验室通过相对较少的成本提供了一个显微镜。

Watch this Scientific Journal Video about Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons at JoVE.com

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

活细胞成像研究细胞器官贩卖的动态时，特别实用。在这里，我们描述了一个致密核心囊泡在培养的神经元，采用宽视场荧光显微镜的实时成像的协议。此协议是灵活的和可适应图像的其它细胞器如线粒体，内体，和过氧化物酶。

在原代培养海马神经元的密集核心囊泡的实时成像

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

live-imaging-dense-core-vesicles-primary-cultured-hippocampal

Research

JoVE Journal

Biology

12.5K 次观看.  Simon Fraser University. 活细胞成像研究细胞器官贩卖的动态时，特别实用。在这里，我们描述了一个致密核心囊泡在培养的神经元，采用宽视场荧光显微镜的实时成像的协议。此协议是灵活的和可适应图像的其它细胞器如线粒体，内体，和过氧化物酶。

视频: Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Gene-gun Transfection of Hippocampal Neurons

海马神经元的基因枪转

科研

生物学

Primary Culture of Hippocampal Neurons from P0 Newborn Rats

The physiological properties of hippocampal neurons are commonly investigated, especially because of the involvement of the hippocampus in learning and memory. Primary hippocampal cell culturing allows neuroscientists to examine the activity and properties of neurons at the individual cell and single synapse level. In this video, we will demonstrate how to isolate and grow primary hippocampal cells from newborn rats. The hippocampus may be isolated from each newborn animal in as short as 2 to 3 minutes, and the cultures can be maintained for up to two weeks. We will also briefly demonstrate how to use these hippocampal neurons for ratiometric calcium imaging. While this protocol describes the process for the hippocampus, with little to no modification, it can be applied to other regions of the brain.

The dissection and growth of cells from an individual brain area facilitates investigation of cellular and physiological parameters. We describe a method for primary cell culturing that produces neuron-enriched cultures in a serum-free environment.

从P0新生大鼠海马神经元的原代培养

The physiological properties of hippocampal neurons are commonly investigated, especially because of the involvement of the hippocampus in learning and ...

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

在斑马鱼胚胎的单个神经元和神经嵴细胞的细胞运动和肌动蛋白细胞骨架的实时成像

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Adenovirus-mediated Genetic Removal of Signaling Molecules in Cultured Primary Mouse Embryonic Fibroblasts

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction analysis. One particular method to achieve this conditional deletion is via the use of the Cre-loxP system. This method involves flanking the gene of interest with loxP sites, which are specific recognition sequences for the Cre recombinase protein. Exposure of the so-called floxed (flanked by loxP site) DNA to this enzyme results in a Cre-mediated recombination event at the loxP sites, and subsequent excision of the intervening gene3. Several different methods exist to administer Cre recombinase to the site of interest. In this video, we demonstrate the use of an adenovirus containing the Cre recombinase gene to infect primary mouse embryonic fibroblasts (MEFs) obtained from embryos containing a floxed Rac1 allele1. Our rationale for selecting Rac1 MEFs for our experiments is that clear morphological changes can be seen upon deletion of Rac1, due to alterations in the actin cytoskeleton2,5. 72 hours following viral transduction and Cre expression, cells were stained using the actin dye phalloidin and imaged using confocal laser scanning microscopy. It was observed that MEFs which had been exposed to the adeno-Cre virus appeared contracted and elongated in morphology compared to uninfected cells, consistent with previous reports2,5. The adenovirus method of Cre recombinase delivery is advantageous as the adeno-Cre virus is easily available, and gene deletion via Cre in nearly 100% of the cells can be achieved with optimized adenoviral infection.

In this video we use an adenovirus carrying the Cre recombinase gene to infect primary mouse embryonic fibroblasts carrying a floxed Rac1 allele.

腺病毒介导的基因去除信号分子在原代培养的小鼠胚胎成纤维

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction ...

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 ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

凋亡细胞清除过程中实时成像<em&gt;果蝇</em&gt;胚胎发育

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

细胞器运输的培养<em&gt;果蝇</em&gt;细胞：S2细胞系和原代神经元。

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

Visualization of G3BP Stress Granules Dynamics in Live Primary Cells

SGs can be visualized in cells by immunostaining of specific protein components or polyA+ mRNAs. SGs are highly dynamic and the study of their assembly and fate is important to understand the cellular response to stress. The deficiency in key factors of SGs like G3BP (RasGAP SH3 domain Binding Protein) leads to developmental defects in mice&#160;and alterations of the Central Nervous System. To study the dynamics of SGs in cells from an organism, one can culture primary cells and follow the localization of a transfected tagged component of SGs. We describe time-lapse experiment to observe G3BP1-containing SGs in Mouse Embryonic Fibroblasts (MEFs). This technique can also be used to study G3BP-containing SGs in live neurons, which is crucial as it was recently shown that these SGs are formed at the onset of neurodegenerative diseases like Alzheimer&#39;s disease. This approach can be adapted to any other cellular body and granule protein component, and performed with transgenic animals, allowing the live study of granules dynamics for example in the absence of a specific factor of these granules.

Stress granules (SGs) are cytoplasmic RNA granules containing stalled ribonucleoprotein particles (RNPs), and important in cellular response to various stresses. Dynamics of SGs can be followed in live cells by visualizing the localization of a tagged component of SGs in transfected primary cells after stress.

中G3BP应力颗粒动力学在Live原代细胞可视化

SGs can be visualized in cells by immunostaining of specific protein components or polyA+ mRNAs. SGs are highly dynamic and the study of their assembly ...

Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single procedure. Due to advances in microscope technology it is relatively easy to capture live cell images for the purpose of investigating cellular events in real time with minimal concern regarding phototoxicity to the cells. This protocol describes how to take live cell timelapse images of primary rat neonatal cardiomyocytes using a confocal spinning disk microscope following lentiviral and adenoviral transduction to modulate properties of the cell. The application of two different types of viruses makes it easier to achieve an appropriate transduction rate and expression levels for two different genes. Well focused live cell images can be obtained using the microscope&#8217;s autofocus system, which maintains stable focus for long time periods. Applying this method, the functions of exogenously engineered proteins expressed in cultured primary cells can be analyzed. Additionally, this system can be used to examine the functions of genes through the use of siRNAs as well as of chemical modulators.

This protocol describes a method of live cell imaging using primary rat neonatal cardiomyocytes following lentiviral and adenoviral transduction using confocal spinning disk microscopy. This enables detailed observations of cellular processes in living cardiomyocytes.

原代大鼠乳鼠心肌细胞的活细胞成像继腺病毒和慢病毒转导利用共聚焦旋转盘显微镜

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single ...