Name: time-lapse video microscopy for assessment of eyfp-parkin aggregation as a marker for cellular mitophagy
Uploaded: 2016-05-04T13:00:00.000+00:00
Duration: 9 min 29 s
Description: Watch this Scientific Journal Video about Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy at JoVE.com

This work was supported in part by NIH grants (1R01CA137494, R01CA132115, R01CA086072 to R.G.P.), the Kimmel Cancer Center NIH Cancer Center Core grant P30CA056036 (R.G.P.), a grant from the Breast Cancer Research Foundation, generous grants from the Dr. Ralph and Marian C. Falk Medical Research Trust (R.G.P.) and a grant from the Pennsylvania Department of Health (R.G.P.). In part this work was supported by an American Italian Cancer Foundation postdoctoral fellowship (G.D.) and Bioimaging Shared Resource of the Sidney Kimmel Cancer Center (NCI 5 P30 CA-56036).The Department specifically disclaims responsibility for an analysis, interpretations or conclusions. There are no conflicts of interest associated with this manuscript.

1.成纤维细胞的电穿孔用两种表达载体EYFP - 帕金和pDsRed2-美图<ol><li>生长在10厘米组织培养板使用的DMEM（Dulbecco氏改进的Eagle培养基）补充有10％胎牛血清，2毫摩尔/升L-谷氨酰胺，100U / ml青霉素介质永生化小鼠胚胎成纤维细胞，和100毫克/毫升链霉素在含有5％的CO 2，在37℃潮湿气氛。 <ol><li>在80％的细胞汇合，通过无菌吸弃完整的DMEM培养基并加入10毫升的无菌1×磷酸盐缓冲液（PBS）（80克NaCl，2.0g的氯化钾，14.4克Na 2 HPO 4 2.4克KH的2 PO 4和1升蒸馏水2 O，PH值:7.4）。 </li><li>无菌吸弃PBS，并加入1毫升胰蛋白酶EDTA 0.25％。在37℃下孵育该板直至细胞被分离（2 - 3分钟）。加入4 ml完全DMEM培养基，重悬细胞，并取出10 _6;细胞悬浮液的血球计数的湖 注:血细胞计数被设计为使得在一组的16角正方形的细胞数量相当于细胞×10 4个 / ml的数。 </li></ol></li><li>种子1×10 6细胞到10厘米组织培养皿前24小时到电穿孔过程。 </li><li>无菌吸弃完整的DMEM中，加入10毫升无菌PBS的。无菌吸弃PBS，并加入1毫升胰蛋白酶EDTA 0.25％。在37℃下孵育该板直至细胞被分离（2 - 3分钟）。加入4 ml完全DMEM培养基和悬浮细胞在一个15毫升管。 </li><li>使用冷冻离心机（4℃）纺细胞向下，在250×g离心5分钟。丢弃无菌吸上清，悬浮颗粒在1毫升无菌PBS。旋细胞向下，在250×g离心5分钟，4℃。 </li><li>丢弃无菌吸上清，加100微升Ø˚F解决方案组合为电（82微升电解决方案V +18μl的补充解决方案1）的细胞沉淀和移液器轻轻悬浮颗粒（欲了解更多详情，请参阅用户指南）。在细胞悬液中加入2微克EYFP - 帕金（激发/发射527分之514）和1微克pDsRed2-美图（激发/发射六百二十〇分之五百六十五）的。 </li><li>转移溶液至用一次性巴斯德灭菌反应杯（这两个工具都随试剂盒提供），使用预先设定的程序的NIH / 3T3的U-030（单脉冲，电压200伏，电容960μF，脉冲时间20毫秒电穿孔，脉冲数:1）。 </li><li>电穿孔后，立即在37℃下在含有5％的CO 2的潮湿气氛中加入500μl新鲜预热完全DMEM培养基和种子上将6cm组织培养板活成像同类细胞孵育它们24小时。 </li></ol> 2.时间推移​​视频显微镜<ol><l我&gt;设置在显微镜的腔室在37℃至使用前的温度。 </li><li>在这一点上，准备一个特定的实时成像介质进行与实验协议。准备实时成像介质如下;混合DMEM苯酚 - 自由补充有10％胎牛血清，2毫摩尔/升L-谷氨酰胺，100U / ml青霉素，和100毫克/毫升链霉素。在37℃的水浴中预暖实时成像介质。 </li><li>丢弃电穿孔的细胞无菌抽吸培养基并加入预热实时成像介质1毫升，使用P1000吸管，并在37℃下孵育该板30分钟。 </li><li>在板潜伏期，流入5％的CO 2在显微镜的腔室已经在37℃的稳定温度。 </li><li>放置板的电穿孔细胞进入显微镜的房间，避免重大变动或振荡。 </li><li>使用软件接口，以便同时检测荧光设置显微镜从由共转染载体编码的融合蛋白刘哲民信号，EYFP-帕金（励磁范围495〜510 nm，发射范围520至550nm，绿色）和pDsRed2-三刀（励磁最大558纳米和最大发射583纳米，红）。 <ol><li>打开时间推移视频显微镜软件。在上面的菜单中选择EYFP - 帕金和罗丹明对pDsRed2-三刀FITC。在上面的菜单中选择放大倍率（20X）。 </li></ol></li><li>使用软件界面，寻找既表达共转染载体的单细胞和注册的位置。直到最少10个细胞的收集对于每个实验条件（实验组）重复此步骤。 <ol><li>选择菜单"应用程序"，然后单击多维采集。在多维采集窗口选择所需的参数，如收购的数量，每个采集和记录单元的位置之间的时间间隔。点击"采集"开始采集过程。 </li></ol></li><li>启动基底采集既EYFP-帕金和采集图像每5分钟的15分钟的总时间间隔的DsRed2-三刀荧光信号。 </li><li>制备预热实时成像介质用浓度FCCP的比最终工作浓度高两倍（0.1 - 10μM，细胞类型依赖性的）。 </li><li>中断采集过程，并添加轻轻地加入1ml用FCCP预热实时成像介质的成用吸管P1000显微镜的室内板。 </li><li>如前所述，为3小时的总时间，使用所记录的位置重新开始获取过程。保存所有采集到的图像，以分析它们，当收购是在创建过程mitophagic的视频。 </li></ol> 3.分析<ol><li>收集的个人计算机上的所有在".TIFF"格式所获取的图像</li><li>与图形软打开图片洁具及定义时间间隔由线粒体引起的去极化与FCCP以显示进入线粒体膜帕金招募的第一个形象出现（图像采集每5分钟）。重复此分析在实验组中的每个小区。 </li><li>标签上与实验组名的电子表格的列的第一个单元。对于每个实验组，测量为帕金招募最少10个细胞的的时间间隔。作出帕金招募所计算的时间间隔的平均值，并定义为每个实验组的标准偏差（平均值±SD，N ​​= 10）。 <ol><li>整理成单个计算时间间隔列。每列包含N =实验组10次测量。 </li><li>选择从公式生成器菜单功能"平均"。选择列中的数据。选择从公式生成器菜单功能"标准差"。塞莱克拉列中的数据。 </li></ol></li><li>使用统计方法，比较以确定在实验组之间mitophagy的动态一个显著差施加适当的统计实验组。 （例如，两组=学生t-检验 ，三个或更多个基团= ANOVA加一个事后检验 ） </li></ol>

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The authors have nothing to disclose.

时间推移显微镜可以被定义为，能在一次观测中时间延长活细胞成像到细胞动力学的过长时间的观察的技术。这种方法是从简单的共焦或活细胞显微镜分辨，因为它允许观察者能够实时识别单个荧光标记的蛋白质，并按照其一个单一的活细胞内的动态。事实上，共焦显微镜可以使用荧光抗体容易识别免疫标记的蛋白质，但它不允许观察细胞，并在其实际环境中的分子事件。 ?...

Macro autophagy is an intracellular process that involves the catabolic degradation of both damaged and dysfunctional cellular components, such as organelles and proteins for the purpose of either recycling or energy production. To initiate this metabolic process, the cell engulfs the damaged cellular components into a double-membrane structure, known as an autophagosome, which fuses with a lysosome and its content is degraded and recycled 1,2. There are two major types of autophagy, the non-selective and selective. The non-selective autophagy process occurs when the cell is under nutrient deprivation conditions and needs to scavenge for both essential nutrients and energy. However, selective autophagy occurs to mediate the removal of both dysfunctional/damaged organelles and proteins that otherwise could be toxic. One of the most studied selective autophagy process is the removal of mitochondria, termed mitophagy 1,3-5.
Mitochondria are the central organelles for cell metabolism and the primary source of adenosine triphosphate (ATP) via oxidative phosphorylation through the electron transport chain, fatty acid oxidation, and tricarboxylic acid (TCA) cycle. Moreover, mitochondria regulate reactive oxygen species (ROS) production and release proteins that participate in cell death pathways 6-8.
PTEN-induced putative kinase 1 (PINK1) and Parkin RBR E3 ubiquitin ligase (Parkin) are the key proteins implicated in the mitophagy process. Parkin can protect against cell death by keeping the cell healthy through mitochondrial quality control9. Upon the loss of mitochondrial membrane potential, cytosolic Parkin is recruited to the mitochondria by PINK1. This recruitment triggers the sequential events of mitophagy 10. There is a broad range of evidence that mitophagy is a fundamental mitochondria quality control process and abnormalities in this process drive disease 7. For instance, autosomal recessive Parkinson's disease has been associated with mutations in the genes that encode for Parkin and PINK1 (PARK2 and PINK1, respectively) 11. The quality control of mitochondrial health is essential for the removal of mitochondria that contribute to the accumulation of ROS12. Excessive presence of intracellular ROS can lead to damage of both nuclear and mitochondrial DNA (DNA and mt DNA, respectively).
Herein, we show a time-lapse video microscopy approach to follow the aggregation of Parkin after the induction of Parkin-mediated mitophagy in immortalized mouse embryonic fibroblasts via in vitro administration of carbonyl cyanide 4-(trifluoromethoxy)-phenylhydrazone (FCCP), an uncoupling agent. FCCP disrupts ATP synthesis by short circuiting protons across the outer mitochondria membrane and hence uncoupling oxidative phosphorylation from the electron transport chain 13. Triggering the depolarization of the mitochondrial membrane leads to the disruption of mitochondria and selective Parkin-dependent removal. Therefore, transfecting the cells of interest with an expression vector encoding Parkin fused with a fluorescent marker (enhanced yellow fluorescent protein, EYFP) can be used as a fluorescent tag to follow the recruitment of Parkin during the mitophagic process. In order to visualize the mitochondria, we co-transfected pDsRed2-Mito, which encodes red fluorescent protein (DsRed2) that contains a mitochondrial targeting sequence of cytochrome c oxidase subunit VIII (Mito). pDsRed2-Mito is designed for fluorescent labeling of mitochondria14. The time required for Parkin translocation into the mitochondrial membrane can be measured and gives an indirect measure of cellular health. For example, we can say that if a cell line knocked-out for a particular gene of interest shows either a faster or slower recruitment of Parkin after the induction of mitophagy by FCCP, that gene product would be a key player in order to keep the metabolic rates of the cell at the physiological status and prevent the development of diseases. Therefore, the time-lapse video microscopy provides a very powerful tool for both basic and clinical research applications in following the dynamic of labeled proteins during their molecular processes and understanding how these processes are affected during a pathological condition.

<table><tbody><tr><td>DMEM</td><td>Corning Life Science</td><td>10-013-CV</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>Phenol-free DMEM</td><td>Corning Life Science</td><td>17-205-CV</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>Fetal Bovine Serum</td><td>Sigma-Aldrich</td><td>F2442</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>L-Glutamine</td><td>Gibco</td><td>25030</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>Penicillin/streptomycin</td><td>Corning Life Science</td><td>30-002-CI</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>EYFP-PARKIN expression vector</td><td>Addgene</td><td>23955</td><td></td></tr><tr><td>pDsRed2-Mito expression vector</td><td>Clontech</td><td>632421</td><td></td></tr><tr><td>Nucleofector 2B device</td><td>LONZA</td><td>AAD-1001S</td><td></td></tr><tr><td>Nucleofector for kit R NIH/3T3</td><td>LONZA</td><td>VCA-1001</td><td></td></tr><tr><td>ZEISS AXIOVERT 200M inverted microscope</td><td>CARL ZEISS</td><td></td><td></td></tr><tr><td>Carbonyl Cyanide 4-(trifluoromethoxy)-Phenylhydrazone (FCCP)</td><td>Sigma-Aldrich</td><td>C2920</td><td></td></tr><tr><td>MetaMorph</td><td>Molecular Devices</td><td>Experimental Builder</td><td></td></tr><tr><td>ImageJ</td><td>National Institute of Health</td><td>Experimental Builder</td><td></td></tr></tbody></table>

time-lapse video microscopy for assessment of eyfp-parkin aggregation as a marker for cellular mitophagy

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time points. Time intervals can be set through a computer interface that controls the microscope-integrated camera. This kind of microscopy requires both the ability to acquire very rapid events and the signal generated by the observed cellular structure during these events. After the images have been collected, a movie of the entire experiment is assembled to show the dynamic of the molecular events of interest. Time-lapse video microscopy has a broad range of applications in the biomedical research field and is a powerful and unique tool for following the dynamics of the cellular events in real time. Through this technique, we can assess cellular events such as migration, division, signal transduction, growth, and death. Moreover, using fluorescent molecular probes we are able to mark specific molecules, such as DNA, RNA or proteins and follow them through their molecular pathways and functions. Time-lapse video microscopy has multiple advantages, the major one being the ability to collect data at the single-cell level, that make it a unique technology for investigation in the field of cell biology. However, time-lapse video microscopy has limitations that can interfere with the acquisition of high quality images. Images can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. Herein, we describe a protocol for the dynamic acquisition of a specific protein, Parkin, fused with the enhanced yellow fluorescent protein (EYFP) in order to track the selective removal of damaged mitochondria, using a time-lapse video microscopy approach.

这里，我们显示的时间推移的视频显微镜如何是一个强大的技术，可用于遵循荧光标记的蛋白质的分子事件在单个小区。代表结果还表明，该技术是如何允许采集高质量的图像。当分子过程的图象，而得到的，我们有机会来分析他们以不同的方式。这里，我们分析的感应mitophagy过程，这是至关重要的分子过程，维持细胞稳态选择性地除去受损的线粒体的起始之间的时间间?...

Watch this Scientific Journal Video about Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy at JoVE.com

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

时间推移视频显微镜对EYFP-Parkin的聚集性评估作为蜂窝Mitophagy一个标记

Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time ...

time-lapse-video-microscopy-for-assessment-eyfp-parkin-aggregation-as

Research

JoVE Journal

Biology

7.1K 次观看.  Thomas Jefferson University, SKCC. Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

视频: Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

Live-cell Video Microscopy of Fungal Pathogen Phagocytosis

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of phagocytes to the site where pathogens are located; (ii) recognition of pathogen-associated molecular patterns (PAMPs) through pattern recognition receptors (PRRs); (iii) engulfment of microorganisms bound to the phagocyte cell membrane, and (iv) processing of engulfed cells within maturing phagosomes and digestion of the ingested particle. Studies that assess phagocytosis in its entirety are informative1, 2, 3, 4, 5 but are limited in that they do not normally break the process down into migration, engulfment and phagosome maturation, which may be affected differentially. Furthermore, such studies assess uptake as a single event, rather than as a continuous dynamic process. We have recently developed advanced live-cell imaging technologies, and have combined these with genetic functional analysis of both pathogen and host cells to create a cross-disciplinary platform for the analysis of innate immune cell function and fungal pathogenesis. These studies have revealed novel aspects of phagocytosis that could only be observed using systematic temporal analysis of the molecular and cellular interactions between human phagocytes and fungal pathogens and infectious microorganisms more generally. For example, we have begun to define the following: (a) the components of the cell surface required for each stage of the process of recognition, engulfment and killing of fungal cells1, 6, 7, 8; (b) how surface geometry influences the efficiency of macrophage uptake and killing of yeast and hyphal cells7; and (c) how engulfment leads to alteration of the cell cycle and behavior of macrophages 9, 10.
In contrast to single time point snapshots, live-cell video microscopy enables a wide variety of host cells and pathogens to be studied as continuous sequences over lengthy time periods, providing spatial and temporal information on a broad range of dynamic processes, including cell migration, replication and vesicular trafficking. Here we describe in detail how to prepare host and fungal cells, and to conduct the video microscopy experiments. These methods can provide a user-guide for future studies with other phagocytes and microorganisms.

We describe methods for live-cell video microscopy of Candida albicans phagocytosis by macrophages. These methods enable stage-specific analysis of macrophage migration, recognition, engulfment and phagosome maturation and reveal novel aspects of phagocytosis.

活细胞视频显微镜病病原菌的吞噬作用

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of ...

科研

免疫与感染

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

在酵母用于监测Mitophagy荧光显微镜检测酿酒酵母

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

生物学

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

单细胞荧光显微镜定量 Efferocytosis 的研究

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

A Faster, High Resolution, mtPA-GFP-based Mitochondrial Fusion Assay Acquiring Kinetic Data of Multiple Cells in Parallel Using Confocal Microscopy

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in living cells by photo-conversion of matrix targeted photoactivatable GFP (mtPAGFP) in a subset of mitochondria. The rate at which the photoconverted molecules equilibrate across the entire mitochondrial population is used as a measure of fusion activity. Thus far measurements were performed using a single cell time lapse approach, quantifying the equilibration in one cell over an hour. Here, we scale up and automate a previously published live cell method based on using mtPAGFP and a low concentration of TMRE (15 nm). This method involves photoactivating a small portion of the mitochondrial network, collecting highly resolved stacks of confocal sections every 15 min for 1 hour, and quantifying the change in signal intensity. Depending on several factors such as ease of finding PAGFP expressing cells, and the signal of the photoactivated regions, it is possible to collect around 10 cells within the 15 min intervals. This provides a significant improvement in the time efficiency of this assay while maintaining the highly resolved subcellular quantification as well as the kinetic parameters necessary to capture the detail of mitochondrial behavior in its native cytoarchitectural environment.
Mitochondrial dynamics play a role in many cellular processes including respiration, calcium regulation, and apoptosis1,2,3,13. The structure of the mitochondrial network affects the function of mitochondria, and the way they interact with the rest of the cell. Undergoing constant division and fusion, mitochondrial networks attain various shapes ranging from highly fused networks, to being more fragmented. Interestingly, Alzheimer's disease, Parkinson's disease, Charcot Marie Tooth 2A, and dominant optic atrophy have been correlated with altered mitochondrial morphology, namely fragmented networks4,10,13. Often times, upon fragmentation, mitochondria become depolarized, and upon accumulation this leads to impaired cell function18. Mitochondrial fission has been shown to signal a cell to progress toward apoptosis. It can also provide a mechanism by which to separate depolarized and inactive mitochondria to keep the bulk of the network robust14. Fusion of mitochondria, on the other hand, leads to sharing of matrix proteins, solutes, mtDNA and the electrochemical gradient, and also seems to prevent progression to apoptosis9. How fission and fusion of mitochondria affects cell homeostasis and ultimately the functioning of the organism needs further understanding, and therefore the continuous development and optimization of how to gather information on these phenomena is necessary.
Existing mitochondrial fusion assays have revealed various insights into mitochondrial physiology, each having its own advantages. The hybrid PEG fusion assay7, mixes two populations of differently labeled cells (mtRFP and mtYFP), and analyzes the amount of mixing and colocalization of fluorophores in fused, multinucleated, cells. Although this method has yielded valuable information, not all cell types can fuse, and the conditions under which fusion is stimulated involves the use of toxic drugs that likely affect the normal fusion process. More recently, a cell free technique has been devised, using isolated mitochondria to observe fusion events based on a luciferase assay1,5. Two human cell lines are targeted with either the amino or a carboxy terminal part of Renilla luciferase along with a leucine zipper to ensure dimerization upon mixing. Mitochondria are isolated from each cell line, and fused. The fusion reaction can occur without the cytosol under physiological conditions in the presence of energy, appropriate temperature and inner mitochondrial membrane potential. Interestingly, the cytosol was found to modulate the extent of fusion, demonstrating that cell signaling regulates the fusion process 4,5. This assay will be very useful for high throughput screening to identify components of the fusion machinery and also pharmacological compounds that may affect mitochondrial dynamics. However, more detailed whole cell mitochondrial assays will be needed to complement this in vitro assay to observe these events within a cellular environment.
A technique for monitoring whole-cell mitochondrial dynamics has been in use for some time and is based on a mitochondrially-targeted photoactivatable GFP (mtPAGFP)6,11. Upon expression of the mtPAGFP, a small portion of the mitochondrial network is photoactivated (10-20%), and the spread of the signal to the rest of the mitochondrial network is recorded every 15 minutes for 1 hour using time lapse confocal imaging. Each fusion event leads to a dilution of signal intensity, enabling quantification of the fusion rate. Although fusion and fission are continuously occurring in cells, this technique only monitors fusion as fission does not lead to a dilution of the PAGFP signal6. Co-labeling with low levels of TMRE (7-15 nM in INS1 cells) allows quantification of the membrane potential of mitochondria. When mitochondria are hyperpolarized they uptake more TMRE, and when they depolarize they lose the TMRE dye. Mitochondria that depolarize no longer have a sufficient membrane potential and tend not to fuse as efficiently if at all. Therefore, active fusing mitochondria can be tracked with these low levels of TMRE9,15. Accumulation of depolarized mitochondria that lack a TMRE signal may be a sign of phototoxicity or cell death. Higher concentrations of TMRE render mitochondria very sensitive to laser light, and therefore great care must be taken to avoid overlabeling with TMRE. If the effect of depolarization of mitochondria is the topic of interest, a technique using slightly higher levels of TMRE and more intense laser light can be used to depolarize mitochondria in a controlled fashion (Mitra and Lippincott-Schwartz, 2010). To ensure that toxicity due to TMRE is not an issue, we suggest exposing loaded cells (3-15 nM TMRE) to the imaging parameters that will be used in the assay (perhaps 7 stacks of 6 optical sections in a row), and assessing cell health after 2 hours. If the mitochondria appear too fragmented and cells are dying, other mitochondrial markers, such as dsRED or Mitotracker red could be used instead of TMRE.
The mtPAGFP method has revealed details about mitochondrial network behavior that could not be visualized using other methods. For example, we now know that mitochondrial fusion can be full or transient, where matrix content can mix without changing the overall network morphology. Additionally, we know that the probability of fusion is independent of contact duration and organelle dimension, is influenced by organelle motility, membrane potential and history of previous fusion activity8,15,16,17.
In this manuscript, we describe a methodology for scaling up the previously published protocol using mtPAGFP and 15nM TMRE8 in order to examine multiple cells at a time and improve the time efficiency of data collection without sacrificing the subcellular resolution. This has been made possible by the use of an automated microscope stage, and programmable image acquisition software. Zen software from Zeiss allows the user to mark and track several designated cells expressing mtPAGFP. Each of these cells can be photoactivated in a particular region of interest, and stacks of confocal slices can be monitored for mtPAGFP signal as well as TMRE at specified intervals. Other confocal systems could be used to perform this protocol provided there is an automated stage that is programmable, an incubator with CO2, and a means by which to photoactivate the PAGFP; either a multiphoton laser, or a 405 nm diode laser.

Mitochondrial fusion was measured by tracking the equilibration of photoconverted matrix-targeted GFP across the mitochondrial network over time. Thus far, only one cell could be subjected to an hour long kinetic analysis at a time. We present a method that simultaneously measures multiple cells, thereby speeding up the data collection process.

速度更快，高分辨率，MTPA-GFP基于线粒体融合含量的收购多个单元格的动力学数据，在并行使用共聚焦显微镜

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in ...

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a wide variety of disorders including metabolic diseases, premature aging syndromes, and neurodegenerative diseases such as Alzheimer's disease (AD) and Parkinson's disease (PD). Maintenance of mitochondrial health depends on mitochondrial biogenesis and the efficient clearance of dysfunctional mitochondria through mitophagy. Experimental methods to accurately detect autophagy/mitophagy, especially in animal models, have been challenging to develop. Recent progress towards the understanding of the molecular mechanisms of mitophagy has enabled the development of novel mitophagy detection techniques. Here, we introduce several versatile techniques to monitor mitophagy in human cells, Caenorhabditis elegans (e.g., Rosella and DCT-1/ LGG-1 strains), and mice (mt-Keima). A combination of these mitophagy detection techniques, including cross-species evaluation, will improve the accuracy of mitophagy measurements and lead to a better understanding of the role of mitophagy in health and disease.

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitophagy, the process of clearing damaged mitochondria, is necessary for mitochondrial homeostasis and health maintenance. This article presents some of the latest mitophagy detection methods in human cells, Caenorhabditis elegans, and mice.

体外和体内检测人类细胞中的 Mitophagy,线虫和小鼠

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a ...

医学

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

荧光标记蛋白 ImageJ 的无偏分析在果蝇变性定量细胞生物学中的应用

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...

神经科学

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

活细胞成像早期自噬活动：Omegasomes与超越

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

Visualization of Endogenous Mitophagy Complexes In Situ in Human Pancreatic Beta Cells Utilizing Proximity Ligation Assay

Mitophagy is an essential mitochondrial quality control pathway, which is crucial for pancreatic islet beta cell bioenergetics to fuel glucose-stimulated insulin release. Assessment of mitophagy is challenging and often requires genetic reporters or multiple complementary techniques not easily utilized in tissue samples, such as primary human pancreatic islets. Here we demonstrate a robust approach to visualize and quantify formation of key endogenous mitophagy complexes in primary human pancreatic islets. Utilizing the sensitive proximity ligation assay technique to detect interaction of the mitophagy regulators NRDP1 and USP8, we are able to specifically quantify formation of essential mitophagy complexes in situ. By coupling this approach to counterstaining for the transcription factor PDX1, we can quantify mitophagy complexes, and the factors that can impair mitophagy, specifically within beta cells. The methodology we describe overcomes the need for large quantities of cellular extracts required for other protein-protein interaction studies, such as immunoprecipitation (IP) or mass spectrometry, and is ideal for precious human islet&#160;samples generally not available in sufficient quantities for these approaches. Further, this methodology obviates the need for flow sorting techniques to purify beta cells from a heterogeneous islet population for downstream protein applications. Thus, we describe a valuable protocol for visualization of mitophagy highly compatible for use in heterogeneous and limited cell populations.

This protocol outlines a method for quantitative analysis of mitophagy protein complex formation specifically in beta cells from primary human islet samples. This technique thus allows analysis of mitophagy from limited biological material, which are crucial in precious human pancreatic beta cell samples.

利用邻近结扎测定法在人类胰腺β细胞中原位内源性米托巴结物的可视化

Mitophagy is an essential mitochondrial quality control pathway, which is crucial for pancreatic islet beta cell bioenergetics to fuel glucose-stimulated ...

Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between defective mitophagy and various human diseases, including neurodegenerative diseases, cancer, and metabolic diseases. In addition, recent studies have shown that mitophagy is involved in normal cellular processes, such as differentiation and development. However, the precise role of and molecular mechanisms underlying mitophagy require further study. Therefore, it is critical to develop a robust and convenient method for measuring changes in mitophagy activity. Here, we describe a detailed protocol for quantitatively assessing mitophagy activity through flow cytometry using the mitochondria-targeted fluorescent protein Keima (mt-Keima). This flow cytometry assay can analyze mitophagy activity more rapidly and sensitively than conventional microscopy- or immunoblotting-based methods. This protocol can be applied to analyze mitophagy activity in various cell types.

Mitophagy, the selective degradation of mitochondria, has been implicated in mitochondrial homeostasis and is deregulated in various human diseases. However, convenient experimental methods for measuring mitophagy activity are very limited. Here, we provide a sensitive assay for measuring mitophagy activity using flow cytometry.

pH 依赖性荧光 Keima 对流式细胞术 Mitophagy 的敏感性测定

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between ...

Protection of H9c2 Myocardial Cells from Oxidative Stress by Crocetin via PINK1/Parkin Pathway-Mediated Mitophagy

This study aimed to explore the oxidative stress-protective effect of crocetin on H2O2-mediated H9c2 myocardial cells through in vitro experiments, and further explore whether its mechanism is related to the impact of mitophagy. This study also aimed to demonstrate the therapeutic effect of safflower acid on oxidative stress in cardiomyocytes and explore whether its mechanism is related to the effect of mitophagy. Here, an H2O2-based oxidative stress model was constructed and assessed the degree of oxidative stress injury of cardiomyocytes by detecting the levels of lactate dehydrogenase (LDH), creatine kinase (CK), malondialdehyde (MDA), superoxide dismutase (SOD), catalase (CAT), and glutathione peroxidase (GSH Px). Reactive oxygen species (ROS)-detecting fluorescent dye DCFH-DA, JC-1 dye, and TUNEL dye were employed to assess mitochondrial damage and apoptosis. Autophagic flux was measured by transfecting Ad-mCherry-GFP-LC3B adenovirus. Mitophagy-related proteins were then detected via western blotting and immunofluorescence. However, crocetin (0.1-10 &#181;M) could significantly improve cell viability and reduce apoptosis and oxidative stress damage caused by H2O2. In cells with excessive autophagic activation, crocetin could also reduce autophagy flow and the expression of mitophagy-related proteins PINK1 and Parkin, and reverse the transfer of Parkin to mitochondria. Crocetin could reduce H2O2-mediated oxidative stress damage and the apoptosis of H9c2 cells, and its mechanism was closely related to mitophagy.

Protection of H9c2 Myocardial Cells from Oxidative Stress by Crocetin via PINK1/Parkin Pathway-Mediated Mitophagy

Based on in vitro experiments, this study revealed the mechanism of crocetin in repairing oxidative stress damage of cardiomyocytes by influencing mitophagy, in which the PINK1/Parkin signaling pathway plays an important role.

通过PINK1 / Parkin途径介导的线粒体自噬 保护 H9c2心肌细胞免受藏红花素氧化应激

This study aimed to explore the oxidative stress-protective effect of crocetin on H2O2-mediated H9c2 myocardial cells through in vitro experiments, and ...

时间推移视频显微镜对EYFP-Parkin的聚集性评估作为蜂窝Mitophagy一个标记

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活细胞视频显微镜病病原菌的吞噬作用

在酵母用于监测Mitophagy荧光显微镜检测酿酒酵母

单细胞荧光显微镜定量 Efferocytosis 的研究

速度更快，高分辨率，MTPA-GFP基于线粒体融合含量的收购多个单元格的动力学数据，在并行使用共聚焦显微镜

体外和体内检测人类细胞中的 Mitophagy,线虫和小鼠

荧光标记蛋白 ImageJ 的无偏分析在果蝇变性定量细胞生物学中的应用

活细胞成像早期自噬活动：Omegasomes与超越

利用邻近结扎测定法在人类胰腺β细胞中原位内源性米托巴结物的可视化

pH 依赖性荧光 Keima 对流式细胞术 Mitophagy 的敏感性测定

通过PINK1 / Parkin途径介导的线粒体自噬保护 H9c2心肌细胞免受藏红花素氧化应激

时间推移视频显微镜对EYFP-Parkin的聚集性评估作为蜂窝Mitophagy一个标记

摘要

探索更多视频

此视频中的章节

活细胞视频显微镜病病原菌的吞噬作用

在酵母用于监测Mitophagy荧光显微镜检测酿酒酵母

单细胞荧光显微镜定量 Efferocytosis 的研究

速度更快，高分辨率，MTPA-GFP基于线粒体融合含量的收购多个单元格的动力学数据，在并行使用共聚焦显微镜

体外和体内检测人类细胞中的 Mitophagy,线虫和小鼠

荧光标记蛋白 ImageJ 的无偏分析在果蝇变性定量细胞生物学中的应用

活细胞成像早期自噬活动：Omegasomes与超越

利用邻近结扎测定法在人类胰腺β细胞中原位内源性米托巴结物的可视化

pH 依赖性荧光 Keima 对流式细胞术 Mitophagy 的敏感性测定

通过PINK1 / Parkin途径介导的线粒体自噬 保护 H9c2心肌细胞免受藏红花素氧化应激

通过PINK1 / Parkin途径介导的线粒体自噬保护 H9c2心肌细胞免受藏红花素氧化应激