作者感谢勒Scriwanek与图编制的援助。 FR是由ECHO（700.59.003）的支持，ALW打开程序（821.02.017和822.02.014），DFG-NWO合作（DN82-303）和ZonMW VICI（016.130.606）资助。

1，原生质球的制备<ol><li>过夜孵育的酵母在30℃下在10毫升的实验设计来确定适当的介质中。 </li><li>一天后，用光度计测定培养物的光密度在600nm（OD 600），在相同的培养基稀释细胞至OD 600为0.2-0.4，并将它们生长至指数生长期除非的设计实验需要不同的条件。细胞处于指数生长期时，培养具有OD 600为1-2。 </li><li>收集离心10 OD 600细胞等价物，在3,500 xg离心5分钟，在50毫升的试管。离心后，弃上清。 </li><li>重悬细胞沉淀在5毫升100毫摩尔PIPES（pH 9.6）中，10 mM的二硫苏糖醇温育于30℃下10分钟。 </li><li>通过离心再次收集细胞，在3,500 xg离心5分钟。弃上清。 </li><li>悬浮细胞我ñ5毫升培养基（由实验的​​设计确定）含1M山梨醇和5mg溶解酶，并在30℃孵育混合物，轻轻摇动30分钟。 </li><li>离心细胞悬浮液以300×g离心5分钟，收集沉淀级分，其对应于球状体。弃上清。 </li><li>重悬原生质球在960微升冰冷的培养基（培养基由实验的设计确定）含1M山梨糖醇，将混合物转移至冰冷的2 ml离心管。 </li></ol> 2，吸收纳米金<ol><li>使用移液管，轻轻混合的原生质球用4纳摩尔再悬浮于40μl水中带正电的纳米金颗粒。该混合物的终体积必须为1毫升。 </li><li>放置在冰上将得到的细胞悬浮液15分钟。 </li><li>转移的悬浮液在室温下孵育它为所需要的时间，以允许内化纳米金通过电子邮件ndocytosis。 注:一个5分钟的摄取将主要导致内吞囊泡和早期内涵体室的标签，而15分钟的摄取将允许标记整个内体系统（早期和晚期内涵体）。潜伏期较长时间（超过30分钟）也将允许标注液泡。 注:脉冲追踪标记实验中不能用这种方法进行的。因此，LE的分析标记时，纳米金也将在下午和INEE发现。 </li></ol> 3，固定和切片<ol><li>加入1 ml的双重强度固定液停止纳米金摄取[4％多聚甲醛（PFA），0.4％的戊二醛（GA）在0.1M PHEM缓冲液（20mM PIPES，50mM的HEPES，pH值6.9，20mM的EGTA，4毫摩尔MgCl 2）]含1M山梨醇在细胞悬浮液。保持在管中于室温下进行。 </li><li>在30分钟轻轻手动反转micocentrifuge管数次（这将允许保持在原生质球悬浮液），然后离心两次在1700×g离心25秒。 </li><li>通过丢弃上清，加入1ml新鲜的标准强度固定液（2％PFA中，0.2％GA在0.1M PHEM缓冲液）含有1M山梨糖醇，孵育2小时，在室温下在缓慢旋转轮更换固定液。 </li><li>如前面所述6处理细胞冷冻切片。注:对于吸收纳米金，银增强程序，在指定的协议描述的周期性酸处理并没有被执行。周期性酸处理是为了更好地透化细胞壁26，但这种结构也球状体的生成过程中被消除。 </li><li>切50nm的薄的冷冻切片在-120℃下干燥钻石刀使用UCT超薄切片机如前所述8，并将其放置在涂福尔瓦碳50啮合镍网格。 </li></ol> 4，银增强的纳米金粒子Visualization 注意:要准备反应混合物中的程序基本上是由制造商所指示的之一。使用此协议的实际适应性，使得银增强程序，有效和可靠的冰冻切片。 <ol><li>从冰箱中取出银增强套件和解冻它在37℃培养箱或水浴。解冻时，将试剂盒在24℃培养箱放置到一个黑暗的房间里，直到使用（见4.7）。 </li><li>将加热板在黑暗的房间，并加热到24℃，终温</li><li>覆盖加热板的顶部表面的保护，有光泽的一面朝上，并用胶带将其固定。 </li><li>将封口膜对Benchkote并标记边缘用黑色标记，以能够看到的封口膜边在黑暗中。 </li><li>磁带上的​​Benchkote顶部的温度计监测加热板的温度。逐步调整温度，如果不是24°C。 </li><li>解放军CE上的50ml管中用双蒸水和内部的银增强试剂盒。 </li><li>填充小培养皿用双蒸水，预加热至37℃。放置在网格中的水，试样面朝下，持续30分钟。 </li><li>重复此4.7步一个更多的时间。 </li><li>传送网格，储物箱，将他们的黑暗的房间。 </li><li>由上几滴双蒸水在24℃下放置于封口膜已被固定在加热板传递它们（与试样的一面朝下）再次冲洗的网格。 </li><li>确保9个附加双蒸水滴准备上的封口膜，用于冲洗的银增强反应后。 </li><li>关灯，并确保所有灯都熄灭。打开红灯。 </li><li>取出从24℃培养箱中银增强试剂盒的A和B的解决方案。 </li><li>把第一个6滴溶液中，然后加入6滴乙溶液在1.5米升微量离心管中，并用玻璃巴斯德吸管，避免使气泡拌匀。该解决方案是很厚，它有手动，最后才涡旋它简单地用吸管混合。 </li><li>把A和B的解决方案早在24℃培养箱中培养，并采取了C溶液。 </li><li>加入6滴了C解决了A / B混合。然后先混合再一次用巴氏吸管，然后通过涡旋，避免气泡。 </li><li>通过将滴（〜20微升/滴）上的封口膜立即使用最终混合物。 </li><li>用干净，防磁镊子，将网格的混合物（ 如 。，银增强反应）为6〜15 min，取决于希望得到增强（ 即 ，大小的金粒）的。 </li><li>从混合物中取出电网，并通过他们的双蒸水的顶部下降在24℃下洗涤。首先，在使用快速连续6滴，然后3滴，每滴7分钟孵育时间，开灯之前。 </li><li>继续下一个步骤，以可视化的结果为EM调查或者免疫金标或膜染色27。 </li></ol>

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The authors have nothing to disclose.

免疫电子显微术是一种技术，它允许结合蛋白的定位与在这些蛋白质所在的载体和细胞器的超微结构分辨率。研究了酵母核内体系统时，因为它的车厢出现点状结构，在荧光显微镜中，这一点尤其重要。因此它是难以区分它们。由于这个原因，用一个探针通过EM检测并输入的内吞途径中一个时间依赖的方式是非常重要的特异性标记不同内涵体。这种类型的探针是有价值的，以评估细胞内吞作用...

内体系统中起着至关重要的多种细胞的作用包括贩运溶酶体酶分选受体和质膜（PM）的回收受体1,2的主要膜分拣设备。内涵体被划分在三个不同的车厢， 即 。在早期内涵体（EE），晚期内涵体（LE）和回收内涵体。这种分类是基于它需要的内吞物质到达他们，对特定的标记蛋白，并且在其形态的时间。膜， 即蛋白质和脂质双层，从PM内在设计可以通过内体输送到溶酶体中降解或再循环回。膜也被输送到内涵体从高尔基体，同样，要么继续溶酶体或检索返回到高尔基体。此外，蛋白质可以分为管腔囊泡从内体界膜，这一过程导致形成向内出芽子类别LE时，多泡体。 酵母胞内体系统是相对较不复杂的较高的真核细胞中的一个。酵母胞内体分为EE和LE。相反，哺乳动物细胞中不含有回收内涵体，而且组织特异性溶酶体相关细胞器。因此，他们有一个不太复杂的内体的贩运路线3,4的网络。因此，酵母曾代表仍然表示一个有利的实验系统，研究一些基础原则膜交通的内体系统。这个优点是通过以下事实，涉及该内体途径的许多基因已被初步分离与遗传筛选在酵母5强调。 而酵母胞内体系统中的野生型和突变的细胞已被广泛采用的生化和荧光显微镜方法的研究，其在超微结构水平调查至今只有最小的。形态分析是特别相关的酵母，因为大部分核内体的细胞器被检测为通过荧光显微镜圈点结构，这使得他们难以明确识别6。不幸的是抗血清仅有限数量识别酵母胞内蛋白标志物是工作在免疫电子显微镜（IEM）制剂7-10。对于一些蛋白质，这个问题已经绕过由感兴趣的基因的内源性标记和使用的抗体识别，检测它7,11,12和标签。然而，通常蛋白是因为它们的低表达水平的检测不到的IEM。他们的表达是不是一个解决方案，因为这种方法可以诱导误本地化和/或改建中的细胞器形态/功能。因此，内吞区室中与探针可检测通过电子显微镜EM标签是一种有效的选择。这是特别是如果一个最优解的公关OBE是进入细胞内吞途径中以时间依赖性方式，它允许知道什么时候会标记一个特定的细胞器6。 带正电荷的纳米级金的酵母原生质球（ 即 ，酵母，其中细胞壁已酶促去除）的摄取已经被成功地用于识别酵母胞内隔室10。这些颗粒牢固结合到带负电荷的脂质构成的生物膜。因此，带正电荷的纳米级金的联系人与PM，穿透细胞通过内吞作用和通过EE和LE到达液泡之前。这些小的金颗粒，但是，没有一个合适的尺寸由EM待观察。以使它们可见，它们的大小可通过化学反应，导致银或金的周围的黄金探测器13-15的沉积被放大。我们已经开发并成功应用的IEM方法的基础上，得安方法来执行的亚细胞定位研究8,16。这种方法允许进行酵母制剂免疫标记与形态8,17-24一个出色的分辨率。我们还建立了一个程序结合本IEM协议与酵母核内体系统的纳米金标记舱6。使用这种方法，我们已形态学特征内涵体的不同亚类和超微结构研究突变体与内体运输缺陷6,25。此外，我们已经证明，这种纳米金标记可以与免疫标号提供探索对不同亚群内体目标蛋白的分布的可能性相结合。在这里，我们介绍如何在酵母核内体系统的带正电的纳米金标记实际上是执行。

<table><tbody><tr><td>PIPES (piperazin-1,4-bis (2-ethansulfensaure)</td><td>Merck, Darmstadt, Germany</td><td>1,102,200,250</td><td></td></tr><tr><td>HEPES</td><td>Merck, Darmstadt, Germany</td><td>391,340</td><td></td></tr><tr><td>EGTA</td><td>Sigma, St louis, MO</td><td>E4378</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Merck, Darmstadt, Germany</td><td>1,058,330,250</td><td></td></tr><tr><td>DL-Dithiothreitol</td><td>Sigma, St louis, MO</td><td>D0632</td><td></td></tr><tr><td>Sorbitol</td><td>Sigma, St louis, MO</td><td>S1876</td><td></td></tr><tr><td>Positively charged Nanogold</td><td>Nanoprobes, Yaphank, NY</td><td>2022</td><td>store at -20 &amp;deg;C</td></tr><tr><td>HQ-silver&amp;trade; enhancement kit</td><td>Nanoprobes, Yaphank, NY</td><td>2012</td><td>store at -20 &amp;deg;C</td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Sigma, St louis, MO</td><td>441244</td><td></td></tr><tr><td>Glutaraldehyde 8% EM grade</td><td>Polyscience, Inc., Warrington, PA</td><td>216</td><td>store at -20 &amp;deg;C</td></tr><tr><td>Lytic enzyme</td><td>MP Biomedicals, Santa Ana, CA</td><td>153526</td><td>store at -20 &amp;deg;C</td></tr><tr><td>Parafilm M</td><td>Sigma, St louis, MO</td><td>P7793</td><td></td></tr><tr><td>50 meshes nickel grids&amp;nbsp;</td><td>Agar Scientific, Stansted, United Kingdom</td><td>G209N</td><td></td></tr><tr><td>Cryo-immuno diamond knife 35&amp;deg;</td><td>Diatome AG, Biel, Switzerland</td><td>N.A.</td><td></td></tr><tr><td>UCT ultramicrotome&amp;nbsp;</td><td>Leica, Solms, Germany</td><td>N.A.</td><td></td></tr><tr><td>Formvar 15/95 resine</td><td>Electron Microscopy Sciences, Hatfield, PA</td><td>15800</td><td></td></tr><tr><td>50 meshes nickel grids&amp;nbsp;</td><td>Agar Scientific, Stansted, United Kingdom</td><td>AGG2050N</td><td></td></tr><tr><td>Parafilm</td><td>Sigma, St louis, MO</td><td>P7793</td><td></td></tr><tr><td>Grids storage box</td><td>Leica, Solms, Germany</td><td>162-50</td><td></td></tr><tr><td>Falcon 100 mm x 15 mm Petri dish</td><td>Corning, Corning, NY</td><td>351029</td><td></td></tr><tr><td>Pasteur capillary pipettes 150 mm</td><td>Van Bruggen Glas, Rotterdam, The Netherlands</td><td>10216234</td><td></td></tr><tr><td>1.5 ml microcentrifuge&amp;nbsp;</td><td>Sarstedt, N&amp;uuml;mbrecht, Germany</td><td>72690001</td><td></td></tr><tr><td>50 ml tube</td><td>Bio-one, Alphen aan den Rijn, The Netherlands</td><td>210296</td><td></td></tr><tr><td>Benchkote surface protector</td><td>Whatman, Maidstone, United Kingdom&amp;nbsp;</td><td>1159201</td><td></td></tr></tbody></table>

nanogold labeling of the yeast endosomal system for ultrastructural analyses

内涵体是主要的膜排序在真核细胞中的检查点中的一个和它们调节回收的蛋白质或毁坏大多从质膜和高尔基体。其结果是内体系统在维持细胞稳态中发挥中心作用，并在属于该网络的细胞器通过小泡运输相互连接的基因的突变，引起严重的病症，包括癌症和神经生物学障碍。因此，黄金的相关性是理解内涵体系统的生物合成和组织相关的机制。 酿酒酵母中一直举足轻重的完成这项任务。要明确标示，并在超微结构水平这一​​模式生物的内体系统的分析，我们在座的带正电荷的纳米金摄取原生质其次是这些粒子通过银增强反应可视化的详细协议。这种方法也是一种宝贵的太l适用于与内体运输缺陷突变体的形态学检查。此外，它不仅适用于超微结构检查，但它也可与免疫标号用于蛋白质定位的调查相结合。

根据所提出的协议，酵母核内体系统的形态可以是由接入传输的EM。 图1显示了不同类型的内体区室中已达成，因此，标记的纳米金。银增强纳米金可以清楚地看到作为电子致密颗粒。建立了银增强反应的最佳设置允许具有在5至15纳米，不与酵母细胞的超微结构的干扰之间的均匀粒度范围的金颗粒。质膜以及早期内涵体车厢都可以访问的纳米金5分钟孵化在24°C（ 图1A）后?...

Watch this Scientific Journal Video about Nanogold Labeling of the Yeast Endosomal System for Ultrastructural Analyses at JoVE.com

Nanogold Labeling of the Yeast Endosomal System for Ultrastructural Analyses

酵母， 酿酒酵母 ，一直是一个关键模式生物识别和研究基因调节内体系统的生物合成和功能。在这里，我们提出了内体车厢的超微结构研究特定的标签一个详细的协议。

酵母核内体系统的超微结构分析的纳米金标记

Endosomes are one of the major membrane sorting checkpoints in eukaryotic cells and they regulate recycling or destruction of proteins mostly from the ...

nanogold-labeling-yeast-endosomal-system-for-ultrastructural

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

10.2K 次观看.  University Medical Center Utrecht. 酵母， 酿酒酵母 ，一直是一个关键模式生物识别和研究基因调节内体系统的生物合成和功能。在这里，我们提出了内体车厢的超微结构研究特定的标签一个详细的协议。 

视频: Nanogold Labeling of the Yeast Endosomal System for Ultrastructural Analyses

A high-throughput method to globally study the organelle morphology in S. cerevisiae

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help achieve an comprehensive understanding of molecular pathways. In Saccharomyces cerevisiae, with the development of the non-essential gene deletion array, we can globally study the morphology of different organelles like the endoplasmic reticulum (ER) and the mitochondria using GFP (or variant)-markers in different gene backgrounds. However, incorporating GFP markers in each single mutant individually is a labor-intensive process. Here, we describe a procedure that is routinely used in our laboratory. By using a robotic system to handle high-density yeast arrays and drug selection techniques, we can significantly shorten the time required and improve reproducibility. In brief, we cross a GFP-tagged mitochondrial marker (Apc1-GFP) to a high-density array of 4,672 nonessential gene deletion mutants by robotic replica pinning.  Through diploid selection, sporulation, germination and dual marker selection, we recover both alleles. As a result, each haploid single mutant contains Apc1-GFP incorporated at its genomic locus. Now, we can study the morphology of mitochondria in all non-essential mutant background. Using this high-throughput approach, we can conveniently study and delineate the pathways and genes involved in the inheritance and the formation of organelles in a genome-wide setting.

GFP-fusion proteins are widely used to visualize organelles by confocal microscopy. However, screening for mutations that affect the morphology of organelles generally requires individual mutagenesis and is time consuming. Here, we demonstrate a method to simultaneously incorporate organelle-GFP markers in almost 5,000 non-essential genes in yeast.

一种高通量的方法，在全球范围内研究在酿酒酵母中的细胞器形态

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help ...

科研

生物学

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

嵌入后突触蛋白的免疫标记海马切片培养

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

神经科学

Application of Fluorescent Nanoparticles to Study Remodeling of the Endo-lysosomal System by Intracellular Bacteria

Fluorescent nanoparticles (NPs) with desirable chemical, optical and mechanical properties are promising tools to label intracellular organelles. Here, we introduce a method using gold-BSA-rhodamine NPs to label the endo-lysosomal system of eukaryotic cells and monitor manipulations of host cellular pathways by the intracellular pathogen Salmonella enterica. The NPs were readily internalized by HeLa cells and localized in late endosomes/lysosomes. Salmonella infection induced rearrangement of the vesicles and accumulation of NPs in Salmonella-induced membrane structures. We deployed the Imaris software package for quantitative analyses of confocal microscopy images. The number of objects and their size distribution in non-infected cells were distinct from the ones in Salmonella-infected cells, indicating extremely remodeling of the endo-lysosomal system by WT Salmonella.

This article describes methods for the synthesis and fluorescent labeling of nanoparticles (NPs). The NPs were applied in pulse-chase experiments to label the endo-lysosomal system of eukaryotic cells. Manipulation of the endo-lysosomal system by activities of the intracellular pathogen Salmonella enterica were followed by live cell imaging and quantified.

通过胞内细菌，研究内溶酶体系统的改造荧光纳米粒子中的应用

Fluorescent nanoparticles (NPs) with desirable chemical, optical and mechanical properties are promising tools to label intracellular organelles. Here, we ...

免疫与感染

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A mitochondrial genome of baker&#8217;s yeast encodes eight proteins: three subunits of the cytochrome c oxidase (Cox1p, Cox2p, and Cox3p), three subunits of the ATP synthase (Atp6p, Atp8p, and Atp9p), a subunit of the ubiquinol-cytochrome c oxidoreductase enzyme, cytochrome b (Cytb), and mitochondrial ribosomal protein Var1p. The purpose of the method described here is to specifically label these proteins with 35S methionine, separate them by electrophoresis and visualize the signals as discrete bands on the screen. The procedure involves several steps. First, yeast cells are cultured in a galactose-containing medium until they reach the late logarithmic growth stage. Next, cycloheximide treatment blocks cytoplasmic translation and allows 35S methionine incorporation only in mitochondrial translation products. Then, all proteins are extracted from yeast cells and separated by polyacrylamide gel electrophoresis. Finally, the gel is dried and incubated with the storage phosphor screen. The screen is scanned on a phosphorimager revealing the bands. The method can be applied to compare the biosynthesis rate of a single polypeptide in the mitochondria of a mutant yeast strain versus the wild type, which is useful for studying mitochondrial gene expression defects. This protocol gives valuable information about the translation rate of all yeast mitochondrial mRNAs. However, it requires several controls and additional experiments to make proper conclusions.

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Baker&#8217;s yeast mitochondrial genome encodes eight polypeptides. The goal of the current protocol is to label all of them and subsequently visualize them as separate bands.

贝克酵母糖精中线粒体基因组表达产品的标记和可视化

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A ...

生物化学

Visualizing Yeast Organelles with Fluorescent Protein Markers

The budding yeast, Saccharomyces cerevisiae, is a classic model system in studying organelle function and dynamics. In our previous works, we have constructed fluorescent protein-based markers for major organelles and endomembrane structures, including the nucleus, endoplasmic reticulum (ER), Golgi apparatus, endosomes, vacuoles, mitochondria, peroxisomes, lipid droplets, and autophagosomes. The protocol presented here describes the procedures for using these markers in yeast, including DNA preparation for yeast transformation, selection and evaluation of transformants, fluorescent microscopic observation, and the expected outcomes. The text is geared toward researchers who are entering the field of yeast organelle study from other backgrounds. Essential steps are covered, as well as technical notes about microscope hardware considerations and several common pitfalls. It provides a starting point for people to observe yeast subcellular entities by live-cell fluorescent microscopy. These tools and methods can be used to identify protein subcellular localization and track organelles of interest in time-lapse imaging.

Here we describe the use of a set of fluorescent protein-based organelle markers in live-cell imaging of the budding yeast, Saccharomyces cerevisiae.

The budding yeast, Saccharomyces cerevisiae, is a classic model system in studying organelle function and dynamics. In our previous works, we have ...

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 ...

Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal remodeling and vesicular trafficking in living cells. Quantitative methodologies using chimeric GFP fusions have been developed for many applications; however, GFP is somewhat resistant to proteolysis, thus its fluorescence persists in the lysosome/vacuole, which can impede quantification of cargo trafficking in the endocytic pathway. An alternative method for quantifying endocytosis and post-endocytic trafficking events makes use of superecliptic pHluorin, a pH-sensitive variant of GFP that is quenched in acidic environments. Chimeric fusion of pHluorin to the cytoplasmic tail of transmembrane cargo proteins results in a dampening of fluorescence upon incorporation of the cargo into multivesicular bodies (MVBs) and delivery to the lysosome/vacuole lumen. Thus, quenching of vacuolar fluorescence facilitates quantification of endocytosis and early events in the endocytic pathway. This paper describes methods using pHluorin-tagged cargos for quantification of endocytosis via fluorescence microscopy, as well as population-based assays using flow cytometry.

Accurate quantification of vesicular trafficking events often provides key insights into roles for specific proteins and the effects of mutations. This paper presents methods for using superecliptic pHluorin, a pH-sensitive GFP variant, as a tool for quantification of endocytic events in living cells using quantitative fluorescence microscopy and flow cytometry.

pHluorin的申请在芽殖酵母细胞内吞的定量，动力学和高通量分析

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal ...

Plunge Freezing: A Tool for the Ultrastructural and Immunolocalization Studies of Suspension Cells in Transmission Electron Microscopy

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize macromolecular complexes at very high resolution. However, to get as close as possible to the native state, perfect sample preservation is required. Conventional electron microscopy (EM) fixation with aldehydes, for instance, does not provide good ultrastructural preservation. The slow penetration of fixatives induces cell reorganization and loss of various cell components. Therefore, conventional EM fixation does not allow for an instantaneous stabilization and preservation of structures and antigenicity. The best choice for examining intracellular events is to use cryofixation followed by the freeze-substitution fixation method that keeps cells in their native state. High-pressure freezing/freeze-substitution, which preserves the integrity of cellular ultrastructure, is the most commonly used method, but requires expensive equipment. Here, an easy-to-use and low-cost freeze fixation method followed by freeze-substitution for suspension cell cultures is presented.

This manuscript describes an easy-to-use and low-cost cryofixation method for visualizing suspension cells by transmission electron microscopy.

暴跌冻结:在透射电子显微镜的超微结构和免疫组化研究工具悬架细胞

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize ...

4D Microscopy of Yeast

The goal of this protocol is to characterize how membrane compartments form and transform in live cells of budding yeast. Many intracellular compartments in yeast are dynamic, and a full understanding of their properties requires time-lapse imaging. Multi-color 4D confocal fluorescence microscopy is a powerful method for tracking the behavior and composition of an intracellular compartment on a time scale of 5-15 min. Rigorous analysis of compartment dynamics requires the capture of thousands of optical sections. To achieve this aim, photobleaching and phototoxicity are minimized by scanning rapidly at very low laser power, and the pixel dimensions and Z-step intervals are set to the largest values that are compatible with sampling the image at full resolution. The resulting 4D data sets are noisy but can be smoothed by deconvolution. Even with high quality data, the analysis phase is challenging because intracellular structures are often numerous, heterogeneous, and mobile. To meet this need, custom ImageJ plugins were written to array 4D data on a computer screen, identify structures of interest, edit the data to isolate individual structures, quantify the fluorescence time courses, and make movies of the projected Z-stacks. 4D movies are particularly useful for distinguishing stable compartments from transient compartments that turn over by maturation. Such movies can also be used to characterize events such as compartment fusion, and to test the effects of specific mutations or other perturbations.

This protocol describes the analysis of fluorescently labeled intracellular compartments in budding yeast using multi-color 4D (time-lapse 3D) confocal microscopy. The imaging parameters are chosen to capture adequate signals while limiting photodamage. Custom ImageJ plugins allow labeled structures to be tracked and quantitatively analyzed.

酵母的4D显微镜

The goal of this protocol is to characterize how membrane compartments form and transform in live cells of budding yeast. Many intracellular compartments ...

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 ...