The authors thank Ren&#233; Scriwanek for the assistance with the preparation of the Figure. F.R. is supported by ECHO (700.59.003), ALW Open Program (821.02.017 and 822.02.014), DFG-NWO cooperation (DN82-303) and ZonMW VICI (016.130.606) grants.

1. Spheroplast Preparation
<ol>
	<li>Incubate the yeast overnight at 30 &#176;C in 10 ml of the appropriate medium determined by the design of the experiment.</li>
	<li>The day after, measure the optical density of the culture at 600 nm (OD600) using a photometer, Dilute cells in the same culture medium to an OD600 of 0.2-0.4 and grow them to an exponential growth phase unless the design of the experiment requires a different condition. Cells are in the exponential phase when the culture has an OD600 of 1-2.</li>
	<li>Collect 10 OD600 equivalents of cells by centrifugation at 3,500 x g for 5 min in a 50 ml tube. After centrifugation, discard the supernatant.</li>
	<li>Resuspend the cell pellet in 5 ml of 100 mM PIPES (pH 9.6), 10 mM dithiothreitol and incubated at 30 &#176;C for 10 min.</li>
	<li>Collect again the cells by centrifugation at 3,500 x g for 5 min. Discard the supernatant.</li>
	<li>Resuspend the cells in 5 ml of medium (determined by the design of the experiment) containing 1 M sorbitol and 5 mg of lytic enzyme, and incubate the mixture at 30 &#176;C with gentle shaking for 30 min.</li>
	<li>Centrifuge the cell suspension at 300 x g for 5 min to collect the pellet fraction, which corresponds to the spheroplasts. Discard the supernatant.</li>
	<li>Resuspend the spheroplasts in 960 &#181;l of ice cold media (medium determined by the design of the experiment) containing 1 M sorbitol and transfer the mixture into an ice cold 2 ml microcentrifuge tube.</li>
</ol>
2. Nanogold Uptake
<ol>
	<li>Using a pipette, gently mix the spheroplast with 4 nmol of positively charged nanogold particles resuspended in 40 &#181;l of water. The final volume of the mixture has to be of 1 ml.</li>
	<li>Place the obtained cell suspension on ice for 15 min.</li>
	<li>Transfer the suspension at room temperature and incubate it for the required time to allow nanogold internalization by endocytosis. 
	NOTE: A 5 min uptake will principally lead to the labelling of endocytic vesicles and early endosomal compartments, while a 15 min uptake will permit to mark the entire endosomal system (early and late endosomes). Longer incubation times (more than 30 min) will also permit to label the vacuole. 
	NOTE: Pulse-chase labelling experiments cannot be performed with this method. Therefore, when labelling for the analysis of LE, nanogold will also be found at the PM and inEE.</li>
</ol>
3. Fixation and Sectioning
<ol>
	<li>Stop the nanogold uptake by adding 1 ml of double strength fixative [4% paraformaldehyde (PFA), 0.4% glutaraldehyde (GA) in 0.1 M PHEM buffer (20 mM PIPES, 50 mM HEPES, pH 6.9, 20 mM EGTA, 4 mM MgCl2)] containing 1 M sorbitol to the cell suspension. Kept the tube at room temperature.</li>
	<li>Gently invert manually the micocentrifuge tubes several times during 30 min (this will permit to keep the spheroplasts in suspension) and then centrifuge it twice at 1,700 x g for 25 sec.</li>
	<li>Replace the fixative by discarding the supernatant and by adding 1 ml of fresh standard strength fixative (2% PFA, 0.2% GA in 0.1 M PHEM buffer) containing 1 M sorbitol and incubate for 2 hr at room temperature on a slowly rotating wheel.</li>
	<li>Process the cells for cryo-sectioning as described previously6. NOTE: For the nanogold uptake and silver enhancement procedures, the periodic acid treatment described in the indicated protocol has not to be performed. The periodic acid treatment is to better permeabilize the cell wall26 but this structure has been eliminated during the generation of spheroplasts.</li>
	<li>Cut 50 nm thin cryosections at -120 &#176;C with dry diamond knife using an UCT ultramicrotome as previously described8 and place them on Formvar carbon coated 50 meshes nickel grids.</li>
</ol>
4. Silver Enhancement for Nanogold Particle Visualization
NOTE: The procedure to prepare the reaction mixtures is basically the one indicated by the manufacturer. Practical adaptations using this protocol, makes the silver enhancement procedure effective and reliable on cryosections.
<ol>
	<li>Remove the silver enhancement kit from the freezer and thaw it in a 37 &#176;C incubator or water bath. When thawed, place the kit in a 24 &#176;C incubator placed into a dark room until use (see 4.7).</li>
	<li>Place the heating plate in the dark room and warm it to the final temperature of 24 &#176;C.</li>
	<li>Cover the top of heating plate with the surface protector, shiny side up, and secure it with a tape.</li>
	<li>Place the Parafilm on the Benchkote and mark the edges with a black marker to be able to see the Parafilm edges in the dark.</li>
	<li>Tape a thermometer on the top of the Benchkote to monitor the temperature of the heating plate. Gradually adjust the temperature if not 24 &#176;C.</li>
	<li>Place the 50 ml tube with double distilled water and the silver enhancement kit inside.</li>
	<li>Fill the small Petri dishes with double distilled water, pre-warmed at 37 &#176;C. Place the grids in the water, specimen side down, for 30 min.</li>
	<li>Repeat this 4.7 step one more time.</li>
	<li>Transfer the grids in the storage box and bring them to the dark room.</li>
	<li>Rinse the grids again by passing them (with the specimen side down) on several drops of double distilled water at 24 &#176;C placed on the Parafilm that has been fixed on the heating plate.</li>
	<li>Make sure that 9 additional double distilled water drops are ready on the Parafilm, for the rinsing after the silver enhancement reaction.</li>
	<li>Turn the light off and make sure that all lights are off. Turn the red light on.</li>
	<li>Take out the A and B solutions of the silver enhancement kit from the 24 &#176;C incubator.</li>
	<li>Put first 6 drops of the A solution and then 6 drops of the B solution in a 1.5 ml microcentrifuge tube, and mix well with a glass Pasteur pipette avoiding to make bubbles. The solution is quite thick and it has to be manually mixed with a pipette before finally vortexing it briefly.</li>
	<li>Put the A and B solutions back in the 24 &#176;C incubator and take out the C solution.</li>
	<li>Add 6 drops of the C solution to the A/B mix. Then mix again first with a Pasteur pipette and then by vortexing, avoid making bubbles.</li>
	<li>Use immediately the final mixture by placing drops (~20 &#181;l/drop) on the Parafilm.</li>
	<li>With a clean, anti-magnetic tweezers, place the grids on the mixture (e.g., silver enhancing reaction) for 6 to15 min depending of the enhancement (i.e. size of the gold particles) wished to be obtained.</li>
	<li>Remove the grids from the mixture and pass them on the top of the double distilled water drops at 24 &#176;C for washes. First, use in rapidly succession 6 drops and then 3 drops with 7 min incubation time per drop, before turning the lights on.</li>
	<li>Proceed to the next step, either immuno-gold labeling or membrane staining27 in order to visualize the result for an EM investigation.</li>
</ol>

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The authors have nothing to disclose.

Immuno-electron-microscopy is a technique that allows combining localization of proteins with the ultrastructural resolution of the carriers and organelles where these proteins reside. This is particularly crucial when studying the yeast endosomal system because its compartments appear as punctuate structures in fluorescence microscopy. It is therefore difficult to distinguish them. For this reason the use of a probe detectable by EM and entering the endocytic route in a time dependent manner is crucial to specifically m...

The endosomal system is a major membrane sorting apparatus that plays multiple crucial cellular roles including trafficking of lysosomal enzyme sorting receptors and the recycling of plasma membrane (PM) receptors1,2. Endosomes are divided in three different compartments, i.e. the early endosomes (EE), the late endosomes (LE) and the recycling endosomes. This classification is based on the time it takes for endocytosed material to reach them, on specific marker proteins and on their morphology. Membranes, i.e. protein and lipid bilayers, internalized from the PM can either be delivered to lysosomes via endosomes for degradation or be recycled back. Membranes are also transported to endosomes from the Golgi and similarly, either continue to lysosomes or be retrieved back to the Golgi. Furthermore, proteins can be sorted into luminal vesicles budding inward from the endosomal limiting membrane, a process that leads to the formation of a subcategory of LE, the multivesicular bodies.
The yeast endosomal system is relatively less complex than the one of high eukaryotic cells. Yeast endosomes are divided into EE and LE. In contrast to mammalian cells they do not contain recycling endosomes but also tissue-specific lysosome-related organelles. Consequently they have a less complex network of endosomal trafficking routes3,4. Therefore yeast has represented and still represents an advantageous experimental system to study some of the principles underlying membrane traffic in the endosomal system. This advantage is emphasized by the fact that numerous genes involved in the endosomal pathways have been initially isolated with genetic screens in yeast5. While the yeast endosomal system in wild type and mutant cells has been extensively studied using biochemical and fluorescence microscopy approaches, its investigation at the ultrastructural level has only been minimal. Morphological analyses are particularly relevant in yeast because most of the endosomal organelles are detected as punctuate structures by fluorescence microscopy, which make difficult their unequivocal identification6. Unfortunately only a limited number of antisera recognizing yeast endosomal protein markers is working in immuno-electron-microscopy (IEM) preparations7-10. For some proteins, this problem has been circumvented by the endogenous tagging of the gene of interest and the use of an antibody recognizing the tag to detect it7,11,12. Often, however, proteins are undetectable by IEM because of their low expression levels. Their overexpression is not a solution because this approach can induce mis-localizations and/or alterations in the organelle morphology/functions. Thus the labelling of the endocytic compartments with a probe detectable by electron microscopy EM is an effective option. This is an optimal solution especially if the probe is entering the endocytic route in a time-dependent manner, which allows to know when it will mark a specific organelle6.
The uptake of positively charged nanogold by yeast spheroplasts (i.e. yeast where the cell wall has been enzymatically removed) has successfully been used to identify the yeast endosomal compartments10. These particles strongly bind to the negatively charged lipids composing the biological membranes. Thus the positively charged nanogold associates with the PM, penetrates the cell by endocytosis and passes through the EE and LE before reaching the vacuole. These small gold particles, however, do not have an appropriate size to be seen by EM. To render them visible, their size can be enlarged by chemical reactions that lead to the deposition of silver or gold around the gold probe13-15. We have developed and successfully applied an IEM approach based on the Tokuyasu method to perform subcellular localization studies8,16. This method allows performing immunogold labelling on yeast preparations with an excellent resolution of the morphology8,17-24. We have also established a procedure combining this IEM protocol with the nanogold labelling of the yeast endosomal system compartments6. Using this approach we have morphologically characterized different subclasses of endosomes and ultrastructurally examined mutants with an endosomal trafficking defect6,25. Moreover, we have demonstrated that this nanogold labelling can be combined with immunogold labelings providing the possibility to explore the distribution of a protein of interest on the different endosome subpopulations. Here we present how the labelling of the yeast endosomal system with positively charged nanogold is practically performed.

<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

Endosomes are one of the major membrane sorting checkpoints in eukaryotic cells and they regulate recycling or destruction of proteins mostly from the plasma membrane and the Golgi. As a result the endosomal system plays a central role in maintaining cell homeostasis, and mutations in genes belonging to this network of organelles interconnected by vesicular transport, cause severe pathologies including cancer and neurobiological disorders. It is therefore of prime relevance to understand the mechanisms underlying the biogenesis and organization of the endosomal system. The yeast Saccharomyces cerevisiae has been pivotal in this task. To specifically label and analyze at the ultrastructural level the endosomal system of this model organism, we present here a detailed protocol for the positively charged nanogold uptake by spheroplasts followed by the visualization of these particles through a silver enhancement reaction. This method is also a valuable tool for the morphological examination of mutants with defects in endosomal trafficking. Moreover, it is not only applicable for ultrastructural examinations but it can also be combined with immunogold labelings for protein localization investigations.

According to the presented protocol, the morphology of the yeast endosome system can be access by transmission EM. Figure 1 shows different types of endosomal compartments that have been reached and therefore labeled with nanogold. The silver-enhanced nanogold can be clearly seen as electron dense particles. The optimal settings established for the silver enhancement reaction allows having gold particles in a homogenous size range between 5 and 15 nm, which does not interfere with the ultrastructure of t...

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

Yeast, Saccharomyces cerevisiae, has been a key model organism to identify and study genes regulating the biogenesis and functions of the endosomal system. Here we present a detailed protocol for the specific labeling of the endosomal compartments for ultrastructural studies.

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 Views.  University Medical Center Utrecht. Yeast, Saccharomyces cerevisiae, has been a key model organism to identify and study genes regulating the biogenesis and functions of the endosomal system. Here we present a detailed protocol for the specific labeling of the endosomal compartments for ultrastructural studies.

Video: 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.

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

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.

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

Neuroscience

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.

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

Immunology and Infection

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.

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

Biochemistry

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.

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.

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.

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.

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.

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 ...

Nanogold Labeling of the Yeast Endosomal System for Ultrastructural Analyses

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

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

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

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

Visualizing Yeast Organelles with Fluorescent Protein Markers

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

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

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

4D Microscopy of Yeast

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