This study is supported by an operating grant from the Canadian Institutes for Health Research to M.L.D. and P.L. The work presented here is supported by a John R. Evans Leader Fund award from the Canadian Foundation for Innovation and a matching fund from the Ontario Research Fund to P.L. Y.J. is supported by an MSc to PhD transfer scholarship from the Dean of the Schulich School of Medicine and Dentistry at The University of Western Ontario. S.D.G. is supported by a PhD Scholarship from ALS Canada.

1. Generation of New Fluorescently Tagged Httex1 Reporters for an Expression in Yeast
Note: This section has been modified from the protocol by Jiang et al.16 and Albakri et al.19.
<ol>
	<li>Design primers to amplify the sequence encoding the fluorescent protein or interest by PCR. The forward primer should include a leader sequence to assist the restriction enzyme during digestion (GATC), followed by a SpeI restriction site (ACTAGT) and 20 bases downstream of the ATG (excluding ATG) of the fluorescent protein gene of interest. The reverse primer should include the leader sequence (GATC), followed by a SalI restriction site (GTCGAC) and the reverse complement of 20 bases upstream of the stop codon of the FP sequence (including stop).</li>
	<li>Using the primers designed in step 1.1, perform the PCR reaction using a thermocycler with the following settings: heat to 95 &#176;C for 1 min and cycle at 95 &#176;C for 30 s, 60 &#176;C for 30 s, and 72 &#176;C for 2 min per kB of PCR product. Performing 18 cycles is ample.</li>
	<li>Run the PCR reaction on an agarose gel (0.5% in Tris-acetate-EDTA). There should a single band corresponding to the expected product size. Isolate the fragment using a gel purification kit.</li>
	<li>The protocol employs an Httex1 vector carrying 25 (nontoxic) and 72 (HD-associated, displaying a strong aggregation) polyQ repeats. Digest both the PCR fragments and the vector with SpeI and SalI restriction enzymes for 3 h at 37 &#176;C.</li>
	<li>Purify the digested vector by running it on an agarose gel as in step 1.3.</li>
	<li>Purify the digested PCR fragment using a PCR purification kit.</li>
	<li>Ligate the resulting digested PCR fragment and p415-GAL1-FLAG-25/72QpolyQ plasmids1 using T4 ligase (1 h at room temperature). Use a 10 &#181;L reaction (1 &#181;L of T4 enzyme, 1 &#181;L of 10X buffer, 6 &#181;L of PCR fragment, and 2 &#181;L of vector).</li>
	<li>Transform 2 &#181;L of the ligation reaction into 50 &#181;L of Escherichia coli-competent cells and incubate them on ice for 30 min. Then, heat-shock the cells at 42 &#176;C for 30 s. Add 1 mL of SOC outgrowth media and incubate at 37 &#176;C for 1 h in a shaker. Plate 200 &#181;L of the reaction on an LB-agar plate containing 100 &#181;g/mL ampicillin. Incubate the plate at 37 &#176;C overnight.</li>
	<li>Select three individual bacterial colonies, grow them overnight in 3 mL of LB-broth containing 100 &#181;g/mL ampicillin at 37 &#176;C in a shaker and extract the plasmid DNA using a plasmid purification kit.</li>
	<li>Check the plasmid by digesting 500 ng of DNA using SpeI and SalI restriction enzymes for 1 h at 37 &#176;C and run the reaction on an agarose gel (0.5% in Tris-acetate-EDTA). There should be two bands at the right sizes of the vector (~7 kb) and the insert (size varies according to the gene of interest). Then, verify the plasmid by sequencing.</li>
	<li>Transform the p415-GAL1-FLAG-polyQ-FP plasmids into the yeast strain W303 following a standard yeast transformation protocol2.</li>
</ol>
2. Spotting Assay
<ol>
	<li>Streak the yeast clones carrying 25Q/72Q tagged with the FP of interest on an agar plate containing yeast selection media (synthetic complete-SC without leucine) with glucose as the carbon source. At the same time, also streak 25Q/72Q-ymsfGFP to serve as a positive control. 
	Note: 25Q/72Q constructs that do not contain a fluorescent tag are not toxic and can serve as negative control.</li>
	<li>Incubate the plates at 30 &#176;C for 2 - 3 d.</li>
	<li>Select up to three single colonies from the plate.</li>
	<li>Inoculate 5 mL of SC supplemented with 2% glucose as the carbon source.</li>
	<li>Pellet 200 &#181;L of each overnight culture and wash it 3x with sterile distilled water.</li>
	<li>Resuspend the cells in SC media containing 2% galactose as the carbon source to induce the expression of polyQ fusions. Incubate the galactose media overnight at 30 &#176;C in a tube rotator. As a control, repeat this step by using glucose-containing media.</li>
	<li>The next morning, equalize the cell densities to optical density at 600 nm (OD600) of 0.2 in 100 &#181;L of SC media in a sterile 96-well plate.</li>
	<li>Prepare four fivefold dilutions of each sample with sterile water by pipetting 20 &#181;L of the sample from the previous well into 80 &#181;L of media in the next well.</li>
	<li>Use a yeast pinning tool to spot the cells onto selective plates (containing glucose or galactose) and incubate at 30 &#176;C for 2 d.</li>
	<li>Image the plates with an image documentation device.</li>
</ol>
3. Quantification of Cell Growth in Liquid Culture
<ol>
	<li>Prepare the cell cultures, following steps 2.1 - 2.5 of this protocol.</li>
	<li>Measure the OD600 using a spectrophotometer.</li>
	<li>Dilute the cells to an OD600 of 0.1 in 300 &#181;L of media in a 96-well plate.</li>
	<li>Run each sample in triplicate.</li>
	<li>Incubate the plate in a plate reader/incubator with shaking capabilities. Set the number of samples, the temperature at 30 &#176;C, the absorbance at 600 nm, the length of the experiments to 24 h, and the measurement intervals to 15 min, and select the continuous shaking mode.</li>
	<li>Create the growth curve and quantify the area under the curve using scientific graphing software. The GraphPad Prism 7 is recommended. Paste the data into an XY table with three replicate values. The growth curve will be shown under the Graphs folder at the left side. To quantify the area under the curve, select Analyze at the top left and click Area under curve in XY analyses.</li>
</ol>
4. Fluorescent Microscopy
<ol>
	<li>Prepare the cell cultures, following steps 2.1 - 2.5 of this protocol.</li>
	<li>Dilute the cells 10x in growth media and transfer 200 &#181;L of each sample to 8-well imaging chambers.</li>
	<li>Image the cells using a confocal microscope equipped with a 63X Plan Aprochromoat objective (1.4 NA) at room temperature. 
	Note: The usage of a confocal microscope is optional. A standard wide-field fluorescent microscope can also be employed.</li>
	<li>Adjust the pinhole and laser power for optimal image acquisition. Since the 72Q aggregates are much brighter than the diffuse 25Q signal, it is often required to use a different acquisition setting between the different plasmids in order to avoid saturation of the fluorescent signal.</li>
	<li>Process the images using ImageJ20 or another image-processing software. At this step, the percentage of cells that display aggregate can be calculated manually is desired.</li>
</ol>
5. Dot Blot
Note: In this protocol, dot blot is used to examine the protein expression levels. Prepare the cell cultures, following steps 2.1 - 2.5 of this protocol.
<ol>
	<li>Generate protein lysates using glass beads in lysis buffer (100 mM Tris, pH 7.5; 200 mM NaCl; 1 mM EDTA; 5% glycerol, 1 mM dithiothreitol [DTT]). Add protease inhibitors, 4 mM phenylmethylsulfonyl fluoride (PSMF) and protease inhibitor cocktail, directly before use. Pellet 5 mL of the overnight culture and resuspend it in 200 &#181;L of glass beads and 200 &#181;L of lysis buffer. Vortex 30 s for 12 rounds. Centrifuge at 12,000 x g at 4 &#176;C for 10 min and collect the supernatant.</li>
	<li>Spot an equal amount of total proteins on a nitrocellulose membrane using a microfiltration apparatus. Prewet the membrane with PBS and assemble the apparatus. Connect to a vacuum source and make sure the screws are tightened. Turn on the vacuum and let the sample filter through the membrane by gravity.</li>
	<li>Block the membrane in PBS- 0.05% Tween/5% fat-free milk.</li>
	<li>Incubate the membrane with primary anti-FLAG antibody overnight at 4 &#176;C. The monoclonal anti-FLAG M1 is recommended.</li>
	<li>Wash the membrane 3x for 10 min each with PSB- 0.05% Tween.</li>
	<li>Incubate the membrane with a fluorescently labeled secondary antibody (anti-mouse IgG) for 1 h at room temperature in PBS- 0.05% Tween/5% fat-free milk.</li>
	<li>Wash membrane 3x 10 min with PSB- 0.05% Tween.</li>
	<li>Image-blot using an immunoblot documentation system.</li>
</ol>

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The authors have nothing to disclose.

In this article, various assays to measure the aggregation of Httex1 polyQ expansions and their effect on yeast growth were employed as a model to study how different fluorescent proteins alter their fusion partners in the context of fluorescent reporters. Using a GFP variant (ymsfGFP) as a positive control, we showed that this detects significant changes in polyQ toxicity and aggregation between different fluorescent tags and allows for a direct and rapid comparison of the polyQ-FP fusion performance against ...

Since the initial characterization of the green fluorescent protein (GFP) from Aequorea victoria1, a wide palette of genetically encoded FPs have been developed, allowing cell biologists to simultaneously localize and track multiple cellular events/proteins in living cells2,3. FPs are derived from multiple organisms, from jellyfish to coral, and therefore, display specific biophysical properties that divert extensively beyond their respective fluorescent spectrum. These properties include brightness, photostability, and a tendency to oligomerize among others2,4. Selecting monomeric FPs is an important aspect in the selection of a suitable tag when designing a fluorescent reporter, in order to minimize inappropriate interactions and alterations of the fusion partner&#39;s function and maximize the reporter efficiency for a given cellular compartment4,5,6. While GFP has, over time, been evolved to minimize the effect of adding the fluorescent tag to the fusion partner5,7,8, how new FP variants perform compared to GFP remains difficult to assess.
Few methods exist to characterize the behavior of FPs. Most of them involve testing biophysical properties of FPs using biochemical approaches, such as ultracentrifugation and gel filtration protocols9,10,11,12. Such methods have the caveat of using purified FPs in solution, offering little insight into their behavior in intact cells. The development of the organized smooth endoplasmic reticulum (OSER) assay offers a quantifiable assessment of FPs&#39; tendency to oligomerize in living cells13 by testing the ability of overexpressed FPs to reorganize endoplasmic reticulum tubules into OSER whorls14. This technique can successfully detect changes between monomeric and oligomeric variants of GFP and other FPs. However, it relies mostly on overexpression in transiently transfected cells, and the quantitation and image analysis can be time-consuming unless the technique is adopted as an automated data collection and analysis workflow.
In order to complement these approaches, we established an assay that takes advantage of the effect of fluorescent tags on the toxicity and aggregation of polyQ expansions in yeast15,16. The expansion of the polyQ stretch with more than 36 repeats within the first exon of the gene encoding the huntingtin protein (Htt) is associated with Huntington&#39;s disease17,18. The expression of expanded Httex1 in yeast results in a strong aggregation of the misfolded Htt protein coupled to a severe growth defect. Interestingly, these phenotypes are strongly influenced by the sequences flanking the polyQ stretch, including FPs15,16. It was rationalized that the different properties of FPs can differentially affect polyQ toxicity in yeast. Indeed, compared to GFP-like FPs, red fluorescent proteins and their evolved forms have shown a reduced toxicity and aggregation16. This manuscript provides a detailed protocol to assess the effect of the next generation of FPs on polyQ toxicity and aggregation in yeast. This assay allows for a rapid and potentially high-content analysis of FP variants that can be used in parallel with previously characterized techniques for the optimal characterization of new FPs and can assess how they perform compared to GFP.

<table><tbody><tr><td>5-alpha Competent E. coli (High efficiency)</td><td>New Englanfd Biolab</td><td>C2987</td><td></td></tr><tr><td>SpeI-HF</td><td>New Englanfd Biolab</td><td>R3133</td><td>High fidelity enzymes are preferred</td></tr><tr><td>SalI-HF</td><td>New Englanfd Biolab</td><td>R0138</td><td>High fidelity enzymes are preferred</td></tr><tr><td>Agarose</td><td>Fisher Scientific</td><td>BP160</td><td></td></tr><tr><td>LB-Agar</td><td>Fisher Scientific</td><td>BP1425</td><td></td></tr><tr><td>LB-Broth</td><td>Fisher Scientific</td><td>BP1426</td><td></td></tr><tr><td>Ampicilin</td><td>Fisher Scientific</td><td>BP1760</td><td></td></tr><tr><td>PfuUltra High-fidelity DNA Polymerase</td><td>Agilent Technologies</td><td>600382</td><td></td></tr><tr><td>EPOCH2 microplate spectrophotometer</td><td>BioTek Instruments inc</td><td>EPOCH2TC</td><td></td></tr><tr><td>Yeast Pin Replicator</td><td>V&amp;amp;P Scientific inc.</td><td>VP407AH</td><td></td></tr><tr><td>SPI imager</td><td>S&amp;amp;P Robotics inc.</td><td>spImager-M</td><td></td></tr><tr><td>Zeiss LSM 800 confocal with AryScan</td><td>Carl Zeiss Microscopy</td><td>LSM 800</td><td></td></tr><tr><td>8 well Lab-Tek imaging chambers</td><td>Fisher Scientific</td><td>12565470</td><td></td></tr><tr><td>Bio-Dot apparatus</td><td>Bio-Rad</td><td>1706545</td><td></td></tr><tr><td>Chemi Doc XRS+</td><td>Bio-Rad</td><td>1708265</td><td></td></tr><tr><td>anti-FLAG M1 antibody</td><td>Sigma-Aldrich</td><td>F3040</td><td></td></tr><tr><td>Goat anti-mouse IgG alexa 555 secondary antibody</td><td>Thermo&amp;nbsp;</td><td>A32727</td><td></td></tr><tr><td>Plasmid MiniPrep Kit</td><td>Fisher Scientific</td><td>K0503</td><td></td></tr><tr><td>Plasmid Gel extraction Kit</td><td>Fisher Scientific</td><td>K0831</td><td></td></tr><tr><td>PCR Purification Kit</td><td>Fisher Scientific</td><td>K0702</td><td></td></tr><tr><td>Prizm</td><td>GraphPad</td><td>N/A</td><td></td></tr><tr><td>TAE (Tris-Acetate-EDTA)</td><td>Fisher Scientific</td><td>BP13354</td><td></td></tr></tbody></table>

effect of fluorescent proteins on fusion partners using polyglutamine toxicity assays in yeast

For the investigation of protein localization and trafficking using live cell imaging, researchers often rely on fusing their protein of interest to a fluorescent reporter. The constantly evolving list of genetically encoded fluorescent proteins (FPs) presents users with several alternatives when it comes to fluorescent fusion design. Each FP has specific optical and biophysical properties that can affect the biochemical, cellular, and functional properties of the resulting fluorescent fusions. For instance, several FPs tend to form nonspecific oligomers that are susceptible to impede on the function of the fusion partner. Unfortunately, only a few methods exist to test the impact of FPs on the behavior of the fluorescent reporter. Here, we describe a simple method that enables the rapid assessment of the impact of FPs using polyglutamine (polyQ) toxicity assays in the budding yeast Saccharomyces cerevisiae. PolyQ-expanded huntingtin proteins are associated with the onset of Huntington&#39;s disease (HD), where the expanded huntingtin aggregates into toxic oligomers and inclusion bodies. The aggregation and toxicity of polyQ expansions in yeast are highly dependent on the sequences flanking the polyQ region, including the presence of fluorescent tags, thus providing an ideal experimental platform to study the impact of FPs on the behavior of their fusion partner.

FPs have different biophysical properties, including their tendency to oligomerize, that can affect the behavior of their fusion partners in the context of fluorescent reporters. This protocol describes a simple method where multiple FPs can be fused to toxic polyQ expansions. Since polyQ toxicity is highly dependent on the sequences flanking the polyQ stretch15, this assay allows a rapid and direct comparison of fluorescent polyQ fusion reporters (<strong class="x...

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Effect of Fluorescent Proteins on Fusion Partners Using Polyglutamine Toxicity Assays in Yeast

This article describes protocols to assess the effect of fluorescent proteins on the aggregation and toxicity of misfolded polyglutamine expansion for the rapid evaluation of a newly uncharacterized fluorescent protein in the context of fluorescent reporters.

For the investigation of protein localization and trafficking using live cell imaging, researchers often rely on fusing their protein of interest to a ...

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Research

JoVE Journal

Biology

6.8K Views.  University of Western Ontario. Deze methode kan helpen bij het beantwoorden van belangrijke vragen op het gebied van fluorescerende eiwitten, zoals hoe fluorescerende eiwitten hun fusiepartners kunnen beïnvloeden. Het belangrijkste voordeel van deze techniek is dat het een snelle en gemakkelijk schaalbare beoordeling biedt van de effecten van de fluorescerende eiwitten en hun fusiepartners. Hoewel deze methode inzicht kan geven in fluorescerend eiwitgedrag, kan het ook worden toegepast op andere genetisch gecodeerde tags....

Video: Effect of Fluorescent Proteins on Fusion Partners Using Polyglutamine Toxicity Assays in Yeast

Generation of Fluorescent Protein Fusions in Candida Species

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. Thus, molecular and genetic tools are needed to facilitate the study of their pathogenesis mechanisms. PCR-mediated gene modification is a straightforward and quick approach to generate epitope-tagged proteins to facilitate their detection. In particular, fluorescent protein (FP) fusions are powerful tools that allow visualization and quantitation of both yeast cells and proteins by fluorescence microscopy and immunoblotting, respectively. Plasmids containing FP encoding sequences, along with nutritional marker genes that facilitate the transformation of Candida species, have been generated for the purpose of FP construction and expression in Candida. Herein, we present a strategy for constructing a FP fusion in a Candida species. Plasmids containing the nourseothricin resistance transformation marker gene (NAT1) along with sequences for either green, yellow, or cherry FPs (GFP, YFP, mCherry) are used along with primers that include gene-specific sequences in a polymerase chain reaction (PCR) to generate a FP cassette. This gene-specific cassette has the ability to integrate into the 3'-end of the corresponding gene locus via homologous recombination. Successful in-frame fusion of the FP sequence into the gene locus of interest is verified genetically, followed by analysis of fusion protein expression by microscopy and/or immuno-detection methods. In addition, for the case of highly expressed proteins, successful fusions can be screened for primarily by fluorescence imaging techniques.

Generation of Fluorescent Protein Fusions in Candida Species

PCR-mediated gene modification can be used to generate fluorescent protein fusions in Candida species, which facilitates visualization and quantitation of yeast cells and proteins. Herein, we present a strategy for constructing a fluorescent protein fusion (Eno1-FP) in Candida parapsilosis.

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. ...

Genetics

Mating-based Overexpression Library Screening in Yeast

Budding yeast has been widely used as a model in studying proteins associated with human diseases. Genome-wide genetic screening is a powerful tool commonly used in yeast studies. The expression of a number of neurodegenerative disease-associated proteins in yeast causes cytotoxicity and aggregate formation, recapitulating findings seen in patients with these disorders. Here, we describe a method for screening a yeast model of the Amyotrophic Lateral Sclerosis-associated protein FUS for modifiers of its toxicity. Instead of using transformation, this new screening platform relies on the mating of yeast to introduce an arrayed library of plasmids into the yeast model. The mating method has two clear advantages: first, it is highly efficient; second, the pre-transformed arrayed library of plasmids can be stored for long-term as a glycerol stock, and quickly applied to other screens without the labor-intensive step of transformation into the yeast model each time. We demonstrate how this method can successfully be used to screen for genes that modify the toxicity of FUS.

This article presents a mating-based method to facilitate overexpression screening in budding yeast using an arrayed plasmid library.

Budding yeast has been widely used as a model in studying proteins associated with human diseases. Genome-wide genetic screening is a powerful tool ...

High-throughput Yeast Plasmid Overexpression Screen

The budding yeast, Saccharomyces cerevisiae, is a powerful model system for defining fundamental mechanisms of many important cellular processes, including those with direct relevance to human disease. Because of its short generation time and well-characterized genome, a major experimental advantage of the yeast model system is the ability to perform genetic screens to identify genes and pathways that are involved in a given process. Over the last thirty years such genetic screens have been used to elucidate the cell cycle, secretory pathway, and many more highly conserved aspects of eukaryotic cell biology 1-5. In the last few years, several genomewide libraries of yeast strains and plasmids have been generated 6-10. These collections now allow for the systematic interrogation of gene function using gain- and loss-of-function approaches 11-16. Here we provide a detailed protocol for the use of a high-throughput yeast transformation protocol with a liquid handling robot to perform a plasmid overexpression screen, using an arrayed library of 5,500 yeast plasmids. We have been using these screens to identify genetic modifiers of toxicity associated with the accumulation of aggregation-prone human neurodegenerative disease proteins. The methods presented here are readily adaptable to the study of other cellular phenotypes of interest.

Here we describe a plasmid overexpression screen in Saccharomyces cerevisiae, using an arrayed plasmid library and a high-throughput yeast transformation protocol with a liquid handling robot.

The budding yeast, Saccharomyces cerevisiae, is a powerful model system for defining fundamental mechanisms of many important cellular processes, ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Growth Assays to Assess Polyglutamine Toxicity in Yeast

Protein misfolding is associated with many human diseases, particularly neurodegenerative diseases, such as Alzheimer&#x2019;s disease, Parkinson's disease, and Huntington's disease 1. Huntington's disease (HD) is caused by the abnormal expansion of a polyglutamine (polyQ) region within the protein huntingtin. The polyQ-expanded huntingtin protein attains an aberrant conformation (i.e. it misfolds) and causes cellular toxicity 2. At least eight further neurodegenerative diseases are caused by polyQ-expansions, including the Spinocerebellar Ataxias and Kennedy&#x2019;s disease 3.
The model organism yeast has facilitated significant insights into the cellular and molecular basis of polyQ-toxicity, including the impact of intra- and inter-molecular factors of polyQ-toxicity, and the identification of cellular pathways that are impaired in cells expressing polyQ-expansion proteins 3-8. Importantly, many aspects of polyQ-toxicity that were found in yeast were reproduced in other experimental systems and to some extent in samples from HD patients, thus demonstrating the significance of the yeast model for the discovery of basic mechanisms underpinning polyQ-toxicity. 
A direct and relatively simple way to determine polyQ-toxicity in yeast is to measure growth defects of yeast cells expressing polyQ-expansion proteins. This manuscript describes three complementary experimental approaches to determine polyQ-toxicity in yeast by measuring the growth of yeast cells expressing polyQ-expansion proteins. The first two experimental approaches monitor yeast growth on plates, the third approach monitors the growth of liquid yeast cultures using the BioscreenC instrument. 
Furthermore, this manuscript describes experimental difficulties that can occur when handling yeast polyQ models and outlines strategies that will help to avoid or minimize these difficulties. The protocols described here can be used to identify and to characterize genetic pathways and small molecules that modulate polyQ-toxicity. Moreover, the described assays may serve as templates for accurate analyses of the toxicity caused by other disease-associated misfolded proteins in yeast models.

This manuscript describes three complementary protocols for assessing the toxicity of polyglutamine (polyQ)-expansion proteins in the yeast Saccharomyces cerevisiae. These protocols can easily be modified to monitor the toxicity of other misfolded proteins in yeast.

Protein misfolding is associated with many human diseases, particularly neurodegenerative diseases, such as Alzheimer&#x2019;s disease, Parkinson's ...

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must be maintained to avoid deleterious consequences 1. External or internal factors that disrupt this balance can lead to protein aggregation, toxicity and cell death. In humans this is a major contributing factor to the symptoms associated with neurodegenerative disorders such as Huntington's, Parkinson's, and Alzheimer's diseases 10. It is therefore essential that the proteins involved in maintenance of proteostasis be identified in order to develop treatments for these debilitating diseases. This article describes techniques for monitoring in vivo protein folding at near-real time resolution using the model protein firefly luciferase fused to green fluorescent protein (FFL-GFP). FFL-GFP is a unique model chimeric protein as the FFL moiety is extremely sensitive to stress-induced misfolding and aggregation, which inactivates the enzyme 12. Luciferase activity is monitored using an enzymatic assay, and the GFP moiety provides a method of visualizing soluble or aggregated FFL using automated microscopy. These coupled methods incorporate two parallel and technically independent approaches to analyze both refolding and functional reactivation of an enzyme after stress. Activity recovery can be directly correlated with kinetics of disaggregation and re-solubilization to better understand how protein quality control factors such as protein chaperones collaborate to perform these functions. In addition, gene deletions or mutations can be used to test contributions of specific proteins or protein subunits to this process. In this article we examine the contributions of the protein disaggregase Hsp104 13, known to partner with the Hsp40/70/nucleotide exchange factor (NEF) refolding system 5, to protein refolding to validate this approach.

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

This article describes the use of a firefly luciferase-GFP fusion protein to investigate in vivo protein folding in Saccharomyces cerevisiae. Using this reagent, refolding of a model heat-denatured protein can be monitored simultaneously by fluorescence microscopy and an enzymatic assay to probe the roles of proteostasis network components in protein quality control.

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must ...

Growth-based Determination and Biochemical Confirmation of Genetic Requirements for Protein Degradation in Saccharomyces cerevisiae

Regulated protein degradation is crucial for virtually every cellular function. Much of what is known about the molecular mechanisms and genetic requirements for eukaryotic protein degradation was initially established in Saccharomyces cerevisiae. Classical analyses of protein degradation have relied on biochemical pulse-chase and cycloheximide-chase methodologies. While these techniques provide sensitive means for observing protein degradation, they are laborious, time-consuming, and low-throughput. These approaches are not amenable to rapid or large-scale screening for mutations that prevent protein degradation. Here, a yeast growth-based assay for the facile identification of genetic requirements for protein degradation is described. In this assay, a reporter enzyme required for growth under specific selective conditions is fused to an unstable protein. Cells lacking the endogenous reporter enzyme but expressing the fusion protein can grow under selective conditions only when the fusion protein is stabilized (i.e. when protein degradation is compromised). In the growth assay described here, serial dilutions of wild-type and mutant yeast cells harboring a plasmid encoding a fusion protein are spotted onto selective and non-selective medium. Growth under selective conditions is consistent with degradation impairment by a given mutation. Increased protein abundance should be biochemically confirmed. A method for the rapid extraction of yeast proteins in a form suitable for electrophoresis and western blotting is also demonstrated. A growth-based readout for protein stability, combined with a simple protocol for protein extraction for biochemical analysis, facilitates rapid identification of genetic requirements for protein degradation. These techniques can be adapted to monitor degradation of a variety of short-lived proteins. In the example presented, the His3 enzyme, which is required for histidine biosynthesis, was fused to Deg1-Sec62. Deg1-Sec62 is targeted for degradation after it aberrantly engages the endoplasmic reticulum translocon. Cells harboring Deg1-Sec62-His3 were able to grow under selective conditions when the protein was stabilized.

Growth-based Determination and Biochemical Confirmation of Genetic Requirements for Protein Degradation in Saccharomyces cerevisiae

This article describes a yeast growth-based assay for the determination of genetic requirements for protein degradation. It also demonstrates a method for rapid extraction of yeast proteins, suitable for western blotting to biochemically confirm degradation requirements. These techniques can be adapted to monitor degradation of a variety of proteins.

Regulated protein degradation is crucial for virtually every cellular function. Much of what is known about the molecular mechanisms and genetic ...

Microscopy of Fission Yeast Sexual Lifecycle

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the mechanics of cell division and cell polarity. Furthermore, classical experiments on its sexual reproduction have yielded results pivotal to current understanding of DNA recombination and meiosis. More recent analysis of fission yeast mating has raised interesting questions on extrinsic stimuli response mechanisms, polarized cell growth and cell-cell fusion. To study these topics in detail we have developed a simple protocol for microscopy of the entire sexual lifecycle. The method described here is easily adjusted to study specific mating stages. Briefly, after being grown to exponential phase in a nitrogen-rich medium, cell cultures are shifted to a nitrogen-deprived medium for periods of time suited to the stage of the sexual lifecycle that will be explored. Cells are then mounted on custom, easily built agarose pad chambers for imaging. This approach allows cells to be monitored from the onset of mating to the final formation of spores.

We provide a reproducible basic method for the long-term microscopy of the fission yeast sexual lifecycle. With minor adjustments described, the presented protocol allows research focus on different steps of the reproductive process.

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the ...

A Method to Study &#945;-Synuclein Toxicity and Aggregation Using a Humanized Yeast Model

Parkinson's disease is the second most common neurodegenerative disorder and is characterized by progressive cell death caused by the formation of Lewy bodies containing misfolded and aggregated &#945;-synuclein. &#945;-synuclein is an abundant presynaptic protein that regulates synaptic vesicle trafficking, but the accumulation of its proteinaceous inclusions results in neurotoxicity. Recent studies have revealed that various genetic factors, including bacterial chaperones, could reduce the formation of &#945;-synuclein aggregates in vitro. However, it is also important to monitor the anti-aggregation effect in the cell to apply this as a potential treatment for the patients. It would be ideal to use neuronal cells, but these cells are difficult to handle and take a long time to exhibit the anti-aggregation phenotype. Therefore, a quick and effective in vivo tool is required for the further evaluation of in vivo anti-aggregation activity. The method described here was used to monitor and analyze the anti-aggregation phenotype in the humanized yeast Saccharomyces cerevisiae, which expressed human &#945;-synuclein. This protocol demonstrates in vivo tools that could be used for monitoring &#945;-synuclein-induced cellular toxicity, as well as the formation of &#945;-synuclein aggregates in cells.

An in vivo physiological model of &#945;-synuclein is required to study and understand the pathogenesis of Parkinson's disease. We describe a method to monitor the cytotoxicity and aggregate formation of &#945;-synuclein using a humanized yeast model.

Parkinson's disease is the second most common neurodegenerative disorder and is characterized by progressive cell death caused by the formation of Lewy ...

Identifying Drugs that Affect Polymerization of Bacterial Cytoskeletal Proteins

Fission Yeast as a Platform for Antibacterial Drug Screens Targeting Bacterial Cytoskeleton Proteins

Bacterial cytoskeletal proteins such as FtsZ and MreB perform essential functions such as cell division and cell shape maintenance. Further, FtsZ and MreB have emerged as important targets for novel antimicrobial discovery. Several assays have been developed to identify compounds targeting nucleotide binding and polymerization of these cytoskeletal proteins, primarily focused on FtsZ. Moreover, many of the assays are either laborious or cost-intensive, and ascertaining whether these proteins are the cellular target of the drug often requires multiple methods. Finally, the toxicity of the drugs to eukaryotic cells also poses a problem. Here, we describe a single-step cell-based assay to discover novel molecules targeting bacterial cytoskeleton and minimize hits that might be potentially toxic to eukaryotic cells. Fission yeast is amenable to high-throughput screens based on microscopy, and a visual screen can easily identify any molecule that alters the polymerization of FtsZ or MreB. Our assay utilizes the standard 96-well plate and relies on the ability of the bacterial cytoskeletal proteins to polymerize in a eukaryotic cell such as the fission yeast. While the protocols described here are for fission yeast and utilize FtsZ from Staphylococcus aureus and MreB from Escherichia coli, they are easily adaptable to other bacterial cytoskeletal proteins that readily assemble into polymers in any eukaryotic expression hosts. The method described here should help facilitate further discovery of novel antimicrobials targeting bacterial cytoskeletal proteins.

Author Spotlight: Exploring Cytoskeletal Dynamics to Unveil Novel Antibiotics Through Innovative Cell-Based Assays

Fission yeast is used here as a heterologous host to express bacterial cytoskeletal proteins such as FtsZ and MreB as translational fusion proteins with GFP to visualize their polymerization. Also, compounds that affect polymerization are identified by imaging using a fluorescence microscope.

Bacterial cytoskeletal proteins such as FtsZ and MreB perform essential functions such as cell division and cell shape maintenance. Further, FtsZ and MreB ...

Effect of Fluorescent Proteins on Fusion Partners Using Polyglutamine Toxicity Assays in Yeast

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