This work was supported by the US Defense Advanced Research Projects Agency contract N66001-12-C-4039 to Y.S., a grant from the Alfred P. Sloan Foundation to R.S.L., and US National Institutes of Health grants R01 HG003224 and R21 CA130266 to F.P.R. F.P.R. was also supported by a fellowship from the Canadian Institute for Advanced Research and by the Canada Excellence Research Chairs program.

1. Generation of ProMonsters
<ol>
	<li>Prepare the universal GFP replacement cassette by amplifying a tetO2-GFP marker and the URA3 marker from the plasmid pYOGM012 (ampicillin resistance) using primers with sequences: 
	GGATCCCCGGGTTAATTAAGGCGCGCCAGATCTGTTTAGCTTGCCCAAGCTCCTCGAGTAATTCG and GGCGTTAGTATCGAATCGACAGCAGTATAGCGACCAGCATTCACGTACCGGGTAATAACTGATATAAT (bolded regions provide homology to the flanking regions of the KanMX4 cassette allowing targeted replacement). The PCR reaction should be carried out with Phusion polymerase, HF buffer, and 3% dimethyl sulfoxide (New England BioLabs) starting with the template DNA concentration of 0.5 ng/&mu;l. The PCR program is 98 &deg;C for 30 sec and 35 cycles of 98 &deg;C for 10 sec, 49 &deg;C for 30 sec, and 72 &deg;C for 1.5 min. The reaction volume should be &gt; 50 &mu;l to provide enough DNA for each transformation experiment. We purify the product using a QIAquick PCR purification kit (Qiagen) with ~50 &mu;l of the PCR reaction processed per column, but skipping the purification step may increase the yield of transformants. 
	Alternatively, release a fragment containing the tetO2-GFP and URA3 markers, as well as homologous regions for KanMX4 targeting, by digesting the plasmid pYOGM057 (ampicillin resistance) using the restriction enzyme EcoRI (New England BioLabs). The DNA concentration in this reaction should be ~30 ng/&mu;l. The reaction volume should be &gt; 34 &mu;l. 
	For integrating the GFP cassette into a region other than KanMX4, adjust the homology region of the PCR primers above to the appropriate homologous sequences.</li>
	<li>Transform an a-haploid strain from the KanMX4 yeast knockout collection12 with the universal GFP deletion cassette. Following the protocol by Woods and Gietz13 .34 &mu;l of the DNA sample (purified PCR product or non-purified restriction digest) should be used for each transformation experiment. Plate one fifth of the transformation mixture onto a CAA-Ura plate (0.67% yeast nitrogen base, 2% glucose, 0.6% casamino acid, 0.0025% adenine hemisulfate, 0.005% tryptophan, and 2% agar).</li>
	<li>Streak out the transformants on a fresh CAA-Ura plate to obtain isolated colonies.</li>
	<li>Transfer cells from a colony onto a YPDA plate containing 200 &mu;g/ml G418 to confirm the lack of growth. To confirm successful GFP integration, follow the steps (1.5) - (1.9) to perform colony PCR.</li>
	<li>Add cells taken from a colony into 2 &mu;l of lysis buffer (0.1 M sodium phosphate, pH 7.4, 1 unit zymolyase from ZymoResearch) in a 0.2-ml PCR tube or a well of a 96-well plate. Mix by pipetting the solution in and out of a pipette tip.</li>
	<li>Overlay with one drop (roughly 20 &mu;l) of mineral oil using a P200 Pipetman (Gilson).</li>
	<li>Incubate the mixture at 37 &deg;C for 20 min and then at 95 &deg;C for 10 min.</li>
	<li>Dilute the lysate with 50 &mu;l water. Mix by pipetting the solution in and out ten times.</li>
	<li>Analyze 1 &mu;l of the lysate using 10-&mu;l PCR reactions with (A) a forward primer that anneals to the upstream flanking region paired with a primer of the sequence CCTGAGAAAGCAACCTGACC (285 bp from the beginning of the cassette), (B) a reverse primer that anneals to the downstream region paired with a primer of the sequence GCATTGGGTCAACAGTATAG (375 bp from the end), (C) two primers that anneal to KanMX4 with the sequences CGTACTCCTGATGATGCATG and GACGAAATACGCGATCGCTG (181 bp), and (D) two primers that anneal to the wild-type sequence of the targeted gene (Figure 4). (D) is important because about 8% of the strains in the deletion collections are known to contain aneuploidy14. A correct transformant should be positive with (A) and (B), and negative with (C) and (D).</li>
	<li>To introduce a GMToolkit into the transformant, add roughly 250,000 cells from the transformed strain and 250,000 cells of either RY0146 (containing GMToolkit-a) or RY0148 (containing GMToolkit-&alpha;) into a 1.6-ml Eppendorf tube. Vortex. The genotypes of RY0146 and RY0148 are MAT&alpha; lyp1&Delta; his3&Delta;1 leu2&Delta;0 ura3&Delta;0 met15&Delta;0 can1&Delta;::GMToolkit-a [CMVpr-rtTA KanMX4 STE2pr-Sp-his5] and MAT&alpha; lyp1&Delta; his3&Delta;1 leu2&Delta;0 ura3&Delta;0 met15&Delta;0 can1&Delta;::GMToolkit-&alpha; [CMVpr-rtTA NatMX4 STE3pr-LEU2], respectively. pr denotes promoter. Cell number should be estimated from optical density of the preceding culture.</li>
	<li>Centrifuge at 800 x g for 2 min.</li>
	<li>Remove the supernatant.</li>
	<li>To allow for air exchange, make a hole on the lid using a pushpin treated with 70% ethanol.</li>
	<li>Add 50 &mu;l of YPDA medium (1% yeast extract, 2% peptone, 2% glucose, and 0.003% adenine hemisulfate). Vortex.</li>
	<li>Centrifuge at 800 x g for 5 min.</li>
	<li>Place the tube in a damp box. This can be created with an empty tip box, a small tip stand, and a wet paper towel.</li>
	<li>Incubate the box overnight at 30 &deg;C.</li>
	<li>Select mated diploids by first streaking out the cells on a CAA-Ura plate. Incubate the plate at 30 &deg;C for one day.</li>
	<li>Taking cells from the bulk area, streak them out to make isolated colonies on YPDA plates (1% yeast extract, 2% peptone, 2% glucose, 0.003% adenine hemisulfate, and 2% agar) containing 200 &mu;g/ml G418 for selecting diploids containing GMToolkit-a or 100 &mu;g/ml nourseothricin (Nat)15 for selecting diploids containing GMToolkit-&alpha;.</li>
	<li>Incubate the plate at 30 &deg;C for 3 days.</li>
	<li>Starting from an isolated colony, transfer most of the cells to 500 &mu;l of GNA medium without agar (1% yeast extract, 3% Difco nutrient broth, and 5% glucose; autoclave glucose and the rest of the components separately as 2&times; solutions to avoid caramelization) in a 5-ml culture tube with the lid loosened.</li>
	<li>Rotate the tube horizontally for five hours at 30 &deg;C. The axis of the rotation needs to be parallel to the longitudinal axis of the tube.</li>
	<li>Confirm that colonies used in (1.21) are Ura+ by streaking them out on a CAA-Ura plate.</li>
	<li>Centrifuge the tube at 800 x g for 2 min. Discard the supernatant.</li>
	<li>Add 500 &mu;l of minimal sporulation medium (1% potassium acetate, 0.005% zinc acetate; filter-sterilize). Vortex.</li>
	<li>Centrifuge the tube at 800 x g for 2 min. Discard the supernatant.</li>
	<li>Repeat (1.25) - (1.26) twice.</li>
	<li>Add 1 ml of minimal sporulation medium supplemented with 7.5 &mu;g/ml lysine, 7.5 &mu;g/ml leucine, 5 &mu;g/ml histidine, 5 &mu;g/ml methionine, and 1.25 &mu;g/ml uracil (to enhance the sporulation rate of the auxotrophic strains). Vortex.</li>
	<li>Rotate the tube at room temperature for one day.</li>
	<li>Rotate the tube at 30 &deg;C for 2 days.</li>
	<li>Centrifuge 125 &mu;l of this mixture in a 1.6-ml Eppendorf tube at 800 x g for 2 min. Discard the supernatant.</li>
	<li>Add 50 &mu;l of zymolyase solution (100 mM sodium phosphate buffer, pH 7.4, 1 M sorbitol, and 2 units zymolyase from ZymoResearch). Mix by pipetting the solution in and out of a pipette tip.</li>
	<li>Incubate at 30 &deg;C for 1 hr.</li>
	<li>Add 50 &mu;l of 0.02% NP-40. Mix by pipetting the solution in and out of a pipette tip.</li>
	<li>Leave the tube at room temperature for five minutes.</li>
	<li>Add 500 &mu;l of water.</li>
	<li>Place the tube on ice.</li>
	<li>Sonicate this mixture twice at the output setting of 1 (weakest setting) for 15 sec using a 3.2-mm microtip with Sonifier 450 (Branson). Between the two rounds of sonication, put the tube on ice to return the temperature to ~0 &deg;C.</li>
	<li>Centrifuge the tubes at 800 x g for 5 min. Discard the supernatant.</li>
	<li>Add 200 &mu;l of SC-Ura-His (a) or SC-Ura-Leu (&alpha;) medium, depending on whether the cross is for incorporating GMToolkit-a or GMToolkit-&alpha;. To prepare the media, make SC-Ura-His-Leu medium (0.67% yeast nitrogen base, 2% glucose, 0.01% arginine, 0.01% aspartic acid, 0.01% glutamic acid, 0.01% isoleucine, 0.01% lysine, 0.01% methionine, 0.01% phenylalanine, 0.08% serine, 0.04% threonine, 0.002% tyrosine, 0.03% valine, 0.0025% adenine, 0.005% tryptophan, 0.003% adenine, and 0.005% tryptophan) and supplement it with sterile histidine or leucine before use to the final concentration of 0.01%.</li>
	<li>Make a hole on the lid using a pushpin treated with 70% ethanol.</li>
	<li>Rotate the tube horizontally at 30 &deg;C for 2 days. The axis of the rotation needs to be parallel to the longitudinal axis of the tube.</li>
	<li>Streak out the cells on a SC-Ura-His or SC-Ura-Leu agar plate (the same ingredients as above plus 2% agar; threonine and aspartic acid must be added after autoclaving; add histidine or leucine before pouring) to make isolated colonies for the ProMonster strains.</li>
</ol>
2. Sexual Cycling
<ol>
	<li>Rotate a culture of an established GFP deletion strain in 100 &mu;l of SC-His (a) or SC-Leu (&alpha;) depending on mating type at 30 &deg;C overnight using a 1.6-ml Eppendorf tube. For air exchange, make a hole using a pushpin treated with 70% ethanol. Rotate the tube horizontally. The axis of the rotation needs to be parallel to the longitudinal axis of the tube. It is possible to cross multiple strains via en masse mating. When a sorted sample is directly used for this purpose, culture the cells in 200 &mu;l at 30 &deg;C for 2 days. To prepare the medium, add uracil (final concentration: 0.0025%) and histidine (0.01%) or leucine (0.01%) to SC-Ura-His-Leu medium.</li>
	<li>Mix 250,000 a-haploid cells and 250,000 &alpha;-haploid cells of GFP deletion strains in a 1.6-ml Eppendorf tube. Vortex.</li>
	<li>Mate these cells by following steps (1.11) - (1.17).</li>
	<li>Transfer 10 &mu;l of the mating mixture to 500 &mu;l GNA medium containing 200 &mu;g/ml G418 and 100 &mu;g/ml Nat in a 5-ml culture tube with a loosened lid. The starting OD600 is roughly 0.1.</li>
	<li>Rotate the culture at 30 &deg;C for 24 hr. This allows for the selection of diploids because only diploids containing both KanMX4 in GMToolkit-a and NatMX4 in GMToolkit-&alpha; can grow in the presence of G418 and Nat (Figure 3).</li>
	<li>Transfer 20 &mu;l of the one-day culture to 500 &mu;l of fresh GNA containing G418 and Nat. The starting OD600 is roughly 0.2.</li>
	<li>Rotate the tube at 30 &deg;C for 5 hr to bring cells to the log phase (OD600 of ~1).</li>
	<li>Follow the steps (1.24)-(1.38) to sporulate the diploids and isolate spores.</li>
	<li>Split the suspension of sporulated cells into two 300-&mu;l parts and put each into a separate 1.5-ml Eppendorf tube.</li>
	<li>Centrifuge the tubes at 800 x g for 5 min. Discard the supernatant.</li>
	<li>To the pellet of one tube, add 100 &mu;l of SC-His medium to select a-haploids. To the pellet of the other tube, add 100 &mu;l of SC-Leu medium to select &alpha;-haploids.</li>
	<li>Rotate the tubes at 30 &deg;C overnight. The final OD600 should be less than 0.5. If OD600 tends to be too high, try culturing at room temperature.</li>
	<li>To induce GFP, add 100 &mu;l of fresh SC-His (a) or SC-Leu (&alpha;) medium containing 20 &mu;g/ml doxycycline. The final concentration of doxycycline is 10 &mu;g/ml. Vortex.</li>
	<li>Rotate the tubes at 30 &deg;C for 2 days.</li>
</ol>
3. Flow Cytometry
<ol>
	<li>Add 900 &mu;l of pre-filtered TE buffer (pH 7.5) containing 10 &mu;g/ml doxycycline to a 5-ml polypropylene tube (BD Falcon #352063) with the cap swapped with a cell-strainer cap from a 5-ml polystyrene tube (BD Falcon #352235). Polypropylene tubes are suitable for flow cytometry. The strainer cap is desired for removing large particles.</li>
	<li>Gently place 75 &mu;l of TE buffer with doxycycline on the cell-strainer cap.</li>
	<li>Add 25 &mu;l of the induced cells to the solution on the cap.</li>
	<li>While pressing the tip of Pipetman against the mesh, shoot all of the combined solution through the strainer.</li>
	<li>Press down the strainer cap and vortex the tube.</li>
	<li>Prepare collection tubes (1.6-ml Eppendorf tubes) filled with 200 &mu;l of SC-His or SC-Leu.</li>
	<li>Start the cell sorter and adjust settings according to the manufacturer manual.</li>
	<li>Vortex both the collection tube (to coat the wall with the medium) and the tube containing cells before loading them onto the cell sorter.</li>
	<li>Acquire flow cytometry data of the sample. A strain not containing GFP can be a negative control.</li>
	<li>Draw gates for sorting. (A) To avoid cell aggregates, which could exhibit high GFP intensity, discard outliers with disproportionately large pulse width for forward scatter (FSC) using a plot of pulse width and pulse height for MoFlo sorter or disproportionately small FSC height using a plot of FSC height and FSC area for FACSAria sorter (Figure 5, top-left panel). (B) Since large FSC (area) is expected for cell aggregates, only take cells in the lower 20% in FSC, while avoiding regions with the lowest FSC and side scatter (SSC) values where cell debris is often found (Figure 5, top-right panel). (C) SSC (area) and GFP signal (area) are often positively correlated. Because simply taking the brightest cells given the gates above would also enrich for cells with a high SSC value, draw a gate along the periphery of the population using a plot of GFP intensity and SSC to take a similar percentage of the brightest cells from each point of the SSC axis (Figure 5, bottom-left diagram). The population selected in (C) should be 0.1-1% of the population selected in (B).</li>
	<li>Sort 200 cells into the collection tube given the criteria in (3.10). For an en masse process or for other projects requiring a more complex population, sort more cells as needed.</li>
	<li>Vortex the collection tube to cover the sorted cells with medium.</li>
	<li>Plate a subset of cells (for example, 50 cells) on a YPDA plate for genotyping.</li>
	<li>Incubate the plate at 30 &deg;C for 2 days.</li>
	<li>Make a lysate for ~12 colonies following the steps (1.5) - (1.8).</li>
	<li>Transfer the residual cells on the tip used in (1.5) to a spot on a YPDA plate containing G418 and Nat by gently touching the plate with the tip. Selected haploids should not grow on this plate. Diploids may have more GFP copies and may therefore be brighter. This test can show the efficiency of haploid selection.</li>
	<li>For genotyping, analyze 1 &mu;l of the lysate using a 10-&mu;l PCR reaction with a forward primer that anneals to the upstream flanking region of each deleted gene paired with a primer of the sequence CCTGAGAAAGCAACCTGACC. 
	Multiplex PCR reactions if possible (conditions can be determined using lysates of single mutants). PCR can be done in a 96- or 384-well format. The latter format is suitable for the reaction volume of as little as 5 &mu;l. Arraying of PCR master mixes differing in primers and addition of cell lysates to these mixes can be carried out using a multi-channel pipette or a liquid handling robot. Tip changes are optional when adding the same lysate to different PCR mixes.</li>
	<li>Analyze PCR products. The Gel XL Ultra V-2 electrophoresis system (Labnet) is compatible with sample loading using multi-channel pipettes.</li>
	<li>Based on information from (3.16) and (3.18), identify likely haploids that have desired genotypes.</li>
	<li>Establish these strains on a fresh plate (YPDA, CAA-Ura, or SC-Ura-His/Leu). Also, freeze them in 25% glycerol at -80 &deg;C. Additional tests such as confirmation of the absence of wild-type sequences for deleted genes and pulsed-field gel electrophoresis are recommended.</li>
	<li>Repeat sexual cycling from (2.1) with useful strains.</li>
</ol>

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No conflicts of interest declared.

As we developed the Green Monster approach, we were concerned with the possibility of recombination between different GFP replacement cassettes, leading to genome rearrangement. Mitigating against this possibility is our selection for cells that have successfully undergone multiple rounds of mating and meiosis. Cells bearing rearranged genomes are expected to be less fit after mating to cells without an identical rearrangement. Indeed, we did not observe any genome instability resulting from recombination betwe...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent or instrument</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>G418</td>
 <td>Sigma-Aldrich</td>
 <td>A1720</td>
 <td>Dissolve in water and filter-sterilize (0.2-&mu;m filter). Stock concentration: 200 mg/ml. Store at 4 &deg;C.</td>
 </tr>
 <tr>
 <td>ClonNAT (nourseothricin)</td>
 <td>WERNER BioAgents</td>
 <td>5001000</td>
 <td>Dissolve in water and filter-sterilize (0.2-&mu;m filter). Stock concentration: 100 mg/ml. Store at 4 &deg;C.</td>
 </tr>
 <tr>
 <td>Doxycycline</td>
 <td>Sigma-Aldrich</td>
 <td>D9891</td>
 <td>Dissolve in 50% ethanol and filter-sterilize (0.2-&mu;m filter). Stock concentration: 10 mg/ml. Make fresh every four weeks. Shield from light using aluminum foil and store at 4 &deg;C. </td>
 </tr>
 <tr>
 <td>Zymolyase</td>
 <td>ZymoResearch</td>
 <td>E1005</td>
 <td></td>
 </tr>
 <tr>
 <td>Difco yeast nitrogen base w/o amino acids</td>
 <td>BD</td>
 <td>291940</td>
 <td></td>
 </tr>
 <tr>
 <td>Revolver (rotator for tubes)</td>
 <td>Labnet</td>
 <td>H5600</td>
 <td></td>
 </tr>
 <tr>
 <td>Enduro Gel XL electrophoresis unit</td>
 <td>Labnet</td>
 <td>E0160</td>
 <td></td>
 </tr>
 <tr>
 <td>Sonifier 450 </td>
 <td>Branson</td>
 <td>101-063-198</td>
 <td></td>
 </tr>
 <tr>
 <td>Microtip for Sonifier 450</td>
 <td>Branson</td>
 <td>101-148-062</td>
 <td></td>
 </tr>
 <tr>
 <td>FACSAria cell sorter</td>
 <td>BD</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>MoFlo cell sorter</td>
 <td>Beckman-Coulter</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Biomek FX or equivalent robot </td>
 <td>Beckman Coulter</td>
 <td></td>
 <td>Optional. For setting up genotyping PCRs.</td>
 </tr>
 </tbody>
</table>

the green monster process for the generation of yeast strains carrying multiple gene deletions

Phenotypes for a gene deletion are often revealed only when the mutation is tested in a particular genetic background or environmental condition1,2. There are examples where many genes need to be deleted to unmask hidden gene functions3,4. Despite the potential for important discoveries, genetic interactions involving three or more genes are largely unexplored. Exhaustive searches of multi-mutant interactions would be impractical due to the sheer number of possible combinations of deletions. However, studies of selected sets of genes, such as sets of paralogs with a greater a priori chance of sharing a common function, would be informative.
In the yeast Saccharomyces cerevisiae, gene knockout is accomplished by replacing a gene with a selectable marker via homologous recombination. Because the number of markers is limited, methods have been developed for removing and reusing the same marker5,6,7,8,9,10. However, sequentially engineering multiple mutations using these methods is time-consuming because the time required scales linearly with the number of deletions to be generated.
Here we describe the Green Monster method for routinely engineering multiple deletions in yeast11. In this method, a green fluorescent protein (GFP) reporter integrated into deletions is used to quantitatively label strains according to the number of deletions contained in each strain (Figure 1). Repeated rounds of assortment of GFP-marked deletions via yeast mating and meiosis coupled with flow-cytometric enrichment of strains carrying more of these deletions lead to the accumulation of deletions in strains (Figure 2). Performing multiple processes in parallel, with each process incorporating one or more deletions per round, reduces the time required for strain construction.
The first step is to prepare haploid single-mutants termed 'ProMonsters,' each of which carries a GFP reporter in a deleted locus and one of the 'toolkit' loci&#x2014;either Green Monster GMToolkit-a or GMToolkit-&alpha; at the can1&Delta; locus (Figure 3). Using strains from the yeast deletion collection12, GFP-marked deletions can be conveniently generated by replacing the common KanMX4 cassette existing in these strains with a universal GFP-URA3 fragment. Each GMToolkit contains: either the a- or &alpha;-mating-type-specific haploid selection marker1 and exactly one of the two markers that, when both GMToolkits are present, collectively allow for selection of diploids.
The second step is to carry out the sexual cycling through which deletion loci can be combined within a single cell by the random assortment and/or meiotic recombination that accompanies each cycle of mating and sporulation.

When an a-haploid strain carrying four GFP-marked deletions (ycl033c&Delta; yer042w&Delta; ykl069w&Delta; yol118c&Delta;) was crossed with an &alpha;-haploid strain carrying four deletions (ycl033c&Delta; ydl242w&Delta; ydl227c&Delta; yer042w&Delta;), with two deletions (ycl033c&Delta; yer042w&Delta;) shared by two strains, a representative result was obtained. The mating mixture was cultured in YPDA medium containing G418 and Nat to select diploids. The resulting diploids were...

Watch this Scientific Journal Video about The Green Monster Process for the Generation of Yeast Strains Carrying Multiple Gene Deletions at JoVE.com

The Green Monster Process for the Generation of Yeast Strains Carrying Multiple Gene Deletions

The Green Monster method enables the rapid assembly of multiple deletions marked with a reporter gene encoding green fluorescent protein. This method is based on driving yeast strains through repeated cycles of sexual assortment of deletions and fluorescence-based enrichment of cells carrying more deletions.

Phenotypes  for a gene deletion are often revealed only when the mutation is tested in a  particular genetic background or environmental condition1,2. ...

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14.2K Views.  J. Craig Venter Institute. The Green Monster method enables the rapid assembly of multiple deletions marked with a reporter gene encoding green fluorescent protein. This method is based on driving yeast strains through repeated cycles of sexual assortment of deletions and fluorescence-based enrichment of cells carrying more deletions.

Video: The Green Monster Process for the Generation of Yeast Strains Carrying Multiple Gene Deletions

A Rapid Technique for the Visualization of Live Immobilized Yeast Cells

We present here a simple, rapid, and extremely flexible technique for the immobilization and visualization of growing yeast cells by epifluorescence microscopy. The technique is equally suited for visualization of static yeast populations, or time courses experiments up to ten hours in length. My microscopy investigates epigenetic inheritance at the silent mating loci in S. cerevisiae. There are two silent mating loci, HML and HMR, which are normally not expressed as they are packaged in heterochromatin. In the sir1 mutant background silencing is weakened such that each locus can either be in the expressed or silenced epigenetic state, so in the population as a whole there is a mix of cells of different epigenetic states for both HML and HMR. My microscopy demonstrated that there is no relationship between the epigenetic state of HML and HMR in an individual cell. sir1 cells stochastically switch epigenetic states, establishing silencing at a previously expressed locus or expressing a previously silenced locus. My time course microscopy tracked individual sir1 cells and their offspring to score the frequency of each of the four possible epigenetic switches, and thus the stability of each of the epigenetic states in sir1 cells. See also  Xu et al., Mol. Cell 2006.

A rapid technique for the visualization of growing immobilized yeast cells, here applied to fluorescent reporters at the silent mating loci HML and HMR

Szybkie Technika Wizualizacja żywo unieruchomionych komórek drożdży

We present here a simple, rapid, and extremely flexible technique for the immobilization and visualization of growing yeast cells by epifluorescence ...

The Preparation of Primary Hematopoietic Cell Cultures From Murine Bone Marrow for Electroporation

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser MXcell  electroporation system and Gene Pulser  electroporation buffer were specifically developed to transfect nucleic acids into mammalian cells and difficult-to-transfect cells, such as primary and stem cells.This video demonstrates how to establish primary hematopoietic cell cultures from murine bone marrow, and then prepare them for electroporation in the MXcell system. We begin by isolating femur and tibia. Bone marrow from both femur and tibia are then harvested and cultures are established. Cultured bone marrow cells are then transfected and analyzed.

This procedure describes how to establish primary hematopoietic cell cultures from murine bone marrow and is followed by transfection using the Gene Pulser MXCell electroporation system.

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser ...

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.
Telomerase is a specialized reverse transcriptase1 that adds short DNA repeats, called telomeres, to the 3' end of linear chromosomes2 that serve to protect them from end-to-end fusion and degradation. Following DNA replication, a short segment is lost at the end of the chromosome3 and without telomerase, cells continue dividing until eventually reaching their Hayflick Limit4. Additionally, telomerase is dormant in most somatic cells5 in adults, but is active in cancer cells6 where it promotes cell immortality7.
The minimal telomerase enzyme consists of two core components: the protein subunit (TERT), which comprises the catalytic subunit of the enzyme and an integral RNA component (TER), which contains the template TERT uses to synthesize telomeres8,9. Prior to 2008, only structures for individual telomerase domains had been solved10,11. A major breakthrough in this field came from the determination of the crystal structure of the active12, catalytic subunit of T. castaneum telomerase, TcTERT1.
Here, we present methods for producing large quantities of the active, soluble TcTERT for structural and biochemical studies, and the reconstitution of the telomerase RNP complex in vitro for telomerase activity assays. An overview of the experimental methods used is shown in Figure 1.

In vitro Reconstitution of the Active T. castaneum Telomerase

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more than a decade. Here, we present methods for the isolation of the recombinant Tribolium castaneum TERT (TcTERT) and the reconstitution of the active T. castaneum telomerase ribonucleoprotein (RNP) complex in vitro.

Efforts to isolate the catalytic subunit of telomerase, TERT, in sufficient quantities for structural studies, have been met with limited success for more ...

Genome-wide Gene Deletions in Streptococcus sanguinis by High Throughput PCR

Transposon mutagenesis and single-gene deletion are two methods applied in genome-wide gene knockout in bacteria 1,2. Although transposon mutagenesis is less time consuming, less costly, and does not require completed genome information, there are two weaknesses in this method: (1) the possibility of a disparate mutants in the mixed mutant library that counter-selects mutants with decreased competition; and (2) the possibility of partial gene inactivation whereby genes do not entirely lose their function following the insertion of a transposon. Single-gene deletion analysis may compensate for the drawbacks associated with transposon mutagenesis. To improve the efficiency of genome-wide single gene deletion, we attempt to establish a high-throughput technique for genome-wide single gene deletion using Streptococcus sanguinis as a model organism. Each gene deletion construct in S. sanguinis genome is designed to comprise 1-kb upstream of the targeted gene, the aphA-3 gene, encoding kanamycin resistance protein, and 1-kb downstream of the targeted gene. Three sets of primers F1/R1, F2/R2, and F3/R3, respectively, are designed and synthesized in a 96-well plate format for PCR-amplifications of those three components of each deletion construct. Primers R1 and F3 contain 25-bp sequences that are complementary to regions of the aphA-3 gene at their 5' end. A large scale PCR amplification of the aphA-3 gene is performed once for creating all single-gene deletion constructs. The promoter of aphA-3 gene is initially excluded to minimize the potential polar effect of kanamycin cassette. To create the gene deletion constructs, high-throughput PCR amplification and purification are performed in a 96-well plate format. A linear recombinant PCR amplicon for each gene deletion will be made up through four PCR reactions using high-fidelity DNA polymerase. The initial exponential growth phase of S. sanguinis cultured in Todd Hewitt broth supplemented with 2.5% inactivated horse serum is used to increase competence for the transformation of PCR-recombinant constructs. Under this condition, up to 20% of S. sanguinis cells can be transformed using ~50 ng of DNA. Based on this approach, 2,048 mutants with single-gene deletion were ultimately obtained from the 2,270 genes in S. sanguinis excluding four gene ORFs contained entirely within other ORFs in S. sanguinis SK36 and 218 potential essential genes. The technique on creating gene deletion constructs is high throughput and could be easy to use in genome-wide single gene deletions for any transformable bacteria.

Genome-wide Gene Deletions in Streptococcus sanguinis by High Throughput PCR

An efficient genome-wide single gene mutation method has been established using Streptococcus sanguinis as a model organism. This method has achieved via high throughput recombinant PCRs and transformations.

Transposon  mutagenesis and single-gene deletion are two methods applied in genome-wide  gene knockout in bacteria 1,2. Although transposon mutagenesis is ...

Budding Yeast Protein Extraction and Purification for the Study of Function, Interactions, and Post-translational Modifications

Homogenization by bead beating is a fast and efficient way to release DNA, RNA, proteins, and metabolites from budding yeast cells, which are notoriously hard to disrupt. Here we describe the use of a bead mill homogenizer for the extraction of proteins into buffers optimized to maintain the functions, interactions and post-translational modifications of proteins. Logarithmically growing cells expressing the protein of interest are grown in a liquid growth media of choice. The growth media may be supplemented with reagents to induce protein expression from inducible promoters (e.g. galactose), synchronize cell cycle stage (e.g. nocodazole), or inhibit proteasome function (e.g. MG132). Cells are then pelleted and resuspended in a suitable buffer containing protease and/or phosphatase inhibitors and are either processed immediately or frozen in liquid nitrogen for later use. Homogenization is accomplished by six cycles of 20 sec&#160;bead-beating (5.5 m/sec), each followed by one minute incubation on ice. The resulting homogenate is cleared by centrifugation and small particulates can be removed by filtration. The resulting cleared whole cell extract (WCE) is precipitated using 20% TCA for direct analysis of total proteins by SDS-PAGE followed by Western blotting. Extracts are also suitable for affinity purification of specific proteins, the detection of post-translational modifications, or the analysis of co-purifying proteins. As is the case for most protein purification protocols, some enzymes and proteins may require unique conditions or buffer compositions for their purification and others may be unstable or insoluble under the conditions stated. In the latter case, the protocol presented may provide a useful starting point to empirically determine the best bead-beating strategy for protein extraction and purification. We show the extraction and purification of an epitope-tagged SUMO E3 ligase, Siz1, a cell cycle regulated protein that becomes both sumoylated and phosphorylated, as well as a SUMO-targeted ubiquitin ligase subunit, Slx5.

The preparation of high quality yeast cell extracts is a necessary first step in the analysis of individual proteins or entire proteomes. Here we describe a fast, efficient, and reliable homogenization protocol for budding yeast cells that has been optimized to preserve protein functions, interactions, and post-translational modifications.

Homogenization by bead beating is a fast and efficient way to release DNA, RNA, proteins, and metabolites from budding yeast cells, which are notoriously ...

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.

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 ...

Isolating Potentiated Hsp104 Variants Using Yeast Proteinopathy Models

Many protein-misfolding disorders can be modeled in the budding yeast Saccharomyces cerevisiae. Proteins such as TDP-43 and FUS, implicated in amyotrophic lateral sclerosis, and &#945;-synuclein, implicated in Parkinson&#8217;s disease, are toxic and form cytoplasmic aggregates in yeast. These features recapitulate protein pathologies observed in patients with these disorders. Thus, yeast are an ideal platform for isolating toxicity suppressors from libraries of protein variants. We are interested in applying protein disaggregases to eliminate misfolded toxic protein conformers. Specifically, we are engineering Hsp104, a hexameric AAA+ protein from yeast that is uniquely capable of solubilizing both disordered aggregates and amyloid and returning the proteins to their native conformations. While Hsp104 is highly conserved in eukaryotes and eubacteria, it has no known metazoan homologue. Hsp104 has only limited ability to eliminate disordered aggregates and amyloid fibers implicated in human disease. Thus, we aim to engineer Hsp104 variants to reverse the protein misfolding implicated in neurodegenerative disorders. We have developed methods to screen large libraries of Hsp104 variants for suppression of proteotoxicity in yeast. As yeast are prone to spontaneous nonspecific suppression of toxicity, a two-step screening process has been developed to eliminate false positives. Using these methods, we have identified a series of potentiated Hsp104 variants that potently suppress the toxicity and aggregation of TDP-43, FUS, and &#945;-synuclein. Here, we describe this optimized protocol, which could be adapted to screen libraries constructed using any protein backbone for suppression of toxicity of any protein that is toxic in yeast.

Yeast proteinopathy models are valuable tools to assess the toxicity and aggregation of proteins implicated in disease. Here, we present methods for screening Hsp104 variant libraries for toxicity suppressors. This protocol could be adapted to screen any protein library for toxicity suppressors of any protein that is toxic in yeast.

Many protein-misfolding disorders can be modeled in the budding yeast Saccharomyces cerevisiae. Proteins such as TDP-43 and FUS, implicated in amyotrophic ...

Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific eukaryotic genome engineering. CRISPR/Cas9 is an inexpensive, facile, and efficient genome editing tool that allows genetic perturbation of genes and genetic elements. Here we present a simple methodology for CRISPR design, cloning, and delivery for the production of genomic deletions. In addition, we describe techniques for deletion, identification, and characterization. This strategy relies on cellular delivery of a pair of chimeric single guide RNAs (sgRNAs) to create two double strand breaks (DSBs) at a locus in order to delete the intervening DNA segment by non-homologous end joining (NHEJ) repair. Deletions have potential advantages as compared to single-site small indels given the efficiency of biallelic modification, ease of rapid identification by PCR, predictability of loss-of-function, and utility for the study of non-coding elements. This approach can be used for efficient loss-of-function studies of genes and genetic elements in mammalian cell lines.

CRISPR/Cas9 is a robust system to produce disruption of genes and genetic elements. Here we describe a protocol for the efficient creation of genomic deletions in mammalian cell lines using CRISPR/Cas9.

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific ...

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 ...