We thank Dr. R. Michael Sramkoski for help with flow cytometry, Dr. Purnima Jaiswal and Dr. Di Wu for earlier input and Ilya Aylyarov for help with the illustrations. Supported by NIH Grant R01GM089872 and the Cytometry &amp; Imaging Microscopy Core Facility of the Case Comprehensive Cancer Center (P30 CA43703).

1. Haploid Cell Growth
<ol>
 <li>Grow S. cerevisiae cells from the S288C background (BY4741) under standard conditions 10.</li>
 <li>Transform haploids to express either of two conspicuous fluorescent markers 10. The markers of choice are soluble GFP expressed in the cytoplasm (pEG220, URA3/YIp, linearized with BglII 11) and a single ribosomal protein, RPL25, fused with mCherry (pAJ1661, URA3/CEN, from A. Johnston). We expect that equivalent two-color purification protocols could make use of other brightly fluorescent proteins that are synthesized by the haploid parental cells. Alternatively, the cell wall could be labeled with fluorescent lectins.</li>
 <li>On the day preceding zygote purification, inoculate 5 ml cultures of each cell type and grow them with shaking overnight at 23 &deg;C to reach A600 not exceeding 1.0. Grow GFP transformants (ATY4442) in complete synthetic medium including 2% glucose (CSM-glucose) and RPL25-mCherry transformants (ATY4552) in synthetic selective medium lacking uracil 10. Maintain parallel cultures of each cell type with shaking at room temperature.</li>
</ol>
2. Conducting a Cross
<ol>
 <li>Adjust the optical density (A600) of each sample to 1.0 in fresh medium. [1 Optical Density Unit ~ 3 x 107 cell/ml]</li>
 <li>Mix equal volumes of the two cultures, vortex briefly 3 times, followed by one longer vortex for 10 sec.</li>
 <li>Prepare CSM-glucose plates. The plates are identical to those used for routine growth and have been dried 45 min after pouring and solidification of the agar 10.</li>
 <li>Distribute as many as twenty 40-ml droplets of the mixture 1 cm from the edge of the surface of CSM-glucose plates. When the droplets are first applied to the surface start the timer to measure time elapsed. Once the liquid has been absorbed (approximately 30 min), cover the plates and leave them undisturbed at room temperature.</li>
</ol>
3. Harvesting Preparations
<ol>
 <li>Harvest samples after 1.75-2.5 hr at 23 &deg;C by repeatedly washing individual dried droplets off the plate using 150 ml cold CSM-glucose, keeping the plates on ice. Repeatedly wash the droplet area 10-20 times with the same medium to maximize recovery. Collect in 15 ml Falcon tubes on ice.</li>
 <li>Probe sonicate the samples 10x at 60% amplitude with a Fisher FB120 sonicator (120Watt, 120Volt, Freq 20KHz) on ice.</li>
 <li>Verify in a phase microscope that few or no aggregates remain. 15-50% of the cells are expected to have formed zygotes at this point.</li>
 <li>Leave the tube of cells in a vertical position undisturbed on ice for 10 min as a further precautionary measure (to allow aggregates to settle).</li>
 <li>Recover the topmost 90% of the volume and filter it through a nylon mesh (BD Falcon Polypropylene tube with strainer cap (Ref 352235)). In a typical experiment, 107 cells recovered from one plate were at this point diluted into 4 ml of CSM-glucose.</li>
</ol>
4. Flow Cytometry
<ol>
 <li>Sort cells using the Sony iCyt Reflection Model Flow Cytometer, (Highly Automated Parallel Sort) HAPS1 with a 100 micron tip. For GFP excitation, use the blue argon-ion, 250 milliwatt laser at 488 nm. For mCherry excitation, use the 100 milliwatt laser at 561 nm. Although we do not have experience with other machines, we expect that Becton Dickinson FACSAria, Becton Dickson Influx or Beckman Coulter MoFlo XDP would be fine when equipped with the correct lasers.</li>
 <li>Base sorting parameters on a forward vs. back scatter plot in order to gate on viable cells. Establish an emission plot of 615 +/- 30 nm bandpass filter (mCherry) versus 525 +/- 25 nm bandpass filter (GFP) to divide the events into four quadrants, "Red+ Green-", "Red+ Green+", "Red- Green-", and "Red- Green+" (Figure S2). Use nonfluorescent cells as a standard for determination of the "Red- Green-" quadrant.</li>
 <li>Sort cells with a sheath pressure of 21.5 psi, a droplet frequency of 43,400 drops/sec, and a sample pressure of 20.5 psi.</li>
 <li>Initial Sort ("Recovery Mode"): Collect samples into chilled Eppendorf tubes containing 250 ml of cold CSM-glucose. Sediment 10 sec at top speed in a refrigerated microfuge (Tomy MRX-150, 15,000 rpm) and aspirate the flow solution (Flow cytometry sheath solution (PBS with 1 mM EDTA) BioSure Catalog No. 1027). Resuspend in 500 ml cold CSM-glucose.</li>
 <li>Second Sort ("Purity Mode"): Conduct the sort as for the initial sort, using the "purity mode" option. Sediment the purified cells and resuspend the pellets in 500 ml-1.0 ml CSM-glucose. Shake for 15 min in 24-well plates at 23 &deg;C to allow metabolic recovery.</li>
</ol>
5. DeltaVision Microscopy
<ol>
 <li>Prepare agarose pads as follows: Bring a suspension of 1.5% Agarose (UltraPure Agarose, Invitrogen, Catalog # 15510-07) in CSM-glucose to a boil, vortex briefly, and then apply 0.3 ml samples of the agarose solution to the surface of horizontal standard glass slides and cover them at once with a second slide (without applying pressure).</li>
 <li>After cooling for 10 min, gently remove the upper slide by sliding it horizontally, taking care to leave a smooth surface and not to disturb the Agarose layer. (When the upper slide has been removed, the sample of cells should be applied within 1-2 min. When kept in a humid chamber at room temperature without removing the upper slide, pads can be stored for up to a day before use.)</li>
 <li>Sediment samples and deposit 1 ml aliquots of the pellet on agarose pads (without touching the surface). Immediately cover the sample with a square No. 1 coverslip. Trim away excess agarose with a single-edged razorblade. Seal the preparations with Vaseline dispensed from a syringe.</li>
 <li>Image using Deltavision (Applied Precision) with a 100x oil immersion objective without binning (Olympus UPlanApo 100x/1.40; &infin;/0.17/FN26.5).</li>
 <li>Deconvolve z-stacks using the softWoRx 5.0.0 program (<a href="http://www.api.com/softworx.asp" target="_blank">http://www.api.com/softworx.asp</a>) and process minimally. Successive z-planes were generally collected at 0.2-0.4 micron intervals.</li>
</ol>
6. Representative Results
Fluorescence intensity levels in transformed haploid cells: The level of fluorescence in the haploids is illustrated in Figure S1. Before the cross, epifluorescent examination demonstrated that essentially all GFP transformants were green and that all the mCherry transformants, although often fainter, were red. To ensure that all cells used for the cross were indeed fluorescent, we used the subpopulation with the brightest fluorescent signal.
Yield: After the initial sort, the Red+ Green+ sample (Figure S2) typically includes 8-20% of the input number of cells and an excessive number of haploid cells, perhaps due to persistent cell-cell adhesion. Since high enrichment is not achieved with a single sort, the partially purified sample should be probe sonicated twice at 60% amplitude followed by re-sorting. (Recovery sorts typically last between 1-2 hr, while the second sort requires only 15 min.) Of the cells subjected to the second sort, the Red+ Green+ samples included 23-52% of the input cells.
Phase microscopic examination: Examination by phase contrast microscopy showed that the purity of the final zygote preparation averaged approximately 90% over 21 preparations (ranging from 86% to 91%). The contaminants were either green or red haploids, or pairs of haploids that were in contact with each other. Figure 3 tabulates the percent of zygotes that have no bud, medial buds (Med), lateral buds (Lat) or terminal buds (Term), the number of paired haploids of the same color ("2Same") or different color ("2Hap"), as well as individual haploids (Hap). When cells were harvested after 1.75 hr, more than 50% were unbudded and the other subcategories (with medial, lateral, or terminal buds) were equally represented. After 2.5 hr there were approximately equal numbers of zygotes in each of four categories (unbudded, with medial, lateral or terminal buds).
Microscopic Observations (Figure 4):
The structure of zygotes appears to remain unchanged during purification. According to the duration of the cross, the population had different proportions of unbudded and budded zygotes, as explained above. Since both the green and red haploids showed considerable variation in fluorescent signal intensity, some zygotes were predominantly red while others were predominantly green, as is observed by comparing the single-color images to the combined images in Figure 4. Note also that the GFP (MW 26kDa) enters the nucleus and can be found in the nucleus even when the nuclei have not fused. Vacuoles stand out as black non-fluorescent spheres.
The end product of purification yields a diverse population at various stages of development. Zygotes labeled A have not undergone fusion. Type B zygotes are unbudded. Type C have a small bud. Type D have a larger bud, and Type E zygotes are about to undergo cytokinesis. Note also that many of the earlier zygotes still exhibit a non-fluorescent medial division line (*), and that the vacuolar "inheritance structure" (I) 12 can often be seen to extend from the zygote into the bud. In some cases, the green signal has redistributed into the partner to a much greater extent that the red signal. This reflects the delay of transfer that is characteristic of polysomes (Tartakoff, in preparation).
Starting with a pair of 5 ml logarithmic cultures of haploid cells, we recover approximately 1-2 x 106 zygotes, which is sufficient for many immunoblots, using chemiluminescent detection procedures. Using phenol-chloroform isolation protocols we extract approximately 1.5 mg of RNA from this number of cells.
Since the purified zygotes have been chilled, prior to further analysis we recommend incubation at 23 &deg;C in growth medium.
<img alt="Figure 1" src="/files/ftp_upload/4197/4197fig1.jpg" /> 
Figure 1. Chronology of zygote formation. Zygotes as an intermediate between haploids and diploids. This diagram shows the classical activation (shmooing) of haploids, their fusion, and the formation of the initial bud. Buds can be either medial (M), lateral (L) or terminal (T).
<img alt="Figure 1" src="/files/ftp_upload/4197/4197fig2.jpg" /> 
Figure 2. Schematic of the zygote purification protocol. After incubation on the surface of agar plates in complete medium with glucose, the cell mixture was recovered by washing and then subjected to an initial sort in "recovery mode" followed by a second sort in "purity mode." The yellow color signifies that both mCherry and GFP signals were visible in the zygotes.
<img alt="Figure 3" src="/files/ftp_upload/4197/4197fig3.jpg" /> 
Figure 3. Varieties of zygotes in the purified fractions. Mating mixtures were harvested after 1.75 or 2.5 hr to recover fractions enriched in relatively early vs. relatively more mature zygotes. Eight independent experiments, designated by the distinct colors, were performed for each time point. The graphs indicate the relative numbers of zygotes that have spatially distinct patterns of bud formation. Given the brief duration of the incubation (less than 2.5 hr at room temperature) all buds observed are initial zygotic budding events.
<img alt="Figure 4" src="/files/ftp_upload/4197/4197fig4.jpg" /> 
Figure 4. DeltaVision examination of enriched fractions. A typical field was examined by Deltavision microscopy. The illustrations show either both red and green or single colors. The red corresponds to the polysomal label (Rpl25p-mCherry) and the green to cytosolic GFP. Note the high enrichment of zygotes, and the designated subcellular features: the vacuoles(v), buds(b), nucleus(n), vacuolar "inheritance structure"(i), and the partition(*) which appears to separate the parental domains in relatively early zygotes. Their variable overall color reflects the relative intensity of the haploids that fused. In cases in which the two parental domains retain a distinct color, equilibration of the markers (cytoplasmic GFP, mCherry-tagged polysomes) was incomplete. Time-lapse studies in progress show that soluble GFP redistributes prior to polysomes. <a href="https://www.jove.com/files/ftp_upload/4197/4197fig4large.jpg" target="_blank"> Click here to view larger figure</a>.
Supplementary Legends
<img alt="Figure S1" src="/files/ftp_upload/4197/4197figS1.jpg" /> 
Figure S1. Fluorescence of haploid cells. Far left: Cells expressing mCherry-tagged polysomes were examined to detect a green signal. The rectangular window was set to avoid the negative cells. Middle graph: Cells expressing cytosolic GFP were analyzed, showing the distinctly bright major population of positives. Far right: Cells expressing mCherry-tagged polysomes were analyzed. The rectangle (identical to that on the far left) includes the brightest cells and allows all negatives to be avoided. The designation "R2" indicates the percentage of cells recovered within the indicated rectangle.
<img alt="Figure S2" src="/files/ftp_upload/4197/4197figS2.jpg" /> 
Figure S2. Cell distribution in first and second sorting. These two-color plots indicate the proportion of Green+ Red+ cells in the upper right quadrant (R5). This proportion increases to 52% in the second sort. Horizontal axis: green signal. Vertical axis: red signal.

<ol>
	<li>Herskowitz, I., Oshima, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Control of cell type in Saccharomyces cerevisiae: Mating type and mating-type interconversion. , 181-210 (1981).</li><li>Muller, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parental+age+and+the+life-span+of+zygotes+of+Saccharomyces+cerevisiae.">Parental age and the life-span of zygotes of Saccharomyces cerevisiae.</a> Antonie Van Leeuwenhoek. 51, 1-10 (1985).</li><li>Tartakoff, A. M., Jaiswal, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nuclear+fusion+and+genome+encounter+during+yeast+zygote+formation.">Nuclear fusion and genome encounter during yeast zygote formation.</a> Mol. Biol. Cell. 20, 2932-2942 (2009).</li><li>Azpiroz, R., Butow, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterns+of+mitochondrial+sorting+in+yeast+zygotes.">Patterns of mitochondrial sorting in yeast zygotes.</a> Mol. Biol. Cell. 4, 21-36 (1993).</li><li>Rose, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nuclear+fusion+in+the+yeast+Saccharomyces+cerevisiae.">Nuclear fusion in the yeast Saccharomyces cerevisiae.</a> Annu. Rev. Cell Dev. Biol. 12, 663-695 (1996).</li><li>Dujon, B., Jones, E., Strathern, J., Broach, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Molecular Biology of the Yeast Saccharomyces: Life Cycle and Inheritance. , 505-635 (1981).</li><li>Sprague, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assay+of+yeast+mating+reaction.">Assay of yeast mating reaction.</a> Methods Enzymol. 194, 77-93 (1991).</li><li>Ydenberg, C. A., Rose, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Yeast+mating:+a+model+system+for+studying+cell+and+nuclear+fusion.">Yeast mating: a model system for studying cell and nuclear fusion.</a> Methods Mol. Biol. 475, 3-20 (2008).</li><li>Sena, E. P., Radin, D. N., Fogel, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synchronous+mating+in+yeast.">Synchronous mating in yeast.</a> Proc. Natl. Acad. Sci. U.S.A. 70, 1373-1377 (1973).</li><li>Amberg, D., Burke, D., Strathern, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Methods in Yeast Genetics: A Cold Spring Harbor Laboratory Course Manual, 2005 Edition. , (2005).</li><li>Nolan, S., Cowan, A. E., Koppel, D. E., Jin, H., Grote, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FUS1+regulates+the+opening+and+expansion+of+fusion+pores+between+mating+yeast.">FUS1 regulates the opening and expansion of fusion pores between mating yeast.</a> Mol. Biol. Cell. 17, 2439-2450 (2006).</li><li>Weisman, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organelles+on+the+move:+insights+from+yeast+vacuole+inheritance.">Organelles on the move: insights from yeast vacuole inheritance.</a> Nat. Rev. Mol. Cell Biol. 7, 243-252 (2006).</li></ol>

No conflicts of interest declared.

The availability of purified zygotes should facilitate biochemical studies including transcriptomics, proteomics and lipidomics, as well as investigation of mechanisms by which zygotes undergo cell cycle re-entry, cell wall remodeling, budding and inheritance characteristics, etc. Alternatively, zygotes could be generated starting with strains carrying characteristic mutations in one or both parents. Moreover, the purified zygotes, as for haploid or diploid cells, can be frozen in medium containing 15% glycerol and later...

<table border="1">
 <tbody>
 <tr>
 <td>Reagent/Equipment</td>
 <td>Source</td>
 <td>Catalogue Number</td>
 </tr>
 <tr>
 <td>pEG220, URA3/YIp, encoding cytoplasmic GFP 11)</td>
 <td>E. Grote</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>pAJ1661, URA3/CEN, encoding RPL25 fused to mCherry </td>
 <td>A. Johnston</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fisher FB120 sonicator</td>
 <td>Fisher Scientific</td>
 <td></td>
 </tr>
 <tr>
 <td>Polypropylene tube with strainer cap</td>
 <td>BD Falcon</td>
 <td>Ref 352235</td>
 </tr>
 <tr>
 <td>Refrigerated microfuge </td>
 <td>Tomy</td>
 <td>MRX-150</td>
 </tr>
 <tr>
 <td>Flow cytometry sheath solution</td>
 <td>Biosure</td>
 <td>1027</td>
 </tr>
 <tr>
 <td>iCyt Reflection Model Flow Cytometer</td>
 <td>Sony</td>
 <td></td>
 </tr>
 <tr>
 <td>DeltaVision deconvolution microscope</td>
 <td>Applied Precision</td>
 <td></td>
 </tr>
 <tr>
 <td>Upright epifluorescence microscope</td>
 <td>Leica</td>
 <td></td>
 </tr>
 <tr>
 <td>UltraPure Agarose</td>
 <td>Invitrogen</td>
 <td>15510-07</td>
 </tr>
 <tr>
 <td>CSM glucose formulation Notes: For agar plates, simply include 2% agar (Difco)</td>
 <td>10</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

flow cytometry-based purification of s. cerevisiae zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae zygotes result from the fusion of haploid cells of distinct mating type (MATa, MATalpha) and give rise to corresponding stable diploids that successively generate as many as 20 diploid progeny as a result of their strikingly asymmetric mitotic divisions 2. Zygote formation is orchestrated by a complex sequence of events: In this process, soluble mating factors bind to cognate receptors, triggering receptor-mediated signaling cascades that facilitate interruption of the cell cycle and culminate in cell-cell fusion. Zygotes may be considered a model for progenitor or stem cell function.
Although much has been learned about the formation of zygotes and although zygotes have been used to investigate cell-molecular questions of general significance, almost all studies have made use of mating mixtures in which zygotes are intermixed with a majority population of haploid cells 3-8. Many aspects of the biochemistry of zygote formation and the continuing life of the zygote therefore remain uninvestigated.
Reports of purification of yeast zygotes describe protocols based on their sedimentation properties 9; however, this sedimentation-based procedure did not yield nearly 90% purity in our hands. Moreover, it has the disadvantage that cells are exposed to hypertonic sorbitol. We therefore have developed a versatile purification procedure. For this purpose, pairs of haploid cells expressing red or green fluorescent proteins were co-incubated to allow zygote formation, harvested at various times, and the resulting zygotes were purified using a flow cytometry-based sorting protocol. This technique provides a convenient visual assessment of purity and maturation. The average purity of the fraction is approximately 90%. According to the timing of harvest, zygotes of varying degrees of maturity can be recovered. The purified samples provide a convenient point of departure for "-omic" studies, for recovery of initial progeny, and for systematic investigation of this progenitor cell.

Watch this Scientific Journal Video about Flow Cytometry-based Purification of S. cerevisiae Zygotes at JoVE.com

Flow Cytometry-based Purification of S. cerevisiae Zygotes

To purify zygotes of S. cerevisiae, haploid cells of opposite mating type were engineered to express red or green fluorescent proteins, co-incubated to allow zygote formation, and fractionated using a flow cytometry-based protocol. The highly-enriched fraction enables subsequent "-omic" studies, recovery of initial progeny, and systematic investigation of zygote morphogenesis.

Flow Cytometry-based Purification of S. cerevisiae Zygotes

Zygotes are essential intermediates between haploid and diploid states in the life cycle of many organisms, including yeast (Figure 1) 1. S. cerevisiae ...

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12.7K ビュー.  Case Western Reserve University School of Medicine. の受精卵を精製するために S.セレビ、反対の交配型の半数体細胞をフローサイトメトリーベースのプロトコルを使用して接合体​​形成を可能にするために共インキュベート、赤色または緑色蛍光タンパク質を発現するように操作し、分画した。高濃縮画分は初期の子孫のその後 "-OMIC"研究、リカバリ、および受精卵形態形成の体系的?...

ビデオ: Flow Cytometry-based Purification of S. cerevisiae Zygotes

Assay for Adhesion and Agar Invasion in S. cerevisiae

Yeasts are found in natural biofilms, where many microorganisms colonize surfaces. In artificial environments, such as surfaces of man-made objects, biofilms can reduce industrial productivity, destroy structures, and threaten human life. 1-3 On the other hand, harnessing the power of biofilms can help clean the environment and generate sustainable energy. 4-8
The ability of S. cerevisiae to colonize surfaces and participate in complex biofilms was mostly ignored until the rediscovery of the differentiation programs triggered by various signaling pathways and environmental cues in this organism. 9, 10 The continuing interest in using S. cerevisiae as a model organism to understand the interaction and convergence of signaling pathways, such as the Ras-PKA, Kss1 MAPK, and Hog1 osmolarity pathways, quickly placed S. cerevisiae in the junction of biofilm biology and signal transduction research. 11-20 To this end, differentiation of yeast cells into long, adhesive, pseudohyphal filaments became a convenient readout for the activation of signal transduction pathways upon various environmental changes. However, filamentation is a complex collection of phenotypes, which makes assaying for it as if it were a simple phenotype misleading. In the past decade, several assays were successfully adopted from bacterial biofilm studies to yeast research, such as MAT formation assays to measure colony spread on soft agar and crystal violet staining to quantitatively measure cell-surface adherence. 12, 21 However, there has been some confusion in assays developed to qualitatively assess the adhesive and invasive phenotypes of yeast in agar.
Here, we present a simple and reliable method for assessing the adhesive and invasive quality of yeast strains with easy-to-understand steps to isolate the adhesion assessment from invasion assessment. Our method, adopted from previous studies, 10, 16 involves growing cells in liquid media and plating on differential nutrient conditions for growth of large spots, which we then wash with water to assess adhesion and rub cells completely off the agar surface to assess invasion into the agar. We eliminate the need for streaking cells onto agar, which affects the invasion of cells into the agar. In general, we observed that haploid strains that invade agar are always adhesive, yet not all adhesive strains can invade agar medium. Our approach can be used in conjunction with other assays to carefully dissect the differentiation steps and requirements of yeast signal transduction, differentiation, quorum sensing, and biofilm formation.

We describe a qualitative assay for yeast adhesion and agar invasion as a measure of invasive and pseudohyphal differentiation. This simple assay can be used to assess the invasive phenotype of various mutants as well as the effects environmental cues and signaling pathways on yeast differentiation.

Yeasts are found in natural biofilms, where many microorganisms colonize surfaces. In artificial environments, such as surfaces of man-made objects, ...

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 ...

Identification of protein complexes with quantitative proteomics in S. cerevisiae

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as important intracellular signalling molecules. However, unlike proteins, lipids are small hydrophobic molecules that traffic primarily by poorly described nonvesicular routes, which are hypothesized to occur at membrane contact sites (MCSs). MCSs are regions where the endoplasmic reticulum (ER) makes direct physical contact with a partnering organelle, e.g., plasma membrane (PM). The ER portion of ER-PM MCSs is enriched in lipid-synthesizing enzymes, suggesting that lipid synthesis is directed to these sites and implying that MCSs are important for lipid traffic. Yeast is an ideal model to study ER-PM MCSs because of their abundance, with over 1000 contacts per cell, and their conserved nature in all eukaryotes. Uncovering the proteins that constitute MCSs is critical to understanding how lipids traffic is accomplished in cells, and how they act as signaling molecules. We have found that an ER called Scs2p localize to ER-PM MCSs and is important for their formation. We are focused on uncovering the molecular partners of Scs2p. Identification of protein complexes traditionally relies on first resolving purified protein samples by gel electrophoresis, followed by in-gel digestion of protein bands and analysis of peptides by mass spectrometry. This often limits the study to a small subset of proteins. Also, protein complexes are exposed to denaturing or non-physiological conditions during the procedure. To circumvent these problems, we have implemented a large-scale quantitative proteomics technique to extract unbiased and quantified data. We use stable isotope labeling with amino acids in cell culture (SILAC) to incorporate staple isotope nuclei in proteins in an untagged control strain. Equal volumes of tagged culture and untagged, SILAC-labeled culture are mixed together and lysed by grinding in liquid nitrogen. We then carry out an affinity purification procedure to pull down protein complexes. Finally, we precipitate the protein sample, which is ready for analysis by high-performance liquid chromatography/ tandem mass spectrometry. Most importantly, proteins in the control strain are labeled by the heavy isotope and will produce a mass/ charge shift that can be quantified against the unlabeled proteins in the bait strain. Therefore, contaminants, or unspecific binding can be easily eliminated. By using this approach, we have identified several novel proteins that localize to ER-PM MCSs. Here we present a detailed description of our approach. 

Here we describe a new quantitative proteomics technique for identifying protein complexes in Saccharomyces cerevisiae. In this study, we have used the SILAC method together with affinity purification followed by tandem mass spectrometry to identify with high specificity the binding partners of an ER protein, Scs2p.

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as ...

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

Studying Age-dependent Genomic Instability using the S. cerevisiae Chronological Lifespan Model

Studies using the Saccharomyces cerevisiae aging model have uncovered life span regulatory pathways that are partially conserved in higher eukaryotes1-2. The simplicity and power of the yeast aging model can also be explored to study DNA damage and genome maintenance as well as their contributions to diseases during aging. Here, we describe a system to study age-dependent DNA mutations, including base substitutions, frame-shift mutations, gross chromosomal rearrangements, and homologous/homeologous recombination, as well as nuclear DNA repair activity by combining the yeast chronological life span with simple DNA damage and mutation assays. The methods described here should facilitate the identification of genes/pathways that regulate genomic instability and the mechanisms that underlie age-dependent DNA mutations and cancer in mammals.

Studying Age-dependent Genomic Instability using the S. cerevisiae Chronological Lifespan Model

Here we describe a set of DNA mutation assays that can be combined with the yeast chronological life span model to study the genes/pathways that regulate or contribute to genomic DNA instability during aging.

Studies using the Saccharomyces cerevisiae aging model have uncovered life span regulatory pathways that are partially conserved in higher eukaryotes1-2. ...

Expression and Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein in Saccharomyces cerevisiae

The cystic fibrosis transmembrane conductance regulator (CFTR) is a chloride channel, that when mutated, can give rise to cystic fibrosis in humans.There is therefore considerable interest in this protein, but efforts to study its structure and activity have been hampered by the difficulty of expressing and purifying sufficient amounts of the protein1-3. Like many 'difficult' eukaryotic membrane proteins, expression in a fast-growing organism is desirable, but challenging, and in the yeast S. cerevisiae, so far low amounts were obtained and rapid degradation of the recombinant protein was observed 4-9. Proteins involved in the processing of recombinant CFTR in yeast have been described6-9 .In this report we describe a methodology for expression of CFTR in yeast and its purification in significant amounts. The protocol describes how the earlier proteolysis problems can be overcome and how expression levels of CFTR can be greatly improved by modifying the cell growth conditions and by controlling the induction conditions, in particular the time period prior to cell harvesting. The reagants associated with this protocol (murine CFTR-expressing yeast cells or yeast plasmids) will be distributed via the US Cystic Fibrosis Foundation, which has sponsored the research. An article describing the design and synthesis of the CFTR construct employed in this report will be published separately (Urbatsch, I.; Thibodeau, P. et al., unpublished). In this article we will explain our method beginning with the transformation of the yeast cells with the CFTR construct - containing yeast plasmid (Fig. 1). The construct has a green fluorescent protein (GFP) sequence fused to CFTR at its C-terminus and follows the system developed by Drew et al. (2008)10. The GFP allows the expression and purification of CFTR to be followed relatively easily. The JoVE visualized protocol finishes after the preparation of microsomes from the yeast cells, although we include some suggestions for purification of the protein from the microsomes. Readers may wish to add their own modifications to the microsome purification procedure, dependent on the final experiments to be carried out with the protein and the local equipment available to them. The yeast-expressed CFTR protein can be partially purified using metal ion affinity chromatography, using an intrinsic polyhistidine purification tag. Subsequent size-exclusion chromatography yields a protein that appears to be &gt;90% pure, as judged by SDS-PAGE and Coomassie-staining of the gel.

Expression and Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein in Saccharomyces cerevisiae

Attempts to express the cystic fibrosis transmembrane conductance regulator (CFTR) in Saccharomyces cerevisiae have, until now, yielded relatively low amounts of protein. This protocol and the associated reagents distributed via the Cystic Fibrosis Foundation should allow the preparation of milligram amounts of this 'difficult' eukaryotic membrane protein.

The  cystic fibrosis transmembrane conductance regulator (CFTR) is a chloride  channel, that when mutated, can give rise to cystic fibrosis in  ...

Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of different cell types in vitro and in vivo.1-5 Although there are literally thousands of publications using such dyes, some of the most commonly encountered cell tracking applications include monitoring of:
<ol>
	<li>stem and progenitor cell quiescence, proliferation and/or differentiation6-8</li>
	<li>antigen-driven membrane transfer9 and/or precursor cell proliferation3,4,10-18 and</li>
	<li>immune regulatory and effector cell function1,18-21.</li>
</ol>
Commercially available cell tracking dyes vary widely in their chemistries and fluorescence properties but the great majority fall into one of two classes based on their mechanism of cell labeling. "Membrane dyes", typified by PKH26, are highly lipophilic dyes that partition stably but non-covalently into cell membranes1,2,11. "Protein dyes", typified by CFSE, are amino-reactive dyes that form stable covalent bonds with cell proteins4,16,18. Each class has its own advantages and limitations. The key to their successful use, particularly in multicolor studies where multiple dyes are used to track different cell types, is therefore to understand the critical issues enabling optimal use of each class2-4,16,18,24.
The protocols included here highlight three common causes of poor or variable results when using cell-tracking dyes. These are:
<ol>
	<li>Failure to achieve bright, uniform, reproducible labeling. This is a necessary starting point for any cell tracking study but requires attention to different variables when using membrane dyes than when using protein dyes or equilibrium binding reagents such as antibodies.</li>
	<li>Suboptimal fluorochrome combinations and/or failure to include critical compensation controls. Tracking dye fluorescence is typically 102 - 103 times brighter than antibody fluorescence. It is therefore essential to verify that the presence of tracking dye does not compromise the ability to detect other probes being used.</li>
	<li>Failure to obtain a good fit with peak modeling software. Such software allows quantitative comparison of proliferative responses across different populations or stimuli based on precursor frequency or other metrics. Obtaining a good fit, however, requires exclusion of dead/dying cells that can distort dye dilution profiles and matching of the assumptions underlying the model with characteristics of the observed dye dilution profile.</li>
</ol>
Examples given here illustrate how these variables can affect results when using membrane and/or protein dyes to monitor cell proliferation.

Successful use of cell tracking dyes to monitor immune cell function and proliferation involves several critical steps. We describe methods for: 1) obtaining bright, uniform, reproducible label-ing with membrane dyes; 2) selecting fluorochromes and data acquisition conditions; and 3) choosing a model to quantify cell proliferation based on dye dilution.

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of ...

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...

Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein Expressed in Saccharomyces cerevisiae

Defects in the cystic fibrosis transmembrane conductance regulator (CFTR) protein cause cystic fibrosis (CF), an autosomal recessive disease that currently limits the average life expectancy of sufferers to &#60;40 years of age. The development of novel drug molecules to restore the activity of CFTR is an important goal in the treatment CF, and the isolation of functionally active CFTR is a useful step towards achieving this goal.
We describe two methods for the purification of CFTR from a eukaryotic heterologous expression system, S. cerevisiae. Like prokaryotic systems, S. cerevisiae can be rapidly grown in the lab at low cost, but can also traffic and posttranslationally modify large membrane proteins. The selection of detergents for solubilization and purification is a critical step in the purification of any membrane protein. Having screened for the solubility of CFTR in several detergents, we have chosen two contrasting detergents for use in the purification that allow the final CFTR preparation to be tailored to the subsequently planned experiments.
In this method, we provide comparison of the purification of CFTR in dodecyl-&#946;-D-maltoside (DDM) and 1-tetradecanoyl-sn-glycero-3-phospho-(1&#39;-rac-glycerol) (LPG-14). Protein purified in DDM by this method shows ATPase activity in functional assays. Protein purified in LPG-14 shows high purity and yield, can be employed to study post-translational modifications, and can be used for structural methods such as small-angle X-ray scattering and electron microscopy. However it displays significantly lower ATPase activity.

Purification of the Cystic Fibrosis Transmembrane Conductance Regulator Protein Expressed in Saccharomyces cerevisiae

Heterologous expression and purification of the cystic fibrosis transmembrane conductance regulator (CFTR) are significant challenges and limiting factors in the development of drug therapies for cystic fibrosis. This protocol describes two methods for the isolation of milligram quantities of CFTR suitable for functional and structural studies.&#160;