Name: quantitation and analysis of the formation of ho-endonuclease stimulated chromosomal translocations by single-strand annealing in saccharomyces cerevisiae
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Description: Watch this Scientific Journal Video about Quantitation and Analysis of the Formation of HO-Endonuclease Stimulated Chromosomal Translocations by Single-Strand Annealing in Saccharomyces cerevisiae at

This work was supported by funds from the National Institutes of Health and the Beckman Research Institute of the City of Hope. We would like to thank the reviewers for their constructive comments that added clarity to the manuscript.

1. HO-stimulated translocation frequencies
<ol>
	<li>Inoculate 10-20 independent 1 ml cultures of YPGly/Lac medium (1% yeast extract, 2% peptone, 3% glycerol and 3% lactate) with single colonies of the desired genotype. Incubate the cultures overnight, or for sufficient time to reach a cell density of approximately 5x107-1x108 cell/ml, at 30&deg;C on a rotator, or with gentle agitation. </li>
	<li>Add galactose to the cultures to a final concentration of 2% to induce HO endonuclease-directed DSBs at the his3-&Delta;3' (chromosome III) and his3-&Delta;5' (chromosome XV) translocation substrates. </li>
	<li>Incubate for 4 h at 30&deg;C on a rotator, or with gentle agitation.</li>
	<li>After 4 h, plate appropriate dilutions of the cultures onto YPD (1% yeast extract, 2% peptone, 2% dextrose) to yield approximately 100 to 200 colonies per plate, and a sufficient number of cells onto medium lacking histidine to yield an observable number of His+ recombinant colonies. Incubate plates at 30&deg;C for two to three days. </li>
</ol>
NOTE: Step 1.4 describes the determination of translocation frequencies under selective conditions by plating the cultures onto medium lacking histidine. This assay can also be conducted under non-selective conditions by plating cultures onto YPD, and then replica-plating the colonies that arise onto &#x2013;His plates two to three days later. These methods produce similar frequencies of translocation (Figure 2C). 
<ol start="5">
	<li>Determine the translocation frequency by dividing the number of histidine prototrophic colonies by the total number of viable cells plated (determined by plating dilutions on YPD). Determine the median translocation frequency and 95% confidence interval15.</li>
</ol>
2. Plating efficiencies
<ol>
	<li>Remove an aliquot of cells from each overnight culture, and determine cell number by hemacytometer count (Baxter Healthcare Corporation. Catolog #: B3178-1).</li>
	<li>Plate approximately 100 to 200 cells using an appropriate dilution onto YPD. Incubate for two to three days at 30&deg;C (e.g. For a culture with a cell density of 1x108 cells/ml, one should plate 100-200 &mu;l of a 10-5 dilution per plate).</li>
	<li>Add 20% galactose to the cultures to a final concentration of 2%.</li>
	<li>After 4 h, remove an aliquot of cells from each culture, determine cell number by hemacytometer count, and plate approximately 100 to 200 cells using an appropriate dilution onto YPD. Incubate for two to three days at 30&deg;C.</li>
	<li>Determine the plating efficiency by dividing the number of colonies appearing on YPD by the number of cells plated, and multiplying this quotient by 100. Determine the median percentage with a 95% confidence interval. </li>
</ol>
3. Genomic Southern blot analysis
<ol>
	<li>Select a single His+ recombinant colony from each independent trial and prepare genomic DNA16.</li>
	<li>Digest approximately 4 &mu;g of DNA with BamHI restriction endonuclease.</li>
	<li>Separate BamHI digested fragments on a 0.7% agarose gel, and transfer to a positively-charged nylon membrane (Hybond N+, GE Healthcare. Product Code: RPN303B)17.</li>
	<li>Hybridize with a 32P-labeled probe obtained by random priming (Amersham Biosciences. Product Code: RPN1604) with a 1.8 kb BamHI/BamHI genomic clone containing the HIS3 gene.</li>
	<li>Visualize DNA fragments by autoradiography or phosphorimaging.</li>
</ol>
4. Chromosome blot analysis using chromosomes separated in a contour-clamped homogeneous electric field (CHEF):
<ol>
	<li>Prepare chromosomes from selected His+ recombinants in agarose plugs18:
		<ul>
			<li>Grow a liquid culture of the His+ candidate tXV::III chromosome-containing recombinant in 5 ml of YPD to approximately 1-2 x 108 cell/ml.</li>
			<li>Spin down and wash the cells 2 times with 50 mM EDTA.</li>
			<li>Resuspend cells in approximately 200 &mu;l of 50 mM EDTA, and warm to 50&deg;C.</li>
			<li>Add an equal volume of molten 2% (w/v) low melt agarose brought to 50&deg;C, mix thoroughly.</li>
			<li>Dispense approximately 80 &mu;l aliquots into plug molds, and allow them to cool for 30 min at 4&deg;C.</li>
			<li>Extrude plugs from the mold into a 12-well dish. Up to five plugs can be placed into each well.</li>
			<li>Add 3 ml of freshly prepared spheroplasting solution (14 mM 2-&beta;-mercaptoethanol, 20 mM EDTA, 0.5 mg/ml Zymolyase 20T, 10 mM Tris-HCl, pH 7.5, 1 M Sorbitol) to each well. Incubate at 37&deg;C for 4 hours with gentle agitation.</li>
			<li>Remove spheroplasting solution and replace with 3 ml of LDS solution (10 mM Tris-HCl, pH 8.0, 100 mM EDTA, 1% (w/v) Lithium Dodecyl Sulfate, adjust pH to 8.0). Incubate at 37&deg;C for 15 min with gentle agitation.</li>
			<li>Remove LDS solution and replace with another 3 ml aliquot of LDS. Incubate at 37&deg;C overnight with gentle agitation.</li>
			<li>Remove LDS and replace with 3 ml of 0.2 X NDS (0.6 g Tris Base, 93 g Disodium EDTA Dihydrate, 5 g N-Lauroyl Sarcosine, adjust pH to 8.0, brought to 500 ml with dH2O). Incubate at room temperature for 30 min with gentle agitation. Remove NDS, and repeat 2 times.</li>
			<li>Remove NDS and replace with 3 ml TE. Wash with gentle agitation for 30 min at room temperature. Repeat 4 times.</li>
			<li>Store plugs at 4&deg;C in 2 ml of TE. Plugs can keep for up to a year.</li>
		</ul>
	</li>
	<li>Separate chromosomes on a 1% agarose gel using a Bio-Rad CHEF-DRII apparatus at 14&deg;C (Catalog #: 170-3612).</li>
</ol>
Parameters: 1st Block: 70s switch time, 15h at 6V/cm. 
2nd Block: 120s switch time, 11h at 6V/cm
<ol start="3">
	<li>Visualize chromosomes by staining with 1&mu;g/ml ethidium bromide for 30 min, irradiating with 60 mJ of UV in a UV Stratalinker (Stratagene), and de-staining for 30 min in dH2O. Irradiating ethidium bromide stained chromosomes nicks the DNA to allow for efficient transfer to the membrane.</li>
	<li>Transfer chromosomes to a positively charged membrane (Hybond N+, GE Healthcare. Product Code: RPN303B) by capillary action in denaturing conditions (0.4N NaOH, 1.5M NaCl).</li>
	<li>Hybridize with a 32P-labeled probe obtained by random priming (Amersham Biosciences. Product Code: RPN1604) with a 1.8 kb BamHI/BamHI genomic clone containing the HIS3 gene17.</li>
	<li>Visualize chromosomes by autoradiography or phosphorimaging.</li>
</ol>
5. Representative Results:
A graphical representation of the translocation assay is depicted at the chromosomal level (Figure 1). A schematic of the experimental procedure is also displayed (Figure 2A). Both pre- and post-induction plating efficiencies are determined by dividing the total number of viable, colony forming cells by the total number of cell bodies in the culture determined by hemacytometer count (Figure 4B). Pre- and post-induction plating efficiencies were not significantly different for wild-type cells (p-value= 0.1400) (Table 1).
<img src="/files/ftp_upload/3150/3150table1.jpg" alt="Table 1" /> 
Table 1. Pre- and post-induction plating efficiencies in wild-type cells.
a The number of cell bodies per milliliter are determined by hemacytometer count. 
b The number of viable cells per milliliter are determined by plating appropriate dilutions onto non-selective medium to produce &tilde;100-200 colonies. 
cThe pre- and post-induction plating efficiencies are determined by dividing the total number of viable, colony forming, cells by the total number of cell bodies in the culture, determined by hemacytometer count.
This suggests that the presence or absence of either translocation chromosome does not affect the ability to survive DSB formation. The frequency of chromosomal translocations can be calculated by dividing the number of histidine prototrophic colonies by the total number of viable cells determined by plating onto YPD (Figure 2B). This assay can be conducted using strains of different genotype to identify how loss of protein function affects SSA (i.e. rad51&Delta;-/-). Recombination frequencies determined in different strains can then be graphed to compare differences in the ability of these strains to repair the HO-endonuclease induced DSBs by SSA (Figure 2C). Translocation frequencies obtained with the wild-type diploid strain under selective (2.2x10-2) and non-selective (2.17x10-2) conditions were not statistically different from each other (p-value= 0.9131), while the translocation frequencies obtained selectively (6.0x10-2) and non-selectively (11.9x10-2) with the rad51&Delta;-/- homozygote were similar but statistically different (p-value= 0.0089). The frequencies obtained both selectively (p-value=0.0001) and non-selectively (p-value=0.0002) with the rad51&deg;-/- homozygote were statistically different from those obtained using the corresponding conditions with the wild-type strain. 
Putative translocation-bearing clones can be further examined by genomic Southern blot and chromosomal blot analyses (Figure 3). For Southern analysis, genomic DNA is digested with BamHI endonuclease prior to agarose gel electrophoresis, blotting and hybridization to a 32P-labeled 1.8kb HIS3 probe to visualize the diagnostic 0.8 kb his3&Delta;200, 1.7 kb his3-&Delta;3', 4 kb tIII::XV, 5 kb tXV::III, and 8 kb his3-&Delta;5' fragments (Figure 3B.1). Intact chromosomes can be prepared, separated by CHEF (Figure 3B.2), blotted to nylon and hybridized with the 32P-labeled 1.8 kb HIS3 probe to visualize the 1.1 Mb intact chromosome XV, 0.8 Mb tXV:III translocation chromosome, 0.6 Mb tIII:XV translocation chromosome, and 0.3 Mb intact chromosome III (Figure 3B.3). A graphical map depicting expected BamHI endonuclease-digested genomic DNA fragments, and parent and recombinant chromosomes, is depicted (Figure 3A). 
<img src="/files/ftp_upload/3150/3150fig1.jpg" alt="Figure 1" /> 
Figure 1. Formation of translocation chromosomes by single-strand annealing (SSA). 1) DSBs are created at the his3-&Delta;3' and his3-&Delta;5' substrates (chromosomes XV and III, respectively) by HO endonuclease following the addition of galactose to the cultures. 2) DSBs are processed to generate 3' single-strands at the ends of the broken chromosomes. 3) SSA machinery anneals complementary 311 or 60 nucleotide single-stranded HIS3 sequences formed at each of the recombination substrates. Non-homologous tails formed upon annealing are removed by endonuclease digestion. Complementary four bp overhangs on the remaining chromosomal fragments formed by HO-endonuclease digestion may also anneal. 4) Ligation concludes the creation of an intact HIS3 gene and tXV::III translocation chromosome by SSA. Cells carrying this chromosome can be selected for by their ability to grow on medium lacking histidine. Ligation may also generate the reciprocal tIII::XV translocation chromosome by a NHEJ-like mechanism.
<img src="/files/ftp_upload/3150/3150fig2.jpg" alt="Figure 2" /> 
Figure 2. Assay for determining the frequency of translocation by SSA. A) One ml YPGly/Lac cultures are inoculated with single colonies of cells of a select genotype and grown to an appropriate density of approximately 5x107-1x108 cell/ml. Galactose is added to a final concentration of 2% to create DSBs at the recombination substrates on chromosomes III and XV. To conduct the assay under selective conditions, appropriate dilutions are made such that approximately 100 to 200 cells are plated onto YPD and a sufficient number of cells are plated onto medium lacking histidine to yield an observable number of His+ recombinant colonies. To conduct the assay without selection, approximately 100 to 200 cells are plated onto YPD, grown for two to three days to single colonies and then replica plated onto medium lacking histidine. B) Translocation frequency can be determined by dividing the number of colonies that grow on &#x2013;His plates by the fraction that grow on YPD. C) The translocation frequencies of strains of different genotype (i.e. Wild-type and rad51&Delta;-/-) can be graphed to compare differences in the ability of these strains to repair the DSBs by SSA.
<img src="/files/ftp_upload/3150/3150fig3.jpg" alt="Figure 3" /> 
Figure 3. Determining plating efficiency. (A) An aliquot of cells is taken from the overnight culture prior to and post DSB induction, appropriate dilutions are made, followed by a hemacytometer count to determine the total number of cell bodies per ml of culture. Appropriate dilutions are plated onto non-selective medium to determine the total number of viable cells per ml, by counting the colonies that appear on YPD. (B) The plating efficiency is then determined by dividing the total number of viable cells by the total number of cell bodies, and multiplying this quotient by 100.
<img src="/files/ftp_upload/3150/3150fig4.jpg" alt="Figure 4" /> 
Figure 4. Detection of chromosomal translocation events by genomic Southern blot and chromosome blot analyses. 
A) Graphical representation of relevant chromosomes before (left) and after (right) translocation formation. 
Sizes of parent and recombinant chromosomes are listed in megabase-pairs (Mb). Sizes of restriction fragments containing relevant sequences generated by BamHI digestion of genomic DNA from parent and recombinant strains, and revealed on blots by hybridization with a 1.8 kb BamHI HIS3 genomic clone, are listed in kilobase-pairs (kb). Chromosomes are not drawn to scale. 
B) Physical analysis of putative translocation-bearing clones. 
(1) Genomic Southern blot analysis &#x2013; Genomic DNA was collected and digested with BamHI restriction endonuclease, fractionated by gel electrophoresis, blotted to nylon, and hybridized to a 32P-labeled 1.8 kb HIS3 probe to visualize the following fragments: 0.8 kb his3&Delta;200, 1.7 kb his3-&Delta;3', 4 kb tIII::XV, 5 kb tXV::III , and 8 kb his3-&Delta;5'. Lanes: a) parent diploid, b) His+ non-reciprocal translocation recombinant, c) His+ reciprocal translocation recombinant. 
(2) CHEF gels - Intact chromosomes were prepared in agarose plugs, separated by CHEF, stained with ethidium bromide and visualized under UV light. Lanes: As above. 
(3) Chromosome blots &#x2013; Separated chromosomes were blotted to nylon and hybridized with the 32P-labeled 1.8 kb HIS3 probe to visualize the following chromosomes: 1.1 Mb intact chromosome XV, 0.8 Mb tXV:III translocation chromosome, 0.6 Mb tIII:XV translocation chromosome, and 0.3 Mb intact chromosome III. Lanes: As above.

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No conflicts of interest declared.

High doses of ionizing radiation present an inherent risk of genome instability through the generation of a large number of DSBs19. Eukaryotic genomes are replete with repetitive sequences that are excellent substrates for generating translocations and other genomic rearrangements20,21. Chromosomal translocations by HR are frequently observed when DSBs are introduced between repetitive sequences12,21,22. Overwhelming evidence suggests that much of the genomic instability associated with leukemias and lymphomas can be attributed to chromosomal translocations, highlighting the importance of understanding how this mechanism occurs in eukaryotes22,23. We have developed a system in budding yeast for examining the formation of translocation chromosomes by DSB-induced HR between short regions of homology on different chromosomes that are similar in size to repetitive elements dispersed throughout the yeast and human genomes.
In the assay, the his3-&Delta;3' translocation substrate is located on one copy of chromosome XV. The other his3 allele (his3-&Delta;200) has a ~1kb deletion of the HIS3 promoter and coding sequence that prevents this sequence from being used as a template for repair24. The his3-&Delta;5' substrate is located at the LEU2 locus on one copy of chromosome III, with the other copy of chromosome III containing an unaltered LEU2 allele (Figure 1). A galactose-inducible HO endonuclease expression cassette marked with KAN-MX was inserted into the TRP1 locus of chromosome IV (trp1::GAL-HO-KAN-MX). Each translocation substrate is flanked by a HO-endonuclease recognition sequence that can be targeted for cleavage by inducing expression of the HO gene through the addition of galactose to the medium. After HO-endonuclease induced cleavage at the his3-&Delta;3' and his3-&Delta;5' substrates, cells can efficiently use the shared short tract of HIS3 sequence homology (311 bp or 60 bp) to repair the broken chromosomes by HR, generating a translocation chromosome with an intact HIS3 allele12-14,25. 
Because the parent cells lack an intact copy of the HIS3 gene, they are unable to grow on &#x2013;His medium. Only cells that have undergone the translocation event can be selected for on medium lacking histidine. Therefore, the frequency of chromosomal translocations can be calculated by dividing the number of histidine prototrophic colonies by the total number of viable cells plated, determined by plating onto YPD. Genomic DNA and intact chromosomes can then be isolated from representative His+ colonies, and the presence of a translocation chromosome verified by genomic Southern and chromosome blot analyses.
Careful analysis has allowed us to gather additional important information about the assay12. Genomic Southern blot analysis has provided evidence that there is virtually complete cutting of chromosomes XV and III after 30 minutes of HO-endonuclease induction, and thus there is no significant background of uncut chromosomal substrates in the population (G. Manthey &amp; A. Bailis, unpublished results). Genomic Southern and chromosome blot analyses of His- survivors indicates that cells frequently lose one, the other, or both cut chromosomes and remain viable (L. Liddell &amp; A. Bailis, unpublished results). Importantly, the nearly equivalent plating efficiencies on non-selective medium before and after induction of expression of HO-endonuclease indicates that neither the failure to repair broken chromosomes, nor the failure to retain translocation chromosomes affects the ability to survive DSB formation. Consistent with this, the tXV:III translocation chromosome has been shown to be unstable in mitotic cells in the absence of selection. This was demonstrated by growing tXV::III containing His+ recombinants overnight non-selectively, plating single colonies onto non-selective plates, and replica plating onto selective medium lacking histidine. Ten to 70% of the colonies arising on these plates had lost the tXV::III translocation chromosome (N. Pannunzio &amp; A. Bailis, unpublished results). 
Translocation chromosomes generated by IR exposure in humans exhibit a similar instability26. This suggests that translocation formation may contribute to early events in tumorigenesis by promoting loss of heterozygosity. Second, extensive genetic and molecular analyses suggest that SSA, an efficient and obligatorily non-conservative mechanism of HR, is the primary mechanism of translocation formation by HR following the simultaneous creation of DSBs on two chromosomes12,27,28. This is consistent with the finding that large doses of IR result in a density of DSBs sufficient to create breaks adjacent to multiple repetitive sequences in the yeast genome, and a high frequency of translocation formation by HR. Together, these observations suggest that the oncogenic effect of IR exposure in humans may, in part, result from the repair of DSBs by an efficient mechanism of HR that generates translocations that, through their inherent instability, promote genetic changes that launch tumorigenesis. Because radiation is often used to treat cancer, genome rearrangements that result from repair of radiation-induced DSBs may contribute to the generation of secondary cancers that arise frequently in patients. Thus, this model may help us gain a better understanding of the genetic and molecular basis of an important clinical response to IR treatment.

quantitation and analysis of the formation of ho-endonuclease stimulated chromosomal translocations by single-strand annealing in saccharomyces cerevisiae

Genetic variation is frequently mediated by genomic rearrangements that arise through interaction between dispersed repetitive elements present in every eukaryotic genome. This process is an important mechanism for generating diversity between and within organisms1-3. The human genome consists of approximately 40% repetitive sequence of retrotransposon origin, including a variety of LINEs and SINEs4. Exchange events between these repetitive elements can lead to genome rearrangements, including translocations, that can disrupt gene dosage and expression that can result in autoimmune and cardiovascular diseases5, as well as cancer in humans6-9.
Exchange between repetitive elements occurs in a variety of ways. Exchange between sequences that share perfect (or near-perfect) homology occurs by a process called homologous recombination (HR). By contrast, non-homologous end joining (NHEJ) uses little-or-no sequence homology for exchange10,11. The primary purpose of HR, in mitotic cells, is to repair double-strand breaks (DSBs) generated endogenously by aberrant DNA replication and oxidative lesions, or by exposure to ionizing radiation (IR), and other exogenous DNA damaging agents. 
In the assay described here, DSBs are simultaneously created bordering recombination substrates at two different chromosomal loci in diploid cells by a galactose-inducible HO-endonuclease (Figure 1). The repair of the broken chromosomes generates chromosomal translocations by single strand annealing (SSA), a process where homologous sequences adjacent to the chromosome ends are covalently joined subsequent to annealing. One of the substrates, his3-&Delta;3', contains a 3' truncated HIS3 allele and is located on one copy of chromosome XV at the native HIS3 locus. The second substrate, his3-&Delta;5', is located at the LEU2 locus on one copy of chromosome III, and contains a 5' truncated HIS3 allele. Both substrates are flanked by a HO endonuclease recognition site that can be targeted for incision by HO-endonuclease. HO endonuclease recognition sites native to the MAT locus, on both copies of chromosome III, have been deleted in all strains. This prevents interaction between the recombination substrates and other broken chromosome ends from interfering in the assay. The KAN-MX-marked galactose-inducible HO endonuclease expression cassette is inserted at the TRP1 locus on chromosome IV. The substrates share 311 bp or 60 bp of the HIS3 coding sequence that can be used by the HR machinery for repair by SSA. Cells that use these substrates to repair broken chromosomes by HR form an intact HIS3 allele and a tXV::III chromosomal translocation that can be selected for by the ability to grow on medium lacking histidine (Figure 2A). Translocation frequency by HR is calculated by dividing the number of histidine prototrophic colonies that arise on selective medium by the total number of viable cells that arise after plating appropriate dilutions onto non-selective medium (Figure 2B). A variety of DNA repair mutants have been used to study the genetic control of translocation formation by SSA using this system12-14.

Watch this Scientific Journal Video about Quantitation and Analysis of the Formation of HO-Endonuclease Stimulated Chromosomal Translocations by Single-Strand Annealing in Saccharomyces cerevisiae at 

Quantitation and Analysis of the Formation of HO-Endonuclease Stimulated Chromosomal Translocations by Single-Strand Annealing in Saccharomyces cerevisiae

The HO-stimulated translocation assay monitors single-strand annealing following the creation of DNA double-strand breaks at multiple loci in diploid Saccharomyces cerevisiae. This mechanism may model genome rearrangements in somatic cells of higher eukaryotes following exposure to high doses of ionizing radiation.

Quantitation and Analysis of the Formation of HO-Endonuclease Stimulated Chromosomal Translocations by Single-Strand Annealing in Saccharomyces cerevisiae

Genetic variation is frequently mediated by genomic rearrangements that arise through interaction between dispersed repetitive elements present in every ...

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Research

JoVE Journal

Biology

Dissection of Saccharomyces Cerevisiae Asci

Yeast is a highly tractable model system that is used to study many different cellular processes. The common laboratory strain Saccharomyces cerevisiae exists in either a haploid or diploid state. The ability to combine alleles from two haploids and the ability to introduce modifications to the genome requires the production and dissection of asci. Asci production from haploid cells begins with the mating of two yeast haploid strains with compatible mating types to produce a diploid strain. This can be accomplished in a number of ways either on solid medium or in liquid. It is advantageous to select for the diploids in medium that selectively promotes their growth compared to either of the haploid strains. The diploids are then allowed to sporulate on nutrient-poor medium to form asci, a bundle of four haploid daughter cells resulting from meiotic reproduction of the diploid. A mixture of vegetative cells and asci is then treated with the enzyme zymolyase to digest away the membrane sac surrounding the ascospores of the asci. Using micromanipulation with a microneedle under a dissection microscope one can pick up individual asci and separate and relocate the four ascopores. Dissected asci are grown for several days and tested for the markers or alleles of interest by replica plating onto appropriate selective media.

Dissection of Saccharomyces Cerevisiae Asci

Micromanipulation of yeast cells is needed for meiotic genetic analysis or to select diploid zygotes. These micromanipulations are carried out using the microneedle of a dissection microscope. The microneedle is used to relocate cells and is controlled by a micromanipulator which are available with various degrees of automation.

Yeast is a highly tractable model system that is used to study many different cellular processes. The common laboratory strain Saccharomyces cerevisiae ...

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground using a cryolysis procedure in a ball mill grinder and the resulting grindate brought into solution by urea solubilization. This procedure allows for the lysis of the cells in a metabolically inactive state, preserving phosphorylation and preventing reorientation of the phosphoproteome during cell lysis.  Following reduction, alkylation, trypsin digestion of the proteins, the samples are desalted on C18 columns and the sample complexity reduced by fractionation using hydrophilic interaction chromatography (HILIC). HILIC columns preferentially retain hydrophilic molecules which is well suited for phosphoproteomics.  Phosphorylated peptides tend to elute later in the chromatographic profile than the non phosphorylated counterparts. After fractionation, phosphopeptides are enriched using immobilized metal chromatography, which relies on charge-based affinities for phosphopeptide enrichment. At the end of this procedure the samples are ready to be quantitatively analyzed by mass spectrometry.

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

Description of a quantitative phosphorylation procedure using cryolysis, urea solubilziation, HILIC fractionation and IMAC enrichment of phosphorylated peptides.

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground ...

Quantitation of &gamma;H2AX Foci in Tissue Samples

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) processes or by exogenous factors, particularly ionizing radiation and radiomimetic compounds. Phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX, is an early response to DNA double-strand breaks1. This phosphorylation event is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Overall, DSB induction results in the formation of discrete nuclear &gamma;H2AX foci which can be easily detected and quantitated by immunofluorescence microscopy2. Given the unique specificity and sensitivity of this marker, analysis of &gamma;H2AX foci has led to a wide range of applications in biomedical research, particularly in radiation biology and nuclear medicine. The quantitation of &gamma;H2AX foci has been most widely investigated in cell culture systems in the context of ionizing radiation-induced DSBs. Apart from cellular radiosensitivity, immunofluorescence based assays have also been used to evaluate the efficacy of radiation-modifying compounds. In addition, &gamma;H2AX has been used as a molecular marker to examine the efficacy of various DSB-inducing compounds and is recently being heralded as important marker of ageing and disease, particularly cancer3. Further, immunofluorescence-based methods have been adapted to suit detection and quantitation of &gamma;H2AX foci ex vivo and in vivo4,5. Here, we demonstrate a typical immunofluorescence method for detection and quantitation of &gamma;H2AX foci in mouse tissues.

Quantitation of &#947;H2AX Foci in Tissue Samples

Quantitation of DNA double-strand breaks on the basis of &gamma;H2AX foci has become an invaluable tool, particularly in radiation biology, for the evaluation of tissue radiosensitivity and effects of radiation modifying compounds. Here we demonstrate the use of an immunofluorescence assay for quantitation of &gamma;H2AX foci in tissue samples.

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) ...

Microarray Analysis for Saccharomyces cerevisiae

In this protocol, gene expression in yeast (Saccharomyces cerevisiae) is changed after exposure to oxidative stress induced by the addition of hydrogen peroxide (H2O2), an oxidizing agent. In the experiment, yeast is grown for 48 hours in 1/2X YPD broth containing 3X glucose. The culture is split into a control and treated group. The experiment culture is treated with 0.5 mM H2O2 in Hanks Buffered Saline (HBSS) for 1 hour. The control culture is treated with HBSS only. Total RNA is extracted from both cultures and is converted to a biotin-labeled cRNA product through a multistep process. The final synthesis product is taken back to the UVM Microarray Core Facility and hybridized to the Affymetrix yeast GeneChips. The resulting gene expression data are uploaded into bioinformatics data analysis software.

Microarray Analysis for Saccharomyces cerevisiae

In this protocol, gene expression in yeast (Saccharomyces cerevisiae) is changed after exposure to oxidative stress induced by the addition of hydrogen peroxide (H2O2), an oxidizing agent.

In this protocol, gene expression in yeast (Saccharomyces cerevisiae) is changed after exposure to oxidative stress induced by the addition of hydrogen ...

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

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

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

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

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  ...

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

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 ...

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;

Defects in the cystic fibrosis transmembrane conductance regulator (CFTR) protein cause cystic fibrosis (CF), an autosomal recessive disease that ...

EPR Monitored Redox Titration of the Cofactors of Saccharomyces cerevisiae Nar1

Electron Paramagnetic Resonance (EPR) monitored redox titrations are a powerful method to determine the midpoint potential of cofactors in proteins and to identify and quantify the cofactors in their detectable redox state.
The technique is complementary to direct electrochemistry (voltammetry) approaches, as it does not offer information on electron transfer rates, but does establish the identity and redox state of the cofactors in the protein under study. The technique is widely applicable to any protein containing an electron paramagnetic resonance (EPR) detectable cofactor.
A typical titration requires 2 ml protein with a cofactor concentration in the range of 1-100 &#181;M. The protein is titrated with a chemical reductant (sodium dithionite) or oxidant (potassium ferricyanide) in order to poise the sample at a certain potential. A platinum wire and a Ag/AgCl reference electrode are connected to a voltmeter to measure the potential of the protein solution. A set of 13 different redox mediators is used to equilibrate between the redox cofactors of the protein and the electrodes. Samples are drawn at different potentials and the Electron Paramagnetic Resonance spectra, characteristic for the different redox cofactors in the protein, are measured. The plot of the signal intensity versus the sample potential is analyzed using the Nernst equation in order to determine the midpoint potential of the cofactor.

EPR Monitored Redox Titration of the Cofactors of Saccharomyces cerevisiae Nar1

The goal of this protocol is to use electron paramagnetic resonance (EPR) monitored redox titrations to identify different cofactors of Saccharomyces cerevisiae Nar1. Redox titrations offer a very robust way to obtain midpoint potentials of different redox active cofactors in enzymes and proteins.

Electron Paramagnetic Resonance (EPR) monitored redox titrations are a powerful method to determine the midpoint potential of cofactors in proteins and to ...