This work was supported by an NIH/NHGRI Genome Scholar/ Faculty Transition Award (K22 HG002908) to A.M.D. and a strategic partnership between the Institute for Systems Biology and the University of Luxembourg. Requests for strains or plasmids should be directed to Aim&#233;e Dudley (aimee.dudley@gmail.com).

1. Strain Construction and Characterization
<ol>
	<li>Construct the heterozygous diploid strain to be used in the cross by mating haploid strains and selecting diploids2 or using a naturally occurring heterozygous strain. The strains should not harbor GFP.</li>
	<li>Confirm that the strain is unable to grow on YPD2 + 200 &#181;g/ml G418.</li>
	<li>Confirm that the strain is able to sporulate. BEST is compatible with low sporulation efficiency crosses, but low sporulation frequency increases FACS sorting time required.
	<ol>
		<li>Set up an O/N culture of the strain in 5 ml YPD and grow with agitation at 30 &#176;C.</li>
		<li>Wash 0.5 ml of the O/N culture three times with PBS (pH 7.4).</li>
		<li>Pellet the washed cells, remove the supernatant, and resuspend in 2 ml sporulation medium7.</li>
		<li>Agitate cells at RT and check daily for tetrad formation using a phase contrast light microscope with a 40X&#160;objective.</li>
	</ol>
	</li>
	<li>Estimate the spore viability of the cross by conventional dissection2,3 of a small number (~20) of tetrads. Spore viability in crosses between different strain backgrounds varies widely and unpredictably. However, a priori expectations of spore viability can be used to evaluate the efficiency of a BEST experiment prior to the labor- and cost-intensive steps of saving and genotyping the strains. Therefore, this step is highly recommended.</li>
</ol>
2. Transformation of Diploids with Library of Barcoded Plasmids
<ol>
	<li>Insert the pCL2_BC barcoded plasmid into the heterozygous diploid strain using the transformation protocol described in Gietz and Woods8 with the following modifications.
	<ol>
		<li>After cells have been incubated with the transformation mixture at 30 &#176;C for 20 min, add DMSO to 8% of the volume of the transformation mix. Addition of DMSO has been found to improve transformation efficiency in some strain backgrounds9.</li>
		<li>After the heat shock step, pellet the cells with a brief centrifugation, decant the transformation mixture, and wash cells gently in YPD. Then, resuspend the cells in 1 ml YPD and allow them to recover at RT for 3 hr without agitation.</li>
	</ol>
	</li>
	<li>Select transformants by plating onto YPD + G418 (200 &#181;g/ml) agar plates. To ensure a complex pool of plasmid barcodes, harvest a large number of colonies, e.g. 5 plates with ~2,500 colonies per plate.</li>
	<li>Once colonies are visible (2-3 days of growth at 30 &#176;C), pool the transformants by carefully scraping them off the plate into liquid YPD + G418 (200 &#181;g/ml) + 15% glycerol.&#160; Vortex this cell suspension, transfer to a cryopreservation vial, and freeze at -80&#176; C. Make a separate frozen stock for each plate. These frozen stocks serve as the starting strains for Step 3.1 and can also be used to generate more tetrads at a later date, if desired.</li>
</ol>
3. Sporulating the Library of Transformed Cells to Obtain Barcoded Tetrads
<ol>
	<li>Revive cells from the frozen stocks created in 2.3 by O/N growth in 5 ml YPD + G418 (200 &#181;g/ml) at 30 &#176;C with agitation.</li>
	<li>Wash the cells from the O/N cultures 3x&#160;in PBS (pH 7.4) and then resuspend the cells in sporulation medium7 plus 200 &#181;g/ml G418.</li>
	<li>Agitate sporulation cultures at RT and monitor sporulation progress daily until new tetrads have stopped forming.</li>
	<li>Once the culture contains well-formed tetrads and relatively few dyads, remove cultures from agitation and leave at RT for at least 7 days. While some crosses may be amenable to on plate digestion and disruption as soon as tetrads have formed (assessed as described in 7.3), the additional incubation (7-10 days in our experiments) is essential for other crosses. Therefore, this step is highly recommended.</li>
</ol>
4. Establish the FACS Gates for Tetrad Sorting
The meiotically expressed SPS2-GFP fusion on the pCL2_BC plasmid allows tetrads in the sporulation culture to be detected and isolated from the culture by fluorescence. The following protocol has been developed for a FACSAria II.
<img alt="Figure 2" fo:content-width="6in" src="/files/ftp_upload/51401/51401fig2highres.jpg" width="500" /> 
Figure 2.&#160;FACS gates.&#160; A series of four sequential gates is used to isolate tetrads. The gates shown in (A) and (B) help reduce the number of clumps. Fluorescent events are selected with the gate in (C). The histogram in (D) has two peaks. Events in the left peak are primarily tetrads (left inset image), and events in the right peak are generally tetrads with an attached bud (right inset image). A final gate is applied to this histogram to select events which are mostly tetrads (red bars). This figure has been modified from Ludlow, et al1.
<ol>
	<li>Load the sporulated culture onto a FACS sorter and set up two forward scatter and side scatter gates to exclude debris, clumped cells, and multiple cells per droplet (Figures 2A and 2B).</li>
	<li>There should be a population of fluorescent cells, which when using the SPS2-GFP reporter construct include dyads and tetrads10. Gate the GFP vs. FSC to include these fluorescent events (Figure 2C).</li>
	<li>Depending on the strain background it may be necessary to include an additional gate to remove clumps that include tetrads and other cells. If this is necessary, inspect an FSC histogram of the fluorescent events and specify a gate that includes events with lower FSC (Figure 2D, gated events are in red).</li>
	<li>Check the efficiency of the gating regime by sorting ~1,500 tetrads onto a microscope slide and counting the proportion of tetrads to non-tetrads using a phase contrast microscope at 400X&#160;magnification. The percent of tetrads can be accurately estimated by counting only ~200 sorted events, but sorting ~1,500 reduces the amount of time spent searching for cells on the microscope slide. Alternatively, the ratio of tetrads to non-tetrads can be assessed during Step 4.5.</li>
	<li>Periodically check that the sorter is accurately reporting the number of events by sorting 25 tetrads onto a microscope slide and examining the cells contained in the droplet using a phase contrast microscope at 400X&#160;magnification. There should be 25 tetrads in the droplet. Repeat 3-4x&#160;to assess the variance of the FACS instrument.</li>
</ol>
5. Sorting Tetrads Onto an Agar Plate
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/51401/51401fig3highres.jpg" width="500" /> 
Figure 3.&#160;Placement of the agar plate on the FACS sorter. To sort tetrads into a small drop of the zymolyase solution placed in the middle of a standard (circular) agar plate, a standard (100&#160;mm rectangular) 96-well plate is first placed on the stage of the cell sorter and the sorter is programmed to deposit tetrads into a specific well (in this case D5). &#160;Then, the circular agar plate is placed on top of the 96-well plate and aligned so that the drop of zymolyase is centered over the specified well.
<ol>
	<li>Warm the desired number of YPD + G418 (200 &#181;g/ml) plates for at least 1.5 hr in a 37 &#176;C incubator positioned near the FACS sorter.</li>
	<li>A key feature of the sorting protocol is the ability to accurately target FACS sorted tetrads to a specific location on an agar plate that contains a pool of zymolyase solution. To do this, place a &#8220;landmark&#8221; 96-well plate on the automated cell deposition unit (ACDU) of the sorter. Prepare a sort layout that will sort 25 tetrads into one well of a 96-well format. Choose a well near the middle of the plate, e.g. well D5. Figure 3 shows a Petri dish and 96-well plate loaded onto a cell sorter. Sorting 25 tetrads per plate yields well-separated colonies.</li>
	<li>Bring a stack of 10 agar plates from the 37 &#176;C incubator to the sorter.</li>
	<li>Take one agar plate and pipette 25 &#956;l of zymolyase solution (1 mg/ml in 0.7 M sorbitol) onto the agar near the center of the plate. Then, place the plate on the landmark 96-well plate, aligning the zymolyase droplet over the target well from Step 5.2, e.g. well D5.</li>
	<li>Start the sort and deposit 25 tetrads into the middle of the zymolyase droplet on the agar plate. Then, remove and cover the plate.</li>
	<li>Repeat Steps 5.4 and 5.5 until 25 tetrads have been sorted into zymolyase droplets on each plate in the stack.</li>
	<li>Return the stack of tetrad-containing plates to the 37 &#176;C incubator.</li>
	<li>Repeat Steps 5.3 through 5.7 until all plates have tetrads.</li>
</ol>
6. Separating Spores
<ol>
	<li>Incubate each stack of tetrad-containing agar plates at 37 &#176;C for 1.5 hr.</li>
	<li>Remove plates from the incubator, add 15-25 sterile 3&#160;mm glass beads per plate, and shake the plates as a stack (or as two stacks of 5 plates) for 3-5 min. The best spore separation is achieved by moving the plates rigidly from side to side or front to back. Do not move the plates such that the beads &#34;swirl&#34; around the outside edge of the plate. Leave the beads on the plates when finished.</li>
	<li>Place the stack of plates with beads face up in a 30 &#176;C incubator. Retrieve the next stack plates from the 37 &#176;C incubator, and repeat Step 6.2.</li>
	<li>Incubate plates O/N at 30 &#176;C.</li>
</ol>
7. Harvesting Colonies
<ol>
	<li>After an O/N incubation at 30 &#176;C, small colonies should have formed on the plates. Carefully remove the plates from the incubator without disturbing the glass beads.</li>
	<li>Remove the glass beads by carefully and quickly inverting the plates over a deep container. The container should be deep enough to catch the beads without allowing them to bounce up and hit the agar plate. Tapping the bottom of the plate while it is inverted can help dislodge beads that are stuck to the plate. These beads can be reused by thorough washing with soap, rinsing well with deionized water and sterilizing by autoclaving.</li>
	<li>Count the number of colonies on each plate and use the number of colonies to estimate the efficiency of spore separation and recovery. For example, 25 tetrads per plate for a cross with near 100% viability could yield a total of 100 colonies per plate.</li>
	<li>From plates with sufficient numbers of colonies (&#62;65 for a high-viability crosses), transfer cells from each colony into a separate well of a 96-well plate containing YPD liquid + G418. Also, make note of which wells contain colonies from the same agar plate; this information will be used to aid the tetrad assignment software.</li>
	<li>Use the 96-well liquid plates to seed cultures for genotyping and for saving strains as frozen glycerol stocks. Details on RAD-tag sequencing and sequence analysis can be found in Ludlow&#160;et al1.</li>
</ol>

<ol>
	<li>Ludlow, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+tetrad+analysis.">High-throughput tetrad analysis.</a> Nat Methods. 10, 671-675 (2013).</li><li>Rose, M., Winston, F., Hieter, P. <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 Laboratory Course Manual. , (1990).</li><li>Morin, A., Moores, A. W., Sacher, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissection+of+Saccharomyces+cerevisiae+asci.">Dissection of Saccharomyces cerevisiae asci.</a> J Vis Exp. 27, (2009).</li><li>Winge, O., Laustsen, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=On+two+types+of+spore+germination,+and+on+genetic+segregations+in+Saccharomyces+demonstrated+through+single+spore+cultures.">On two types of spore germination, and on genetic segregations in Saccharomyces demonstrated through single spore cultures.</a> C.R. Trav. Lab. Carlsberg Ser. Physiol. 24, 263-315 (1937).</li><li>Coluccio, A., Neiman, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interspore+bridges:+a+new+feature+of+the+Saccharomyces+cerevisiae+spore+wall.">Interspore bridges: a new feature of the Saccharomyces cerevisiae spore wall.</a> Microbiology. 150, 3189-3196 (2004).</li><li>Baird, N. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+SNP+discovery+and+genetic+mapping+using+sequenced+RAD+markers.">Rapid SNP discovery and genetic mapping using sequenced RAD markers.</a> PLoS One. 3, (2008).</li><li>Tong, A. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systematic+genetic+analysis+with+ordered+arrays+of+yeast+deletion+mutants.">Systematic genetic analysis with ordered arrays of yeast deletion mutants.</a> Science. 294, 2364-2368 (2001).</li><li>Gietz, R. D., Woods, 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=Transformation+of+yeast+by+lithium+acetate/single-stranded+carrier+DNA/polyethylene+glycol+method.">Transformation of yeast by lithium acetate/single-stranded carrier DNA/polyethylene glycol method.</a> Methods Enzymol. 350, 87-96 (2002).</li><li>Hill, J., Donald, K. A., Griffiths, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DMSO-enhanced+whole+cell+yeast+transformation.">DMSO-enhanced whole cell yeast transformation.</a> Nucleic Acids Res. 19, 5791-57 (1991).</li><li>Gerke, J. P., Chen, C. T., Cohen, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Natural+isolates+of+Saccharomyces+cerevisiae+display+complex+genetic+variation+in+sporulation+efficiency.">Natural isolates of Saccharomyces cerevisiae display complex genetic variation in sporulation efficiency.</a> Genetics. 174, 985-997 (2006).</li><li>Bloom, J. S., Ehrenreich, I. M., Loo, W. T., Lite, T. L., Kruglyak, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Finding+the+sources+of+missing+heritability+in+a+yeast+cross.">Finding the sources of missing heritability in a yeast cross.</a> Nature. 494, 234-237 (2013).</li><li>Fogel, S., Mortimer, R., Lusnak, K., Tavares, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Meiotic+gene+conversion:+a+signal+of+the+basic+recombination+event+in+yeast.">Meiotic gene conversion: a signal of the basic recombination event in yeast.</a> Cold Spring Harb Symp Quant Biol. 43 Pt 2, 1325-1341 (1979).</li></ol>

The authors have nothing to disclose.

Many areas of genetics research, ranging from complex trait mapping11 to the study of mechanisms underlying DNA recombination12, require extremely large numbers of progeny and high volumes of DNA sequencing. Techniques that marry the power of conventional approaches with the capacity of high throughput DNA sequencing have the potential to enable studies that were previously intractable. Although several high-throughput yeast genetics methods have been applied effectively to specific problems, each has limitations that fall short of the broad applicability of conventional tetrad analysis. BEST addresses these challenges by employing a high-throughput approach combining tetrad dissection and genotyping the progeny of yeast crosses. In conjunction with multiplexed RAD-tag sequencing, BEST offers an inexpensive way to glean genotype information for each progeny strain while preserving tetrad relationships by means of unique tetrad barcodes.&#160; Of all currently used high throughput methods, BEST most closely recapitulates the information provided by a manually dissected yeast cross.
BEST has two key features. The first is the use of FACS to rapidly isolate tetrads away from unsporulated cells in the culture (Protocol Steps 5-6). The ability to isolate thousands of fluorescent events (tetrads) gives BEST its greatest gain in throughput. This automation provides an even greater advantage over manual dissection for crosses with low sporulation efficiency, due to the increased time needed to visually identify rare events. The second key feature is the use of a highly complex molecular barcode (Protocol Step 2) that is transmitted to each sister spore of a tetrad. Because these barcodes can be used to computationally reconstruct the tetrad relationships between spores that have been randomly distributed across an agar plate, the need to manually array cells in an ordered grid is alleviated. Finally, because the molecular barcode and the sporulation-specific GFP reporter gene are physically linked (on the same plasmid), only tetrads that have retained the molecular barcode are selected by FACS sorting.
There are two considerations that govern the length of time that cultures should be incubated in the sporulation medium. First, because the sporulation specific reporter (SPS2-GFP) is expressed early in meiosis, GFP alone does not distinguish complete 4-spore tetrads from cells that have not completed meiosis (i.e. dyads). While additional FACS gating may decrease the number of fluorescent, incomplete tetrads, the easiest way to decrease these unwanted events is simply monitoring the sporulation culture (Step 3.3) and proceeding once new tetrads have stopped forming. In our experiments, the frequency of dyads sorted is less than 1%. Second, for some crosses additional time spent in sporulation medium at RT without agitation (Step 3.4) significantly improves the separation of tetrads by the glass beads during plating ((Protocol Step 6). In fact, this step is absolutely necessary for the disruption of some crosses.
Like all high throughput methods, BEST is &#8220;noisier&#8221; than the corresponding conventional approach. The modest reduction in efficiency of BEST compared to conventional tetrad dissection could result from combinations of several factors. One factor is the increased loss of otherwise viable spores, which could result from adhesion to the glass beads during spreading or increased spore death due to the mechanical stresses of the process. A second factor could be simple plasmid loss during meiosis. &#160;A third factor could be an effective decrease in usable colonies resulting from colonies with mixed genotypes, an estimated ~5% of colonies in our pilot crosses1. It is likely that these colonies represent failures of sister spore separation. Because the rapid fluorescence sorting and spore separation permit the collection of enormous numbers of tetrads, BEST is well equipped to overcome decreases in efficiency on this scale. While future iterations of BEST could increase efficiencies approaching that of manual dissection, it is perhaps more likely that the current exponential trajectory of DNA sequencing capacity will rapidly compensate for this level of usable strain loss. Regardless, the ability to perform tetrad analysis on the scale to which BEST can be applied will enable studies that are currently unfeasible by manual methods.

Meiotic gene mapping, commonly known as tetrad analysis, is central to yeast genetics. Briefly, two haploid strains, each containing a single copy of each chromosome, are mated to form a heterozygous diploid strain with two copies of each chromosome, one from each parent. Starving diploid cells for nitrogen induces meiosis (sporulation) in which chromosomes duplicate, undergo rearrangement, and divide into four haploid cells (spores or segregants) with new combinations of alleles from each parent. The four spores resulting from a single meiosis can be isolated by a manual process called tetrad dissection.
Conventional tetrad dissection2,3 which has changed relatively little since the original publication in 19374, has three goals. First, cells that have undergone meiosis (tetrads) must be isolated away from a background of vegetative cells. In conventional analysis, this is accomplished by a researcher who, using a microscope, visually identifies tetrads on an agar plate and employs a micromanipulator apparatus visually identifying tetrads on an agar plate and using a micromanipulator to move the tetrad onto a cell-free area of the plate. Next, the four spores in the tetrad must be physically separated to prevent interspore mating.&#160; Spores within a tetrad are held together by both an ascus, the remnant of the cell wall of the original diploid cell, and a set of interspore bridges5. In conventional tetrad analysis the ascus is removed by enzymatic digestion, and a researcher uses the micromanipulator to break the interspore bridges and tease apart the spores. Finally, the individual spores are arrayed in a gridded pattern that maintains the relationship between the spores, i.e. all progeny in a row of four are sister spores of the same original tetrad. An experienced yeast researcher can complete the process of dissecting 10 tetrads (a generally accepted minimum number per cross) in approximately 15 min.
We have developed a new high-throughput tetrad genotyping method, which we call Barcode Enabled Sequencing of Tetrads (BEST)1. BEST expands upon current methods in high-throughput genetics by enabling the generation and genotyping of large numbers of progeny that are isolated, genotyped, and maintained as individuals in a manner that allows the sister spore relationships of all four meiotic products to be recovered. This is accomplished using three main strategies (Figure 1). First is the introduction of a reporter construct that labels cells that have undergone meiosis with GFP and allows them to be isolated by fluorescence-activated cell sorting (FACS). Second is the incorporation of a highly complex DNA barcode (a string of 15 random nucleotides) in a form that is transmitted to all four spores of a tetrad and can be read by DNA sequencing of the recombinant progeny and thus identifies sister spores from the same tetrad. Third is the genotyping step, and BEST is compatible with numerous genotyping platforms that capture both a set of genomic markers as well as the tetrad-specific barcode. We employ a restriction site-associated DNA tag sequencing (RAD-seq)6 method that directs genome sequencing to specific restriction sites1, thereby capturing a consistent 2-3% of the genome of the progeny strains as well as the plasmid-borne barcode. The recovery of tetrad relationships along with the empirically-derived genotyping data from the cross allows the accurate inference of missing information, including the status of markers with low sequence coverage and the complete genotype of inviable (and therefore unrecoverable) individuals. Here, we describe the application of the method in the most commonly used microorganism for meiotic mapping, the yeast S. cerevisiae. However, with minor substitutions of organism-specific reagents, e.g. different sporulation-specific proteins fused to GFP, the method should be readily transferrable to other microorganisms, including organisms in which meiotic mapping is significantly more labor intensive or currently intractable.
<img alt="Figure 1" fo:content-width="4in" src="/files/ftp_upload/51401/51401fig1highres.jpg" width="400" /> 
Figure 1.&#160;BEST method. (A)&#160;The pCL2_BC plasmid is a 2-micron plasmid with a sporulation-specific GFP fusion, resistance to G418, and a 15-mer random barcode. Construction of the plasmid library is described in Ludlow&#160;et al1. (B) A pool of barcoded plasmids is transformed into the diploid strain from a cross. (C) Transformants are then grown on selection and ~10,000 transformed colonies are pooled and sporulated. During meiosis each spore of a tetrad inherits a copy of the barcoded plasmid. (D) Tetrads are separated from unsporulated cells by FACS and collected on agar plates, where they are digested and disrupted to allow each spore to form a colony. (E) Colonies are then picked into 96-well plates, phenotyped, and genotyped. During genotyping the plasmid barcode is read and used to identify the four members of each tetrad. This figure has been modified from Ludlow&#160;et al1. <a href="https://www.jove.com/files/ftp_upload/51401/51401fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="0" cellpadding="0" cellspacing="0" width="726">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:295px;">FACSAria II (Special order unit)</td>
			<td style="width:149px;">BD Biosciences</td>
			<td style="width:127px;">-</td>
			<td style="width:155px;">Other cell sorters should work for this procedure. The key factors are: (1) a laser/detection system for the fluorescent reporter being used (488nm laser to excite EGFP in our case), (2) the ability to sort single events, (3) the ability to sort onto microscope slides for assessing the sort quality and (4) the ability to sort onto petri dishes (which may need</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Automated Cell Deposition Unit (for FACSAria II)</td>
			<td>BD Biosciences</td>
			<td colspan="2">643155 (if purchased separately from the FACSAria instrument)</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Zymolyase 100T</td>
			<td>Seikagaku Biobusiness</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">G418</td>
			<td>A.G. Scientific</td>
			<td></td>
			<td>http://www.agscientific.com/molecular-biology/gene-expression/g-418-sulfate-solid.html</td>
		</tr>
	</tbody>
</table>

best: barcode enabled sequencing of tetrads

Tetrad analysis is a valuable tool for yeast genetics, but the laborious manual nature of the process has hindered its application on large scales. Barcode Enabled Sequencing of Tetrads (BEST)1 replaces the manual processes of isolating, disrupting and spacing tetrads. BEST isolates tetrads by virtue of a sporulation-specific GFP fusion protein that permits fluorescence-activated cell sorting of tetrads directly onto agar plates, where the ascus is enzymatically digested and the spores are disrupted and randomly arrayed by glass bead plating. The haploid colonies are then assigned sister spore relationships, i.e. information about which spores originated from the same tetrad, using molecular barcodes read during genotyping. By removing the bottleneck of manual dissection, hundreds or even thousands of tetrads can be isolated in minutes. Here we present a detailed description of the experimental procedures required to perform BEST in the yeast Saccharomyces cerevisiae, starting with a heterozygous diploid strain through the isolation of colonies derived from the haploid meiotic progeny.

BEST can greatly improve the speed with which spores from tetrads can be isolated. As an example of the throughput this method can deliver, one researcher performed the FACS sorting and spore separation (Steps 5.1 through 6.3) with 149 agar plates in 3 hr1. An equivalent yield (approximately 3,725 tetrads) would require more than 90 hr of manual dissection. However, as with many high throughput methods, the current iteration of the method has some loss of efficiency compared to manual dissection. This loss manifests as a reduced number of colonies recovered compared to the expectation calculated based on the spore viability of the cross, determined by manual dissection (Step 1.4).&#160; Assuming that the causes of spore loss (Discussion) affect spores at random, estimations can be made of how many recovered colonies will be members of tetrads in which 1, 2, 3, or 4 spores have been recovered (Figure 4).
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51401/51401fig4highres.jpg" width="500" /> 
Figure 4.&#160; Simulation of method efficiency at the tetrad level. Failure to successfully recover spores (as colonies) may be due to spore inviability, spore adhesion to the glass beads, failure to separate spores from one another, etc. The Binomial distribution was used to calculate the expected fraction of tetrads (with at least one successfully recovered spore) producing 4, 3, 2 or 1 colonies based on the proportion of all spores successfully recovered. This graph can serve as a rough guide to the number of strains that will ultimately be assembled into 3- or 4- spore tetrads. As the number of colonies per plate increases, so does the number of complete tetrads recovered.
In the experiment described above, 25 tetrads from a cross with spore viability close to 100% (as determined by manual dissection)1 were plated on 149 agar plates. In this experiment, the maximum number of colonies observed per plate was 80, indicating that 20 spores failed to form discrete colonies. The mean number of colonies per plate was 62. Colonies from the 61 plates containing 65 colonies or more were harvested and sequenced. Of the 3,700 strains that passed sequencing quality control, 72% could be computationally assigned into 3- or 4- spore tetrads.

Watch this Scientific Journal Video about BEST: Barcode Enabled Sequencing of Tetrads at JoVE.com

BEST: Barcode Enabled Sequencing of Tetrads

Barcode Enabled Sequencing of Tetrads (BEST) replaces the manual processes of isolating, disrupting and spacing tetrads. BEST isolates tetrads by fluorescence-activated cell sorting onto agar plates, separates the spores by agitation with glass beads, and determines which randomly arrayed colonies were derived from the same original tetrad using molecular barcodes.

Tetrad analysis is a valuable tool for yeast genetics, but the laborious manual nature of the process has hindered its application on large scales. ...

best-barcode-enabled-sequencing-of-tetrads

Research

JoVE Journal

Biology

10.0K Views.  Pacific Northwest Diabetes Research Institute. Barcode Enabled Sequencing of Tetrads (BEST) replaces the manual processes of isolating, disrupting and spacing tetrads. BEST isolates tetrads by fluorescence-activated cell sorting onto agar plates, separates the spores by agitation with glass beads, and determines which randomly arrayed colonies were derived from the same original tetrad using molecular barcodes.

Video: BEST: Barcode Enabled Sequencing of Tetrads

Competitive Genomic Screens of Barcoded Yeast Libraries

By virtue of advances in next generation sequencing technologies, we have access to new genome sequences almost daily. The tempo of these advances is accelerating, promising greater depth and breadth. In light of these extraordinary advances, the need for fast, parallel methods to define gene function becomes ever more important. Collections of genome-wide deletion mutants in yeasts and E. coli have served as workhorses for functional characterization of gene function, but this approach is not scalable, current gene-deletion approaches require each of the thousands of genes that comprise a genome to be deleted and verified. Only after this work is complete can we pursue high-throughput phenotyping. Over the past decade, our laboratory has refined a portfolio of competitive, miniaturized, high-throughput genome-wide assays that can be performed in parallel. This parallelization is possible because of the inclusion of DNA 'tags', or 'barcodes,' into each mutant, with the barcode serving as a proxy for the mutation and one can measure the barcode abundance to assess mutant fitness. In this study, we seek to fill the gap between DNA sequence and barcoded mutant collections. To accomplish this we introduce a combined transposon disruption-barcoding approach that opens up parallel barcode assays to newly sequenced, but poorly characterized microbes. To illustrate this approach we present a new Candida albicans barcoded disruption collection and describe how both microarray-based and next generation sequencing-based platforms can be used to collect 10,000 - 1,000,000 gene-gene and drug-gene interactions in a single experiment.

We have developed comprehensive, unbiased genome-wide screens to understand gene-drug and gene-environment interactions. Methods for screening these mutant collections are presented.

By virtue of advances in next generation sequencing technologies, we have access to new genome sequences almost daily.  The tempo of these advances is ...

Detection of Rare Genomic Variants from Pooled Sequencing Using SPLINTER

As DNA sequencing technology has markedly advanced in recent years2, it has become increasingly evident that the amount of genetic variation between any two individuals is greater than previously thought3. In contrast, array-based genotyping has failed to identify a significant contribution of common sequence variants to the phenotypic variability of common disease4,5. Taken together, these observations have led to the evolution of the Common Disease / Rare Variant hypothesis suggesting that the majority of the "missing heritability" in common and complex phenotypes is instead due to an individual's personal profile of rare or private DNA variants6-8. However, characterizing how rare variation impacts complex phenotypes requires the analysis of many affected individuals at many genomic loci, and is ideally compared to a similar survey in an unaffected cohort. Despite the sequencing power offered by today's platforms, a population-based survey of many genomic loci and the subsequent computational analysis required remains prohibitive for many investigators.
To address this need, we have developed a pooled sequencing approach1,9 and a novel software package1 for highly accurate rare variant detection from the resulting data. The ability to pool genomes from entire populations of affected individuals and survey the degree of genetic variation at multiple targeted regions in a single sequencing library provides excellent cost and time savings to traditional single-sample sequencing methodology. With a mean sequencing coverage per allele of 25-fold, our custom algorithm, SPLINTER, uses an internal variant calling control strategy to call insertions, deletions and substitutions up to four base pairs in length with high sensitivity and specificity from pools of up to 1 mutant allele in 500 individuals. Here we describe the method for preparing the pooled sequencing library followed by step-by-step instructions on how to use the SPLINTER package for pooled sequencing analysis (<a href="http://www.ibridgenetwork.org/wustl/splinter" target="_blank">http://www.ibridgenetwork.org/wustl/splinter</a>). We show a comparison between pooled sequencing of 947 individuals, all of whom also underwent genome-wide array, at over 20kb of sequencing per person. Concordance between genotyping of tagged and novel variants called in the pooled sample were excellent. This method can be easily scaled up to any number of genomic loci and any number of individuals. By incorporating the internal positive and negative amplicon controls at ratios that mimic the population under study, the algorithm can be calibrated for optimal performance. This strategy can also be modified for use with hybridization capture or individual-specific barcodes and can be applied to the sequencing of naturally heterogeneous samples, such as tumor DNA.

Pooled DNA sequencing is a fast and cost-effective strategy to detect rare variants associated with complex phenotypes in large cohorts. Here we describe the computational analysis of pooled, next-generation sequencing of 32 cancer-related genes using the SPLINTER software package. This method is scalable, and applicable to any phenotype of interest.

As DNA sequencing technology has markedly advanced in recent years2, it has become increasingly evident that the amount of genetic variation between any ...

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Phenotypes are determined by a complex series of physical (e.g. protein-protein) and functional (e.g. gene-gene or genetic) interactions (GI)1. While physical interactions can indicate which bacterial proteins are associated as complexes, they do not necessarily reveal pathway-level functional relationships1. GI screens, in which the growth of double mutants bearing two deleted or inactivated genes is measured and compared to the corresponding single mutants, can illuminate epistatic dependencies between loci and hence provide a means to query and discover novel functional relationships2. Large-scale GI maps have been reported for eukaryotic organisms like yeast3-7, but GI information remains sparse for prokaryotes8, which hinders the functional annotation of bacterial genomes. To this end, we and others have developed high-throughput quantitative bacterial GI screening methods9, 10.
Here, we present the key steps required to perform quantitative E. coli Synthetic Genetic Array (eSGA) screening procedure on a genome-scale9, using natural bacterial conjugation and homologous recombination to systemically generate and measure the fitness of large numbers of double mutants in a colony array format. Briefly, a robot is used to transfer, through conjugation, chloramphenicol (Cm) - marked mutant alleles from engineered Hfr (High frequency of recombination) 'donor strains' into an ordered array of kanamycin (Kan) - marked F- recipient strains. Typically, we use loss-of-function single mutants bearing non-essential gene deletions (e.g. the 'Keio' collection11) and essential gene hypomorphic mutations (i.e. alleles conferring reduced protein expression, stability, or activity9, 12, 13) to query the functional associations of non-essential and essential genes, respectively. After conjugation and ensuing genetic exchange mediated by homologous recombination, the resulting double mutants are selected on solid medium containing both antibiotics. After outgrowth, the plates are digitally imaged and colony sizes are quantitatively scored using an in-house automated image processing system14. GIs are revealed when the growth rate of a double mutant is either significantly better or worse than expected9. Aggravating (or negative) GIs often result between loss-of-function mutations in pairs of genes from compensatory pathways that impinge on the same essential process2. Here, the loss of a single gene is buffered, such that either single mutant is viable. However, the loss of both pathways is deleterious and results in synthetic lethality or sickness (i.e. slow growth). Conversely, alleviating (or positive) interactions can occur between genes in the same pathway or protein complex2 as the deletion of either gene alone is often sufficient to perturb the normal function of the pathway or complex such that additional perturbations do not reduce activity, and hence growth, further. Overall, systematically identifying and analyzing GI networks can provide unbiased, global maps of the functional relationships between large numbers of genes, from which pathway-level information missed by other approaches can be inferred9.

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Systematic, large-scale synthetic genetic (gene-gene or epistasis) interaction screens can be used to explore genetic redundancy and pathway cross-talk. Here, we describe a high-throughput quantitative synthetic genetic array screening technology, termed eSGA that we developed for elucidating epistatic relationships and exploring genetic interaction networks in Escherichia coli.

Phenotypes  are determined by a complex series of physical (e.g. protein-protein) and  functional (e.g. gene-gene or genetic) interactions (GI)1. While ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

Next-generation Sequencing of 16S Ribosomal RNA Gene Amplicons

One of the major questions in microbial ecology is &#8220;who is there?&#8221; This question can be answered using various tools, but one of the long-lasting gold standards is to sequence 16S ribosomal RNA (rRNA) gene amplicons generated by domain-level PCR reactions amplifying from genomic DNA. Traditionally, this was performed by cloning and Sanger (capillary electrophoresis) sequencing of PCR amplicons. The advent of next-generation sequencing has tremendously simplified and increased the sequencing depth for 16S rRNA gene sequencing. The introduction of benchtop sequencers now allows small labs to perform their 16S rRNA sequencing in-house in a matter of days. Here, an approach for 16S rRNA gene amplicon sequencing using a benchtop next-generation sequencer is detailed. The environmental DNA is first amplified by PCR using primers that contain sequencing adapters and barcodes. They are then coupled to spherical particles via emulsion PCR. The particles are loaded on a disposable chip and the chip is inserted in the sequencing machine after which the sequencing is performed. The sequences are retrieved in fastq format, filtered and the barcodes are used to establish the sample membership of the reads. The filtered and binned reads are then further analyzed using publically available tools. An example analysis where the reads were classified with a taxonomy-finding algorithm within the software package Mothur is given. The method outlined here is simple, inexpensive and straightforward and should help smaller labs to take advantage from the ongoing genomic revolution.

Characterizing microbial community has been a longstanding goal in environmental microbiology. Next-generation sequencing methods now allow for the characterization of microbial communities at an unprecedented depth with minimal cost and labor. We detail here our approach to sequence bacterial 16S ribosomal RNA genes using a benchtop sequencer.

One of the major questions in microbial ecology is &#8220;who is there?&#8221; This question can be answered using various tools, but one of the ...

Targeted DNA Methylation Analysis by Next-generation Sequencing

The role of epigenetic processes in the control of gene expression has been known for a number of years. DNA methylation at cytosine residues is of particular interest for epigenetic studies as it has been demonstrated to be both a long lasting and a dynamic regulator of gene expression. Efforts to examine epigenetic changes in health and disease have been hindered by the lack of high-throughput, quantitatively accurate methods. With the advent and popularization of next-generation sequencing (NGS) technologies, these tools are now being applied to epigenomics in addition to existing genomic and transcriptomic methodologies. For epigenetic investigations of cytosine methylation where regions of interest, such as specific gene promoters or CpG islands, have been identified and there is a need to examine significant numbers of samples with high quantitative accuracy, we have developed a method called Bisulfite Amplicon Sequencing (BSAS). This method combines bisulfite conversion with targeted amplification of regions of interest, transposome-mediated library construction and benchtop NGS. BSAS offers a rapid and efficient method for analysis of up to 10 kb of targeted regions in up to 96 samples at a time that can be performed by most research groups with basic molecular biology skills. The results provide absolute quantitation of cytosine methylation with base specificity. BSAS can be applied to any genomic region from any DNA source. This method is useful for hypothesis testing studies of target regions of interest as well as confirmation of regions identified in genome-wide methylation analyses such as whole genome bisulfite sequencing, reduced representation bisulfite sequencing, and methylated DNA immunoprecipitation sequencing.

Bisulfite amplicon sequencing (BSAS) is a method for quantifying cytosine methylation in targeted genomic regions of interest. This method uses bisulfite conversion paired with PCR amplification of target regions prior to next-generation sequencing to produce absolute quantitation of DNA methylation at a base-specific level.

The role of epigenetic processes in the control of gene expression has been known for a number of years. DNA methylation at cytosine residues is of ...

Amplification, Next-generation Sequencing, and Genomic DNA Mapping of Retroviral Integration Sites

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of diverse libraries of retroviral integration sites using ligation-mediated PCR (LM-PCR) amplification and next-generation sequencing (NGS), (2) mapping the genomic location of each virus-host junction using BEDTools, and (3) analyzing the data for statistical relevance. Genomic DNA extracted from infected cells is fragmented by digestion with restriction enzymes or by sonication. After suitable DNA end-repair, double-stranded linkers are ligated onto the DNA ends, and semi-nested PCR is conducted using primers complementary to both the long terminal repeat (LTR) end of the virus and the ligated linker DNA. The PCR primers carry sequences required for DNA clustering during NGS, negating the requirement for separate adapter ligation. Quality control (QC) is conducted to assess DNA fragment size distribution and adapter DNA incorporation prior to NGS. Sequence output files are filtered for LTR-containing reads, and the sequences defining the LTR and the linker are cropped away. Trimmed host cell sequences are mapped to a reference genome using BLAT and are filtered for minimally 97% identity to a unique point in the reference genome. Unique integration sites are scrutinized for adjacent nucleotide (nt) sequence and distribution relative to various genomic features. Using this protocol, integration site libraries of high complexity can be constructed from genomic DNA in three days. The entire protocol that encompasses exogenous viral infection of susceptible tissue culture cells to integration site analysis can therefore be conducted in approximately one to two weeks. Recent applications of this technology pertain to longitudinal analysis of integration sites from HIV-infected patients.

We describe a protocol for amplifying retroviral integration sites from the genomic DNA of infected cells, sequencing the amplified virus-host junctions, and then mapping these sequences to a reference genome. We also describe techniques to quantify the distribution of integration sites relative to various genomic annotations using BEDTools.

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of ...

3' End Sequencing Library Preparation with A-seq2

Studies in the last decade have revealed a complex and dynamic variety of pre-mRNA cleavage and polyadenylation reactions. mRNAs with long 3&#39; untranslated regions (UTRs) are generated in differentiated cells whereas proliferating cells preferentially express transcripts with short 3&#39;UTRs. We describe the A-seq protocol, now at its second version, which was developed to map polyadenylation sites genome-wide and study the regulation of pre-mRNA 3&#39; end processing. Also this current protocol takes advantage of the polyadenylate (poly(A)) tails that are added during the biogenesis of most mammalian mRNAs to enrich for fully processed mRNAs. A DNA adaptor with deoxyuracil at its fourth position allows the precise processing of mRNA 3&#39; end fragments for sequencing. Not including the cell culture and the overnight ligations, the protocol requires about 8 h hands-on time. Along with it, an easy-to-use software package for the analysis of the derived sequencing data is provided. A-seq2 and the associated analysis software provide an efficient and reliable solution to the mapping of pre-mRNA 3&#39; ends in a wide range of conditions, from 106 or fewer cells.

3&#039; End Sequencing Library Preparation with A-seq2

This protocol describes a method for mapping pre-mRNA 3&#39; end processing sites.

Studies in the last decade have revealed a complex and dynamic variety of pre-mRNA cleavage and polyadenylation reactions. mRNAs with long 3&#39; ...

Visualization of 3D White Adipose Tissue Structure Using Whole-mount Staining

Adipose tissue is an important metabolic organ with high plasticity and is responsive to environmental stimuli and nutrient status. As such, various techniques have been developed to study the morphology and biology of adipose tissue. However, conventional visualization methods are limited to studying the tissue in 2D sections, failing to capture the 3D architecture of the whole organ. Here we present whole-mount staining, an immunohistochemistry method that preserves intact adipose tissue morphology with minimal processing steps. Hence, the structures of adipocytes and other cellular components are maintained without distortion, achieving the most representative 3D visualization of the tissue. In addition, whole-mount staining can be combined with lineage tracing methods to determine cell fate decisions. However, this technique has some limitations to providing accurate information regarding deeper parts of adipose tissue. To overcome this limitation, whole-mount staining can be further combined with tissue clearing techniques to remove the opaqueness of tissue and allow for complete visualization of entire adipose tissue anatomy using light-sheet fluorescent microscopy. Therefore, a higher resolution and more accurate representation of adipose tissue structures can be captured with the combination of these techniques.

The focus of the present study is to demonstrate the whole-mount immunostaining and visualization technique as an ideal method for 3D imaging of adipose tissue architecture and cellular component.

Adipose tissue is an important metabolic organ with high plasticity and is responsive to environmental stimuli and nutrient status. As such, various ...

Assessment of the Metabolic Effects of Isocaloric 2:1 Intermittent Fasting in Mice

Intermittent fasting (IF), a dietary intervention involving periodic energy restriction, has been considered to provide numerous benefits and counteract metabolic abnormalities. So far, different types of IF models with varying durations of fasting and feeding periods have been documented. However, interpreting the outcomes is challenging, as many of these models involve multifactorial contributions from both time- and calorie-restriction strategies. For example, the alternate day fasting model, often used as a rodent IF regimen, can result in underfeeding, suggesting that health benefits from this intervention are likely mediated via both caloric restriction and fasting-refeeding cycles. Recently, it has been successfully demonstrated that 2:1 IF, comprising 1 day of fasting followed by 2 days of feeding, can provide protection against diet-induced obesity and metabolic improvements without a reduction in overall caloric intake. Presented here is a protocol of this isocaloric 2:1 IF intervention in mice. Also described is a pair-feeding (PF) protocol required to examine a mouse model with altered eating behaviors, such as hyperphagia. Using the 2:1 IF regimen, it is demonstrated that isocaloric IF leads to reduced body weight gain, improved glucose homeostasis, and elevated energy expenditure. Thus, this regimen may be useful to investigate the health impacts of IF on various disease conditions.

The current article describes a detailed protocol for isocaloric 2:1 intermittent fasting to protect and treat against obesity and impaired glucose metabolism in wild-type and ob/ob mice.

Intermittent fasting (IF), a dietary intervention involving periodic energy restriction, has been considered to provide numerous benefits and counteract ...