The Molecular Genetics Part of the Yeast Project

Prepared by Jon Schrock

The Theory

Before starting the molecular genetics part of the yeast project, we were working on the first part, using our three-cross strategy to analyze the K+ mutants. Once we had a group of seven mutants that all had a mutation in the same gene, we started the second part of the project. The objective of this part is to figure out which one gene in the yeast genome has mutated to transport potassium in this group of strains. At the bottom of this page, you can find a chart showing the genotypes of strains mentioned on this page.

The first step was to verify that the mutation did not occur in the HXT6 or HXT7 gene. These genes code for hexose transporters, which have already been shown to have the ability to mutate and transport potassium. We suppressed most of the hexose transporters in our yeast by selecting the mutants on glucose deficient media. However, HXT6 and HXT7 are constitutive, that is, they are present whether or not glucose is present in the media (Liang and Gaber, 1996). To make sure that these genes did not mutate to transport potassium, we transformed x111-4D with the pJO167 plasmid. x111-4D is a spore we obtained from one of the seven mutants in the group we are examining. The plasmid contains wild type HXT6 and HXT7 genes as well as a uracil gene, which confers the ability to grow on uracil deficient media. We isolated the plasmids from E. coli using Wizard® Plus SV Minipreps DNA isolation and used them to transform x111-4D. Since x111-4D is unable to grow on -ura media, any colonies that grew on the uracil deficient plates would have received a piece of DNA isolated from the plasmid. We replica plated these colonies onto 7K media and found that all of the colonies grew, which is what we were hoping for. Had the mutation in x111-4D been in either the HXT6 or HXT7 gene, the wild type DNA obtained from the pJO167 plasmid would have suppressed the K+ phenotype when it recombined with the chromosomal DNA in the transformants. Now that the HXT6 and HXT7 genes have been ruled out, we know that we have a new, previously unidentified K+ mutation. (See July 6, 1998 current development)

Now that all possibilities for previously identified K+ mutants have been ruled out, the second step is to figure out which new gene has mutated to transport potassium. The procedure for this step is similar to that of the previous step except that we are transforming a large population of mutant yeast with all of the yeast genes, not just HXT6 and HXT7, and looking for the one transformant that does not grow.

The first part of this transformation procedure is to isolate DNA from the Snyder library. The Snyder library is the entire yeast genome cut into pieces and ligated into anE. coli plasmid along with a transposon which contains the lacZ gene, a leu2 gene, and a gene conferring resistance to ampicillin. Again, we isolate the plasmids from E. coli, growing the E. coli in media containing ampicillin so that only cells that have a transposon would grow. The next step is to cut the DNA and transform x212-9B with cut DNA from the Snyder library. x212-9B is a strain we engineered by crossing one of the seven mutants from the group with HY480. We select our transformants on leucine deficient media, since the transposon confers the ability to grow on -leu plates.The wild type fragments then recombine with complementary bases on the yeast chromosome. Crossing over occurs, and the yeast cell expresses the piece of DNA from the library rather than its own DNA. If a piece of library DNA inserts into our mutant gene, that yeast cell would lose its ability to grow on 7K media, since the library DNA is wild type. Once the transformant is found that loses the mutant phenotype, we will rescue the gene containing the mutation. In this step, we isolate another plasmid from E. coli called pRSQ. It contains the ura3 gene, allowing the previously ura- cells to grow on uracil deficient media. We go through a similar transformation procedure, cutting the plasmid and causing the pRSQ-ura3 portion of the DNA to enter the K- transformant. At each end of this insert, this piece of DNA has a portion which is complementary to the lacZ gene, so when the pRSQ-ura3 insert enters the K- transformant, the insert will replace the lacZ gene. On the diagram to the left, the blue portions on the pRSQ-ura3 insert represent DNA that is complementary to the lacZ gene in the original transposon, also shown in blue. After we isolate the double-transformed yeast cells, we will isolate this DNA from them and then cut it, religating it into a plasmid. Once the DNA is in plasmid form, we will clone it in an E. coli vector. We will transform E. coli cells with our plasmids and grow the cells on media that selects for those cells which have received the new plasmid.The E. coli cells replicate the plasmid along with their own DNA, producing many copies of our plasmid. We will then isolate our plasmids from these E. coli cells and begin the sequencing procedure. We will construct a primer which will attach to the left end of the transposon. After sequencing about 30-35 base pairs from that point, we will send that base pair sequence into the yeast genome on the internet, and the database will tell us which gene that sequence belongs to.

To learn more about the Snyder library, follow these links:

mTn-lacZ/LEU2

The Practice

This section is a more detailed explanation of what has already been done on this part of the project, including the problems we have had and how we solved them. Hopefully this will be an aid for future yeast lab workers so that they don't have to struggle with the same problems.

Transforming x111-4D with pJO167

The procedure where we transformed x111-4D with pJO167 took much longer than it should have. The major stumbling block was getting the DNA to cut. We would run gels with DNA we had isolated cut with NsiI, the enzyme that cuts pJO167, and the cut and uncut lanes would look the same, showing us that the enzyme had not worked. The first thing we tried was to use new restriction enzyme, thinking that the enzyme we were using might have been denatured. When that didn't work, we made all new reagents for the STET prep DNA isolation procedure we were using at the time. When that didn't work, we used a commercial DNA isolation procedure we found in the lab. This kit, the Wizard® Minipreps DNA isolation kit (different from the Wizard® Plus SV Minipreps kit), had all of the necessary reagents with it except for the final solution in which the DNA was dissolved. On the protocol, the previous user had written that it was better to use TE buffer than water to suspend the DNA, so we did that. Still, the DNA would not cut. The final variable was the TE buffer. We used the same isolation kit, but suspended the DNA in nuclease free water that came with a new DNA isolation kit, the Wizard® Plus SV Minipreps kit, instead of the TE buffer we had been using, and the DNA cut perfectly! We still don't know what was wrong with the TE buffer, which was new, but we never again had problems isolating or cutting DNA when using the Wizard® Plus kit.

The next stumbling block in this part of the project was figuring out how long to cut the DNA. In order for the transformation of x111-4D to work, we had to linearize the pJO167 plasmid by cutting it once, but there is more than one NsiI site on pJO167. If we let the enzyme cut too long, it would cut twice and wouldn't be useful for transformations. We first tried cutting the DNA for 30 minutes, 1 hour, and 2 hours and found that most of the plasmids cut twice at 30 minutes. We went through tests like this for a few days and ended up making a 1:50 dilution of NsiI (1 part NsiI, 5 parts NsiI buffer, 45 parts water). Again, we cut the DNA for a range of times and found that cutting pJO167 for one hour with this 1:50 dilution of enzyme produced the most one-cut plasmids. (See June 18, 1998 current development) We also found that the dilution must be made fresh each time you want to cut DNA. A diluted enzyme will not work after being stored in the freezer with the other enzymes.

Once we got the enzyme to cut like we wanted it to, we transformed x111-4D by electroporation. We had contamination problems which we traced to the morning dilution procedure and eventually fixed. At first, we selected the transformants on -ura + sorbitol plates, but we read somewhere that it was actually better not to use sorbitol in the plates, so we then selected our transformants on -ura plates without sorbitol. The transformants that grew were very small and did not grow larger with time, so we had to pick the colonies under a dissecting scope and streak on a new YPD plate. This isolation worked well, and we replica plated the transformants onto 7K media. All of the transformants grew, showing us that our mutation is not in the HXT6 or HXT7 genes.

Transforming x212-9B with the Snyder library

Transforming x212-9B with the Snyder library is the major portion of the second part of the yeast project. The first step was to find a strain from our group of seven strains which had the right mating type and the right amino acid markers. To find this specific strain we made x212, with parents GC604 (one of the seven strains in the group) with HY480. The haploid strain we were looking for needed to be leu-, ura-, K+, and mating type a. We also wanted as many amino acid markers as we could get. Ideally, the spore would have been ade+, trp+, and lys-. Unfortunately, none of the dissected x212 spores were exactly what we were looking for, but x212-9B was the best. It had all of the required characteristics but was ade+, trp+, and lys+. We reasoned that two markers would be sufficient.

The next step was to make x218, with parents x212-9B and HY480. The plan was to transform x218 with the Snyder library; however, x218, which should be ura-, kept reverting to ura+. We tried making new media, making new dropout powders, and using noble agar, but we couldn't get x218 to stay ura-. After some brainstorming, we decided that we could just transform x212-9B, and then cross any K- transformants with a mating strain that we would find later.

We used the electroporation transformation procedure during May term and summer of 1998. The protocols for the electroporation transformation procedure and yeast prep can be found on the lab procedures page. Our first problem was cutting the plasmid DNA that we isolated using the STET prep procedure with NotI, the enzyme we use to cut the Snyder library plasmids. Read the pJO167 section to see how we solved that problem. At first, we had trouble getting good transformation efficiency. When we tried to use more DNA in each transformation, the cuvettes sparked when we zapped them, killing the yeast cells. We reasoned that we needed to use more concentrated DNA in our transformations and set out to test the concentration of DNA obtained from the Wizard® Preps. We did this by comparing the intensity of a band of our Snyder library DNA on a gel with the intensity of a band that came from a known concentration of DNA. It was a rough quantification, but it was good enough for our purposes. The DNA we had been isolating from the Wizard® Preps was about 0.24µg/µl. We did a new Wizard® Prep, but eluted the DNA using only 25µl of nuclease free water instead of the 100µl called for in the protocol. Fortunately, the smaller amount of water was able to dissolve as much DNA as the larger amount, and our DNA concentration was around 1µg/µl. From then on, we used the concentrated Wizard® Plus SV Minipreps DNA isolation protocol. After getting the DNA isolation procedure straightened out, we proceeded with the transformations and started getting relatively good transformation efficiency. (See July 6, 1998 current development) Periodically, we would encounter contamination problems. Some of these were frustrating, but we worked through them all. We continued throughout the summer of 1998 to do transformations and then replica plating onto 7K media. None of the transformants was K-. The electroporation procedure worked well but was time consuming and required two people to do well. We had to grow up yeast each night and then dilute it the next morning. Five hours later, we would start the yeast prep. The yeast prep, transformation, and plating out took about three to four hours, making it a full day. This was fine during the summer with two people but wasn't practical for one person during the school year. Luckily, we found a much simpler and faster transformation procedure which gave as good or better transformation efficiency.

During the 1998-99 school year, we have been using the FastTrack® Yeast Transformation kit. This procedure was simpler for several reasons. First, competent yeast cells could be stored in the deep freeze indefinitely, eliminating the need to do a yeast prep before every transformation. Second, the transformation procedure could take as little as one hour or as long as three hours, giving us a lot of flexibility. Third, timing of individual steps was not as critical as it was in the electroporation procedure, eliminating the need for two people to work together. The FastTrack® transformation procedure uses considerably more DNA than the electroporation procedure, so we began using the Wizard® Plus Midipreps DNA isolation kit, which yields 300µl of DNA at roughly 0.3µl/µg, which is concentrated enough for this procedure. After experimenting with different amounts of NotI, we found that using 1/3 as much enzyme as DNA (divide enzyme amount by five if using 5x NotI) was ideal. All of the 1998-99 school year was spent doing Wizard® Midipreps, transformations, and replica plating onto 7K. There were some problems with the Wizard® Preps procedure, which you can read about in the description on the lab procedures page, but for the most part it worked very well. Until the end of the year, the transformation procedure was also working well, but we hadn't found any K- transformants. We found a lot that looked K-, but after they were transferred to YPD and then replica plated again turned out to be K+. In early March 1999, we started running into problems with the transformation efficiency. It was as if one day the FastTrack® procedure stopped working. At first we weren't too concerned because there aren't many variables in this transformation procedure, and we thought it would be easy to find the problem. We tested the DNA isolation procedure by running some of the isolated DNA on a gel, and the gel showed that we had plenty of DNA and that it cut normally. We did a new yeast prep and tested our media, but still the procedure didn't yield any transformants. Finally, we thought that maybe we had received a defective transformation kit, so we used a new kit but encountered the same problem. At this point we have suspended transformations and really have no idea what is wrong, since we have tested all of the variables. If we can't find out what the problem is, we may need to find a new transformation procedure.

Luckily, one of the last transformation procedures that did work produced five potential K- transformants. These transformants also appeared K- after being grown up on YPD and replica plated again. The five transformants were T107, T110, T111, T112, and T113, and they came from S7. After growing on YPD, T107, T110, and T112 lost their ability to grow on -leu plates, showing us that the transposon isn't very stable. In order to prevent this from happening to T111 and T113, which are still leu+, these strains and any future K- transformants must be kept on -leu media and transferred frequently. The next step was to find a mating strain with which to cross our two remaining K-, leu+ transformants. To do this, we remade x218 with parents x212-9B and HY480. The spore we were looking for needed to be his+, lys+, ura-, trp-, leu-, ade-, K+, and alpha mating type. x218-7B fit all of these requirements. The next step was to cross our K- transformants with this strain. We made x224 with parents x218-7B and T111; and x225 with parents x218-7B and T113. Dissection of these crosses should verify that the K- transformants received only one transposon and that it inserted into our mutant gene.

The following table shows the genotypes of strains mentioned on this page.

Reference

Liang, H. and Gaber, R.F. Dec 1996. A Novel Signal Transduction Pathway in Saccharomyces cerevisiae Defined by Snf3-regulated Expression of HXT6. Molecular Biology of the Cell 7: 1953-1966

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