New & Noteworthy

Mother Yeast Keeps Daughters Tidy

December 18, 2014


A mess in a room is annoying, but a mess in a cell may be lethal. Image by Tobin via Flickr

Yeast need translation and organelle tethering to sequester misfolded proteins away from the cytoplasm:

Sometimes parents feel like they’re constantly nagging their kids to clean things up. Eventually, some parents just give in and do it themselves. It turns out that yeast cells aren’t all that different.

And it is even more important for cells to clean up their cellular trash than it is for that sullen teenager. If trash like misfolded or damaged proteins is not sequestered from the rest of the cytoplasm, it can cause other proteins to change their conformation or interfere with metabolic processes. This is obviously much worse than having a smelly room!

One way that cells bin their trash is by compacting it into globs, or aggregates. While this works in the short run, as cells age these aggregates build up, and in human cells, they’re hallmarks of diseases like ALS and Alzheimer’s.

In a new study published in Cell, Zhou and colleagues looked at the process of protein aggregation in S. cerevisiae. A number of other studies had already suggested that just like human parents, mother yeast cells clean up trash for their daughters. The researchers confirmed this and also made a couple of very surprising findings.

Zhou and coworkers discovered that although aggregates are mostly composed of previously translated proteins, they don’t form without new translation. And they found that aggregates aren’t littered around the cytoplasm, but instead are collected in very specific trash bins located on the surface of cellular organelles.

To do these studies, the scientists assembled a toolkit of ways to induce and visualize protein aggregation.  They used stresses like heat shock or various chemicals to stimulate aggregate formation.

They visualized the aggregates by using GFP-labeled Hsp104p, which binds to them specifically. They also had some thermally unstable reporter proteins fused to different colored fluorescent markers that helped them track aggregation. And they created time-lapse videos to watch the whole process.

They first looked in detail at aggregate formation, creating two different kinds of cellular trash (aggregates of distinct sets of marker proteins) at different times to ask whether they would all end up in the same aggregates or in separate ones. These experiments showed that in general, newly aggregated proteins will join existing aggregates rather than creating new ones. And intriguingly, these results also hinted that new translation was needed to start the aggregation process.

Zhou and coworkers tested this directly by adding cycloheximide, an inhibitor of translation, to their aggregation experiments. Sure enough, cycloheximide prevented all of the treatments from causing aggregation, and other treatments and conditions that blocked translation did the same thing.

So just as the threat of taking away the car keys may get that sullen teenager off the couch to start cleaning up, newly synthesized polypeptides are the inducer for the cellular clean-up. Without them, all the trash just stays littered around the cell.

The researchers guessed that if aggregation starts at sites where translation is occurring, it might be concentrated at the surface of the endoplasmic reticulum (ER), where a large proportion of ribosomes are bound. They used several different sophisticated microscopy techniques to confirm that most aggregates were in fact associated with the ER.

But they also got a surprise: in addition to the ER, many aggregates were associated with the mitochondrial surface and some even formed directly on it.* The authors also observed aggregates that formed at the ER but migrated along it to ER-mitochondrial contact points and eventually to mitochondria. So, cells don’t litter their trash just anywhere; they start specific collection points on the surfaces of organelles. And much of the trash seems to end up attached to mitochondria.

The researchers noticed something else when they looked at aggregates in cells that were dividing. When a bud forms, mitochondria are actively transported into it. However, the aggregates that were on mitochondria didn’t go into buds. They stayed on mitochondria that remained in the mother cell. Mom protects her daughter from any trash that mom created!

To figure out what controls this asymmetric segregation, Zhou and colleagues tested a panel of 72 mutant strains, each with a deletion in a mitochondrial outer membrane protein. One strain, the fis1 null mutant, was markedly defective: aggregates often went into the bud. Fis1p is known to be involved in mitochondrial fission, but this result suggests it may have an entirely separate role in making sure that trash-bearing mitochondria stay in the mother cell.

And finally, the authors saw that as mother cells got older, they got less and less able to keep aggregates out of their daughters’ cytoplasm. Towards the end of the mothers’ lives, aggregates were distributed more or less randomly between mother and daughter.

Trash build up is a big problem in teenagers’ rooms and in cells. Just like mom, the cell packs the trash away into bins where it will do less harm. Unfortunately, as the cell (and mom) get older, this gets harder and harder to do. In both cases, the daughter is saddled with more and more trash as mom struggles to keep up. And this is bad for the daughter as well as for the mom.

So, there’s actually a very good reason behind all that nagging to clean up your room. The secret to a long life is to always pick up your trash!

*New data from the Weissman lab, described here in a recent blog post, dovetail nicely with this finding since they establish that a lot of translation takes place on the mitochondrial surface.

by Maria Costanzo, Ph.D., Senior Biocurator, SGD

Categories: Research Spotlight

Tags: mitochondria, protein aggregation, Saccharomyces cerevisiae

New Protein Modification and Abundance Data in YeastMine

December 17, 2014

Have you ever wondered what’s happening to your favorite protein as it’s hanging out in the cell? SGD’s advanced search tool, YeastMine, now includes four new templates that can be used to find protein modification and abundance data.

The Gene -> Protein Modifications template retrieves phosphorylation, ubiquitination, succinylation, acetylation and methylation data, currently curated from the following 11 publications: Peng et al. 2003, Hitchcock et al. 2003, Seyfried et al. 2008, Vogtle et al. 2009, Ziv et al. 2011, Mommen et al. 2012, Henriksen et al. 2012, Swaney et al. 2013, Kolawa et al. 2013, Weinert et al. 2013, and Wang et al. 2014.

The Gene -> Experimental N-termini and N-terminal modifications template retrieves experimentally-determined amino-terminal sequence and acetylation data, currently curated from Vogtle et al. 2009 and Mommen et al. 2012.

Lastly, two new templates pull protein abundance data curated from Ghaemmaghami et al. 2003. Gene -> Protein Abundance retrieves molecules/cell counts for a gene or list of genes. The same data can be quickly filtered using the Retrieve -> Proteins in a given molecules/cell abundance range template.

Please explore these new YeastMine protein data templates, and send us your feedback.

Categories: New Data, Data updates

Telomerator is the Chromosomal Terminator

December 10, 2014


A simple tool for adding telomeres to linear DNA:

First there was the Terminator. Now, in a new study published in PNAS by Mitchell and Boeke, we have the telomerator.

When he cuts you up, you won’t survive. But when the telomerator cuts your circular DNA, it will survive fine as a stable, linearized piece of DNA. Image via Wikimedia Commons

Instead of being a homicidal robot from the future bent on killing Sarah Connor, the telomerator is a tool that lets scientists easily turn circular DNA into stable chromosomes in the yeast Saccharomyces cerevisiae. While less splashy, this bit of synthetic biology is definitely cool in its own way (and much less dangerous!).

The system that Mitchell and Boeke created is very clever. They first inserted the intron from the ACT1 gene into the middle of the URA3 gene. The URA3 gene was still functional, as ura3 mutants could use it to grow on medium lacking uracil.  

They next inserted a sequence into the middle of the intron that consisted of an 18 base pair I-SceI cleavage site flanked on each side by around 40 base pairs of yeast telomere repeats (called Telomere Seed Sequences or TeSSs). This construct still allowed ura3 mutants to grow in the absence of added uracil.

The final step was to introduce the homing endonuclease I-SceI to the cell so that it cut the circular DNA precisely between the two TeSSs. The idea is that when you add the homing endonuclease, the newly linearized piece of DNA ends up with the telomere seeds on each end. Telomerase adds more repeats to the seeds until the DNA has proper telomeres. Voilà, a chromosome is born.

The URA3 gene part of the plasmid is important for selecting cells with the linearized DNA. Basically a circularized DNA will grow on medium lacking uracil but fail to grow on medium with 5-FOA, while the linearized DNA will do the opposite. In other words, the process of linearization should destroy the URA3 gene. And that’s just what they found.

Previous work had shown that to be stable in yeast, a chromosome needs to be at least 90 kilobases (kb) or so long. This is why they tested their new telomerator in synIXR, a synthetic yeast chromosome that is about 100 kb in length. This chromosome has 52 genes from the right arm of chromosome 9, two genes from the left arm, around 10 kb of nonessential BAC DNA, the native centromere CEN9, and a LEU2 marker. 

Mitchell and Boeke inserted the telomerator sequence into two different locations in the BAC part of the circularized synIXR and found that adding I-SceI appeared to linearize the DNA. In both cases they found that around 100 out of 200 cells were resistant to 5-FOA and unable to grow in the absence of uracil but could still grow in the absence of leucine. This is just what we would predict if we cut the DNA in the middle of the URA3 gene and created a stable piece of linear DNA.

They next wanted to use this tool to study the effects of telomeric DNA on nearby genes. We would predict that because of telomeric silencing, genes near a telomere will be downregulated. Any genes that affect growth when turned down should quickly become evident.

To accomplish this they inserted the telomerator three base pairs downstream of each of the 54 genes on synIXR, generating 54 new plasmids. After activating the telomerator by expressing the I-SceI nuclease, they used pulsed field gel electrophoresis to confirm that 51 of the 54 synthetic chromosomes had indeed been linearized.

As expected, they found that putting a telomere near a gene sometimes has profound effects. For example, when they linearized DNA where the telomerator was 3’ of either YIR014W, MRS1, or YIR020C-B, they got no growth. They also found many more effects on the growth rate at both 30° C and 37° C at many different, “telomerized” genes. The implication is that when these genes are near telomeric DNA, they no longer function at a high enough level for the yeast to grow well or in some cases to even survive.

To confirm that the effects they saw were due to telomeric silencing, Mitchell and Boeke tested each linearized DNA in a sir2 mutant, a key player in this form of silencing. Mutating sir2 reversed the effects of placing a telomere near the gene, further supporting the idea that the newly created chromosome ends are like normal telomeres because they undergo the same Sir2-mediated silencing.

Finally, the researchers tested the stability of the newly created chromosomes by selecting for Ura+ revertants from six individual cultures with different linearized molecules. They failed to select any revertants in which the DNA had recircularized, showing that the linear chromosomes are stable.

So in contrast to the Terminator, who sliced and diced his victims randomly, the telomerator will allow synthetic biologists to create linear chromosomes with precisely positioned telomeres. This study proved the concept, and this tool will be incredibly useful in the future, both in yeast and potentially in other eukaryotes. Both the Terminator and the telomerator can say, “I’ll be back”!

by D. Barry Starr, Ph.D., Director of Outreach Activities, Stanford Genetics

Categories: Research Spotlight

Tags: synthetic biology, Saccharomyces cerevisiae, telomere

New Alternative Reference Genomes

December 08, 2014

At SGD, we are expanding our scope to provide annotation and comparative analyses of all major budding yeast strains, and are making progress in our move toward providing multiple reference genomes. To this end, the following new S. cerevisiae genomes have been incorporated into SGD as “Alternative References”: CEN.PK, D273-10B, FL100, JK9-3d, RM11-1a, SEY6210, SK1, Sigma1278b, W303, X2180-1A, Y55. These genomes are accessible via Sequence, Strain, and Contig pages, and are the genomes for which we have curated the most phenotype data, and for which we aim to curate specific functional information. It is important to emphasize that we are not abandoning a standard sequence; S288C is still in place as “The Reference Genome”. However, we do recognize that it is helpful for students and researchers to be able to ‘shift the reference’, selecting the genome that is most appropriate and informative for a specific area of study.

These new genome sequences have been also been added to SGD’s BLAST datasets, multiple sequence alignments, the Pattern Matching tool, and the Downloads site. Please explore these new genomes, and send us your feedback.

Categories: New Data, Data updates, Sequence

Tags: strains, Saccharomyces cerevisiae, reference genome

A Lot Hinges on Ndc80

December 04, 2014


The ability to fold tent poles can help you get to beautiful places, but the ability to fold the Ndc80 complex is essential to life.
Image by Maria Ly

A collapsible protein complex makes sure that dividing cells get the right number of chromosomes:

The invention of collapsible tent poles was a boon to backpackers everywhere. These long, rigid poles provide strong support for a tent, but when it’s time to pack up and go, they fold up into a short package. The secret? Flexible regions in between the rigid sections.

Although the inventors of these poles couldn’t have known it, something similar already existed in nature. In a new paper in GENETICS, Tien and colleagues used the awesome power of yeast genetics, along with some cool biochemistry, to look at the shape of the S. cerevisiae Ndc80 complex in vivo. They found that, just like a tent pole, it bends sharply at a flexible region to fold two rigid sections next to each other. And this ability to fold is critical for accurate chromosome segregation.

During cell division, chromosomes must be correctly attached to the mitotic spindle so that the mother and daughter cells each get one, and only one, copy of each. If a yeast cell doesn’t get this process right, it can become sick or even die. And if it happens in an animal cell, it can lead to cancer.

The Ndc80 complex is shaped something like a dumbbell.

The Ndc80 complex, which is conserved from yeast to humans, is an integral part of this process because it connects the chromosomes to the spindle during mitosis. It consists of four subunits and has an elongated middle and globular parts on each end, sort of like a dumbbell. The Ndc80 protein, one of the subunits, has an unstructured  “loop” region in the middle of its elongated section.

Previous work had shown that the loop region of Ndc80 is flexible in vitro, and in vivo experiments had shown that the whole complex can change its conformation. Tien and colleagues wanted to know whether the Ndc80 loop region was important for the shape of the complex during mitosis, and whether flexibility in this region was important for function.

They started with a genetic approach, and isolated mutations in NDC80 that caused heat sensitivity. One particular allele, ndc80-121, was especially interesting. The mutant protein had two amino acid changes, near each other and near the loop region. The Ndc80 complex containing the mutant protein was just as stable, and bound to microtubules just as tightly, as the wild-type complex. So why did the cells die at higher temperatures?

Tien and colleagues visualized mitosis in the mutant cells using fluorescence microscopy. They could see that when they raised the temperature, dividing mutant cells had lots of aberrant attachments between chromosomes and the spindle. Because of these attachments, proceeding through mitosis caused their spindles to break—a lethal event.

However, if they timed the temperature shift to happen later in the cell cycle, the ndc80-121 mutant cells were fine. If chromosomes had already been lined up correctly on the spindle before the temperature was raised, then the rest of mitosis could go on without a problem.

Tien and coworkers wondered whether the mutation might disrupt the binding of some other protein to the complex at high temperatures. To look for interactions, they selected mutations that suppressed the heat-sensitive phenotype of ndc80-121. But they didn’t find any suppressor mutations in other genes. However, they did find an intragenic suppressor mutation within the ndc80-121 gene.

Interestingly, this mutation affected a residue that was on the other side of the loop relative to the original two changes. If the Ndc80 complex is a dumbell, imagine that the dumbell is collapsible like a two-segment tent pole, with the loop region of Ndc80 as the elastic between the sections. If you folded the complex in this way, the amino acids changed in the ndc80-121 mutant protein would be positioned close to the amino acid that the suppressor mutation affected—an intriguing explanation for how these mutations might affect each other.

The flexible loop region of Ndc80 allows it to fold tightly, like a tent pole.

Of course, genetic interactions don’t prove a direct physical interaction. So the researchers looked to see whether they could detect physical interactions between these regions. They treated the complex with a reagent that would permanently cross-link amino acids that were close to each other. Then they chopped the complex into smaller peptides using a protease, and analyzed the cross-linked peptides using mass spectrometry to locate the linked residues.

Sure enough, they were able to detect multiple cross-links within the complex, and their locations confirmed that the complex folds much like a tent pole. Based on their mutant phenotypes, the researchers think it’s likely that the original ndc80-121mutation destabilizes folding of the complex and that the intragenic suppressor mutation makes folding tighter. Consistent with this idea, the intragenic suppressor mutation alone confers a slow-growth phenotype, as if it makes the complex fold just a little too tightly to support vigorous growth.

These experiments as a whole establish that the Ndc80 complex folds tightly early in mitosis. So, creative inventors and Mother Nature have arrived at similar solutions for the tent pole and for this important complex. And just as collapsible tent poles have become ubiquitous in the backpacking world, so too has the collapsible Ndc80 complex been conserved throughout evolution: even the specific residues that mediate the folding are highly conserved. Since this work has shown that correct folding of the yeast complex is necessary for its role in helping chromosomes to line up accurately on the spindle, the same is almost certainly true in mammalian cells.

by Maria Costanzo, Ph.D., Senior Biocurator, SGD

Categories: Research Spotlight

Tags: protein structure, mitosis, Ndc80 complex, Saccharomyces cerevisiae

Ribosomes Caught in the Act

November 24, 2014


If you want to see what animals really do out in the wild, first you need to hide a camera and a trip-wire so well that the jungle seems totally undisturbed. Then, if you’re lucky, you’ll be able to catch them in the middle of the night as they pass by. Now you can surprise that tiger and find out what he is doing at that specific spot.

The Weissman group has developed a technique for catching ribosomes as they go about their normal business, just like this tiger making his nightly rounds of the jungle. Image from Wikimedia Commons

In two companion Science articles from the Weissman group at UCSF, Jan et al. and Williams et al. did essentially the same thing to S. cerevisiae ribosomes. They hid a molecular tag and the enzyme that recognizes it at various interesting places within yeast cells, so cleverly that the cells had no idea anything was different. Instead of a flash of light, they used a pulse of the small molecule biotin to find out which mRNAs were being translated at specific locations in the cell.

What they found was that when ribosomes are translating proteins that are targeted to a particular organelle, they hang around the surface of that organelle—way more frequently than was previously thought. And the exquisite specificity of this technique, allowing them to pinpoint one particular mRNA within the cell, uncovered a fascinating case of dual protein localization.

Pinpointing Translation Locations

The researchers needed to develop a technique for catching ribosomes in the act of translation. One part of this had already been worked out in the same group: ribosomal profiling, a method that allows you to map very precisely the positions of ribosomes on mRNAs.

Briefly, cells are lysed and translating ribosomes are treated with nucleases that nibble away mRNAs, except for the 30 nucleotides or so that are protected within the ribosome. Then those protected fragments are analyzed by deep sequencing. This shows, at the single nucleotide level, where ribosomes are sitting on each individual mRNA.

Ribosomal profiling tells us where translating ribosomes are in relation to mRNAs, but not where they are in relation to the rest of the cell. To get this location information, the researchers came up with a clever tagging strategy.

They started with a bacterial gene, E. coli BirA, that encodes a biotin ligase—an enzyme that can attach biotin to specific acceptor peptides. They fused BirA to various yeast genes in order to target biotin ligase to different places in the cell.

Next they tagged ribosomes by putting a biotin acceptor, called the AviTag, on ribosomal proteins such that the tag would be sticking out on the ribosomal outer surface. They tested both the BirA and AviTag fusions to make sure that they didn’t interfere with the functions of any proteins. Just like the camera hidden in the jungle, the tags didn’t perturb yeast cells in the least.

Now the researchers were set to surprise ribosomes with a pulse of biotin. Any ribosomes that were close to BirA would become biotinylated. The tagged ribosomes could then be isolated, and the mRNA sequences being translated in those ribosomes could be identified. The method as a whole is termed proximity-specific ribosomal profiling.

A translating ribosome. Image from Wikimedia Commons

Jan and coworkers set up and validated this method in their paper, and used it to look at translation of secretory proteins at the surface of the endoplasmic reticulum (ER), while Williams and colleagues used the method to look closely at translation at the mitochondrial surface. Import into both of those organelles has previously been studied intensively, but often in vitro and mostly for just a few model protein substrates. In contrast, proximity-specific ribosomal profiling gives us the ability to look at translation of the entire proteome in vivo.

While it was known before that proteins targeted towards a certain organelle tended to be translated near that organelle, these researchers found that it was much more common than previously believed. For example, they found that most mitochondrial inner membrane proteins were translated at the mitochondrial surface and imported cotranslationally, in contrast to the previous view that mitochondrial import is predominantly posttranslational.

Both studies discovered many more details than we can summarize here. But the comparison between the ER and mitochondrial studies led to a special insight about one protein.

Osm1p Goes Both Ways

Osm1p, fumarate reductase, was thought to be a mitochondrial protein (although results from a few high-throughput studies had hinted at a link to the ER). But proximity-specific ribosomal profiling showed very clearly that it was translated at both the ER and mitochondrial surfaces. Williams and coworkers went on to confirm by fluorescence microscopy of an Osm1p-GFP fusion that Osm1p is indeed present in both ER and mitochondria.

Both of these organelles have pretty strict criteria for the signal sequences of proteins they import, so how could it be possible that the same protein goes to both locations? The researchers found that in fact, it’s not! They repeated the ribosomal profiling on the OSM1 mRNA, this time adding the drug lactimidomycin which makes ribosomes pile up at translational start sites. This showed that OSM1 actually has two start codons and produces two different proteins targeted to the two locations.

The OSM1 methionine codon currently annotated as the start would produce a protein with an ER targeting signal. Ribosomes piled up there, but also at another methionine codon 32 codons downstream. Starting translation at this codon would produce a protein with a mitochondrial targeting signal. Williams and colleagues confirmed this idea by showing that mutating the first Met codon made all of the Osm1p go to mitochondria, while mutating the Met codon at position 32 sent all of it to the ER.

The mutant form of Osm1p that couldn’t go to the ER conferred an intriguing phenotype: the inability to grow in the absence of oxygen. Osm1p generates oxidized FAD, which is necessary for oxidative protein folding, and it also interacts genetically with ERO1, which is involved in this process. Taken together, this all suggests that Osm1p activity drives oxidative protein folding in the ER.

The traditional ways of determining where a protein is in the cell, microscopic visualization or physical fractionation, can both be difficult and imprecise. Proximity-specific ribosomal profiling gets around those challenges, and gives a very precise picture of exactly where proteins are being created and how ribosomes are oriented with respect to organelles.

The example of Osm1p localization gives just a hint of the insights that are waiting for scientists who exploit this technique further. And we’re not just talking about yeast: the authors tested and validated the method in mammalian cells. Just like that tiger, surprised ribosomes in many different cell types will be giving up their secrets about where they roam and what they do.

by Maria Costanzo, Ph.D., Senior Biocurator, SGD

Categories: Research Spotlight

Tags: protein targeting, translation, mitochondria, endoplasmic reticulum, Saccharomyces cerevisiae

Better Performance for SGD’s Locus Summary Pages

November 20, 2014

 

We heard from many of you about recent performance problems with SGD web pages, and we greatly appreciate your feedback. 

The new features and new information recently added to SGD’s Locus Summary pages affected the performance of the web pages, particularly when viewed with the Google Chrome browser. We’ve been working very hard to implement solutions to address these issues. The fix is now in place and the pages now open and respond much more quickly. Please stay in touch with us as we work together to make SGD increasingly informative and useful.

Categories: Website changes

Lots of Ways to Get to the Same Place

November 13, 2014

 

An advantage of taking one route to the post office might be that you get to see the elephant topiary in front of the zoo. For a yeast cell, taking a roundabout route to a wild-type phenotype might confer a big advantage in a different environment. Image from Wikimedia Commons

How Bad Mutations Can Help Yeast Thrive in New Environments:

If you’ve ever asked for directions from more than one person, you know there are many ways to get to the same place. But not all routes are created equal. You might be trying to get to the post office, but on some routes you’ll pass by the zoo, while on others you might pass the museum or the bus station.

Turns out that something like this may be happening with individuals in a population too. Each may be well adapted for its environment, but each may have arrived there in different ways. And although they may seem similar, they might actually be more different than they look on the surface.

In a new study in PLOS biology, Szamecz and coworkers show that a lot of these different routes to the same place happen in yeast because of bad mutations. They found that when a yeast gets a deleterious mutation, it is sometimes able to mutate its way back to being competitive with wild type again. But the evolved strain is genetically distinct from the original wild type strain. Similar phenotype, distinct genotype.

And this isn’t just an interesting academic exercise either. As any biologist knows, bad mutations are much more common than are good ones. This means that populations may often be evolving to overcome the effects of these bad mutations. This process may help to explain the wide range of diverse genotypes seen in any wild population. 

The authors started out by focusing on 187 yeast strains in which a single gene had been deleted. Each strain grew more poorly than wild type under the tested conditions.

They then took 4 replicates of each mutant along with the wild type strain and grew them for 400 or so generations. They looked for strains that had evolved to overcome the growth defect caused by the mutation. 

To take into account the fact that every strain would probably evolve a bit to grow better in the environment, they only looked for those that had gained more growth advantage than the wild type had. Around 68% of the strains showed at least one replicate that met this criterion.

So as we might expect, it is possible for a strain that grows poorly to mutate its way closer to a wild type growth rate. The next question was whether these mutant strains had mutated back to something close to wild type or to something new.

The authors decided to answer this question by doing a gene expression analysis of the wild type, the eight mutant strains, and a corresponding evolved line from each of these eight. After doing transcriptome analysis, they found that for the most part the evolved lines did not simply revert back to the original gene expression pattern of the wild type strain. Instead, they generated a novel gene expression pattern to deal with the consequences of having lost the original gene. And in the next set of experiments, the authors showed that this matters when the evolved strains are put in a new environment.

The researchers took 237 evolved lines that grew nearly as well as the original wild type strain and tested how well they each did in 14 different environments. In other words, they tested genetically distinct, phenotypically similar strains in new environments.

They found that even though the original mutant strains grew poorly in all the environments tested, the evolved ones sometimes did better. Fitness improved in 52% of the strains and declined in 8%. What is even more interesting is that a few stumbled upon genotypes that were significantly better than the evolved wild type in a particular environment.

A couple of great examples are the rpl6b or atp11 deletion mutant strains. Strains evolved from either mutant did around 25% better than the evolved wild type strain in high salt, even though both of the original mutants did significantly worse than the wild type strain. By suffering a bad mutation, the evolved strain had been rerouted so that it now grew better than wild type. 

So it looks like getting a bad mutation may not be all bad after all. It might just give you that competitive edge you need when things change. Sometimes the best way to get from point A to point B is not a straight line. 

by D. Barry Starr, Ph.D., Director of Outreach Activities, Stanford Genetics

Categories: Research Spotlight

Tags: evolution, Saccharomyces cerevisiae

Finding the Right Tools for the Job

November 06, 2014

 

Researchers Create a New Gene Editing Tool for Prototrophs and Diploids:

If you like spending your weekends tinkering with your beautiful 1957 Ford Thunderbird that only leaves the garage on sunny days, then you probably own a set of “standard” wrenches, sized in inches.  But if you wanted to tune up the trusty 2008 Subaru that gets you to work every day, you’d be out of luck. The standard wrenches are no use and you’d need a set of metric wrenches to fit the Subaru parts.

Whether you want to tinker with a classic car or a yeast genome, it’s important to use the right tools. Image from Wikimedia Commons

Molecular tools can be like that too. The genetic engineering toolkit that has been used on Saccharomyces cerevisiae for many years has been terrifically handy, but it only works for strains that have specific nutritional requirements, termed auxotrophs. It’s not so useful for so-called prototrophic strains that are already able to make all the compounds that they need. And some of the non-cerevisiae Saccharomyces strains that are being studied more and more these days happen to be prototrophic.

But in a new GENETICS paper, Alexander and colleagues describe a new toolkit that works on the prototrophic wild and industrial Saccharomyces species and even works efficiently on diploid strains. This sets the stage for faster and more versatile modification of these strains that are becoming useful models for molecular evolution, as well as helping us to make wine, brew beer, and produce chemicals. 

The “standard” toolkit for S. cerevisiae is based on nutritional markers that can be selected. For example, a ura3 mutant strain can’t grow without added uracil in the medium, but when it is transformed with the wild-type URA3 gene it doesn’t need uracil any more.

Importantly, URA3 can also be selected against: ura3 mutants can survive in the presence of 5-fluoroorotic acid (5-FOA), but wild-type URA3 strains cannot. So, starting with a ura3 mutant you can replace any desired sequence with the URA3 gene, selecting for growth in the absence of uracil; then you can tinker with a gene of interest and add the modified version back into the cell to replace URA3, now selecting for 5-FOA resistance. 

Another tool in the classic toolkit is a site-specific nuclease like SceI that can encourage any added fragments to end up in the right spot in the yeast genome. Free DNA ends at a chromosomal break stimulate integration of a transformed fragment if its ends are homologous to the chromosome near the break.

To do this kind of tinkering with prototrophic strains, the researchers needed a marker that could be selected both positively and negatively like URA3. Using a gene that has no equivalent in yeast would be an added plus, since it would be easy to detect and follow. They turned to thymidine kinase (TK), which was lost from the fungal lineage a billion years ago.

TK from Herpes simplex virus had already been expressed in yeast and was known to confer resistance to antifolate drugs. And it had already been shown in other organisms that TK makes cells sensitive to 5-fluorodeoxyuridine (FUdR). The researchers tried it in yeast, and sure enough, only cells that had lost the added TK gene were able to grow in the presence of FUdR.

Alexander and colleagues next created a gene cassette, which they named HERP (Haploid Engineering and Replacement Protocol). The cassette contained the TK gene, a galactose-inducible version of the SCEI gene, and an SceI cleavage site.

As a test case, they decided to replace the S. cerevisiae ADE2 gene with the ADE2 orthologs from seven Saccharomyces species. Transforming with a mixture of seven different sequences and retrieving all seven desired constructs would be a proof of concept showing that this procedure could be used to transform with pools of different sequences and generate a whole library of different strains. And it worked.

They first replaced the ADE2 gene in S. cerevisiae with the HERP cassette, selecting for resistance to antifolates. Next they transformed the HERP-containing strain with a mixture of seven fragments, in the presence of galactose (to turn on SceI and cut the chromosome at that location) and FUdR (to select against cells in which the HERP cassette was not replaced with ADE2). The strategy worked perfectly and efficiently: they got transformants carrying each of the genes, integrated at the correct location.

This work in haploids was all well and good, but most wild type and industrial strains are diploid. And these sorts of strategies tend to work badly in diploids because the cell uses the other gene copy for homologous recombination instead of the DNA fragments scientists put in. To be really useful, HERP would need to work in diploids too.

Alexander and coworkers managed to get HERP to work efficiently in diploids by setting up a situation where both homologous chromosomes had HERP cassettes with SceI recognition sites. Now, when they induced SceI with galactose, both chromosomes of the diploid strain were cut. This dramatically increased the efficiency of transformation: 14 out of 15 strains carried the transformed sequence on both chromosomes instead of the more typical 4% seen with other methods.

So now we have a versatile toolkit that can be used on many different makes and models of yeasts. With a little modification, it could also be applied to many other fungi. And although you can’t use the metric wrench set on your Thunderbird, these HERP cassettes work just as well on S. cerevisiae as on other Saccharomyces species.

The ability to work with prototrophic strains could be a big advantage even in lab strains of S. cerevisiae, since auxotrophic strains sometimes grow slower than wild type even when supplemented with the nutrients they need. So now we have an even larger set of tools to choose from when we want to take that old Thunderbird out for a spin. 

by Maria Costanzo, Ph.D., Senior Biocurator, SGD

Categories: Research Spotlight

Tags: genetic engineering, Saccharomyces cerevisiae, selectable marker

Taking the Pulse of a Yeast Cell

October 30, 2014

Yeast cells have a pulse of sorts, although it’s not the kind that would interest Dracula. Image from Wikimedia Commons

Halloween is the time of year when we are all reminded of vampires. And if our favorite yeast Saccharomyces cerevisiae isn’t careful, it might be a vampire’s next target.

Vampires are drawn to living things with pulses so they can suck out their blood. And in a new study in Current Biology, Dalal and colleagues have found a pulse of sorts in yeast cells.

Now of course the yeast aren’t pumping blood. Instead, hordes of proteins are pulsing from place to place within the cell. Dracula might be attracted by these pulses, but would be very disappointed indeed to find that there’s no blood involved…

It’s a pretty recent discovery that some proteins move around in a coordinated way. They come together in one location at the same time, and then later all move to another location in response to changes in environmental conditions, or even under constant conditions.  This phenomenon of “pulsatile dynamics” has been seen in organisms ranging from bacteria to human.

Dalal and colleagues harnessed the awesome power of yeast genomics, in combination with sophisticated imaging equipment, to ask a question that was unthinkable before these resources were available.  How many yeast proteins, out of the entire proteome, show pulsatile dynamics?

This was obviously an ambitious goal. The researchers started with a library of 4159 strains, each containing a different yeast open reading frame fused to the gene for green fluorescent protein.  Rather than following the location of each protein over time in great detail, which would have been a huge amount of work, they devised an ingenious scheme to narrow down the possibilities and focus on potentially pulsing proteins.

In the first phase, they looked at the library at fairly low resolution, following individual cells by taking pictures once an hour over about 10 hours.  This improved their focus right away: most proteins just stayed put, and only 170 showed any hint of pulsing.

In the next phase, Dalal and coworkers looked more closely at those 170 strains, taking a picture every 4 minutes over a 4-hour span.  This eliminated another large group and left them with 64 proteins that still seemed to show pulsatile dynamics.

Another two steps cut the list of candidates way down. Other scientists had shown previously that some yeast proteins show cell cycle-dependent pulsatile movement. Since the researchers were less interested in these, they looked at protein movement during the cell cycle and eliminated 25 proteins whose movement followed it.

They also wondered whether some of the apparent movement they saw was simply due to small shifts in the focal plane during the experiment. To control for this, they examined their candidate proteins in three focal planes, spaced 0.5 um apart.

After all of these steps, Dalal and colleagues were left with just 9 proteins. All of the proteins showed pulsing into the nucleus; seven were known transcription factors, and two were subunits of the RPD3L histone deacetylase complex.

The researchers wondered whether they might have missed any other transcription factors because they hadn’t used the right conditions to see pulsing. So they looked specifically at 122 known transcription factors, testing each under conditions known to induce their activity. This analysis confirmed the previous 9 proteins found, and added just one more.

Interestingly, eight of the ten were members of paralog pairs. In three of these pairs, both members showed pulsing. Looking in detail at the MSN2-MSN4 paralog pair, the researchers found that the pulsing of Msn2p was correlated with that of Msn4p. It isn’t yet clear whether the pulsing phenomenon evolved before the whole-genome duplication that created the paralog pairs, or alternatively whether regulators shared between paralogs later became pulsatile.

Since all of the yeast pulsing proteins are involved in transcriptional regulation, and pulsing brings them into the nucleus where they are active, it’s very likely that the pulsing is an important part of their regulatory role. And since this regulatory mechanism is conserved across species, yeast will provide a great model for studying and understanding it. Dracula might be disappointed, but the rest of us will benefit.

by Maria Costanzo, Ph.D., Senior Biocurator, SGD

Categories: Research Spotlight

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