New & Noteworthy

Ribosomes Caught in the Act

November 24, 2014

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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.

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.

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

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.

Lots of Ways to Get to the Same Place

November 13, 2014

 

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

Finding the Right Tools for the Job

November 6, 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.

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