New & Noteworthy

Red Ink for the S. cerevisiae Genome

April 02, 2015

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Editing is an essential part of producing good writing. You might cringe when your masterpiece comes back covered in red ink, but in the end your paper is better for it.

These days, computers have made editing much, much faster and easier than using ink on paper. Some of us remember when cutting and pasting literally mean cutting the piece of paper on which your manuscript was typed and pasting the sections in a different order!

The same kind of revolution has happened when it comes to editing genomes. A seemingly obscure system used by bacteria to defend against invading phages, discovered in the 1950s, led to the development of restriction enzymes as the reagents that have enabled virtually all modern molecular biology. And now, an equally “obscure” system of bacterial immunity has opened the door to genome editing that is as precise and nearly as fast as your thesis advisor using Microsoft Word to edit your dissertation (maybe faster!).

This bacterial system is called CRISPR/Cas. Bacteria use it to defend against the foreign DNA—from infecting phages, bacteria that transfer plasmids via conjugation, or other sources—that constantly assaults them. And now scientists are using it to edit the genomes of most any beast they want, including our favorite yeast.

What makes this technique so powerful is that it is easily programmable. You can make virtually any sequence change that you wish, and even more impressively, make lots of changes all at once. CRISPR/Cas may be one of those technological leaps that changes everything.

The incredible potential of this approach, and the ethics of its use in humans, are hot topics right now. But while researchers and philosophers are hammering out guidelines for using the CRISPR/Cas system in larger organisms, yeast researchers are free to forge ahead and edit the S. cerevisiae genome to their hearts’ content. This has now been made much easier with the availability of a whole toolkit, created at the Delft University of Technology and described in a new paper by Mans et al. 

The yeast system has three essential components. The first is a plasmid that will express a specific guide RNA (gRNA). The gRNA leads a nuclease to the right place in the genome.  Mans and colleagues made a whole set of plasmids, with different nutritional and antibiotic resistance markers, that can express one or more gRNAs.

The second component of the system is the nuclease that binds to the gRNA and makes a double-stranded cut in the target DNA. The researchers used the Cas9 nuclease from Streptococcus pyogenes, and engineered a set of yeast strains that had the cas9 gene stably integrated into a chromosome and expressed from a strong yeast promoter.

The third component is one or more repair fragments: pieces of DNA that specify the modified sequence that the researcher wants to engineer into the genome.

So, for example, if a scientist wants to delete a gene precisely, she can create a gRNA that targets the gene, and then co-transform a Cas9-expressing yeast strain with both the plasmid expressing the gRNA and a repair fragment that corresponds to the gene’s upstream and downstream flanking sequences, fused together. When the gRNA is expressed in the transformant, it leads Cas9 to the gene, where it makes a double-stranded cut at a precise position in the DNA.

Now the researcher can let yeast do the rest of the work. S. cerevisiae has a powerful homologous recombination system, and it’s greatly stimulated by double-stranded breaks in DNA.

After Cas9 cuts the gene, yeast will repair the break, using as a template the repair fragment that the researcher designed. In this case, the upstream and downstream sequences will recombine with the homologous sequences in the chromosome, but since the coding sequence is missing from the repair fragment, the resulting strain will have a precise deletion of the gene of interest.

This example illustrates one of the simplest uses of the technique. Mans and colleagues tried successively more complicated tasks and were able to accomplish some amazing feats.

They were able to precisely delete six genes in one step by transforming with repair fragments for all six, along with three plasmids that each expressed two gRNAs. Also in one step, they replaced one yeast gene with six Enterococcus faecalis genes encoding subunits of the pyruvate dehydrogenase complex and other enzymes in the pathway. The E. faecalis genes were specified on six overlapping repair fragments that were cotransformed into the strain.

The CRISPR/Cas9 system designed by Mans and colleagues can do much more than gene deletions and replacements. By designing repair fragments specifying particular mutations, the method can also be used to create point mutations or other modifications.

For this technique to work, it’s important that there are no mismatches between the gRNA sequence and the chromosomal target sequence, and other sequence characteristics can influence the efficiency of the method.  So the authors created an online tool that helps researchers select optimal Cas9 targets in regions of interest and design gRNA sequences. Since they incorporated the sequences of 33 different S. cerevisiae strains into the tool, researchers can specify a strain and retrieve information on the best sequences for targets and gRNAs for their gene(s) of interest based on sequences found in that particular strain.

Importantly, the CRISPR/Cas9 technique allows researchers to make multiple changes in a single step. This is a big advantage, since transformation itself can be mutagenic. For example, a strain that has been commonly used to investigate the function of hexose transporters was engineered to carry multiple deletions in the conventional manner, using successive rounds of transformation and selection.  Its genomic sequence, which was recently determined, reveals that its genome is a complete mess, with many rearrangements and deletions. 

With the development of this toolkit, editing the S. cerevisiae genome is beginning to be almost as easy as editing a text document. And since Mans and colleagues have made all of the strains, plasmids, and online tool freely available to the world, everyone will be able to take advantage of them. Just think of the stories that yeast researchers will be able to write!

CRISPR/Cas in Bacteria

Way before this became a powerful tool for researchers, it was a very cool immune system for bacteria, allowing them to defend against assaults from foreign invaders. 

Bacteria collect foreign DNA sequences from invaders that threaten them, just like collecting mug shots of notorious criminals. They store these mug shots in a special place in their genome, integrated between blocks of a repeated sequence. (The CRISPR acronym refers to these repeats.) This region is transcribed, and the RNA is chopped into pieces containing individual mug shots. Because these mug shots, called crRNAs, are complementary to the DNA sequences of invaders, they can recognize and hybridize with those invading sequences.

Bacteria have an additional small RNA, the tacrRNA, that binds to both the crRNA and to a CRISPR-associated (Cas) nuclease. This forms a RNA-DNA-protein complex on the foreign DNA and allows the nuclease to do its work, cutting both strands of the DNA and neutralizing the invader.

To use this system for genome engineering, scientists have fused the two RNAs of the bacterial system into a single RNA, the guide RNA (gRNA).  It contains both the mug shot (with sequences complementary to the target) and the RNA sequence that recruits the nuclease. The gRNA, a Cas nuclease, and a repair fragment are the essential components of the system.

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

Categories: Research Spotlight

Tags: CRISPR/Cas, genome engineering, Saccharomyces cerevisiae, synthetic biology

Conformity Preferred in Yeast

March 25, 2015

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In a classic Apple ad, the world is a gray, dreary place where everyone is the same. In charges a woman dressed in brightly colored clothing who hurls a hammer into a big black and white screen, smashing the old conformist world order. Individuality can now blossom.

This is a powerful story for people, but it turns out that if Mother Nature had her druthers, she would like that woman to either stay at home or dress and act like everyone else. At least this is true if we are talking about individuals that are genetically identical. In this case, alleles that cut down on cell to cell variation tend to be the ones that prosper.

This is confirmed in a new study in Nature in which Metzger and colleagues in the Wittkopp lab showed that there is a selection against mutations that cause increased variation between individuals in the S. cerevisiae TDH3 gene. In other words, mutations that cause more “noise” are selected against. The squeaky wheel is eliminated.

The TDH3 gene encodes glyceraldehyde-3-phosphate dehydrogenase (GAPDH), an important metabolic enzyme. Yeast cells can survive a deletion of this gene, but their fitness is greatly reduced. Overexpression of the gene also has noticeable effects. This suggested to the researchers that the level of TDH3 expression would be under selection pressure during evolution.

Metzger and coworkers compared the promoters of the TDH3 gene from 85 different strains of S. cerevisiae and found that the promoter had undergone selection out in the wild. The authors were interested in why certain polymorphisms were selected for and why others were selected against. To try to tease this out, they compared the activities of evolved changes to randomly selected ones.

The authors first used the sequences of the 27 haplotypes they saw in the 85 strains to predict what the original, ancestral promoter probably looked like. They then re-created this sequence and also sequences that represented the most likely intermediates on the way to the current promoters. They linked each of these promoter sequences to a yellow fluorescent protein (YFP) reporter and looked at mean activity and expression noise. In other words, they looked at how much promoter activity there was in aggregate and how much it varied between individual cells for the 10,000 cells in each culture.

They next set out to generate a pool of polymorphisms that didn’t make the cut during evolution, so they could compare these to the successful ones. To do this, they individually mutated 236 G:C to A:T transitions throughout the promoter region. They chose this transition because these are the most common spontaneous mutations seen in yeast and the most common SNP seen in this promoter out in the wild.

Now they were ready to do their experiment! Comparing the randomly created mutations to the evolved changes, they looked at both the overall level of expression and how much variation there was between each of the 10,000 individual cells in the tested culture.

What they found was that the effects on the mean level of activity were pretty comparable between the “selected” mutations and the random ones. But the same was not true for individual variation. The random mutations were much more likely to increase expression noise compared to the “selected” mutations.

From these results the authors conclude that there is a selection against mutations that increase the level of noise. In fact, they go a step further and conclude that at least for the TDH3 promoter, there was more of a selection against noise than there was a selection for a particular level of activity. It was more important that individuals had consistent activity than it was to have some mean level of activity.

This makes some sense, as a cell is a finely tuned machine where all the parts need to work in harmony together to succeed. If one part is erratic and shows different levels of activity in different individuals, then some of those individuals won’t do as well and so won’t survive.

This also means that certain paths to a more fit organism will be selected over others. And it could be that organisms miss out on some potential fitter states because they can’t survive the dangerous evolutionary journey that would be needed to get there.

So the cell prefers that all the parts work together in a predictable way. When you’re a population of individuals that are more or less genetically identical, nonconformists are dangerous.  The gray sameness of the Nineteen Eighty-Four world is preferable to a more bohemian atmosphere where diversity is celebrated.

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

Categories: Research Spotlight

Tags: evolution, promoter, Saccharomyces cerevisiae

From Sourdough Bread to Chemotherapeutic Drugs

March 18, 2015

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Microbes can achieve great things when they work together:

Microbiologists in the lab spend a lot of time and effort keeping each microbial strain and species separate. The conventional wisdom is that if you want to really understand an organism, you need to study it in isolation, as a pure culture. If contaminating colonies of some other bug appear on your Petri dishes, you’d better melt down those plates in the autoclave and trash them!

On the other hand, amateur microbiologists have known for centuries that mixtures of microbes can do great things. Sourdough bread, for example, is made using a culture of Lactobacilli and yeasts. They complement each other during the fermentation: the bacteria metabolize sugars that the yeast can’t use, and make new compounds that can be fermented by the yeast. The result is extremely tasty. 

A new study from Zhou and colleagues brings a microbial community into the lab to make a medicine called paclitaxel that is used to treat cancer. Yes, bread bowls for your clam chowder are cool, but this is obviously way more important for human health.

Paclitaxel is an incredibly successful drug for treating breast and ovarian cancer, but unfortunately there isn’t an easy way to make it.  It can be purified from the bark of the Pacific yew (killing the trees in the process), synthesized by plant cells cultured in vitro, or synthesized chemically. But all of these processes are expensive and complicated, which means this life saving drug is always in short supply.

The researchers wanted to produce it more cheaply and easily, and an obvious solution was to let microbes do most of the work. But neither of the most commonly used microbial workhorses, S. cerevisiae and E. coli, was exactly right for the job.

E. coli had already been engineered to overproduce the compound taxadiene. The taxadiene then needs to be oxidized to create oxygenated taxanes, which are paclitaxel precursors. This oxidation can be done by membrane-bound oxidoreductase enzymes called cytochrome P450s. But these enzymes are not found naturally in bacteria, and getting them expressed and functional in E. coli is challenging.

The researchers decided to see whether they could coax these two microbes into cooperating to produce oxygenated taxanes. After creating an E. coli strain that produced taxadiene and an S. cerevisiae strain that produced a P450 oxidoreductase, they grew them together in the same culture, with glucose as the carbon source. 

As planned, the E. coli pumped out taxadiene and it was able to diffuse into the yeast cells, where it became oxygenated. However, the two species weren’t as happy together as the researchers had hoped. 

One of the things that humans love about yeast is that when it grows on glucose, it produces ethanol. However, the E. coli cells didn’t love being bathed in ethanol: their yield of taxadiene went way down as the ethanol levels in the culture went up.

So Zhou and coworkers switched the carbon source to xylose. S. cerevisiae cannot consume xylose, but E. coli can. When growing on xylose, E. coli produces acetate, which the yeast can use—and they don’t produce ethanol under these conditions.

Growing the microbes in xylose doubled the yield of oxygenated taxanes over that of the glucose-grown culture. But still, only 8% of the taxadiene that was produced was getting oxygenated.

To be sure that the yeast cells were producing the P450 enzyme as efficiently as possible, the researchers tried driving transcription of the gene using several different promoters. Using the promoter that was strongest in the co-culture conditions made a significant difference in the proportion of taxadiene that was oxygenated.

The researchers guessed that another factor limiting the final yield was that the yeast cells were not growing as well as they could. Tweaking the ratio of the two species in culture and the contents of the media resulted in a three-fold increase in oxygenated taxanes. But Zhou and colleagues hoped to improve things even more.

Thinking that the limiting step in yeast growth might be the supply of acetate, the researchers tried to beef up the acetate synthesis pathway in E. coli. They engineered the bacteria to overproduce several of the enzymes in the acetate biosynthesis pathway, but this didn’t make a large difference. 

They reasoned that if they forced the E. coli to rely on the acetate biosynthesis pathway for energy, the cells might ramp up their acetate production. To do this they blocked oxidative phosphorylation by deleting the the atpFH gene that encodes a subunit of ATP synthase. Now more acetate was produced, the yeast cells grew better, and 75% of the taxadiene that was produced got oxygenated. They were in business!

Zhou and coworkers went on to show that the co-culture environment could be modified to generate several other isoprenoids. This class of naturally-occurring molecules includes some that are in use, or in development, as pharmaceuticals (paclitaxel, and others currently in clinical trials) and compounds that have other applications, from fragrances to fuels. 

There’s much more work to be done, and the potential of microbial communities is just beginning to be realized. Harnessing the power of multiple organisms means that different steps of pathways can be optimized separately and then mixed and matched for a desired result. This approach could turn out to be the best thing since sliced bread! But then again, sourdough bakers already knew that.

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

Categories: Research Spotlight

Tags: paclitaxel, Saccharomyces cerevisiae, synthetic biology

Those Yeast Got Talent

March 11, 2015

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The winners of American Idol go through quite a selection process. They start out as one of tens of thousands of people who audition, and survive each cut until they are finally crowned.

At the first cuts, those with any sort of advantage are kept in the pool and the others dropped. As the cuts continue, contestants not only need to have had that stronger initial advantage (or a bit of luck), but they also need to have picked up some new skills from all of those off- and on-air performances.

Some of these contestants start with a lot of raw talent but then progress only a little, while others are able to hone their weaker initial talent with lots of practice. Once their numbers are winnowed down to a handful, it gets close to being anyone’s game because the remaining contestants are so talented.

A new study by Levy and coworkers paints a similar sort of picture for evolving populations of yeast. Very early on a whole lot of yeast stumble upon weak, beneficial mutations that keep them going in the population. These are the yeast that make the initial cut in the hurly-burly world of the Erlenmeyer flask.

At later times a few yeast end up with strongly beneficial mutations that allow them to start to dominate. These are the pool of yeast that are the finalists of the flask.

Of course a big difference (among many) between American Idol and the yeast in this experiment is that the pool of contestants in the flask hangs around—they are not thrown off the show. This means that some cell that didn’t do too well early on can suddenly gain a strongly beneficial mutation and begin to dominate. Until, of course, that cell is usurped by another more talented yeast, in which case that finalist will fade away unless it can adapt.

And this study isn’t just a fascinating dissection of evolutionary population dynamics either. It might also have implications for treating bacterial infections and even cancer.

Bacteria and cancer cells live in large populations with each cell trying to outcompete the others. By understanding the set of mutations that allow some cells to succeed against the others and become more harmful, researchers may be able to come up with new ways to treat these devastating diseases.

One of the trickiest parts of this experiment was figuring out how to follow lots of yeast lineages all at once in a growing culture. Levy and coworkers accomplished this by adding 500,000 unique DNA barcodes to a yeast population and using high-throughput DNA sequencing to follow the lineages in real time.

They set up two replicate cultures and followed them for around 168 generations. In both cultures the researchers saw that while most of the lineages became much less common, around 5% happened upon a beneficial mutation that allowed them to increase in number by generation 112.

In other words, around 25,000 lineages ended up with beneficial mutations that let them make the first cut in both cultures. This translates to a beneficial mutation rate of around 1 X 10^-6 per cell per generation and means that around 0.04% of the yeast genome (around 5000 base pairs) can change in a way that confers a growth advantage.

But of course not all mutations are the same. Weakly beneficial mutations are very common, which means both cultures have plenty of these early on. This is why the replicate cultures behave so similarly up to around generation 80.

Eventually, though, a few yeast stumble upon stronger, more beneficial mutations. Since these are rarer and harder to get, each replicate culture gets them at different generations. This is why the cultures begin to diverge as the 100 or so of the strongest beneficial mutations begin to dominate.

The experiment did not go on for long enough to see many double mutations. In other words, it was very rare in this experiment to see a yeast lineage succeed because it had developed additive beneficial mutations. This is because there simply wasn’t enough time for a yeast cell to get a beneficial mutation and establish itself and then have one of its lineage gain and establish a second beneficial mutation. There was no Jennifer Hudson who came in 7^th but then went on to win a Grammy and an Oscar.

When a cancerous tumor is developing, however, there is plenty of time for multiple “beneficial” mutations to be established. These mutations are only beneficial for the tumor; they are devastating for the person with cancer. This is why it is so critically important to understand not only which mutations are implicated in cancer, but also the dynamics of how they accumulate in the cancer cell population during progression of the disease. Talented yeast in the hands of talented researchers are helping us figure this out.

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

Categories: Research Spotlight

Tags: cancer, evolution, Saccharomyces cerevisiae

Sweet or Salty? It’s Hard to Tell Just By Looking

March 05, 2015

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If you have ever accidentally added salt to your coffee, you know that sugar and salt are very different things even though they look pretty much the same. Turns out that genes can sometimes be this way too. They can look similar at the DNA level but have very different functions.

A great example of this can be found in a new study in GENETICS by Varshney and coworkers. They found that a protein kinase in Candida albicans, Sch9, is important for ensuring that chromosomes end up in the right place when this yeast reproduces by budding.

Turns out that the same is not true for the Sch9 ortholog in our favorite yeast Saccharomyces cerevisiae. There is no evidence that Sch9 has anything to do with chromosome segregation there, even though the Sch9 sequences in these two yeasts look very similar.

C. albicans Sch9 is very important for keeping filamentous growth at bay under certain conditions (hypoxia and high levels of carbon dioxide). To understand better how Sch9 does this, Varshney and coworkers used chromatin immunoprecipitation (ChIP) to figure out where the protein binds in the genome. They were surprised when they found that it bound mostly to centromeres.

Despite this binding, the authors saw no evidence that Sch9 was involved in stabilizing the kinetochore, the protein structure that forms at the spindle of sister chromatids. When a kinetochore is destabilized, a cell’s nuclear morphology changes, its centromeres decluster during the cell cycle, and the centromeric histone Cse4 delocalizes away from its centromeres. The authors saw none of these things in a C. albicans strain in which the SCH9 gene was deleted.

They did, however, find that C. albicans cells lacking Sch9 had anywhere from a 150 to a 750-fold increase in chromosome loss. They found this by using a strain of C. albicans that had an arginine marker on one copy of its chromosome 7 and a histidine marker on the other, and looking for how often cells lost one of the two markers. From this the authors concluded that like many other kinetochore associated proteins, Sch9 is involved in chromosome segregation.

As a final experiment, Varshney and coworkers used ChIP to see if the Sch9 protein bound to centromeres in S. cerevisiae. It did not. While the authors did not directly test whether Sch9 had any effect on chromosome segregation in S. cerevisiae, the presumption is that it didn’t, as it doesn’t appear to interact with centromeres and no such effect has been seen previously.

But Sch9 isn’t completely different in the two yeasts. A close look at the ChIP data showed that Sch9 bound the rDNA locus in both C. albicans and S. cerevisiae.

How did orthologous proteins in two budding yeasts end up with such different functions? One idea is that the ancestral gene to Sch9 was important for rDNA regulation and that it later gained a function in chromosome segregation in C. albicans. Another possibility is that the ancestral gene had both functions and that centromere binding was lost in S. cerevisiae. More work will need to be done to tell the difference.

Whichever explanation is correct, this study reminds us that, just like sugar and salt, even if two genes look similar they may have quite different functions. Assuming that similar appearance means identical function may lead to an experimental result that is just as unpleasant as salty coffee!

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

Categories: Research Spotlight

Tags: Candida albicans, chromosome segregation, ortholog, Saccharomyces cerevisiae

Message in a Bubble

February 18, 2015

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If you’re shipwrecked on a desert island, writing a message on a scrap of paper, sealing it in a bottle, and flinging it into the ocean could be your only chance at communication. As the song goes (see below), the message in a bottle is an S.O.S. to the world.

It turns out that cells may do something very similar. But instead of using a bottle, they enclose their messages in membrane-bound bubbles.

Many different mammalian cell types have been seen to form these extracellular vesicles (EVs). In mammalian cells, it’s known that EVs are used for cell-to-cell communication. They contain signals that allow cells to influence their neighbors, both for good (for example, regulating the immune response) and for bad (transmitting viruses or toxic peptides). In most cases these signals aren’t well-characterized, but the EVs may include DNAs and proteins, and they’re rich in RNAs.

Fungi have been found to produce EVs too, but they’ve been much less studied. In a new paper in Scientific Reports, da Silva and colleagues looked at the extracellular vesicles (EVs) produced by four different fungal species, including S. cerevisiae, and found that among other things, the vesicles actually include at least parts of many RNAs, both protein-coding and non-coding.

The scientists decided to look at S. cerevisiae and three species of fungal pathogens that infect humans: Cryptococcus neoformans, Paracoccidioides brasiliensis, and Candida albicans. (S. cerevisiae can be pathogenic too, but isn’t as virulent as any of those species.) They isolated EVs from each and treated the unbroken EVs with RNase to get rid of any RNA that might be contaminating their surfaces.

Then they broke open the vesicles to see what was inside. They found that the EVs contained many small RNAs, most less than 250 nucleotides in length. The scientists used RNA-seq analysis to determine the sequences of these small RNAs, and compared them to the genomic sequences that were already known for these organisms.

Many of the sequences corresponded to noncoding RNAs. For S. cerevisiae, the RNA sequences identified included the mitochondrial small and large ribosomal RNAs, RNA components of RNase enzyme complexes, a variety of small nuclear and small nucleolar RNAs, and tRNAs.

Sequences corresponding to several dozen S. cerevisiae mRNAs were also detected. There wasn’t much rhyme or reason to the kinds of proteins they encoded.  But the set of mRNA fragments didn’t correspond simply to the set of most abundant mRNAs in the cell. So it seemed like the vesicles didn’t just contain random samples of the cytoplasm, but instead had been loaded selectively with particular mRNAs.

This study raises as many questions as it answers, and there is a lot of work to be done before fungal EVs will be understood. The intriguing discovery of RNA in EVs suggests the possibility that the RNAs could influence gene expression in cells that take up the EVs, either by regulating processes like splicing or translation, or even by encoding a protein that gets translated in the recipient cell.

Being able to influence neighboring fungal cells via EVs could be an advantage for fungi in the fierce competition for biological resources. Or perhaps EVs are used to subvert gene expression in host tissues during a fungal infection. Far from being an S.O.S., these messages could be threatening.

These speculations all need much more research. Fungi are an integral part of our world, and we need to pay careful attention to the messages that they send us!

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

“Message in a Bottle,” The Police, 1979

Categories: Research Spotlight

Tags: cell-cell communication, extracellular vesicles, Saccharomyces cerevisiae

Yeast Finds Needles in a Haystack to Combat Malaria

February 11, 2015

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Finding a needle in a haystack would take a long time and would be very tedious (although it’s been done!) Finding a specific drug to fight malaria by testing the effect of each drug, one at a time, on a purified protein in vitro would be at least as tedious and maybe even more so.

Luckily, we don’t have to sift through a haystack. In a new study in ACS Chemical Biology, Frame and colleagues used our friend S. cerevisiae to find nine drugs out of a collection of more than 64,000 that are promising candidates for stopping the malaria parasite in its tracks. It is as if yeast allowed them to set fire to the haystack and see nine needles gleaming in the ashes.

Malaria is a huge problem for global health. Plasmodium falciparum, the organism that causes malaria, is fast developing resistance to the few effective drugs that we have left.

But P. falciparum has an Achilles heel—it can’t make its own purine nucleotides! Since these are the building blocks of DNA and, obviously, essential for life, if we can keep P. falciparum from being able to take them up, we can kill it. 

P. falciparum imports purines via a major transporter protein, called PfENT1, located in the plasma membrane. So a drug that specifically inhibited this transporter could be a good way to attack the pathogen.

It’s possible to assay the activity of the transporter in vitro, adding different drugs one at a time and seeing which inhibits transport. But doing this for thousands of drugs might make you wish you were looking for a needle in a haystack. Frame and colleagues decided to harness the awesome power of yeast genetics to test a very large set of drugs more quickly.

The toxic nucleoside analog 5-fluorouridine (5-FUrd) is taken into yeast cells by the high-affinity uridine transporter Fui1. It kills normal yeast cells, but fui1 null mutant yeast can survive in the presence of 5-FUrd.

The researchers engineered a yeast codon-optimized version of pfENT1 and expressed it in the mutant, restoring 5-FUrd uptake. The nucleoside analog was again toxic to this strain, and the only way the yeast could survive was if the transporter activity of pfENT1 was inhibited.

This system allowed a simple and powerful screen for pfENT1 inhibitors. The yeast strain expressing pfENT1 would be able to grow in the presence of 5-FUrd only if pfENT1 transporter activity was blocked by the drug that was being tested.

Setting up the screen on a large scale, the scientists were able to test 64,560 compounds. They initially found 171 compounds that allowed the yeast to grow. They narrowed these down to 9 compounds that worked well and belonged to different structural classes of chemicals.

Because of the way the study was designed, it was likely that these compounds allowed yeast to grow because they prevented PfENT1 from pumping the toxic 5-FUrd into the cell. But what if the compounds were actually doing something different, and unexpected? To rule out this possibility, the researchers designed a secondary screen for the 9 top candidate drugs.

They used ade2 mutant yeast, which can’t make their own adenine and need to be fed it in order to survive.  These mutants can make do with the related compound adenosine, but it can’t normally get inside the cell; yeast doesn’t have a transporter that will take it up. However, PfENT1 can transport adenosine, so ade2 mutants can grow on it if they are expressing PfENT1.

With this system, if the candidate drugs are working as expected, they should prevent yeast growth. And that is exactly what the researchers found. This confirmed that the drugs are working because they specifically inhibit PfENT1 and do not allow growth by some other, indirect mechanism.

To be completely sure of the mechanism, the scientists did a direct test. They found that the nine drugs prevented PfENT1-expressing cells from taking up radiolabeled adenosine.

This was all fine, but the ultimate goal of the study was to affect growth of the malaria parasite. So Frame and colleagues tested the drugs on P. falciparum. 

In the presence of any of the nine drugs, the parasite couldn’t take up adenosine and also failed to grow. This even happened when the parasites were grown in medium containing  much higher purine concentrations than found in human blood.

Even though PfENT1 was targeted by the drugs, all nine of the drugs also killed Pfent1 null mutants. This suggested that the drugs have a secondary target or targets in addition to PfENT1. This could be a real advantage, because it could help prevent the parasites from developing resistance to the drugs.

All nine of these diverse drugs are promising candidates for the treatment of malaria. And the same approach could be used to find chemicals that affect the function of other transporters from various organisms. 

As usual, yeast is providing scientists with streamlined ways to find new treatments for serious human diseases. Instead of tediously rummaging about in a haystack, yeast lets us quickly and easily find the needles we need. 

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

Categories: Research Spotlight

Tags: high throughput screen, malaria, Saccharomyces cerevisiae, transporters

Ticket to Transcribe

February 04, 2015

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tRNA_CUG may not just be for translation anymore:

Back before airplanes and cars, when times got tough people would often take trains to what they hoped were greener pastures.  And to hitch a ride on a train, they’d usually need to have a ticket. Turns out the same is true for Gln3, a transcription factor in yeast.

Basically, Gln3 stays in the cytoplasm as long as there are good sources of nitrogen available to the cell.  When these sources run out, Gln3 moves from the cytoplasm to the nucleus where it can turn on genes that can help the yeast cope with its new situation.

In a new study in GENETICS, Tate and coworkers have identified one of the tickets that lets Gln3 take the trip to the nucleus. And it was totally unexpected. To get to the nucleus, Gln3 needs a fully functional glutamine tRNA_CUG. No, really.

To get this evidence, Tate and coworkers used a reporter in which Gln3 was linked to GFP (green fluorescent protein). They tracked the location of Gln3 in the cell using fluorescence microscopy.

Using a temperature-sensitive mutant of tRNA_CUG, sup70-65, the authors showed that at the nonpermissive temperature of 30 degrees C, Gln3 could not translocate to the nucleus under a wide variety of conditions in which nitrogen was limiting. Gln3 had no problems translocating at the permissive temperature of 22 degrees C, and in wild-type cells Gln3 translocated at both temperatures. Clearly tRNA_CUG is doing something important in this process!

The next experiment showed that tRNA_CUG was more like a one-way ticket. Once Gln3 entered the nucleus under nitrogen starvation conditions at the permissive temperature, switching to the nonpermissive temperature had little effect. Gln3 stayed put.

A possible wrinkle in these experiments was that cells harboring sup70-65 formed chains reminiscent of pseudohyphae at the nonpermissive temperature no matter what the nitrogen conditions. One possible explanation for the results seen here was that many of these cells lacked nuclei. In this case, they might not see nuclear translocation because there was no nucleus to translocate to.

In the course of these studies, Tate and coworkers showed that adding rapamycin mimicked the effects of nitrogen starvation with one big difference—nuclear localization happened much more rapidly than with nitrogen starvation. This fast response allowed the authors to look at Gln3 localization while visualizing nuclei by staining DNA with DAPI (which gives a short-lived signal). They were able to use the DAPI to see that these cells did indeed have nuclei and that when they raised the temperature, Gln3 did not colocalize with the DAPI stained nuclei.  Gln3 was being kept out of nuclei at the nonpermissive temperature.

So it really looks like Gln3 needs a working tRNA_CUG to get into the nucleus. There are a couple of possible ways that this tRNA could be needed for Gln3 to make the trip.

In the first model, the tRNA is part of a complex that allows Gln3 to make the trip to the nucleus. In this model, it is almost as if Gln3 (or one of its compatriots) is clutching its ticket, tRNA_CUG. In the second, less fun model, the tRNA is required to translate a protein involved in Gln3’s transit. Which model is the correct one is still up in the air, but it will be interesting to see which is the right one.

This was the most astonishing finding in the article, but it was by no means the only one. We don’t have the time to go into the other experiments, which, among other things, teased apart differences in the four or five distinguishable pathways that work to turn on the cell’s nitrogen response.

This work highlights a recurring theme in basic research: we may think we know everything that’s going on (tRNAs just help to translate proteins, right?) but just about every time we look more closely, there is much more to see than first meets the eye. Being in the right place at the right time is essential, whether you’re escaping the Dust Bowl in The Grapes of Wrath or a transcription factor responding to the lack of a nutrient. It’s not so surprising that the cell has drafted every possible player into this process, even a lowly tRNA. 

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

Since the title has this song stuck in our heads, we thought you might want to hear it too. Enjoy!

Categories: Research Spotlight

Tags: nitrogen utilization, Saccharomyces cerevisiae, transcription factor localization, tRNA

Species Can’t Risk the New Coke

January 29, 2015

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Genome organization may protect key genes from the ravages of increased mutation rate during meiosis:

Back in 1985, Coca Cola decided to completely rejigger the flavor of their flagship soft drink, calling it the New Coke. This radical change to the product was a colossal failure. Toying with such an essential part of a key product was simply too risky a move. If only they had learned from our favorite beast, Saccharomyces cerevisiae.

In a new study in PLOS Genetics, Rattray and coworkers show that the mutation rate is higher during meiosis in yeast because of the double-strand breaks associated with recombination. This makes sense, because any new mutations need to be passed on to the next generation for evolution to happen, and germ cells are made by meiosis. But their results also bring up the possibility that key genes might be protected from too many mutations by being in recombination cold spots. Unlike the Coca Cola company, yeast (and everything else) may protect essential genes from radical change.

Previous work in the Strathern lab had suggested that when double strand breaks (DSBs) in the DNA are repaired, one result is an increased mutation rate in the vicinity. The major culprit responsible for the mutations appeared to be DNA polymerase zeta (Rev3p and Rev7p).

To test whether the same is true for the DSBs that happen during the first meiotic prophase, Rattray and coworkers created a strain that contained the CAN1 gene linked to the HIS3 gene. The idea is that mutants in the CAN1 gene can be identified as they will be resistant to canavanine. The HIS3 gene is included as a way to rule out yeast that have become canavanine resistant through a loss of the CAN1 gene. So the authors were looking for strains that were both resistant to canavanine and could grow in the absence of histidine.

The first things the authors found was that the mutation rate during meiosis was indeed increased as compared to mitosis in diploids. For example, when the reporter cassette was inserted into the BUD5 gene, the mitotic mutation rate was 5.7 X 10^-8 while the meiotic mutation rate was 3.7 X 10^-7, a difference of around 6.5 fold.

This effect was dependent on the DSBs associated with recombination, since the increased mutation rate wasn’t seen in a spo11 mutant; the SPO11 gene is required for these breaks. Using a rev3 mutant, the authors could also conclude that at least half of the increased mutation rate is due to DNA polymerase zeta. This all strongly suggests that the act of recombination increases the local mutation rate.

If recombination is associated with the mutation rate, then areas on the genome that recombine more frequently should have a higher rate of mutation during meiosis. And they do. The authors inserted their cassette into a known recombination hotspot between the BUD23 and the ARE1 genes and saw a meiotic mutation rate of 1.77 X 10^-6  as compared to a rate of 4.9 X 10^-7 when inserted into a recombination coldspot. This 3.6 fold increase provides additional evidence that recombination is an important factor in meiotic recombination.

This may be more than just an unavoidable side effect of recombination. It could be that yeast and perhaps other beasts end up with their genes arrayed in such a way as to protect important genes by placing them in recombination dead zones.

And perhaps genes where lots of variation is tolerated or even helpful are placed in active recombination areas. In keeping with this, recent studies have shown that essential S. cerevisiae genes tend to be located in recombination cold spots, and that this arrangement is conserved in other yeasts.

It is too early to tell yet how pervasive this sort of gene placement is.  But if this turns out to be a good way to protect essential genes, Coca Cola should definitely have left the Coke formula in a part of its genome with little or no recombination. Mutating that set of instructions was as disastrous as mutating an essential gene!

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

Categories: Research Spotlight

Tags: evolution, meiosis, recombination, Saccharomyces cerevisiae

Not Lost Without Translation

January 22, 2015

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We all learned in biology class that nucleotides get added to an mRNA using a DNA template. And that amino acids get added to proteins using an mRNA template. But as with most everything in biology, there are exceptions.

For example, a long string of A’s gets added to mRNAs in eukaryotes without a DNA template of T’s. And now, in a new study published in Science, Shen and co-workers have shown that in certain cases amino acids can be added to proteins without translating an mRNA template.

Specifically, these authors showed that threonines and alanines that are not encoded by mRNA can be added to polypeptide chains stalled on the ribosome and that a key protein in this process is Rqc2p. It makes sense that Rqcp is on the spot to do this job, as this protein is part of the ribosome quality control (RQC) complex whose job it is to ubiquinate proteins stalled at the ribosome to target them for destruction.    

These added amino acids aren’t the result of some glitch of a misbehaving cell. They appear to be critical for the cells to mount a response to a situation where ribosomes are failing to complete normal translation.

Working with our favorite model organism, S. cerevisiae, the researchers started out to investigate the ribosome quality control (RQC) complex. This complex is like a cleanup crew for stalled ribosomes. If something goes wrong during translation and the ribosome stops elongating the nascent protein chain, the RQC complex steps in and tags the partially synthesized protein with ubiquitin, marking it for degradation.

The scientists were hoping to figure out how the RQC complex finds and recognizes stalled ribosomes. So they immunoprecipitated RQC and used cryo-electron microscopy to look at the structure of the complex bound to stalled ribosomes.

After a ribosome stops translating, it falls apart into its large and small subunits. Shen and colleagues found that one component of the RQC complex, Rqc2p, binds to the large (60S) ribosomal subunit after dissociation. They also found something unanticipated: tRNAs were present in the 60S ribosomal subunit at the A and P sites. This is where the tRNAs normally reside during translation, but translation obviously couldn’t be happening, since there was no mRNA present.

The presence of a tRNA at the ribosomal A site was especially surprising because tRNAs don’t bind there stably; they need mRNA and elongation factors to stabilize the interaction. It turned out that what was keeping the tRNAs on the ribosomal large subunit was Rqc2p, which bound to both of them and stabilized them. The researchers used a new thermostable reverse transcriptase and deep sequencing to find that the bound tRNAs were two specific alanine and threonine tRNA species. Why these specific tRNAs?

Pursuing this question, Shen and colleagues made another unexpected discovery: nascent polypeptide chains from stalled ribosomes were smaller in the rqc2 null mutant than in wild type.  This suggested that Rqc2p was adding something extra to the unfinished, stalled proteins.

Putting these observations together, the researchers formulated the hypothesis that Rqc2p mediates the addition of extra alanine and threonine residues to the C termini of proteins whose translation has been stalled. They created an ingenious set of reporter constructs to test this hypothesis.

The basic reporter contained the green fluorescent protein (GFP) gene fused to a coding sequence containing multiple “difficult” codons that would cause the ribosome to stall. The researchers found that most of the strains that were mutant for different subunits of the RQC complex contained a smear of variably sized protein products, the size of GFP and larger. But the rqc2 mutant only contained unmodified GFP; it failed to add anything. This confirmed the earlier suggestion that Rqc2p was responsible for adding the mysterious extra mass.

Next, they added a protease cleavage site at various locations in the reporter gene, and found that the extra mass was added at or downstream of the stalling sequence. In other words, it was added to the C terminus of the nascent polypeptide.

To be completely sure that translation was not involved, the researchers put stop codons in every frame after the stalling sequence. They had no effect, so the extra mass couldn’t be attributed to translation in any frame.

Finally, the scientists analyzed the C-terminal extensions by total amino acid analysis, Edman degradation, and mass spectrometry. They found that the extensions consisted of between 5 and 19 alanine and threonine residues, in no defined sequence. They named them Carboxy-terminal Ala and Thr extensions, or CAT tails.

This is all very cool, but do the tails actually do anything? Yes, it looks like they do!

When translation stalls, the cell responds to this stress using the transcription factor Hsf1p. By mutating three conserved residues in the Rqc2p NFACT nucleotide-binding domain, Shen and colleagues were able to generate a protein that could still recognize the plugged-up ribosome, but couldn’t add the alanines or threonines. It still worked fine to clean up stalled proteins, but the stalled proteins had no CAT tails. And sure enough, there was no Hsf1p-mediated heat shock response either.

So the CAT tails are part of the signal that tells the cell it had better start a stress response because things aren’t looking too good at the ribosome. It’s still not obvious exactly how the CAT tails participate in this process. But this isn’t some peculiarity of yeast: the genes are conserved, and mutations in homologs of some of the RQC genes and other genes involved in translation quality control cause neurodegeneration in mice.

The ribosome is one of the best-studied molecular machines, and you might have thought we already knew just about everything there was to know about it. This work reminds us that no matter how familiar something seems, there is always more to learn when we pay attention to unexpected results.

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

Categories: Research Spotlight

Tags: ribosome quality control, Saccharomyces cerevisiae, stress response

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