Research Area: Genetics

Making medicine runs in the family
Greta Friar | Whitehead Institute
May 5, 2020

What do the painkillers morphine and codeine, the cancer chemotherapy drug vinblastine, the popular brain health supplement salidroside, and a plethora of other important medicines have in common? They are all produced in plants through processes that rely on the same family of enzymes, the aromatic amino acid decarboxylases (AAADs). Plants, which have limited ability to physically react to their environments, have instead evolved to produce a stunning array of chemicals that allow them to do things like deter pests, attract pollinators, and adapt to changing environmental conditions. A lot of these molecules have also turned out to be useful in medicine—but it’s unusual for one family of enzymes to be responsible for so many different molecules of importance to both plants and humans. New research from Whitehead Institute Member Jing-Ke Weng, who is also an associate professor of biology at the Massachusetts Institute of Technology, and postdoctoral researcher Michael Torrens-Spence delves into the science behind the AAADs’ unusual generative capacity.

Plants create their useful molecules through biochemical pathways made up of chains of enzymes. Each enzyme acts as an assembly worker, taking in a molecule—starting with a basic building block like an amino acid—and performing biochemical modifications in sequence. The altered molecules get passed down the line until the last enzyme creates the final natural product. Once the pathway enzymes for a molecule of interest have been identified, researchers can copy their corresponding genes into organisms like yeast and bacteria that are capable of producing the molecules at scale more easily than the original plants. The AAAD family of enzymes function as gatekeepers to plants’ specialized molecule production because they operate at the beginning of many of the enzyme assembly lines; they take various amino acids, molecules that are widely available in nature, and direct them into different enzymatic pathways that produce unique molecules that only exist in plants. When an AAAD evolves to perform a new function, as has occurred frequently in their evolutionary history, this change high up in the assembly lines can cascade into the development of new biochemical pathways that create new natural products—leading to the diversity of medicines that stem from AAAD-gated pathways.

Due to the AAADs’ prominent role in the production of medically important molecules, Weng and Torrens-Spence decided to investigate how the AAADs came to be so prolific. In research published in the journal PNAS on May 5, the researchers illuminate the structural and functional underpinnings of the AAADs’ diversity. They also demonstrate how their detailed knowledge of the enzymes can be used to engineer novel enzymatic pathways to produce important molecules of interest from plants.

“We characterized these enzymes very thoroughly, which is a great starting place for manipulating the system and engineering it to do something new. That’s particularly exciting when you’re dealing with enzymes at the interface between primary and specialized plant metabolism; it can apply to a lot of downstream drugs,” Torrens-Spence says.

The AAAD family evolved from one ancestral enzyme into a diverse set of related enzymes over a relatively short period of time. This sort of diversification occurs when an enzyme gets accidentally duplicated, after which one copy has evolutionary pressure on it to maintain the same function, but the other copy suddenly has free range to evolve. If the superfluous enzyme mutates to do something new that is useful to the organism, from then on both enzymes, with their distinct roles, are likely to be maintained. In the case of the AAADs, this process occurred many times, leading to a large number of enzymes that appear almost exactly alike, yet can do very different things.

In order to explain the AAADs’ successful rate of diversification, the researchers took a close look at four enzymes in the AAAD family with different roles, and discovered the composition and three-dimensional shape—the crystal structure—of each. The crystal structure allowed the researchers to see how these molecular machines hold and modify specific molecules; this meant that they could understand why some AAADs initiate certain specialized-molecule production lines while other AAADs initiate alternative production lines. The researchers next used genetics and biochemistry to pinpoint the differences between the enzymes and how small genetic variations enact very major changes to the enzyme’s underlying machinery. This detailed analysis explained, among others things, how a subset of enzymes that evolved out of the AAADs, the aromatic acetaldehyde synthases (AASs), came to perform a completely different action on molecules while still being so similar to true AAADs that the two types of enzymes are often mistaken for each other.

After the researchers developed this thorough understanding of the AAAD family of enzymes, as well as knowledge of the AAAD-containing pathways that create useful medicinal molecules, they applied this knowledge by engineering an entirely new pathway to create a molecule of interest, (S)-norcoclaurine, a precursor molecule for morphine and other poppy-based painkillers. Torrens-Spence combined enzymes from pathways in different species to invent a novel chain of enzyme reactions that can produce (S)-norcoclaurine in fewer steps than is seen in nature. This experiment was a proof of concept that Torrens-Spence says shows the potential for such biosynthetic engineering, for example as a method to produce plant-based drugs more easily.

“Often with these molecules of interest, you figure out the pathway in plants and copy-paste it into a more scalable system, like yeast, that will produce larger quantities of the molecule,” Torrens-Spence says. “Here we’re applying engineering principles to biology, so that we can innovate and build something new.”

Written by Greta Friar

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Jing-Ke Weng’s primary affiliation is with Whitehead Institute for Biomedical Research, where his laboratory is located and all his research is conducted. He is also an associate professor of biology at Massachusetts Institute of Technology.

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Citation:

“Structural basis for divergent and convergent evolution of catalytic machineries in plant aromatic amino acid decarboxylase proteins”

PNAS, May 5, 2020

DOI: https://doi.org/10.1073/pnas.1920097117

Michael P. Torrens-Spence (1), Ying-Chih Chiang (2†), Tyler Smith (1,3), Maria A. Vicent (1,4), Yi Wang (2), and Jing-Ke Weng (1,3)

1 Whitehead Institute for Biomedical Research, Cambridge, Massachusetts 02142, USA.

2 Department of Physics, the Chinese University of Hong Kong, Shatin, N.T., Hong Kong.

3 Department of Biology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, USA.

4 Department of Biology, Williams College, Williamstown, Massachusetts 01267, USA.

† Present address: School of Chemistry, University of Southampton, Southampton, SO17 1BJ, UK.

Harnessing the moonseed plant’s chemical know-how
Eva Frederick | Whitehead Institute
April 20, 2020

In overgrown areas from Canada to China, a lush, woody vine with crescent-shaped seeds holds the secret to making a cancer-fighting chemical. Now, Whitehead Institute researchers in Member Jing-Ke Weng’s lab have discovered how the plants do it.

Plants in the family Menispermaceae, from the Greek words “mene” meaning “crescent moon,” and “sperma,” or seed, have been used in the past for a variety of medicinal purposes. Native Americans used the plants to treat skin diseases, and would ingest them as a laxative. Moonseed was also used as an ingredient in curare, a muscle relaxant used on the tips of poison arrows.

But the plants also may have a use in modern-day medicine: a compound called acutumine shown to have anti-cancer properties (although not tested specifically against cancer cells, the chemical has been shown to kill human T-cells, an important quality for leukemia and lymphoma treatments). Acutumine is a halogenated product, which means the molecule is capped on one end by a halogen atom — a group that includes fluorine, chlorine and iodine, among others. In this case, the halogen is chlorine.

Halogenated compounds like acutumine can be useful in medicinal chemistry — their unusual chemical appendages mean they react in interesting ways with other biomolecules, and drug designers can put them to use in creating compounds to complete specific tasks in the body. Today, 20% of pharmaceutical compounds are halogenated. “However, chemists’ ability to efficiently install halogen atoms to desirable positions of starting compounds has been quite limited,” Weng says.

Most natural halogenated products come from microorganisms such as algae or bacteria, and acutumine is one of the only halogenated products made by plants. Chemists finally succeeded in synthesizing the compound in 2009, although the reaction is time-consuming and expensive (10 mg of synthesized acutumine can cost around $2,000).

Colin Kim, a graduate student in the Weng lab at Whitehead Institute, wanted to know how these plants were completing this tricky reaction using only their own genetic material. “We thought, why don’t we ask how the plants make it and then upscale the reaction [to produce it more efficiently]?” Kim says.

“By understanding how living organisms such as the moonseed plant perform chemically challenging halogenation chemistry, we could devise new biochemical approaches to produce novel halogenated compounds for drug discovery,” Weng says.

Kim knew that for every halogenated molecule in an organism, there is an enzyme called a halogenase that catalyzes the reaction that sticks on that halogen. Halogenases are useful in creating pharmaceuticals – a well-placed halogen can help fine-tune the bioactivities of various drugs. So Weng, who is also an associate professor of biology at Massachusetts Institute of Technology, and Kim, who spearheaded the project, began working to identify the helper molecule responsible for creating acutumine in moonseed plants.

First, the scientists obtained three species of Menispermaceae plants. Two of them, common moonseed (Menispermum canadense) and Chinese moonseed (Sinomenium acutum), were known to produce acutumine. They also procured one plant in the same family called snake vine (Stephania japonica) which did not produce the compound.

They began their investigation by using mass spectrometry to look for acutumine in all three plants, and then find out exactly where in the plants it was located. They found the chemical all throughout the first two — and some extra in the roots of common moonseed. As expected, the third plant, snake vine, had none, and could therefore be used as a reference species, since presumably it would not ever express the gene for the halogenase enzyme that could stick on the chlorine molecule.

Next, the researchers started searching for the gene. They began by sequencing the RNA that was being expressed in the plants (RNA serves as a messenger between genomic DNA and functional proteins), and created a huge database of RNA sorted by what tissue it had been identified in.

At this point, the extra acutumine in the roots of common moonseed came in handy. The researchers had some idea of what the enzyme might look like – past research on other halogenases in bacteria suggested that one specific family of enzyme, called Fe(II)/2-oxoglutarate-dependent halogenases, or 2ODHs, for short, was capable of site-specifically adding a halogen in the same way that the moonseed’s mystery enzyme did. Although no 2ODHs had yet been found in plants, the researchers thought this lead was worth a look. So they searched specifically for transcripts similar to 2ODH sequences that were more highly expressed in the roots of common moonseed than in the leaves and stems.

After analyzing the RNA transcripts, Kim and Weng were pretty sure they had found what they were looking for: one gene in particular (which they named McDAH, short for M. canadense dechloroacutumine halogenase) was highly expressed in the roots of common moonseed. Then, in Chinese moonseed, they identified another protein that shared 99.1 percent of McDAH’s sequence, called SaDAH. No similar protein was found in snakevine, suggesting that this protein was likely the enzyme they wanted.

To be sure, the researchers tested the enzyme in the lab, and found that it was indeed the first-ever plant 2ODH, able to stick on the chlorine molecule to the alkaloid molecule dechloroacutumine to form acutumine. Interestingly, the enzyme was pretty picky; when they gave it other alkaloids like codeine and berberine to see if it would install a halogen on those as well, the enzyme ignored them, suggesting it was highly specific toward its preferred substrate, dechloroacutumine, the precursor of acutumine. They compared the enzyme’s activity to other similar enzymes, and found the key to its ability lay in the substitution of one specific amino acid in the active site– aspartic acid — for a glycine.

Now that they had identified the enzyme responsible for the moonseed’s halogenation reactions, Kim and Weng wanted to see what else it could do. A chemical capable of catalyzing such a complex reaction might be useful for chemists trying to synthesize other compounds, they hypothesized.

So they presented the enzyme with some dechloroacutumine and a whole buffet of alternative anions to see whether it might catalyze a reaction with any of these molecules in lieu of chlorine. Of the selection of anions, including bromide, azide, and nitrogen dioxide, the enzyme catalyzed a reaction only with azide, a construct of 3 nitrogen atoms.

“That is super cool, because there isn’t any other naturally occurring azidating enzyme that we know of,” Kim says. The enzyme could be used in click chemistry, a nature-inspired method to create a desired product through a series of simple, easy reactions.

In future studies, Weng and Kim hope to use what they’ve learned about the McDAH and SaDAH enzymes as a starting point to create enzymes that can be used as tools in drug development. They’re also interested in using the enzyme on other plant products to see what happens. “Plant natural products, even without chlorines, are pretty effective and bioactive, so it would be cool to see if you can take those plant natural products and then install chlorines to see what kind of changes and bioactivity it has, whether it develops new-to-nature functions or retain its original bioactivity with enhanced properties,” Kim says. “It expands the biocatalytic toolbox we have for natural product biosynthesis and its derivatization.”

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Written by Eva Frederick

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Citation: Kim, Colin Y. et al. The chloroalkaloid (−)-acutumine is biosynthesized via a Fe(II)- and 2-oxoglutarate-dependent halogenase in Menispermaceae plants. Nature Communications. April 20, 2020. DOI: 10.1038/s41467-020-15777-w

Exploring How Cells Repair and Tolerate DNA Damage
National Institute of Environmental Health Sciences
March 2, 2020

Graham Walker, Ph.D., studies the processes cells use to repair and tolerate DNA damage from environmental pollutants. For more than 40 years, he has worked to understand how cells respond to DNA damage, and how these processes can introduce mutations that lead to cancer and other human diseases.

His current NIEHS-funded work focuses on translesion synthesis (TLS). This damage tolerance process allows specialized enzymes that copy DNA, called TLS DNA polymerases, to replicate past lesions in damaged DNA. The process can help cells tolerate environmental DNA damage, but because TLS polymerases frequently insert the wrong DNA base, they can also lead to DNA mutations.

“The TLS process is critically important to human health because it helps cells survive DNA damage, but it can come at a cost,” said Walker. “It isn’t the kind of repair system you would think we would want because it makes a lot of mistakes. However, as we drill into these details, we are finding that there is so much more to be learned than just the strict biochemistry.”

In 2017, Walker was one of eight environmental health scientists to receive an inaugural Revolutionizing Innovative, Visionary Environmental Health Research (RIVER) Outstanding Investigator Award from NIEHS. The grant, which funds researchers rather than specific projects, provides Walker with flexibility to explore novel directions in his research.

From the Ames Test to TLS

Walker was drawn into the world of DNA repair and mutagenesis as a postdoctoral fellow at the University of California, Berkeley, under the guidance of Bruce Ames, Ph.D. Ames’ group created the Ames test, still used today, to determine whether a given chemical is likely to cause cancer. The Ames test uses bacterial strains that include a derivative of a naturally occurring drug-resistant plasmid, a small circular DNA molecule, known as pKM101. This molecule significantly increases the mutation rate of bacterial genes in response to chemical exposures, playing an important role in this quick and convenient test to estimate carcinogenic potential.

“I decided there must be something really interesting on that plasmid because it led to much higher mutation rates in bacteria for the same amount of damage,” said Walker.

After arriving at the Massachusetts Institute of Technology, his current employer, Walker continued to study the mechanisms behind these mutations.

Walker and his research team discovered the specific genes of pKM101 that are needed for it to produce more mutations. They showed that these genes are orthologs, or genes that evolved from a common ancestral gene, in the Escherichia coli (E. coli) chromosome that are required for the bacteria to mutate in response to DNA damage. This work helped lay the groundwork for the discovery of TLS DNA polymerases and how they are controlled.

“When we first sequenced these genes, nothing like them had been previously reported, but subsequently more and more related genes were discovered in all domains of life,” said Walker. “After decades of work by many labs, we now know that these are all TLS DNA polymerases and that the pKM101 plasmid encodes a polymerase that is responsible for the increased mutations.”

Using Bacteria to Understand DNA Damage

Walker’s prior research on the mutagenesis-enhancing function of pKM101 also led him to analyze E. coli’s SOS system, a set of biological responses that are activated to rescue cells from severe DNA damage. Walker and his team identified genes turned on by DNA damage that are regulated as part of E. coli’s SOS response. Many of the genes encode functions involved in DNA repair or mutagenesis. This work on the SOS response of E. coli was the first to directly demonstrate, in any organism, that DNA damage from environmental sources can change gene expression.

By further exploring TLS DNA polymerases in E. coli, he also identified the biological role of one of the most conserved DNA-damage response enzymes, DinB, which encodes a TLS DNA polymerase, and reported that the gene is required for resistance to some DNA-damaging agents. His work on DinB also suggested an additional mechanism by which antibiotics can become toxic to bacterial cells.

Blocking TLS in Cancer

“While a postdoc in the mid 1970’s with Bruce Ames, my ambitious hope was that by studying pKM101, I would learn something about the fundamental mechanism of how mutations arose in bacteria and humans, and might even learn how to control it,” said Walker. “That is now happening with my current, NIEHS-funded work.”

Some tumors can withstand damage from chemotherapy drugs by relying on TLS, which allows them to survive by replicating past damaged DNA caused by the drugs. In eukaryotes, including humans, mutagenic TLS is carried by two TLS DNA polymerases known as Rev1 and Pol zeta.

In addition to his innovative research, Walker is devoted to improving education and helping undergraduate students. In 2002, Walker became a Howard Hughes Medical Institute Professor and used his funding to establish a science education group modeled on his laboratory research group.

“I feel that training the next generations of scientists is as important as the science itself, and I have been incredibly lucky to have a spectacular set of grad students and post docs work with me over the years,” said Walker. “I have tried to focus as much on training, through teaching and mentoring, as on advancing the science.”

“Not only are these TLS polymerases responsible for introducing a lot of mutations that cause cancer, they also help cancer cells survive in the face of chemotherapy drugs that introduce DNA damage that would otherwise kill them,” said Walker.

Recently, Walker and his colleagues discovered that a small molecule and compound known as JH-RE-06 can block the Rev1-Pol zeta mutagenic TLS pathway by interfering with the ability of the Rev1 domain to recruit Pol zeta. The researchers tested the molecule in human cancer cell lines and showed that it enhanced the ability of several different types of chemotherapy to kill cancer cells, while also suppressing their ability to mutate in the presence of DNA-damaging drugs. In a mouse model of human melanoma, they found that not only did the tumors stop growing in mice treated with a combination of the chemotherapy drug cisplatin and JH-RE-06, those mice also survived longer.

“I am able to take more chances and try more high-risk experiments with the RIVER award,” said Walker. “The flexibility and extra resources are now allowing me to identify TLS inhibitors, which are offering startlingly unexpected mechanistic insights and also show potential to improve chemotherapy.”

Researchers discover an RNA-related function for a DNA repair enzyme
Raleigh McElvery
February 26, 2020

After decades of speculation, researchers have demonstrated that a classical DNA repair enzyme also binds to RNA, affecting blood cell development.

The DNA-dependent protein kinase, otherwise known as DNA-PK, is one of the most important enzymes that binds DNA and repairs double-stranded breaks. This mode of repair is essential for generating receptors that help the immune system fight off intruders. But DNA-PK doesn’t just bind DNA; it also binds RNA. Although researchers have known this for decades, they didn’t fully understand what kinds of RNAs DNA-PK bound in mammalian cells, or the physiological consequences of this binding.

In a new study published on February 26 in Nature, researchers from MIT and Columbia University have uncovered a mechanism whereby DNA-PK binds to the RNA involved in ribosome assembly. Ribosomes — the cell’s protein synthesis machinery — ensure that stem cells give rise to enough red blood cells. The researchers found that mutating DNA-PK prevents the ribosomes from being built properly, which prevents blood cells from doing their job and leads to blood disorders.

“This is the first biochemical evidence of DNA-PK assembly and activation by RNA inside cells,” says Eliezer Calo, a co-senior author and assistant professor in MIT’s Department of Biology. “We’re still trying to determine the mechanisms that regulate protein synthesis in stem cells, and this study reveals one of them.”

Co-senior author, Shan Zha from Columbia University, had previously studied DNA-PK’s role in DNA repair by generating a mouse model that carried enzymatically-dead versions of DNA-PK. While using this model to investigate tumorigenesis, Zha’s lab found these mutant mice developed a form of blood cancer known as myeloid disease. At the same time, another research group showed that mutations in DNA-PK also led to anemia, which occurs when the body does not have enough healthy red blood cells

Neither myeloid disease nor anemia could be easily explained by DNA repair defects alone. However, the two blood disorders did share some similarities to diseases caused by ribosome defects. Because DNA-PK resides in the same organelle where ribosomes are made, the Zha and Calo labs began to wonder whether DNA-PK could bind to the RNA there and control ribosome biogenesis.

In this new study, the Zha lab found that DNA-PK mutations impaired protein translation in red blood cell progenitors, which might contribute to anemia. In parallel, the Calo lab was investigating ribosomal RNA processing and was surprised to find that DNA-PK seemed to be implicated in ribosome assembly. The Calo lab then mapped all the RNAs in cells that bind DNA-PK. The enzyme unexpectedly attached to U3, a small RNA that helps assemble one of the subunits comprising the ribosome. Once it binds U3, DNA-PK can transfer a phosphate group to several specific sites on one of its own subunits. If DNA-PK is defective and cannot transfer the phosphate group, protein synthesis in blood stem cells is impaired, eventually causing anemia.

DNA-PK is essential for cellular viability in nearly all human cell lines, including cancer cell lines, while many other proteins involved in same DNA repair pathway are dispensable. Several studies, including one published by the Zha lab, showed that DNA-PK protein levels are 50-fold higher in common human cell lines than in rodent cell lines. The researchers do not yet know why the enzyme is so critical, but they suspect it might have to do with its ability to bind RNA. “We are interested in exploring whether this new role for DNA-PK could provide clues to this puzzle,” Zha says.

Calo says their findings could also have important implications for cancer treatment, because DNA-PK has emerged as a promising target for cancer therapy. Drugs that inhibit DNA-PK could prevent cancer cells from repairing their DNA and replicating successfully, but he warns these same remedies could also impact stem cell function. The next step is to explore DNA-PK’s other RNA binding targets and the related molecular pathways.

“We’ve demonstrated that DNA-PK has an entirely separate role that has nothing to do with DNA repair,” Calo says. “In the future, we’re excited to learn what additional RNA-related duties it may have beyond stem cell maintenance.”

Top Image: Ribosomes are assembled in the nucleoli (shown here in human cells).

Citation:
“DNA-PKcs has KU-dependent function in rRNA processing and haematopoiesis”
Nature, online February 26, 2020, DOI: 10.1038/s41586-020-2041-2
Zhengping Shao, Ryan A. Flynn, Jennifer L. Crowe, Yimeng Zhu, Jialiang Liang, Wenxia Jiang, Fardin Aryan, Patrick Aoude, Carolyn R. Bertozzi, Verna M. Estes, Brian J. Lee, Govind Bhagat, Shan Zha, and Eliezer Calo

To be long-lived or short-lived?
Nicole Davis | Whitehead
February 20, 2020

Genes are often imagined as binary actors: on or off. Yet such a simple view ignores the fact that genes’ activities, exerted by their corresponding proteins, can run the gamut from barely perceptible to off the charts. This rheostat-like range is due in part to molecular controls that determine how long the protein-making instructions for any given gene — known as messenger RNA (mRNA) — can persist before being destroyed.

Now, in a pair of papers published online in Molecular Cell, Whitehead Institute member David Bartel and his colleagues take a deep and systematic look at the dynamics of mRNA decay across thousands of genes. Their analysis — the most extensive to date — reveals surprising variability in the rate at which the ends (or “tails”) of mRNAs are shortened. In addition, the researchers uncover a link between this rate of shortening and how quickly the short-tailed mRNAs decay.

“Ultimately, these dynamics are responsible for determining how much mRNA is present for each gene, and that, of course, is really important for determining cell identity — for example, whether a cell is cancerous or a normal, healthy cell,” says Bartel, who is also a professor of biology at the Massachusetts Institute of Technology and an investigator at the Howard Hughes Medical Institute. “There is a thousand-fold difference in how long mRNAs stick around. That has a very profound effect on the amount of protein that gets made.”

TOWARDS A GLOBAL VIEW OF MRNA DEGRADATION

The anatomy of a typical mRNA consists of three key parts: a body, which contains the protein-making instructions; at one end, a string of repeating A’s known as the poly(A) tail; and at the other end, a protective biochemical cap.

Prior to the Molecular Cell studies, the future of a mRNA was known be linked to the length of its poly(A) tail — the longer the string of A’s, the longer the mRNA tends to persist. However, the speed that tails shorten as they age, and the rate at which mRNAs decay when their tails become short was known for just a handful of mRNAs.

To gain a more global picture, Bartel and his team, most recently led by graduate student Timothy Eisen, combined a set of techniques for high-throughput analyses of mRNA. These include a method for chemically modifying mRNAs as they are being made in order to distinguish newly synthesized mRNAs from those that are older, as well as sequencing-based approaches for measuring both the length of poly(A) tails and the amount of mRNA that was recently made. In addition, Eisen used computational methods to model the data they gathered and make predictions about them.

“All of the work in these papers involves time as an axis,” says Eisen. “The power of our approach is that it allowed us to plot and visualize how things change over time — and to infer for mRNAs from thousands of genes the rate at which the tail shortens and the subsequent rate at which the mRNA is destroyed.”

THE TAIL WAGS THE MRNA

By leveraging these techniques, Bartel, Eisen and their colleagues explored the mRNA dynamics for thousands of genes. One key observation is that mRNAs enter the cytoplasm with diverse poly(A) tail lengths. That variability encompasses not only the mRNAs from different genes but even those that correspond to the same gene.

“Previously, there wasn’t any reason to think there would be any differences, so people just assumed that the initial tail lengths would be the same,” says Bartel. “But it turns out there’s quite a bit of variability there.”

The Whitehead team also uncovered a striking amount of variation in the rate at which poly(A) tails are shortened. For some mRNAs, the tail shortens at a rate of about 30 nucleotides per minute. With an average tail length of around 200 nucleotides, that translates to the tail lasting just a few minutes. Other mRNAs have much more durable tails, with shortening rates of just a nucleotide or two an hour.

“That’s a thousand-fold difference,” says Eisen. Previously, researchers had shown that tail-shortening rates could vary, but they had observed only a 60-fold difference.

Bartel and his colleagues also found some striking differences among mRNAs once their poly(A) tails became short. “If we consider just those mRNA molecules that have tails of only 20 nucleotides, the ones that come from certain genes disappear much more rapidly than those coming from other genes — again spanning a thousand-fold range,” says Bartel.

That finding challenges long-held views about mRNA stability, as it had been generally assumed that short tails equaled short lives, and that all mRNAs whose tails had been shortened decay at the same rate. But it turns out that both processes are important: the rate at which mRNA tails are shortened (a process known as deadenylation), and the rate at which mRNAs decay after this shortening. Moreover, Bartel and his colleagues find that these two processes are coupled —  the more rapidly deadenylated mRNAs also degrade more rapidly once they have short tails.

“This coupling between rate of decay of short-tailed mRNAs and the rate of deadenylation is important because it prevents a large build-up of short-tailed versions of mRNAs that had undergone rapid deadenylation,” says Bartel. “Because these short-tailed versions do not build up, the thousand-fold difference that we observe in deadenylation rates can impart a thousand-fold difference in mRNA stabilities.”

SHINING A LIGHT ON MICRORNAS

MicroRNAs are small, regulatory RNA molecules that play critical roles in human biology. Their primary job is to recruit molecular machinery that shortens the poly(A) tails of mRNAs, thereby accelerating mRNA degradation, which reduces gene activity.

But strikingly, when Eisen and his colleagues harnessed their elegant system to examine microRNA activity, it appeared that these regulatory RNAs were leaving the tails of their targets completely unaltered — despite the fact that those mRNAs were being more rapidly degraded.

“That really left us scratching our heads wondering, ‘How could this be?’” adds Eisen. “It’s been known for quite some time that microRNAs operate by influencing poly(A) tail length.”

The team decided to look at the dynamics of this process, focusing on newly generated mRNAs. In this context, they observed that microRNAs accelerate both tail-shortening of target mRNAs and the subsequent decay of those mRNAs once their tails become short. “This second aspect of microRNA activity really hadn’t been appreciated before,” says Bartel. “But it’s a critical part of the story because it helps explain why we don’t see a build-up of short-tailed mRNAs.”

These findings, as well as the other results described here, significantly enhance what is known about mRNA decay and the factors that can influence it. With this expanded knowledge, Bartel and his colleagues, together with other research teams can work to uncover the molecular components and cellular contexts that cause mRNAs to have such drastically different lifetimes.

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Written by Nicole Davis

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Citations:

Eisen T, et al. The Dynamics of Cytoplasmic mRNA Metabolism. Molecular Cell. Published online January 2, 2020.

Eisen T, et al. MicroRNAs Cause Accelerated Decay of Short-Tailed Target mRNAs. Molecular Cell. Published online January 2, 2020.

Mary Gehring: Using flowering plants to explore epigenetic inheritance

Biologist’s studies illuminate a control system that influences how traits are passed along to new generations.

Anne Trafton | MIT News Office
December 16, 2019

Genes passed down from generation to generation play a significant role in determining the traits of every organism. In recent decades, scientists have discovered that another layer of control, known as epigenetics, is also critically important in shaping those characteristics.

Those added controls often work through chemical modifications of genes or other sections of DNA, which influence how easily those genes can be expressed by a cell. Many of those modifications are similar across species, allowing scientists to use plants as an experimental model to uncover how epigenetic processes work.

“Many of the epigenetic phenomena we know about were first discovered in plants, and in terms of understanding the molecular mechanisms, work on plants has also led the way,” says Mary Gehring, an associate professor of biology and a member of MIT’s Whitehead Institute for Biomedical Research.

Gehring’s studies of the small flowering plant Arabidopsis thaliana have revealed many of the mechanisms that underlie epigenetic control, shedding light on how these modifications can be passed from generation to generation.

“We’re trying to understand how epigenetic information is used during plant growth and development, and looking at the dynamics of epigenetic information through development within a single generation, between generations, and on an evolutionary timescale,” she says.

Seeds of discovery

Gehring, who grew up in a rural area of northern Michigan, became interested in plant biology as a student at Williams College, where she had followed her older sister. During her junior year at Williams, she took a class in plant growth and development and ended up working in the lab of the professor who taught the course. There, she studied how development of Arabidopsis is influenced by plant hormones called auxins.

After graduation, Gehring went to work for an environmental consulting company near Washington, but she soon decided that she wanted to go to graduate school to continue studying plant biology. She enrolled at the University of California at Berkeley, where she joined a lab that was studying how different genetic mutations affect the development of seeds.

That lab, led by Robert Fischer, was one of the first to discover an epigenetic phenomenon called gene imprinting in plants. Gene imprinting occurs when an organism expresses only the maternal or paternal version of particular gene. This phenomenon has been seen in flowering plants and mammals.

Gehring’s task was to try to figure out the mechanism behind this phenomenon, focusing on an Arabidopsis imprinted gene called MEDEA. She found that this type of imprinting is achieved by DNA demethylation, a process of removing chemical modifications from the maternal version of the gene, effectively turning it on.

After finishing her PhD in 2005, she worked as a postdoc at the Fred Hutchinson Cancer Research Center, in the lab of Steven Henikoff. There, she began doing larger, genome-scale studies in which she could examine epigenetic markers for many genes at once, instead of one at a time.

During that time, she began studying some of the topics she continues to investigate now, including regulation of the enzymes that control DNA methylation, as well as regulation of “transposable elements.” Also known as “jumping genes,” these sequences of DNA can change their position within the genome, sometimes to promote their own expression at the expense of the organism. Cells often use methylation to silence these genes if they generate harmful mutations.

Patterns of inheritance

After her postdoc, Gehring was drawn to MIT by “how passionate people are about what they’re working on, whether that’s biology or another subject.”

“Boston, especially MIT and Whitehead, is a great environment for science,” she says. “It seemed like there were a lot of opportunities to get really smart and talented students in the lab and have interesting colleagues to talk with.”

When Gehring joined the Whitehead Institute in 2010, she was the only plant biologist on the faculty, but she has since been joined by Associate Professor Jing-Ke Weng.

Her lab now focuses primarily on questions such as how maternal and paternal parents contribute to reproduction, and how their differing interests can lead to genetic conflicts. Gene imprinting is one way that this conflict is played out. Gehring has also discovered that small noncoding RNA molecules play an important role in imprinting and other aspects of inheritance by directing epigenetic modifications such as DNA methylation.

“One thing we’ve found is that this noncoding RNA pathway seems to control the transcriptional dosage of seeds, that is, how many of the transcripts are from the maternally inherited genome and how many from the paternally inherited genome. Not just for imprinted genes, but also more broadly for genes that aren’t imprinted,” Gehring says.

She has also identified a genetic circuit that controls an enzyme that is required to help patterns of DNA methylation get passed from parent to offspring. When this circuit is disrupted, the methylation state changes and unusual traits can appear. In one case, she found that the plants’ leaves become curled after a few generations of disrupted methylation.

“You need this genetic circuit in order to maintain stable methylation patterns. If you don’t, then what you start to see is that the plants develop some phenotypes that get worse over generational time,” she says.

Many of the epigenetic phenomena that Gehring studies in plants are similar to those seen in animals, including humans. Because of those similarities, plant biology has made significant contributions to scientists’ understanding of epigenetics. The phenomenon of epigenomic imprinting was first discovered in plants, in the 1970s, and many other epigenetic phenomena first seen in plants have also been found in mammals, although the molecular details often vary.

“There are a lot of similarities among epigenetic control in flowering plants and mammals, and fungi as well,” Gehring says. “Some of the pathways are plant-specific, like the noncoding RNA pathway that we study, where small noncoding RNAs direct DNA methylation, but small RNAs directing silencing via chromatin is something that happens in many other systems as well.”

A new way to regulate gene expression

Biologists uncover an evolutionary trick to control gene expression that reverses the flow of genetic information from RNA splicing back to transcription.

Raleigh McElvery | Department of Biology
December 9, 2019

Sometimes, unexpected research results are simply due to experimental error. Other times, it’s the opposite — the scientists have uncovered a new phenomenon that reveals an even more accurate portrayal of our bodies and our universe, overturning well-established assumptions. Indeed, many great biological discoveries are made when results defy expectation.

A few years ago, researchers in the Burge lab were comparing the genomic evolution of several different mammals when they noticed a strange pattern. Whenever a new nucleotide sequence appeared in the RNA of one lineage, there was generally an increase in the total amount of RNA produced from the gene in that lineage. Now, in a new paper, the Burge lab finally has an explanation, which redefines our understanding of how genes are expressed.

Once DNA is transcribed into RNA, the RNA transcript must be processed before it can be translated into proteins or go on to serve other roles within the cell. One important component of this processing is splicing, during which certain nucleotide sequences (introns) are removed from the newlymade RNA transcript, while others (the exons) remain. Depending on how the RNA is spliced, a single gene can give rise to a diverse array of transcripts.

Given this order of operations, it makes sense that transcription affects splicing. After all, splicing cannot occur without an RNA transcript. But the inverse theory — that splicing can affect transcription — is now gaining traction. In a recent study, the Burge lab showed that splicing in an exon near the beginning of a gene impacts transcription and increases gene expression, offering an explanation for the patterns in their previous findings.

“Rather than Step A impacting Step B, what we found here is that Step B, splicing, actually feeds back to influence Step A, transcription,” says Christopher Burge, senior author and professor of biology. “It seems contradictory, since splicing requires transcription, but there is actually no contradiction if — as in our model — the splicing of one transcript from a gene influences the transcription of subsequent transcripts from the same gene.”

The study, published on Nov. 28 in Cell, was led by Burge lab postdoc Ana Fiszbein.

Promoting gene expression

In order for transcription to begin, molecular machines must be recruited to a special sequence of DNA, known as the promoter. Some promoters are better at recruiting this machinery than others, and therefore initiate transcription more often. However, having different promoters available to produce slightly different transcripts from a gene helps boost expression and generates transcript diversity, even before splicing occurs mere seconds or minutes later. ​

At first, Fiszbein wasn’t sure how the new exons were enhancing gene expression, but she theorized that new promoters were involved. Based on evolutionary data available and her experiments at the lab bench, she could see that wherever there was a new exon, there was usually a new promoter nearby. When the exon was spliced in, the new promoter became more active.

The researchers named this phenomenon “exon-mediated activation of transcription starts” (EMATS). They propose a model in which the splicing machinery associated with the new exon recruits transcription machinery to the vicinity, activating transcription from nearby promoters. This process, the researchers predict, likely helps to regulate thousands of mammalian genes across species.

A more flexible genome

Fiszbein believes that EMATS has increased genome complexity over the course of evolution, and may have contributed to species-specific differences. For instance, the mouse and rat genomes are quite similar, but EMATS could have helped produce new promoters, leading to regulatory changes that drive differences in structure and function between the two. EMATS may also contribute to differences in expression between tissues in the same organism.

“EMATS adds a new layer of complexity to gene expression regulation,” Fiszbein says. “It gives the genome more flexibility, and introduces the potential to alter the amount of RNA produced.”

Juan Valcárcel, a research professor at the Catalan Institution for Research and Advanced Studies in the Center for Genomic Regulation in Barcelona, Spain, says understanding the mechanisms behind EMATS could also have biotechnological and therapeutic implications. “A number of human conditions, including genetic diseases and cancer, are caused by a defect or an excess of particular genes,” he says. “Reverting these anomalies through modulation of EMATS might provide innovative therapies.”

Researchers have already begun to tinker with splicing to control transcription. According to Burge, pharmaceutical companies like Ionis, Novartis, and Roche are concocting drugs to regulate splicing and treat diseases like spinal muscular atrophy. There are many ways to decrease gene expression, but it’s much harder to increase it in a targeted manner. “Tweaking splicing might be one way to do that,” he says.

“We found a way in which our cells change gene expression,” Fiszbein adds. “And we can use that to manipulate transcript levels as we want. I think that’s the most exciting part.”

This research was funded by the National Institutes of Health and the Pew Latin American Fellows Program in the Biomedical Sciences.

The surprising individuality of miRNAs
Greta Friar | Whitehead Institute
December 5, 2019

In order for the instructions contained within a gene to ultimately execute some function in the body, the nucleotides, or letters, that make up the gene’s DNA sequence must be “read” and used to produce a messenger RNA (mRNA). This mRNA must then be translated into a functional protein. A number of different pathways within the cell influence this essential biological process, informing whether, when, and to what extent a gene is expressed. A major class of such regulators are microRNAs (miRNAs). These minute RNAs—they are, on average, 22 nucleotides long—join with a protein called Argonaute to cause certain mRNAs to be degraded, which in turn decreases the amount of translation of those mRNAs into their functional protein forms. Scientists have identified hundreds of miRNAs that are common amongst mammals and other vertebrate animals, and most mammalian mRNAs are targeted by at least one of these miRNAs—an indication of their pervasive importance to our biology. Accurately predicting how any particular miRNA will affect gene expression in a cell is important for understanding our own biology, and might facilitate the design of therapeutic drugs that affect or utilize miRNAs, but the complexity of the miRNA pathway makes this sort of prediction difficult.

The success rate with which a miRNA is able to repress a specific gene (by degrading its mRNA) is called its targeting efficacy, and researchers have used a variety of models to calculate it, with mixed results. In the past, researchers have treated miRNAs as a group and looked at average behavior in order to make predictions, because there simply wasn’t enough data specific to individual miRNAs available to do otherwise. However, Whitehead Institute Member David Bartel, who is also a professor of biology at the Massachusetts Institute of Technology and a Howard Hughes Medical Institute investigator, graduate student Sean McGeary, and former graduate student Kathy Lin collected a massive amount of data on six miRNAs, and from that foundation developed an improved predictive model for all individual miRNAs. Their findings, published online in Science on December 5, provide unprecedented accuracy and granularity in miRNA targeting prediction.

“We used to focus our attention on microRNA targeting patterns that were consistent, because that consistency gave us confidence in what we were seeing,” Bartel says, “but with the robust results of this research, we can now pay attention to differences between individual miRNAs.”

Bartel and the Whitehead Institute Bioinformatics and Research Computing group operate one of the go-to resources for prediction of miRNAs’ targets and target efficacy, known as TargetScan. This latest research will be used to update TargetScan, giving scientists around the world an even more useful reference tool for research involving miRNA-mediated regulation of gene expression.

To understand miRNA targeting, researchers need to identify the particular sites within an mRNA sequence where the miRNA can bind, and they additionally need to know how strong the interaction will be at each site—the binding affinity. In general, a miRNA will bind to an mRNA when there is a match between at least six of the first eight nucleotides of the miRNA and a complementary sequence of nucleotides somewhere on the mRNA. The two sequences are like rows of puzzle pieces being pushed together: if each puzzle piece slots into the corresponding piece, the rows combine into one locked puzzle—the miRNA binds its target. If the pieces don’t fit together, the rows can’t connect. These sorts of binding sites, perfect matches within the first eight nucleotides of the miRNA, are called canonical site types, and researchers used to think that there was a clear hierarchy between them, with each individual site type conferring a similar amount of repression regardless of the miRNA identity. But that’s not what McGeary observed.

McGeary looked at six miRNAs and developed a method to measure, for each miRNA, relative binding affinities to a massive collection of RNA sequences.

“I performed experiments that provide vast numbers of measurements, which collectively inform us on how well a miRNA will bind to an mRNA,” McGeary says.

These measurements, as well as further calculations that McGeary made from them, formed a novel, rich pool of data with which to improve miRNA targeting prediction. From their experiments, the researchers found that the expected targeting hierarchy of canonical sites did not apply to all miRNAs. An individual miRNA might actually have a stronger affinity to one of the canonical sites lower in the expected hierarchy than another. Furthermore, the group discovered that the miRNAs each had unique noncanonical binding sites, some of which were sites that contained at least one mismatch but were still able to bind miRNA. The researchers found many instances in which a miRNA bound more strongly to one of its noncanonical sites than to some of its canonical sites, despite the imperfect or unusual pairing of the noncanonical sites.

“As humans, we like to classify things into discrete buckets with discrete characteristics,” Lin says. “But to build a model that is quantitative, you have to recognize that each miRNA and target interaction is different.”

Factors in a target site’s environment contribute to the individuality of target interactions, as they can affect the structural accessibility of the site for binding. In particular, the researchers found that the four nucleotides closest to a target site could have a huge, even 100-fold combined impact on affinity.

With their high-resolution data, the researchers were able to rigorously verify a supposition within the miRNA research community: that the strength with which a miRNA binds to a target site is the major determinant for how effective that miRNA will be at degrading that mRNA. This striking correlation between site affinity and targeting efficacy also allowed them to create a biochemical model of miRNA targeting that used the vast collection of affinity measurements to predict the efficacy of repression of every mRNA in cell, significantly out-performing all existing models of miRNA targeting. They then used machine learning, in the form of a convolutional neural network developed by Lin, to extend the improved predictions to all miRNAs without the need to generate additional data.

Altogether, these findings paint a much richer picture of miRNA-mediated gene repression. The new level of specificity in miRNA targeting prediction will provide all researchers working on the subject with better information about the impact of a given miRNA in a cell.

This work was supported by the NIH and Howard Hughes Medical Institute.

Written by Greta Friar

***

David Bartel’s primary affiliation is with Whitehead Institute for Biomedical Research, where his laboratory is located and all his research is conducted. He is also a professor of biology at Massachusetts Institute of Technology and investigator with the Howard Hughes Medical Institute.

***

Citation:

“The biochemical basis of microRNA targeting efficacy”

Science, online December 5, 2019, DOI: 10.1126/science.aav1741

Sean E. McGeary (1,2,3†), Kathy S. Lin (1,2,3,4†), Charlie Y. Shi (1,2,3), Thy Pham (1,2,3), Namita Bisaria (1,2,3), Gina M. Kelley (1,2,3), and David P. Bartel (1,2,3,4)

  1. Howard Hughes Medical Institute, Cambridge, MA, 02142, USA
  2. Whitehead Institute for Biomedical Research, Cambridge, MA, 02142, USA
  3. Department of Biology, Massachusetts Institute of Technology, Cambridge, MA 02139, USA
  4. Computational and Systems Biology Program, Massachusetts Institute of Technology, Cambridge, MA, 02139, USA

†These authors contributed equally to this work.

MicroRNAs work together to tune gene expression in the brain
Raleigh McElvery
November 4, 2019

A new study from the MIT Department of Biology suggests we may need to re-think how certain RNAs operate to impact development and disease.

According to the “central dogma” of biology, DNA is converted into messenger RNA (mRNA) before being expressed as a protein. However, not all RNAs are destined to become proteins. MicroRNAs (miRNAs) are small, non-coding RNAs, which regulate a variety of cellular processes by binding to mRNAs and destabilizing them to reduce their expression.

A single miRNA can target hundreds of different mRNAs. And yet, on its own, an individual miRNA only represses the expression of each mRNA target by about 10-20%. Given that the effects of a single miRNA are so mild, researchers couldn’t understand how they could exert such powerful control over so many processes. One theory is that, rather than acting alone, perhaps multiple miRNAs bind to the same target mRNA in concert to exert enhanced repression. However, few studies have explored this idea in-depth, or identified examples of such co-regulation.

In a new study published in Genome Research on October 24, MIT biologists were able to pinpoint specific miRNAs that collaborate with one another to repress mRNA expression in the brain — adding credence to the notion that miRNAs often collaborate with one another.

“The idea that miRNAs may work by co-targeting sets of transcripts together has been around for a while,” says Jennifer Cherone, the study’s lead author. “But it’s only recently that certain key advances — like better annotations of where transcripts end and more accurate predictions of miRNA target sites — have allowed us to uncover these relationships and rigorously test them in the lab.”

Using powerful computational analyses to compare target sets of different miRNAs, Cherone was able to identify hundreds of distinct miRNAs, which — despite their sequence differences — bound many of the same mRNAs. Of all the tissues she examined, the brain appeared to have the most co-targeting. So she narrowed her focus to explore the overlapping functions of just two miRNAs that worked together there: miR-138 and miR-137.

“That was a really interesting observation and a functional demonstration of the overlap between these two miRNAs,” she says. “One miRNA can rescue the loss of a completely different miRNA if they share targets.”If she deleted miR-138 from her cells, they could no longer differentiate and become neurons. However, when she added miR-138’s co-targeting partner, miR-137, the cells were once again able to differentiate.

Cherone went on to identify an entire group of miRNAs within the brain, nine in total, that also shared similar targets. She selected several genes targeted by three or more of these miRNAs, and mutated every possible combination of the miRNA sites to determine their individual contributions. She ultimately established that subsets of the miRNAs could repress gene expression between five- and tenfold if they were expressed at the same time and bound close together.

According to Cherone, “seeing a tenfold repression by miRNAs is unheard of.” Such strong repression can have serious phenotypic consequences. She attributes this finding to the lab’s advanced computational strategies, which allowed them to systematically and unbiasedly identify the miRNAs that work together and their gene targets.

Why might a single gene be regulated by so many different miRNAs? There are more evolutionary paths to acquire sites for many different miRNAs than paths to acquire sites for the same miRNA. And, the authors explain, this arrangement may allow more precise control of cell type-specific expression.

Given that their miRNAs of interest primarily worked in the brain, the researchers wondered why this tissue might require so much co-targeting. One idea is that mRNAs in the brain tend to have longer regions where more miRNAs can bind to exert their effects. Another possibility is that mRNA expression in the brain must be especially fine-tuned, because too much or too little expression could have severe ramifications for neuronal function and development. For instance, fragile X-associated tremor/ataxia syndrome (FXTAS) can result from fairly subtle changes in proteins levels.

“Co-targeting appears to be widespread in many tissues, not just the brain,” says senior author Christopher Burge, a professor of biology at MIT. “This means that strategies to modulate the activity of a miRNA in a genetic or therapeutic context will be most effective when they take into account the levels of the other miRNAs that frequently partner with the miRNA of interest.”

“It’s time to start thinking of miRNAs as working together in networks, rather than functioning as individual units,” Cherone says. “If you want to know the function of a given miRNA, you have to understand the group it’s collaborating with, and explore its function within that group.”

Top image: Graphical illustration of co-targeting by miRNAs. Credit: Jennifer Cherone.

Citation:
“Cotargeting among microRNAs in the brain.”
Genome Research, online October 24, 2019, DOI: 10.1101/gr.249201.119
Jennifer M. Cherone, Vjola Jorgji, and Christopher B. Burge.