Research Area: Cell Biology

New gene regulation model provides insight into brain development

A well-known protein family binds to many more RNA sequences than previously thought to help neurons grow.

Raleigh McElvery
August 17, 2020

In every cell, RNA-binding proteins (RBPs) help tune gene expression and control biological processes by binding to RNA sequences. Researchers often assume that individual RBPs latch tightly to just one RNA sequence. For instance, an essential family of RBPs, the Rbfox family, was thought to bind one particular RNA sequence alone. However, it’s becoming increasingly clear that this idea greatly oversimplifies Rbfox’s vital role in development.

Members of the Rbfox family are among the best-studied RBPs and have been implicated in mammalian brain, heart, and muscle development since their discovery 25 years ago. They influence how RNA transcripts are “spliced” together to form a final RNA product, and have been associated with disorders like autism and epilepsy. But this family of RBPs is compelling for another reason as well: until recently, it was considered a classic example of predictable binding.

More often than not, it seemed, Rbfox proteins bound to a very specific sequence, or motif, of nucleotide bases, “GCAUG.” Occasionally, binding analyses hinted that Rbfox proteins might attach to other RNA sequences as well, but these findings were usually discarded. Now, a team of biologists from MIT has found that Rbfox proteins actually bind less tightly — but no less frequently — to a handful of other RNA nucleotide sequences besides GCAUG. These so-called “secondary motifs” could be key to normal brain development, and help neurons grow and assume specific roles.

“Previously, possible binding of Rbfox proteins to atypical sites had been largely ignored,” says Christopher Burge, professor of biology and the study’s senior author. “But we’ve helped demonstrate that these secondary motifs form their own separate class of binding sites with important physiological functions.”

Graduate student Bridget Begg is the first author of the study, published on August 17 in Nature Structural & Molecular Biology.

“Two-wave” regulation

After the discovery that GCAUG was the primary RNA binding site for mammalian Rbfox proteins, researchers characterized its binding in living cells using a technique called CLIP (crosslinking-immunoprecipitation). However, CLIP has several limitations. For example, it can indicate where a protein is bound, but not how much protein is bound there. It’s also hampered by some technical biases, including substantial false-negative and false-positive results.

To address these shortcomings, the Burge lab developed two complementary techniques to better quantify protein binding, this time in a test tube: RBNS (RNA Bind-n-Seq), and later, nsRBNS (RNA Bind-n-Seq with natural sequences), both of which incubate an RBP of interest with a synthetic RNA library. First author Begg performed nsRBNS with naturally-occurring mammalian RNA sequences, and identified a variety of intermediate-affinity secondary motifs that were bound in the absence of GCAUG. She then compared her own data with publicly-available CLIP results to examine the “aberrant” binding that had often been discarded, demonstrating that signals for these motifs existed across many CLIP datasets.

To probe the biological role of these motifs, Begg performed reporter assays to show that the motifs could regulate Rbfox’s RNA splicing behavior. Subsequently, computational analyses by Begg and co-author Marvin Jens using mouse neuronal data established a handful of secondary motifs that appeared to be involved in neuronal differentiation and cellular diversification.

Based on analyses of these key secondary motifs, Begg and colleagues devised a “two-wave” model. Early in development, they believe, Rbfox proteins bind predominantly to high-affinity RNA sequences like GCAUG, in order to tune gene expression. Later on, as the Rbfox concentration increases, those primary motifs become fully occupied and Rbfox additionally binds to the secondary motifs. This results in a second wave of Rbfox-regulated RNA splicing with a different set of genes.

Begg theorizes that the first wave of Rbfox proteins binds GCAUG sequences early in development, and she showed that they regulate genes involved in nerve growth, like cytoskeleton and membrane organization. The second wave appears to help neurons establish electrical and chemical signaling. In other cases, secondary motifs might help neurons specialize into different subtypes with different jobs.

John Conboy, a molecular biologist at Lawrence Berkeley National Lab and an expert in Rbfox binding, says the Burge lab’s two-wave model clearly shows how a single RBP can bind different RNA sequences — regulating splicing of distinct gene sets and influencing key processes during brain development. “This quantitative analysis of RNA-protein interactions, in a field that is often semi-quantitative at best, contributes fascinating new insights into the role of RNA splicing in cell type specification,” he says.

A binding spectrum

The researchers suspect that this two-wave model is not unique to Rbfox. “This is probably happening with many different RBPs that regulate development and other dynamic processes,” Burge says. “In the future, considering secondary motifs will help us to better understand developmental disorders and diseases, which can occur when RBPs are over- or under-expressed.”

Begg adds that secondary motifs should be incorporated into computer models that predict gene expression, in order to probe cellular behavior. “I think it’s very exciting that these more finely-tuned developmental processes, like neuronal differentiation, could be regulated by secondary motifs,” she says.

Both Begg and Burge agree it’s time to consider the entire spectrum of Rbfox binding, which are highly influenced by factors like protein concentration, binding strength, and timing. According to Begg, “Rbfox regulation is actually more complex than we sometimes give it credit for.”

Citation:
“Concentration-dependent splicing is enabled by Rbfox motifs of intermediate affinity”
Nature Structural & Molecular Biology, online August 17, 2020, DOI: 10.1038/s41594-020-0475-8
Bridget E. Begg, Marvin Jens, Peter Y. Wang, Christine M. Minor, and Christopher B. Burge

Top illustration: Some RNA-binding proteins like Rbfox (gold ellipses) help tune gene expression and control biological processes by latching onto more RNA sequences (black and gold lines) as their concentration increases (teal shading). Credit: Bridget Begg
Posted: 8.17.20
This molecule helps sweet-toothed protein complex sense sugar
Eva Frederick | Whitehead Institute
July 28, 2020

In order to grow and thrive, cells need sugar. A repertoire of cellular mechanisms turn unwieldy molecules of glucose and fructose into versatile building blocks for making useful molecules such as lipids, and energy to fuel necessary processes in the cell. But for any of these things to happen, the cells need to sense when sugars are present in the first place — and scientists are still unraveling how they do it.

Now, in a new paper online July 27 in Nature Metabolism, researchers in the lab of Whitehead Institute Member David Sabatini, identify a key molecule that signals to the cell’s growth-triggering complex mTORC1 when there is sugar to be had, leading to a metabolic response. “This discovery puts us another step closer to understanding the biology of mTORC1 and its effects on cellular growth and metabolism,” said Sabatini, who is also a professor of biology at Massachusetts Institute of Technology and investigator with the Howard Hughes Medical Institute.

mTORC1 — short for “mechanistic target of rapamycin complex 1” — is a complex of proteins involved in regulating cell growth and metabolism. Jose Orozco, a fifth-year M.D./Ph.D student in Sabatini’s lab, describes mTORC1 as a sort of cellular licensing board. In order for other parts of the cell to grow and create new products, they must first be “approved” by mTORC1. If there are enough building blocks in the cell to create a certain product, mTORC1 will add a phosphate group to the appropriate “builders” — a signal that allows the building to begin.

“The builders in this case are metabolic pathways responsible for the creation of proteins, the regulation of nucleotides, regulation of glycolysis, regulation of fatty acid synthesis,” he says. “None of these builders can sense everything. But mTORC1 can, and it makes this sort of unified decision for the cell, ‘Yes, we have everything we need to grow.’”

One essential component for cellular building is glucose. That means mTORC1 has a sweet tooth by necessity: the complex is only active when there is enough glucose in the cell. When there’s glucose to go around, mTORC1 is “on” and binds to a lysosome, a structure that serves as the cell’s “digestive system”, where it perches to perform its phosphorylation duties. When a cell is starved for glucose, the complex falls off the lysosome, inactive.

Since the early 2010s, scientists have known one way that mTOR proteins sense glucose: when there is no glucose available, the cell inhibits the action of mTORC1 through a pathway involving the protein AMPK. But another study suggested that even without AMPK, mTORC1 can still sense an absence of glucose. “I think a lot of people had written it off as ‘Oh, [the signal] must just be AMPK,’” Orozco says. “But when we tested that hypothesis, we showed that even cells that didn’t have any AMPK were still able to sense glucose availability. That was the observation that started this project.”

To find the mysterious second sugar-sensing process, Orozco and colleagues created cells in which the known signalling protein AMPK was out of the picture. Using these modified cells, they began looking for specific traits of the glucose molecule that might be triggering the response. The team found that sugars that could be broken down by the cell, such as mannose, glucosamine and fructose, were able to activate mTORC1. Non-metabolizable sugars had no effect.

This suggested that the signaling molecule was not glucose itself, but something produced when glucose is taken apart during glycolysis — the biochemical process that breaks down the sugar into usable building blocks. With this in mind, the researchers next combed step by step through glycolysis products to see which ones could be the signal molecule.

The team identified a step of glycolysis that seemed to be key, zeroing  in on a glycolysis product called dihydroxyacetone phosphate, or DHAP. Even in the complete absence of glucose, the researchers could turn on mTORC1 by adding DHAP.

It is difficult to prove exactly why the cell relies on DHAP as a signal, but Orozco has some ideas. For one thing, DHAP later goes on to serve as the backbone of lipids, which are built by a process controlled by mTORC1 — so it would make sense that mTORC1 would respond to its presence or absence. Also, DHAP levels are extremely sensitive to changes in the amount of cellular glucose, more so than any other glycolysis intermediary. Also, DHAP is a product of both glucose and fructose, which are both important sugars in the human diet.

In the future, the team hopes to understand more. “We don’t know the biochemical details of how DHAP [conveys its message],” Orozco says. “We don’t know the sensor, we don’t know what proteins bind it, and we don’t know if that causes conformational changes in [associated proteins]. That we sort of leave as the enticing next question that we want to tackle.”

At the moment, studying the glucose sensing pathway is purely foundational research. But while there are no clear applications yet, surprises could lurk just around the corner. “Targeting nutrient sensing in mTOR has shown some promise in, of all things, regulating depression and mood,” Orozco says. “That’s interesting, and we don’t really understand why that is the case. How is glucose targeting going to be important? We don’t know yet. But we think it has a lot of potential.”

***

Written by Eva Frederick

Citation:

Orozco, J.M., Krawczyk, P.A., Scaria, S.M. et al. Dihydroxyacetone phosphate signals glucose availability to mTORC1. Nat Metab (2020). https://doi.org/10.1038/s42255-020-0250-5

Bringing RNA into genomics

ENCODE consortium identifies RNA sequences that are involved in regulating gene expression.

Anne Trafton | MIT News Office
July 29, 2020

The human genome contains about 20,000 protein-coding genes, but the coding parts of our genes account for only about 2 percent of the entire genome. For the past two decades, scientists have been trying to find out what the other 98 percent is doing.

A research consortium known as ENCODE (Encyclopedia of DNA Elements) has made significant progress toward that goal, identifying many genome locations that bind to regulatory proteins, helping to control which genes get turned on or off. In a new study that is also part of ENCODE, researchers have now identified many additional sites that code for RNA molecules that are likely to influence gene expression.

These RNA sequences do not get translated into proteins, but act in a variety of ways to control how much protein is made from protein-coding genes. The research team, which includes scientists from MIT and several other institutions, made use of RNA-binding proteins to help them locate and assign possible functions to tens of thousands of sequences of the genome.

“This is the first large-scale functional genomic analysis of RNA-binding proteins with multiple different techniques,” says Christopher Burge, an MIT professor of biology. “With the technologies for studying RNA-binding proteins now approaching the level of those that have been available for studying DNA-binding proteins, we hope to bring RNA function more fully into the genomic world.”

Burge is one of the senior authors of the study, along with Xiang-Dong Fu and Gene Yeo of the University of California at San Diego, Eric Lecuyer of the University of Montreal, and Brenton Graveley of UConn Health.

The lead authors of the study, which appears today in Nature, are Peter Freese, a recent MIT PhD recipient in Computational and Systems Biology; Eric Van Nostrand, Gabriel Pratt, and Rui Xiao of UCSD; Xiaofeng Wang of the University of Montreal; and Xintao Wei of UConn Health.

RNA regulation

Much of the ENCODE project has thus far relied on detecting regulatory sequences of DNA using a technique called ChIP-seq. This technique allows researchers to identify DNA sites that are bound to DNA-binding proteins such as transcription factors, helping to determine the functions of those DNA sequences.

However, Burge points out, this technique won’t detect genomic elements that must be copied into RNA before getting involved in gene regulation. Instead, the RNA team relied on a technique known as eCLIP, which uses ultraviolet light to cross-link RNA molecules with RNA-binding proteins (RBPs) inside cells. Researchers then isolate specific RBPs using antibodies and sequence the RNAs they were bound to.

RBPs have many different functions — some are splicing factors, which help to cut out sections of protein-coding messenger RNA, while others terminate transcription, enhance protein translation, break down RNA after translation, or guide RNA to a specific location in the cell. Determining the RNA sequences that are bound to RBPs can help to reveal information about the function of those RNA molecules.

“RBP binding sites are candidate functional elements in the transcriptome,” Burge says. “However, not all sites of binding have a function, so then you need to complement that with other types of assays to assess function.”

The researchers performed eCLIP on about 150 RBPs and integrated those results with data from another set of experiments in which they knocked down the expression of about 260 RBPs, one at a time, in human cells. They then measured the effects of this knockdown on the RNA molecules that interact with the protein.

Using a technique developed by Burge’s lab, the researchers were also able to narrow down more precisely where the RBPs bind to RNA. This technique, known as RNA Bind-N-Seq, reveals very short sequences, sometimes containing structural motifs such as bulges or hairpins, that RBPs bind to.

Overall, the researchers were able to study about 350 of the 1,500 known human RBPs, using one or more of these techniques per protein. RNA splicing factors often have different activity depending on where they bind in a transcript, for example activating splicing when they bind at one end of an intron and repressing it when they bind the other end. Combining the data from these techniques allowed the researchers to produce an “atlas” of maps describing how each RBP’s activity depends on its binding location.

“Why they activate in one location and repress when they bind to another location is a longstanding puzzle,” Burge says. “But having this set of maps may help researchers to figure out what protein features are associated with each pattern of activity.”

Additionally, Lecuyer’s group at the University of Montreal used green fluorescent protein to tag more than 300 RBPs and pinpoint their locations within cells, such as the nucleus, the cytoplasm, or the mitochondria. This location information can also help scientists to learn more about the functions of each RBP and the RNA it binds to.

“The strength of this manuscript is in the generation of a comprehensive and multilayered dataset that can be used by the biomedical community to develop therapies targeted to specific sites on the genome using genome-editing strategies, or on the transcriptome using antisense oligonucleotides or agents that mediate RNA interference,” says Gil Ast, a professor of human molecular genetics and biochemistry at Tel Aviv University, who was not involved in the research.

Linking RNA and disease

Many research labs around the world are now using these data in an effort to uncover links between some of the RNA sequences identified and human diseases. For many diseases, researchers have identified genetic variants called single nucleotide polymorphisms (SNPs) that are more common in people with a particular disease.

“If those occur in a protein-coding region, you can predict the effects on protein structure and function, which is done all the time. But if they occur in a noncoding region, it’s harder to figure out what they may be doing,” Burge says. “If they hit a noncoding region that we identified as binding to an RBP, and disrupt the RBP’s motif, then we could predict that the SNP may alter the splicing or stability of the gene.”

Burge and his colleagues now plan to use their RNA-based techniques to generate data on additional RNA-binding proteins.

“This work provides a resource that the human genetics community can use to help identify genetic variants that function at the RNA level,” he says.

The research was funded by the National Human Genome Research Institute ENCODE Project, as well as a grant from the Fonds de Recherche de Québec-Santé.

A recipe for cell fitness

Researchers determine how much of an enzyme is ‘just enough’ to keep a cell healthy and growing.

Raleigh McElvery
July 28, 2020

What ratio of ingredients makes a healthy cell? Researchers know which components are required for proper function, but they’re still working to understand what happens when there’s too much of one protein or not enough of another. As a graduate student in Gene-Wei Li’s lab, Darren Parker PhD ’20 spent years tweaking the recipe for a bacterial cell, adding more or less of one enzyme, aminoacyl-tRNA synthetase (aaRS). He wanted to know: How much aaRS is “just right” for bacterial cells? His findings were published in Cell Systems on July 28.

tRNAs, or transfer RNAs, carry amino acids to the ribosome to help produce proteins. But first, aaRSs must “charge” the tRNAs by attaching an amino acid to them. In doing so, aaRSs not only help the cell make proteins and grow; they also ensure the levels of “uncharged” tRNAs lacking amino acids don’t rise too high, as too many of them can trigger stress responses that slow cell growth. Parker and his collaborators predicted that tinkering with aaRS levels would uncover one of two possible scenarios. Perhaps cells tune their aaRS production to minimize the amount of uncharged tRNAs present. Alternatively, aaRS production could be dictated by the rate of protein synthesis necessary for cell growth — even if that means accumulating uncharged tRNAs.

The researchers determined the latter was true: cells make “just enough” aaRSs to optimize protein production and cell growth. This delicate balance was easily upset when too few aaRSs were produced, cueing the stress responses to kick in and slow growth. Although excess aaRSs reduced the amount of uncharged tRNA, it also hindered cell growth. The researchers determined that the cellular circuits in charge of controlling and sensing tRNA charging are collectively tuned to optimize bacterial growth.

“These results demonstrate that cells have delicately balanced the costs and benefits of producing their proteins,” Parker says. “Understanding the driving forces behind protein production is important for better understanding disease processes, and engineering cells to perform new functions.”

3 Questions: Ibrahim Cissé on using physics to decipher biology

A biophysicist employs super-resolution microscopy to peer inside living cells and witness never-before-seen phenomena.

Raleigh McElvery | Department of Biology
July 23, 2020

How do cells use physics to carry out biological processes? Biophysicist Ibrahim Cissé explores this fundamental question in his interdisciplinary laboratory, leveraging super-resolution microscopy to probe the properties of living matter. As a postdoc in 2013, he discovered that RNA polymerase II, a critical protein in gene expression, forms fleeting (“transient”) clusters with similar molecules in order to transcribe DNA into RNA. He joined the Department of Physics in 2014, and was recently granted tenure and a joint appointment in biology. He sat down to discuss how his physics training led him to rewrite the textbook on biology.

Q: How does your work revise conventional models describing how RNA polymerases carry out their cellular duties?

A: My interest in biology has always been curiosity-driven. As a physicist reading biology textbooks, I thought that transcription — the process by which DNA is made into RNA — was fully understood. It’s so basic, and the textbooks write about it with such confidence. Come to find out, most of what we know about the cell nucleus, where gene expression starts, comes from people studying these processes outside the cell, inside a test tube. I started to wonder: Do we actually know how they work in a living cell?

The textbook models say that when a specific gene is being activated, RNA polymerase and dozens of other molecules are recruited to the DNA to begin transcription. If you don’t look closely enough, the polymerases appear to be uniformly distributed and acting randomly throughout the nucleus. However, my single-molecule and “super-resolution” microscopy methods allowed me to see something different when I looked inside live cells: polymerase clusters, which are very dynamic. In the mid-’90s, people had observed similar clusters in so-called “fixed” cells that were chemically frozen. But these findings were dismissed as possible artifacts of the fixation procedure. However, when we saw these same protein clusters in living cells that were not treated with harsh chemicals, it suggested that the textbook explanation may be incomplete.

Q: How has your background in physics given you a unique perspective on the mechanics of living cells?

A: When I arrived at the University of Illinois at Urbana-Champaign to begin my PhD in physics, I hadn’t enrolled in a biology class since high school. I was really taken with the interdisciplinary work of one physics professor, Taekjip Ha, who became my PhD mentor. He had developed single-molecule fluorescence resonance energy transfer techniques, to study with unprecedented sensitivity when two biomolecules are close to each other and monitor the distance between them in real time.

Taekjip graciously accepted me into his lab despite my limited biology background, and I never looked back. His work mirrored my interest in condensed matter physics, but the material we were looking at wasn’t from the inanimate world, it was living matter.

Between 2006 and 2008, as I was working on my PhD, super-resolution microscopy really took off from the single-molecule microscopes I used in grad school. It was a natural progression, in my mind, to learn cell biology during my postdoc fellowship at École Normale Supérieure in Paris, and to try to visualize weak and transient interactions directly in living cells using single-molecule and super-resolution imaging. You could now pinpoint molecules with nanometer accuracy; you could “turn on” and “off” molecules to observe them individually and ensure there was no overlap between those that were side-by-side.

Thanks to these new techniques, we saw clusters of RNA polymerases in living cells for the first time during my postdoc, and I pushed the technique further to reveal the cluster dynamics. But the fact that you had to turn individual molecules on and off made it really hard to see these clusters assembling or disassembling. I didn’t want to trade temporal resolution for spatial resolution. So I came up with an approach called Time-Correlated Photoactivated Localization Microscopy (tcPALM). It allowed us to measure the lifetimes of these ephemeral polymerase clusters, and we found that they last just a few seconds.

Once I arrived at MIT, we wanted to test whether the clusters could be fleeting but still biologically relevant. We pushed a dual-color super-resolution technique where we correlated the clusters with gene activity. With RNA live-imaging experts at Howard Hughes Medical Institute’s Janelia Research Campus, Brian English and Tim Lionnet, and my postdoc, Wonki Cho, we found that roughly 80 to 100 polymerases form a cluster on a gene where transcription is about to start. Although the cluster is only there for a few seconds, that’s enough time to load a handful of polymerases and generate “bursts” of RNA transcription. In fact, there was a linear correlation between the clusters’ transient dynamics and the number of messenger RNAs made in each burst.

Q: What is it like to be a physicist working with biologists?

A: Even though I joined MIT as a physics hire, I was lucky enough to get lab space in Building 68 alongside amazing biologists. They were the perfect people to talk to about my crazy ideas. And it turned out that renowned researchers like Rick Young and Phil Sharp actually had similar theories. They had genomic evidence for clusters of gene regulators, which they call “super enhancers,” that we all thought could relate to what my lab was seeing. That’s led to hours of exciting discussions between our labs, and has evolved into one of my most rewarding collaborations — and revealed that clusters associate as tiny transcriptional condensates with properties of liquid droplets.

Now, students and postdocs in my lab are wondering about the clusters’ functions and mechanisms of action, and whether protein clustering extends beyond transcription. For instance, clustering could explain some aspects of neurodegeneration. One perplexing idea that came out of this work is that perhaps it gets harder for our cells to clear protein condensates as we age, leading to Parkinson’s, Alzheimer’s, and other diseases. It’s becoming clearer that physics may be just as important as biology for understanding how cells work. The physics of how condensates and droplets form in the inanimate world is increasingly helpful in determining how living cells can evolve to regulate the same process for specific biological functions like transcription. Nature uses physics in much more elaborate ways than we initially anticipated.

Lindsay Case

Education

  • PhD, 2014, University of North Carolina at Chapel Hill
  • BA, 2008, Biology, Franklin and Marshall College

Research Summary

We study how cells regulate the spatial organization of signaling molecules at the plasma membrane to control downstream signaling. For example, receptor clustering and higher-order assembly with cytoplasmic proteins can create compartments with unique biochemical and biophysical properties. We use quantitative experimental approaches from biochemistry, molecular biophysics, and cell biology to study transmembrane signaling pathways and how they are misregulated in diseases like cancer.

Awards

  • NSF Career Award, 2025
  • Searle Scholar, 2022
  • NIH Director’s New Innovator Award, 2022
  • AFOSR Young Investigator Award, 2021
  • Brown-Goldstein Award, 2020
  • Damon Runyon-Dale F. Frey Breakthrough Scientist, 2020
Yukiko Yamashita

Education

  • PhD, 1999, Kyoto University
  • BS, Biology, 1994, Kyoto University

Research Summary

Two remarkable feats of multicellular organisms are generation of many distinct cell types via asymmetric cell division and transmission of the germline genome to the next generation, essentially in eternity. Studying these processes using the Drosophila male germline as a model system has led us to venture into new areas of study, such as functions of satellite DNA, a ‘genomic junk,’ and how they might be involved in speciation.

Awards

  • National Academy of Sciences, 2025
  • Tsuneko and Reiji Okazaki Award, 2016
  • Howard Hughes Medical Institute, Investigator, 2014
  • MacArthur Fellow, 2011
  • Women in Cell Biology Early Career Award, American Society for Cell Biology, 2009
  • Searle Scholar, 2008
Seemingly similar, two neurons show distinct styles as they interact with the same muscle partner
Picower Institute
July 7, 2020

A new study by MIT neuroscientists into how seemingly similar neuronal subtypes drive locomotion in the fruit fly revealed an unexpected diversity as the brain’s commands were relayed to muscle fibers. A sequence of experiments revealed a dramatic difference between the two nerve cells – one neuron scrambled to adjust to different changes by the other, but received no requital in response when circumstances were reversed.

The findings published in the Journal of Neuroscience suggest that these subclasses of neurons, which are also found abundantly in people and many other animals, exhibit a previously unappreciated diversity in their propensity to respond to changes, a key property known as “synaptic plasticity.” Synaptic plasticity is considered an essential mechanism of how learning and memory occur in the brain, and aberrations in of the process are likely central to disorders such as autism.

“By seeing that these two different types of motor neurons actually show very distinct types of plasticity, that’s exciting because it means it’s not just one thing happening,” said senior author Troy Littleton, a member of The Picower Institute for Learning and Memory and Menicon Professor of Neuroscience in MIT’s Departments of Biology and of Brain and Cognitive Sciences. “There’s multiple types of things that can be altered to change connectivity within the neuromuscular system.”

Tonic and phasic neurons

Both of the neurons work in the same way, by emitting the neurotransmitter glutamate onto their connections, or synapses, with the muscles. But these two neurons do so with different styles. The “tonic” neuron, which connects only to a single muscle, emits its glutamate at a constant but low rate while the muscle is active. Meanwhile, the “phasic” neuron connects to a whole group of muscles and jumps in with a strong quick pulse of activity to spring the muscles into action.

Heading into the study Littleton and lead author Nicole Aponte-Santiago were curious to explore whether these different neurons compete or cooperate to drive the muscle fibers, and if they exhibited different plasticity when their functions were altered. To get started on what ultimately became her dissertation research, Aponte-Santiago developed the means to tailor genetic alterations specifically in each of the two neurons.

“The reason we were able to answer these questions in the first place was because we produced tools to start differentially manipulating one neuron versus the other one, or label one versus the other one,” said Aponte-Santiago, who earned her PhD in Littleton’s lab earlier this spring and is now a postdoc at the University of California at San Francisco.

With genetic access to each neuron, Aponte-Santiago distinctly labeled them to watch each one grow in fly larvae as they developed. She saw that the tonic neuron reached the muscle first and that the phasic one connected to the muscle later. She also observed that unlike in mammals, the neurons did not compete to control the muscle but remained side by side, each contributing in its characteristic way to the total electrical activity needed to drive movement.

To study the neurons’ plasticity, Aponte-Santiago employed two manipulations of each neuron. She either wiped them out completely by making them express a lethal protein called “reaper” or she substantially tamped down their glutamate activity via expression of tetanus toxin.

When she wiped out the phasic neuron with reaper, the tonic neuron quickly stepped up its signaling, attempting to compensate as much as it could. But in flies where she wiped out the tonic neuron, the phasic neuron didn’t budge at all, continuing as if nothing had changed.

Similarly when Aponte-Santiago reduced the activity of the phasic neuron with the toxin, the tonic neuron increased the number of boutons and active zone structures in its synapses to respond to the loss of its partner. But when she reduced the activity of the tonic neuron the phasic neuron again didn’t appear to respond.

In all the experiments, the muscle received less overall drive from the neurons than when everything was normal. And while the phasic neuron  apparently didn’t bother to make up for any loss on the part of the tonic neuron, the tonic neuron employed different means to compensate – either increasing its signaling or by enhancing the number of its connections on the muscle – depending on how the phasic neuron was diminished.

“It was quite intriguing that Nicole found that when the phasic input wasn’t there, there was a unique form of plasticity that the tonic neuron showed,” Littleton said, “but if the phasic neuron was there and wasn’t working, the tonic neuron behaved in a very different way.”

Another intriguing aspect of the study is the role of the muscle itself, which may be an active intermediary of the plasticity, Littleton said. The neurons may not sense when each other have been wiped out or inactivated. Instead the muscle appears to call for those changes.

“Even though a muscle has two distinct inputs, it can sort of uniquely control those two,” Littleton said. “When the muscle is getting glutamate, does it know whether it is coming from the tonic or the phasic neuron and does it care? It appears that it does care, that it really needs the tonic more than the phasic. When the phasic is gone it shifts some of the plasticity, but when the tonic is gone the phasic can’t do much about it.”

In new work, the lab is now looking at differences in gene expression between the two neurons to identify which proteins are responsible for the unique properties and plasticity of the tonic and phasic neurons. By defining the genetic underpinnings of their unique properties, the lab hopes to begin to get a handle on the molecular underpinnings of neuronal diversity in the brain.

In addition to Aponte-Santiago and Littleton, the paper’s other authors are Kiel Ormerod and Yulia Akbergenova.

The National Institutes of Health and the JPB Foundation supported the study.

Parasite research heats up
Greta Friar | Whitehead Institute
July 7, 2020

Apicomplexan parasites infect hundreds of millions of people around the world each year. Several species of apicomplexan parasites in the Plasmodium genus cause malaria, while another apicomplexan species, Toxoplasma gondii (T. gondii), causes toxoplasmosis, a disease with flu-like symptoms that can be lethal for people with weakened immune systems. In spite of their impact, the biology of these disease-causing parasites is not very well understood and treatment options for infection are limited.

One potential approach to treat infection could be drugs that disrupt the parasites’ calcium signaling, which they rely on to spread from cell to cell in their hosts. The parasites need an influx of calcium in order to burst out of an infected host cell—a process called egress—and move through the host’s body and invade other cells. In previous work, a researcher from Whitehead Institute Member Sebastian Lourido’s lab, Saima Sidik, had tested a large collection of molecules and identified one called enhancer 1 (ENH1), which perturbed the parasites’ calcium levels and prevented egress, as a promising anti-parasitic lead. However, the original experiments did not determine how ENH1 acts. In research published in the journal ACS Chemical Biology on June 29, Alice Herneisen, a graduate student in Lourido’s lab, and Lourido, who is also an assistant professor of biology at the Massachusetts Institute of Technology, used an approach called thermal proteome profiling to discover how ENH1 prevents T. gondii parasites from egress. They identified the main target of ENH1 as a calcium-dependent molecule called CDPK1 that parasites use to prepare for egress, moving between cells, and invasion of host cells. ENH1 binds to and prevents CDPK1 from functioning.

“Advances over the past few decades have made discovering a molecule’s potentially therapeutic activity much easier, but the next step of figuring out how the molecule works is often still a challenge,” Lourido says. “By applying newer expansive approaches, we are starting to build a more holistic picture of the parasites’ cell biology.”

Understanding the biology responsible for a potential drug’s observed effects is important because most drugs require modification before they are ready for human use—they may need to be made less toxic, more potent, or more amenable to the environment of the human body—and these sorts of modifications cannot be made until the molecule and its activity are understood.

Herneisen decided to use a relatively new approach in parasites, thermal proteome profiling, to discover the targets of ENH1—the molecules it binds to, leading to its therapeutic effects. The approach works by graphing how each of the proteins inside the parasite reacts to changes in heat with and without being exposed to ENH1. One advantage of this approach is that it is unbiased, meaning that instead of researchers picking likely targets up front to test, they investigate as many molecules as possible, which can lead to unexpected findings. For example, Lourido has been investigating CDPK1 in other contexts for many years, and based on his lab’s previous understanding of its role would not have expected it to be a main target of ENH1—such surprises can direct research in exciting new directions.

Although CDPK1 is ENH1’s main target, the investigations did not uncover the target that allows ENH1 to cause oscillations in the parasites’ calcium levels. Finding this missing target is one of the lab’s next goals.

“The fact that ENH1 affects multiple aspects of calcium signaling may be what makes it such an effective antiparasitic agent,” Herneisen says. “It’s messing with the parasites on several levels.”

Translation of the research for clinical testing is a long way off, but there are multiple indicators that this is a promising direction for investigation. Not only is calcium signaling key to the parasites’ life cycle and ability to spread inside of a host, but the molecules and mechanisms that the parasites use to modulate calcium levels are very different from the ones found in mammals. This means that a drug that disrupts the parasites’ calcium signaling is unlikely to interfere with calcium signaling in human patients, and so could be deadly to the parasites without harming the patients’ cells.

Written by Greta Friar

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Sebastian Lourido’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 assistant professor of biology at the Massachusetts Institute of Technology.

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Herneisen, Alice L. et al. “Identifying the target of an antiparasitic compound in Toxoplasma using thermal proteome profiling.” ACS Chemical Biology, June 29, 2020. https://doi.org/10.1021/acschembio.0c00369

Ruth Lehmann

Education

  • Dr. rer. nat., 1985, University of Tübingen
  • MS, 1981, Biology, University of Freiburg

Research Summary

We study germ cells, the only cells in the body naturally able to generate completely new organisms. In addition to the nuclear genome, cytoplasmic information is passed though the egg cell to the next generation. We analyze the organization and regulation of germ line specific RNA-protein condensates, and explore mechanisms used by endosymbionts such as mitochondria and the intracellular bacterium, Wolbachia, to propagate through the cytoplasm of the female germ line.

Awards

  • Vanderbilt Prize in Biomedical Science, 2022
  • Gruber Genetics Prize, 2022
  • Thomas Hunt Morgan Medal, Genetics Society of America, 2021
  • Francis Amory Prize in Reproductive Medicine and Reproductive Physiology, American Academy of Arts and Sciences, 2020
  • Vilcek Prize in Biomedical Science, 2020
  • Keith R. Porter Award, American Society for Cell Biology, 2018
  • Inaugural Klaus Sander Prize, German Society for Developmental Biology, 2017
  • European Molecular Biology Organization, Foreign Associate, 2012
  • Conklin Medal of the Society of Developmental Biology, 2011
  • National Academy of Sciences, Foreign Associate, 2005; Member, 2008
  • American Academy of Arts and Sciences, Member, 1998
  • Howard Hughes Medical Institute, Investigator, 1990 and 1997