From The University of California-Santa Cruz (US) : “Mirror-image peptides form ‘rippled sheet’ structure predicted in 1953”

From The University of California-Santa Cruz (US)

December 17, 2021
Tim Stephens
stephens@ucsc.edu

A UCSC team obtained an x-ray ‘snapshot’ of a novel protein structure with potential applications in biomedicine and materials science.

This illustration shows the “left-handed” and “right-handed” triphenylalanine peptides which bond together to form a rippled beta sheet. Illustration by Jevgenij Raskatov.

The dimeric rippled sheets assembled into a layered crystal structure with a herringbone pattern. Image credit: Kuhn et al., Chemical Science 2021.

By mixing a small peptide with equal amounts of its mirror image, a team of scientists at UC Santa Cruz has created an unusual protein structure known as a “rippled beta sheet” and obtained images of it using x-ray crystallography. They reported their findings in a paper published December 8 in Chemical Science.

The rippled sheet is a distinctive variation on the pleated beta sheet, which is a well-known structural motif found in thousands of proteins, including important disease-related proteins. Linus Pauling and Robert Corey described the rippled beta sheet in 1953, two years after introducing the concept of the pleated beta sheet.

While the pleated beta sheet (often called simply the beta sheet) quickly became a textbook example of a common protein structure, the rippled sheet has languished in obscurity as a rarely studied and largely theoretical structure. Previous studies have found experimental evidence of rippled sheet formation, but none using x-ray crystallography, which is the gold standard for determining protein structures.

“Now, for the first time, we have the crystal structure of a rippled sheet, which is like a snapshot of it, and the structure closely matches the predictions of Pauling and Corey,” said Jevgenij Raskatov, associate professor of chemistry and biochemistry at UC Santa Cruz and corresponding author of the paper.

“The rippled sheet paradigm may have significance for both materials research and biomedical applications, and having the crystal structure is important for the rational design of rippled sheet materials,” Raskatov noted.

Proteins consist of long chains of amino acids folded into complex three-dimensional shapes that enable them to carry out a huge variety of functions in all living things. A pleated beta sheet is composed of linear strands (called beta strands) bonded together side by side to form a 2-dimensional sheet-like structure. A rippled beta sheet is similar except that alternate strands are mirror images of each other.

The amino acids that make up proteins can have either a “left-handed” (L) or “right-handed” (D) orientation in the arrangement of their atoms—the same in all respects but mirror images, like left and right hands. All natural proteins are made with left-handed amino acids, but synthetic proteins can be made with either L or D amino acids.

In the new study, the researchers used mirror-image forms of triphenylalanine, a short peptide consisting of three phenylalanine amino acids. When mixed in equal amounts, the mirror-image peptides joined in pairs, which then packed together into herringbone layer structures.

“They pack together to form a crystal, so we could use x-ray crystallography to see that rippled sheet structure,” said coauthor Timothy Johnstone, assistant professor of chemistry and biochemistry. “It’s a highly enabling discovery that opens up new avenues for exploration, because it gives us a new building block, or a new way to put building blocks together, for creating novel polypeptide structures with desirable properties.”

Having determined the crystal structure, the researchers then searched the Protein Data Bank, an online archive of structural data, for other proteins involving mirror-image peptides. They found three additional crystal structures containing rippled sheets that had not been recognized when the structures were originally analyzed.

The co-first authors of the paper are Ariel Kuhn, a Ph.D. student in Raskatov’s lab, and Beatriz Ehlke, a Ph.D. student in the lab of coauthor Scott Oliver, professor of chemistry and biochemistry.

“It was a great collaborative effort between the three labs, as well as demonstrating the incredible capabilities of our new single crystal XRD instrument for x-ray crystallography,” Kuhn said.

This work was supported by The National Institutes of Health (US) and The National Science Foundation (US).

See the full article here .

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UC Santa Cruz (US) Lick Observatory Since 1888 Mt Hamilton, in San Jose, California, Altitude 1,283 m (4,209 ft)

UC Observatories Lick Automated Planet Finder fully robotic 2.4-meter optical telescope at Lick Observatory, situated on the summit of Mount Hamilton, east of San Jose, California, USA.

The UCO Lick C. Donald Shane telescope is a 120-inch (3.0-meter) reflecting telescope located at the Lick Observatory, Mt Hamilton, in San Jose, California, Altitude 1,283 m (4,209 ft).
UC Santa Cruz (US) campus.

The University of California-Santa Cruz (US) , opened in 1965 and grew, one college at a time, to its current (2008-09) enrollment of more than 16,000 students. Undergraduates pursue more than 60 majors supervised by divisional deans of humanities, physical & biological sciences, social sciences, and arts. Graduate students work toward graduate certificates, master’s degrees, or doctoral degrees in more than 30 academic fields under the supervision of the divisional and graduate deans. The dean of the Jack Baskin School of Engineering oversees the campus’s undergraduate and graduate engineering programs.

UCSC is the home base for the Lick Observatory.

UCO Lick Observatory’s 36-inch Great Refractor telescope housed in the South (large) Dome of main building.

Search for extraterrestrial intelligence expands at Lick Observatory
New instrument scans the sky for pulses of infrared light
March 23, 2015
By Hilary Lebow

The NIROSETI instrument saw first light on the Nickel 1-meter Telescope at Lick Observatory on March 15, 2015. (Photo by Laurie Hatch.)

Astronomers are expanding the search for extraterrestrial intelligence into a new realm with detectors tuned to infrared light at UC’s Lick Observatory. A new instrument, called NIROSETI, will soon scour the sky for messages from other worlds.

Alumna Shelley Wright, now an assistant professor of physics at UC San Diego (US), discusses the dichroic filter of the NIROSETI instrument, developed at the U Toronto Dunlap Institute for Astronomy and Astrophysics (CA) and brought to UCSD and installed at the UC Santa Cruz (US) Lick Observatory Nickel Telescope (Photo by Laurie Hatch).

“Infrared light would be an excellent means of interstellar communication,” said Shelley Wright, an assistant professor of physics at UC San Diego (US) who led the development of the new instrument while at the U Toronto Dunlap Institute for Astronomy and Astrophysics (CA).

Shelley Wright of UC San Diego with (US) NIROSETI, developed at U Toronto Dunlap Institute for Astronomy and Astrophysics (CA) at the 1-meter Nickel Telescope at Lick Observatory at UC Santa Cruz

NIROSETI team from left to right Rem Stone UCO Lick Observatory Dan Werthimer, UC Berkeley; Jérôme Maire, U Toronto; Shelley Wright, UCSD; Patrick Dorval, U Toronto; Richard Treffers, Starman Systems. (Image by Laurie Hatch).

Wright worked on an earlier SETI project at Lick Observatory as a UC Santa Cruz undergraduate, when she built an optical instrument designed by University of California-Berkeley (US) researchers. The infrared project takes advantage of new technology not available for that first optical search.

Infrared light would be a good way for extraterrestrials to get our attention here on Earth, since pulses from a powerful infrared laser could outshine a star, if only for a billionth of a second. Interstellar gas and dust is almost transparent to near infrared, so these signals can be seen from great distances. It also takes less energy to send information using infrared signals than with visible light.

Frank Drake, professor emeritus of astronomy and astrophysics at UC Santa Cruz and director emeritus of the SETI Institute, said there are several additional advantages to a search in the infrared realm.

Frank Drake with his Drake Equation. Credit Frank Drake.

Drake Equation, Frank Drake, Seti Institute (US).

“The signals are so strong that we only need a small telescope to receive them. Smaller telescopes can offer more observational time, and that is good because we need to search many stars for a chance of success,” said Drake.

The only downside is that extraterrestrials would need to be transmitting their signals in our direction, Drake said, though he sees this as a positive side to that limitation. “If we get a signal from someone who’s aiming for us, it could mean there’s altruism in the universe. I like that idea. If they want to be friendly, that’s who we will find.”

Scientists have searched the skies for radio signals for more than 50 years and expanded their search into the optical realm more than a decade ago. The idea of searching in the infrared is not a new one, but instruments capable of capturing pulses of infrared light only recently became available.

“We had to wait,” Wright said. “I spent eight years waiting and watching as new technology emerged.”

Now that technology has caught up, the search will extend to stars thousands of light years away, rather than just hundreds. NIROSETI, or Near-Infrared Optical Search for Extraterrestrial Intelligence, could also uncover new information about the physical universe.

“This is the first time Earthlings have looked at the universe at infrared wavelengths with nanosecond time scales,” said Dan Werthimer, UC Berkeley SETI Project Director. “The instrument could discover new astrophysical phenomena, or perhaps answer the question of whether we are alone.”

NIROSETI will also gather more information than previous optical detectors by recording levels of light over time so that patterns can be analyzed for potential signs of other civilizations.

“Searching for intelligent life in the universe is both thrilling and somewhat unorthodox,” said Claire Max, director of UC Observatories and professor of astronomy and astrophysics at UC Santa Cruz. “Lick Observatory has already been the site of several previous SETI searches, so this is a very exciting addition to the current research taking place.”

NIROSETI will scan the skies several times a week on the Nickel 1-meter telescope at Lick Observatory, located on Mt. Hamilton east of San Jose.

From DOE’s Brookhaven National Laboratory: “Cell Membrane Proteins Imaged in 3-D”

From DOE’s Brookhaven National Laboratory

April 13, 2020 [From Year End Wrap-up]
Stephanie Kossman
skossman@bnl.gov
(631) 344-8671

Peter Genzer
genzer@bnl.gov
(631) 344-3174

Scientists used lanthanide-binding tags to image proteins at the level of a cell membrane, opening new doors for studies on health and medicine.

Ultrabright x-rays revealed the concentration of erbium (yellow) and zinc (red) in a single E.coli cell expressing a lanthanide-binding tag and incubated with erbium.

A team of scientists including researchers at the National Synchrotron Light Source II (NSLS-II) [below]—a U.S. Department of Energy (DOE) Office of Science User Facility at DOE’s Brookhaven National Laboratory—have demonstrated a new technique for imaging proteins in 3-D with nanoscale resolution. Their work, published in the Journal of the American Chemical Society, enables researchers to identify the precise location of proteins within individual cells, reaching the resolution of the cell membrane and the smallest subcellular organelles.

In the structural biology world, scientists use techniques like x-ray crystallography and cryo-electron microscopy to learn about the precise structure of proteins and infer their functions, but we don’t learn where they function in a cell,” said corresponding author and NSLS-II scientist Lisa Miller. “If you’re studying a particular disease, you need to know if a protein is functioning in the wrong place or not at all.”

The new technique developed by Miller and her colleagues is similar in style to traditional methods of fluorescence microscopy in biology, in which a molecule called green fluorescent protein (GFP) can be attached to other proteins to reveal their location. When GFP is exposed to UV or visible light, it fluoresces a bright green color, illuminating an otherwise “invisible” protein in the cell.

“Using GFP, we can see if a protein is in subcellular structures that are hundreds of nanometers in size, like the nucleus or the cytoplasm,” Miller said, “but structures like a cell membrane, which is only seven to 10 nanometers in size, are difficult to see with visible light tags like GFP. To see structures the size of 10 nanometers in a cell, you benefit greatly from the use of x-rays.”

To overcome this challenge, researchers at NSLS-II teamed up with scientists at the Massachusetts Institute of Technology (MIT) and Boston University (BU) who developed an x-ray-sensitive tag called a lanthanide-binding tag (LBT). LBTs are very small proteins that can bind tightly to elements in the lanthanide series, such as erbium and europium.

Part of the research team is shown at NSLS-II’s Hard X-ray Nanoprobe. Pictured from left to right are Xiaojing Huang, Randy Smith, Yong Chu, Hanfei Yan, Tiffany Victor, and Lisa Miller.

“Unlike GFP, which fluoresces when exposed to UV or visible light, lanthanides fluoresce in the presence of x-rays,” said lead author Tiffany Victor, a research associate at NSLS-II. “And since lanthanides do not occur naturally in the cell, when we see them with the x-ray microscope, we know the location of our protein of interest.”

The researchers at NSLS-II, MIT, and BU worked together to combine LBT technology with x-ray-fluorescence.

“Although LBTs have been used extensively over the last decade, they’ve never been used for x-ray fluorescence studies,” Miller said.

Beyond obtaining higher resolution images, x-ray fluorescence simultaneously provides chemical images on all trace elements in a cell, such as calcium, potassium, iron, copper, and zinc. In other studies, Miller’s team is researching how trace elements like copper are linked to neuron death in diseases like Alzheimer’s. Visualizing the location of these elements in relation to specific proteins will be key to new findings.

In addition to their compatibility with x-rays, LBTs are also beneficial for their relatively small size, compared to visible light tags.

“Imagine you had a tail attached to you that was the size of your whole body, or bigger,” Miller said. “There would be a lot of normal activities that you’d no longer be able to do. But if you only had to walk around with a tiny pig’s tail, you could still run, jump, and fit through doorways. GFP is like the big tail—it can be a real impediment to the function of a many proteins. But these little lanthanide-binding tags are almost invisible.”

To demonstrate the use of LBTs for imaging proteins in 3-D with nanoscale resolution, the researchers at MIT and BU tagged two proteins in a bacterial cell—one cytoplasmic protein and one membrane protein. Then, Miller’s team studied the sample at the Hard X-ray Nanoprobe (HXN) beamline at NSLS-II and the Bionanoprobe beamline at the Advanced Photon Source (APS)—a DOE Office of Science User Facility at DOE’s Argonne National Laboratory.

ANL Advanced Photon Source.

“HXN offers the world-leading x-ray focus size, which goes down to about 12 nanometers. This was critical for imaging the bacterial cell in 3-D with nanoscale resolution,” said Yong Chu, lead beamline scientist at HXN. “We also developed a new way of mounting the cells on a specialized sample holder in order to optimize the efficiency of the measurements.”

By coupling the unparalleled resolution of HXN with the capabilities of LBTs, the team was able to image both of the tagged proteins. Visualizing the cell membrane protein proved LBTs can be seen at a high resolution, while imaging the cytoplasmic protein showed LBTs could also be visualized within the cell.

“At high concentrations, lanthanides are toxic to cells,” Victor said, “so it was important for us to show that we could treat cells with a very low lanthanide concentration that was nontoxic and substantial enough to make it past the cell membrane and image the proteins we wanted to see.”

Now, with this new technique demonstrated successfully, scientists hope to be able to use LBTs to image other proteins within the cell at a resolution of 10 nanometers.

This study was supported by the U.S. Department of Energy and the National Science Foundation. Operations at NSLS-II and APS are supported by DOE’s Office of Science.

See the full article here .

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Brookhaven Campus.

BNL Center for Functional Nanomaterials.

BNL NSLS-II.

BNL NSLS II.

BNL RHIC Campus.

BNL/RHIC Star Detector.

BNL/RHIC Phenix.

One of ten national laboratories overseen and primarily funded by the Office of Science of the U.S. Department of Energy (DOE), Brookhaven National Laboratory conducts research in the physical, biomedical, and environmental sciences, as well as in energy technologies and national security. Brookhaven Lab also builds and operates major scientific facilities available to university, industry and government researchers. The Laboratory’s almost 3,000 scientists, engineers, and support staff are joined each year by more than 5,000 visiting researchers from around the world. Brookhaven is operated and managed for DOE’s Office of Science by Brookhaven Science Associates, a limited-liability company founded by Stony Brook University, the largest academic user of Laboratory facilities, and Battelle, a nonprofit, applied science and technology organization.

From UC Santa Cruz: “New cryo-electron microscopy facility a boon for UCSC structural biologists”

From UC Santa Cruz

December 14, 2020
Tim Stephens
stephens@ucsc.edu

Melissa Jurica, Harry Noller, and Sarah Loerch are among the users of UCSC’s new cryo-EM facility. Credit: C. Lagattuta.

A new cryo-electron microscopy facility is nearing completion at UC Santa Cruz, giving researchers powerful tools for studying the structures and functions of complex molecules involved in human health and disease.

Grants from the National Institutes of Health (NIH) funded the acquisition of a new electron microscope and camera needed to perform cryogenic electron microscopy (cryo-EM), a revolutionary technique for determining the 3-dimensional shapes of large proteins and nucleic acids and the complex assemblies of those molecules that drive the vital functions of living cells.

“A growing community of structural biologists at UCSC is excited to exploit the potential of this technology,” said Melissa Jurica, professor of molecular, cell, and developmental (MCD) biology at UC Santa Cruz.

Structural biologists in several departments at UCSC are doing important work to understand the molecular machinery that operates in the cells of our bodies and underlies human health and disease. This includes research on structures such as the ribosome, spliceosome, cell-cycle regulators, circadian clocks, immune response regulators, childhood viruses, and more.

High-end instruments

Jurica set up UCSC’s first electron microscope with cryogenic capabilities when she joined the faculty in 2003. Cryo-EM technology has improved greatly since then, and in recent years, UCSC researchers have had to go to other institutions to use the latest high-end instruments. Now, they can do the work here and train their students to use state-of-the-art equipment.

“Cryo-EM has gone through a revolution in the past five years or so, made possible by improvements in the technology of the microscopes and the cameras,” Jurica said. “UCSC has a very strong group of structural biologists who need this technology. We all worked together to build this facility.”

Jurica led the application for the $1.6 million NIH High-End Instrumentation grant that funded the purchase of the new cryo-electron microscope, which includes an “autoloader” device that loads the samples and vastly increases both through-put and stability of the instrument.

Seth Rubin, professor of chemistry and biochemistry, and Harry Noller, professor emeritus of MCD biology, got a separate NIH grant to purchase the specialized camera, a direct electron detector, which is a critical component for achieving high-resolution images of molecular structures. The campus administration, meanwhile, funded renovations for the new facility in the Sinsheimer Laboratories building.

“We’re now in a position to do a lot of really good work,” Jurica said.

Users of the facility will include nearly a dozen faculty members in the Departments of Chemistry and Biochemistry, MCD Biology, Biomolecular Engineering, and Microbiology and Environmental Toxicology. The new facility was crucial to the hiring of a new faculty member, Assistant Professor of Chemistry and Biochemistry Sarah Loerch, who uses cutting-edge cryo-EM methods to study the structure and function of RNA-protein complexes. Focusing on specialized ribosomal complexes, the Loerch lab is working to understand how a cell instructs ribosomes to make the right protein at the right time and in the right place.

Crystallography

Traditionally, the preferred technique for determining the structures of complex biomolecules has been x-ray crystallography. This was the method used by Noller and others to determine the structure of the ribosome, a molecular machine composed of multiple proteins and RNAs that performs protein synthesis in all cells. But x-ray crystallography requires crystallizing the structures first, an extremely challenging step for many biological molecules.

Cryo-EM doesn’t require crystals, and its use has grown dramatically as the methods and equipment have improved. The technique involves flash-freezing samples to capture flexible and dynamic biological molecules in static poses. “With cryo-EM, we can now get direct, high-resolution images of molecules that were not possible to crystallize,” Jurica said.

Noller plans to use cryo-EM images of a ribosome at different steps of protein synthesis to create a molecular movie of the process. With x-ray crystallography, this would require many months of trying to get a crystal for each frame of the movie, but cryo-EM makes it much faster and easier to do. Loerch plans to take the technology one step further and actually image ribosomes while they are inside a neuron to determine how the ribosomes are instructed to make proteins in response to a synapse firing.

Jurica’s lab will be looking at the splicing machinery that edits the RNAs copied from genes; Rebecca Dubois, associate professor of biomolecular engineering, will be looking at the structure of viruses to aid in vaccine development; and Carrie Partch, professor of chemistry and biochemistry, will be using cryo-EM to look at how all the parts of the circadian clock work together to maintain the daily rhythms of our cells.

All of these molecules are large and change their shapes constantly as they do their cellular jobs, which means crystallography would take decades, and might never work. “Cryo-EM lets us take a short-cut straight to the molecules,” Jurica said.

Support from multiple departments and divisions, as well as from campus administration, enabled UCSC to build the new cryo-EM facility, she said. The new facility will open in the new year.

See the full article here .

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The University of California, Santa Cruz, opened in 1965 and grew, one college at a time, to its current (2008-09) enrollment of more than 16,000 students. Undergraduates pursue more than 60 majors supervised by divisional deans of humanities, physical & biological sciences, social sciences, and arts. Graduate students work toward graduate certificates, master’s degrees, or doctoral degrees in more than 30 academic fields under the supervision of the divisional and graduate deans. The dean of the Jack Baskin School of Engineering oversees the campus’s undergraduate and graduate engineering programs.

UCSC Lick Observatory, Mt Hamilton, in San Jose, California, Altitude 1,283 m (4,209 ft).

UC Observatories Lick Automated Planet Finder, fully robotic 2.4-meter optical telescope at Lick Observatory, situated on the summit of Mount Hamilton, east of San Jose, California, USA.

The UCO Lick C. Donald Shane telescope is a 120-inch (3.0-meter) reflecting telescope located at the Lick Observatory, Mt Hamilton, in San Jose, California, Altitude 1,283 m (4,209 ft).

UC Santa Cruz campus.

UCSC is the home base for the Lick Observatory.

Lick Observatory’s 36-inch Great Great Refractor telescope housed in the South (large) Dome of main building.

From Penn Today: “Penn joins ‘cryo revolution’ by adding Nobel-winning microscope”

From Penn Today

November 30, 2020
Writers
Erica K. Brockmeier, From Penn Engineering
Lauren Salig

Photographer
Eric Sucar

The Singh Center’s Krios G3i, an electron microscope for studying samples at extremely low temperatures, allows researchers to look at cells, proteins, and nanoparticles like never before.

The Krios G3i, housed at the Singh Center for Nanotechnology, is a Nobel-winning cryogenic electron microscope that allows researchers to look at cells, proteins, and engineered nanoparticles like never before. Credit: Eric Sucar.

Penn’s Singh Center for Nanotechnology has added the newest cutting-edge technology to its collection of already impressive microscopes: The Krios G3i, a cryogenic electron microscope that will allow researchers to look at cells, proteins, and engineered nanoparticles like never before. The trick is to keep them frozen in time, holding them in their natural environment while bombarding them with ultrafast, ultrahot subatomic particles.

“Cryo-electron microscopy, or cryo-EM, will allow scientists at Penn to understand diseases like cancer, Alzheimer’s, Parkinson’s, and heart and kidney disorders. Biomedical engineers will use cryo-EM to improve nanoparticle technology for drug delivery. Bringing this microscope onto campus is a big deal for everyone,” says Vera Moiseenkova-Bell, associate professor of pharmacology in the Perelman School of Medicine and faculty director of the new Beckman Center for Cryo-Electron Microscopy.

The Arnold and Mabel Beckman Foundation, together with the School of Medicine made this Nobel-winning technology available on Penn’s campus last year, allowing scientists to look at molecules in their natural state at a nearly atomic level.

The ‘cryo revolution’

________________________________________________

Technology definitions

A closer look at the different types of imaging methods available

Cryogenic electron microscopy: Electron microscopes (EM) use beams of accelerated electrons, instead of light, as an illumination source, meaning that they have higher resolution and are able to distinguish very small, atomic-level details. In Cryo-EM, a liquified sample is applied onto a grid mesh and flash-frozen in liquid ethane to plunge the sample down to cryogenic temperatures (around −315 °F).

NMR spectroscopy: A technique that studies local magnetic fields around the nuclei of atoms. NMR can help resolve smaller structures in proteins, such as the presence of different functional groups, but larger complexes are not possible to see using this approach.

X-ray crystallography: An approach for determining the atomic and molecular structure of a crystal using X-rays. This method can be used to study both small and large structures, but samples must first be solidified into a highly structured crystal form, an arduous and time-consuming process.
________________________________________________

Moiseenkova-Bell is already seeing firsthand the advantages of using cryo-EM.

“Getting the structures of biologically important molecules at an atomic resolution is now possible because of the cryo revolution. Compared to other techniques, this is so fast. Crystallography can take sometimes over a year to get one structure. Cryo-EM allows you to get a structure in days.”

Many structural biologists like Moiseenkova-Bell consider cryo-EM to be a revolution in their field because of the ways it both supersedes and complements other imaging methods currently available, namely nuclear magnetic resonance (NMR) and X-ray crystallography. Cryo-EM fills the hole left by NMR and X-ray crystallography: Not only can it image the larger molecules excluded from NMR, but it skips the demanding crystallization process of X-ray crystallography.

“With NMR we can only get a small number of protein structures due to the size limitation, and with crystallography you need a well-ordered crystal to get only one structure [of the molecule]. With cryo-EM, you could visualize pretty much everything that is bigger than hemoglobin without needing to crystalize and solve many structures at a time,” Moiseenkova-Bell says.

An added benefit of cryo-EM is that it allows scientists to look at structures in their natural state without crystallizing or dehydrating them. While crystallization requires scientists to take proteins and unnaturally force their atoms into a perfect crystal lattice before imaging, cryo-EM presses pause on a cell’s natural processes, allowing researchers to look at where proteins are and more accurately infer how they work.

“Because the proteins get flash frozen in liquid ethane, we are able to trap them in their natural conformations. That’s another benefit of cryo: The proteins, or cells, are in their native environment which is impossible in other techniques,” says Moiseenkova-Bell.

How to freeze time

The “cryo” and the “EM” portions of the microscope are actually separate processes. What allows cryo-EM to capture a sample in its native state is the “cryo” freezing that takes place when researchers are preparing a sample. Samples are plunged into liquid ethane at temperatures below -315°F, seven times colder than the average temperature of the North Pole during winter, instantly freezing them solid. This process is known as vitrification: Unlike normal freezing, which might form ice crystals that could hinder the imaging, the liquid ethane fixes the sample in a glass-like block that preserves its structure.

Samples prepared for cryo-EM analysis are plunged into liquid ethane, reaching temperatures below -315°F and instantly freezing the material. Credit: Eric Sucar.

The transmission electron microscope, the “EM” part of cryo-EM, works by sending a beam of energetic electrons through the sample. In other EM imaging methods, the powerful beam of subatomic particles can destroy the sample, but cryo-EM’s vitrification process, along with its low electron dose imaging, protects the sample from damage. The electrons’ interactions with the sample’s atoms are detected by the direct electron camera and used to reconstruct a 3D image of the sample.

This last step is crucial: The detectors and the computer program that collect and create images were the final parts of the cryo-EM technique to fall into place. The individual “cryo” and “EM” technologies have been around for decades, metaphorically collecting dust as some structural biologists dismissed the cryo-EM method entirely. Others, however, waited hopefully for the time when technological conditions would be ripe for the cryo-EM revolution to take off.

3D volumes of reconstructed protein channel structures, using data collected with the cryo-EM. (Image: Ruth Anne Pumroy)

“The modern transmission electron microscopes are very advanced and can produce high-resolution images. It’s the development of the medium that we collect images of the biological samples on and the software to reconstruct 3D structures from these images that revolutionized everything. It was a technological evolution,” says Moiseenkova-Bell.

Prior to these advances, cryo-EM relied on photographic film or charge-coupled device cameras to capture images of molecules and cells. On top of the challenges presented by low-quality film and low electron dose imaging, frozen cells continue to move ever so slightly during imaging. Combined, these factors resulted in blurry, often indecipherable photographs of molecular structures, earning cryo-EM the nickname “blob-ology” among structural biologists.

Now, updated technology allows scientists to capture mini-movies of proteins and remove any motion blur from the images. These crisp images of molecules analyzed by powerful computational methods allow scientists to see protein structures at atomic or near atomic resolution.

Looking ahead

With its newly elevated level of resolution, Moiseenkova-Bell sees cryo-EM as a tool that can benefit Penn researchers across disciplines. On the engineering side, cryo-EM offers scientists the chance to more precisely engineer nanoparticles that require each atom to be placed with purpose. On the medical side, researchers hope to use cryo-EM to better understand protein structure and function and in turn, how to best design effective drugs to treat a variety of conditions—a use that is also attracting the attention of big pharmacological companies in Philadelphia.

“Membrane proteins represent 60 percent of druggable targets. If we can know the structure of each target at atomic resolution, we could understand their function and then start doing novel structure-based drug development for these targets,” says Moiseenkova-Bell. “Or, we could solve the structures of proteins with known therapeutic compounds and see how we can improve their effect. The options are endless for drug discovery.”

Part of the allure of Penn’s cryo-EM is its accessibility. Not only will Penn researchers of many disciplines have access to its capabilities, but the machine could attract collaboration with researchers and companies all over the northeastern United States. The Singh Center is already part of the National Nanotechnology Coordinated Infrastructure, which aims to provide access to specialized resources for nanotechnology researchers, and the addition of cryo-EM expands the resources that Penn can offer to valuable research endeavors.

Eric Stach, professor in the School of Engineering and Appled Science and chair of the Faculty Oversight Committee at the Nanoscale Characterization Facility, sees Penn’s cryo-EM as teeming with potential for insightful research and collaboration.

“The co-location of this forefront instrument in the Singh Center for Nanotechnology will provide unique opportunities at the interface between biological sciences and nanotechnology,” he says. “[Additionally,] Penn recently won a National Science Foundation Major Research Instrumentation grant to install a unique electron microscopy sample preparation instrument—a Cryo Focused Ion Beam Microscope.”

The Cryo Focused Ion Beam Microscope accelerates ions into a sample, rather than using electrons like EM devices. Although the instrument can be used for imaging, the microscope is also useful as a cryo-EM preparation device because, at its highest energy levels, the laser-like beam of ions can cut frozen samples into thin slices that are suitable for subsequent imaging in the Krios G3i machine.

“[Having access to this instrument] will allow researchers to use the cryo-EM to effectively sample the interface between ‘soft’ biological structures and ‘hard’ materials,” says Stach. “This will enable entirely new areas of scientific exploration across multiple research fields.”

See the full article here .

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Academic life at Penn is unparalleled, with 100 countries and every U.S. state represented in one of the Ivy League’s most diverse student bodies. Consistently ranked among the top 10 universities in the country, Penn enrolls 10,000 undergraduate students and welcomes an additional 10,000 students to our world-renowned graduate and professional schools.

Penn’s award-winning educators and scholars encourage students to pursue inquiry and discovery, follow their passions, and address the world’s most challenging problems through an interdisciplinary approach.

From Lawrence Berkeley National Lab: “Showtime for Photosynthesis”

From Lawrence Berkeley National Lab

June 4, 2020
Aliyah Kovner
akovner@lbl.gov
510-486-6376

One of nature’s most important chemical reactions is now being captured in a breakthrough “molecular movie”.

(Credit: Greg Stewart/SLAC National Accelerator Laboratory)

Using a unique combination of nanoscale imaging and chemical analysis, an international team of researchers has revealed a key step in the molecular mechanism behind the water splitting reaction of photosynthesis, a finding that could help inform the design of renewable energy technology.

“Life depends on the oxygen that plants and algae split from water; how they do it is still a mystery, but scientists, including our team, are slowly peeling away the layers to get to the answer,” said Vittal K. Yachandra, co-lead author of a new study published in PNAS and a chemist senior scientist at the Department of Energy’s (DOE) Lawrence Berkeley Laboratory (Berkeley Lab). “If we can understand this step of natural photosynthesis, it would enable us to use those design principles for building artificial photosynthetic systems that produce clean and renewable energy from sunlight and water.”

With an instrument that the team designed and fabricated, they analyzed photosynthetic proteins using both X-ray crystallography and X-ray emission spectroscopy. This dual approach, which the team pioneered and have been refining for the past 10 years, generates chemical and protein structure information from the same sample at the same time. The imaging was performed with the X-ray free-electron laser (XFEL) at the LCLS at SLAC National Laboratory, and at SACLA in Japan.

“With this technique, we get the overall picture of how the entire protein structure dynamically changes and we see the chemical intricacies occurring at the reaction site,” said co-lead author Junko Yano, a chemist senior scientist in Berkeley Lab’s Molecular Biophysics and Integrated Bioimaging (MBIB) Division. “The X-ray free electron laser produces extremely bright, short bursts of X-rays that allow us to not only analyze a protein at room temperature, which is how these reactions occur in nature, but also capture various moments over the reaction time scale.”

Structural changes of Photosystem II and its catalytic center (Mn4Ca cluster) during the water oxidation reaction. The movie shows the S2 to S3 transition step, where the first water (as shown in Ox) comes into the catalytic center after the photochemical reaction at the reaction center. (Credit: Jan Kern and Isabel Bogacz/Berkeley Lab)

Structural changes of Photosystem II and its catalytic center (Mn4Ca cluster) during the water oxidation reaction. The movie shows the S2 to S3 transition step, where the first water (as shown in Ox) comes into the catalytic center after the photochemical reaction at the reaction center. (Credit: Jan Kern and Isabel Bogacz/Berkeley Lab)

Traditional crystallography methods often require the sample proteins to be frozen; consequently, they can only generate snapshots of static proteins. This limitation makes it difficult for scientists to get a handle on how proteins actually behave in living organisms, because the molecules morph between different physical states during chemical reactions.

“The water-splitting reaction in photosynthesis is a cyclical process that needs four photons and cycles between four stable ‘states,’” said Yano. “Previously, we could only take pictures of these four states. But by taking multiple snapshots in time, we now can visualize how one state goes to the other.”

“We saw, really nicely, how the structure changes step-by-step as it transforms from one state to the next state,” said Jan F. Kern, MBIB chemist and co-author. “It is pretty exciting, because we can see the ‘cause and effect’ and the role that each moving atom plays in this transition.”

Nicholas K. Sauter, co-author and MBIB computational senior scientist, added: “Essentially, we’re trying to take a ‘movie’ of a chemical reaction. We made a lot of progress to get to this point, in terms of our technology and our computational analyses. The work of our co-author Paul Adams and others in MBIB was critical to interpreting the XFEL and X-ray data. But we still have to get the other frames to see how the reaction is completed and the enzyme is ready for the next cycle.”

The Berkeley Lab researchers hope to continue the project once the many research sites that the entire international team relies upon – located in the U.S., Japan, Switzerland, and South Korea – are operating normally following the COVID-19 pandemic.

Kern concluded by noting that the technological milestone presented in this paper benefited greatly from the diverse expertise of the authors from SLAC, Uppsala and Umeå Universities in Sweden, Humboldt University in Germany, and from the capabilities of five DOE Office of Science user facilities: the Stanford Synchrotron Radiation Lightsource and LCLS at Stanford University, and the Advanced Light Source, Energy Sciences Network, and National Energy Research Scientific Computing Center at Berkeley Lab.

Other Berkeley Lab scientists who contributed to this work include: Ruchira Chatterjee, Louise Lassalle, Kyle D. Sutherlin, Iris D. Young, Sheraz Gul, In-Sik Kim, Philipp S. Simon, Isabel Bogacz, Cindy C. Pham, Nicholas Saichek, Trent Northen, Asmit Bhowmick, Robert Bolotovsky, Derek Mendez, Nigel W. Moriarty, James M. Holton, Aaron S. Brewster, and David Skinner.

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In the world of science, Lawrence Berkeley National Laboratory (Berkeley Lab) is synonymous with “excellence.” Thirteen Nobel prizes are associated with Berkeley Lab. Seventy Lab scientists are members of the National Academy of Sciences (NAS), one of the highest honors for a scientist in the United States. Thirteen of our scientists have won the National Medal of Science, our nation’s highest award for lifetime achievement in fields of scientific research. Eighteen of our engineers have been elected to the National Academy of Engineering, and three of our scientists have been elected into the Institute of Medicine. In addition, Berkeley Lab has trained thousands of university science and engineering students who are advancing technological innovations across the nation and around the world.

Berkeley Lab is a member of the national laboratory system supported by the U.S. Department of Energy through its Office of Science. It is managed by the University of California (UC) and is charged with conducting unclassified research across a wide range of scientific disciplines. Located on a 202-acre site in the hills above the UC Berkeley campus that offers spectacular views of the San Francisco Bay, Berkeley Lab employs approximately 3,232 scientists, engineers and support staff. The Lab’s total costs for FY 2014 were $785 million. A recent study estimates the Laboratory’s overall economic impact through direct, indirect and induced spending on the nine counties that make up the San Francisco Bay Area to be nearly $700 million annually. The Lab was also responsible for creating 5,600 jobs locally and 12,000 nationally. The overall economic impact on the national economy is estimated at $1.6 billion a year. Technologies developed at Berkeley Lab have generated billions of dollars in revenues, and thousands of jobs. Savings as a result of Berkeley Lab developments in lighting and windows, and other energy-efficient technologies, have also been in the billions of dollars.

Berkeley Lab was founded in 1931 by Ernest Orlando Lawrence, a UC Berkeley physicist who won the 1939 Nobel Prize in physics for his invention of the cyclotron, a circular particle accelerator that opened the door to high-energy physics. It was Lawrence’s belief that scientific research is best done through teams of individuals with different fields of expertise, working together. His teamwork concept is a Berkeley Lab legacy that continues today.

A U.S. Department of Energy National Laboratory Operated by the University of California.

From SLAC National Accelerator Lab: “Researchers reveal how enzyme motions catalyze reactions”

From SLAC National Accelerator Lab

December 16, 2019
Ali Sundermier

What they learned could lead to a better understanding of how antibiotics are broken down in the body, potentially leading to the development of more effective drugs.

This illustration shows how an enzyme moves and changes as it catalyzes complex reactions and breaks down organic compounds. (10.1073/pnas.1901864116)

In a time-resolved X-ray experiment, researchers uncovered, at atomic resolution and in real time, the previously unknown way that a microbial enzyme breaks down organic compounds.

The team, led by Mark Wilson at the University of Nebraska Lincoln (UNL) and Henry van den Bedem at the Department of Energy’s SLAC National Accelerator Laboratory (now at Atomwise Inc.), published their findings last week in the Proceedings of the National Academy of Sciences. What they learned about this enzyme, whose structure is similar to one that is implicated in neurodegenerative diseases such as Parkinson’s, could lead to a better understanding of how antibiotics are broken down by microbes and to the development of more effective drugs.

Previously, the researchers used SLAC’s Stanford Synchrotron Radiation Lightsource (SSRL) to obtain the structure of the enzyme at very low temperatures using X-ray crystallography.

SLAC/SSRL

In this study, Medhanjali Dasgupta, a UNL graduate student who was the study’s first author, used the Linac Coherent Light Source (LCLS), SLAC’s X-ray laser, to watch the enzyme and its substrate within the crystal move and change as it went through a full catalytic cycle at room temperature.

SLAC/LCLS

The scientists used special software, designed by van den Bedem, that is highly sensitive to identifying protein movement from X-ray crystallography data to interpret the results, revealing never-before-seen motions that play a key role in catalyzing complex reactions, such as breaking down antibiotics. Next, the researchers hope to use LCLS to obtain room temperature structures of other enzymes to get a better look at how the motions occurring within them help move along reactions.

SSRL and LCLS are DOE Office of Science user facilities. This work was funded by the DOE Office of Science and the National Institutes of Health, among other sources.

See the full article here .

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SLAC is a multi-program laboratory exploring frontier questions in photon science, astrophysics, particle physics and accelerator research. Located in Menlo Park, California, SLAC is operated by Stanford University for the DOE’s Office of Science.

From Brookhaven National Lab: “Imaging the Chemical Structure of Individual Molecules, Atom by Atom”

From Brookhaven National Lab

July 22, 2019

Ariana Manglaviti
amanglaviti@bnl.gov

Using atomic force microscopy images, scientists at Brookhaven Lab’s Center for Functional Nanomaterials developed a guide for discriminating atoms other than hydrogen and carbon in aromatic molecules—ring-shaped molecules with special bonding properties—to help identify contaminants found in petroleum.

Brookhaven Lab physicist Percy Zahl with the noncontact atomic force microscope he adapted and used at the Center for Functional Nanomaterials (CFN) to image nitrogen- and sulfur-containing molecules in petroleum.

For physicist Percy Zahl, optimizing and preparing a noncontact atomic force microscope (nc-AFM) to directly visualize the chemical structure of a single molecule is a bit like playing a virtual reality video game. The process requires navigating and manipulating the tip of the instrument over the world of atoms and molecules, eventually picking some up at the right location and in the right way. If these challenges are completed successfully, you advance to the highest level, obtaining images that precisely show where individual atoms are located and how they are chemically bonded to other atoms. But take one wrong move, and it is game over. Time to start again.

“The nc-AFM has a very sensitive single-molecule tip that scans over a carefully prepared clean single-crystal surface at a constant height and “feels” the forces between the tip molecule and single atoms and bonds of molecules placed on this clean surface,” explained Zahl, who is part of the Interface Science and Catalysis Group at the Center for Functional Nanomaterials (CFN), a U.S. Department of Energy (DOE) Office of Science User Facility at Brookhaven National Laboratory. “It can take an hour or days to get this sensor working properly. You can’t simply press a button; fine tuning is required. But all of this effort is definitely worthwhile once you see the images appearing like molecules in a chemistry textbook.”

A history of chemical structure determination

Since the beginning of the field of chemistry, scientists have been able to determine the elemental composition of molecules. What has been more difficult is to figure out their chemical structures, or the particular arrangement of atoms in space. Knowing the chemical structure is important because it impacts the molecule’s reactivities and other properties.

Kekulé claims that the idea of the ring structure of benzene came to him in a dream of a snake eating its own tail.

For example, Michael Faraday isolated benzene in 1825 from an oil gas residue. It was soon determined that benzene is composed of six hydrogen and six carbon atoms, but its chemical structure remained controversial until 1865, when Friedrich August Kekulé proposed a cyclic structure. However, his proposal was not based on a direct observation but rather on logic deduction from the number of isomers (compounds with the same chemical formula but different chemical structures) of benzene. The correct symmetric hexagonal structure of benzene was finally revealed through its diffraction pattern obtained by Kathleen Lonsdale via x-ray crystallography in 1929. In 1931, Erich Huckel used quantum theory to explain the origin of “aromaticity” in benzene. Aromaticity is a property of flat ring-shaped molecules in which electrons are shared between atoms. Because of this unique arrangement of electrons, aromatic compounds have a special stability (low reactivity).

Today, x-ray crystallography continues to be a mainstream technique for determining chemical structures, along with nuclear magnetic resonance spectroscopy. However, both techniques require crystals or relatively pure samples, and chemical structure models must be deducted by analyzing the resulting diffraction patterns or spectra.

The first-ever actual image of a chemical structure was obtained only a decade ago. In 2009, scientists at IBM Research–Zurich Lab in Switzerland used nc-AFM to resolve the atomic backbone of an individual molecule of pentacene, seeing its five fused benzene rings and even the carbon-hydrogen bonds. This breakthrough was made possible by selecting an appropriate molecule for the end of the tip—one that could come very close to the surface of pentacene without reacting with or binding to it. It also required optimized sensor readout electronics at cryogenic temperatures to measure small frequency shifts in the probe oscillation (which relates to the force) while maintaining mechanical and thermal stability through vibration damping setups, ultrahigh vacuum chambers, and low-temperature cooling systems.

“Low-temperature nc-AFM is the only method that can directly image the chemical structure of a single molecule,” said Zahl. “With nc-AFM, you can visualize the positions of individual atoms and the arrangement of chemical bonds, which affect the molecule’s reactivity.”

However, currently there are still some requirements for molecules to be suitable for nc-AFM imaging. Molecules must be mainly planar (flat), as the scanning occurs on the surface and thus is not suitable for large three-dimensional (3-D) structures such as proteins. In addition, because of the slow nature of scanning, only a few hundred molecules can be practically examined per experiment. Zahl notes that this limitation could be overcome in the future through artificial intelligence, which would pave the way toward automated scanning probe microscopy.

According to Zahl, though nc-AFM has since been applied by a few groups around the world, it is not widespread, especially in the United States.

“The technique is still relatively new and there is a long learning curve in acquiring CO tip-based molecular structures,” said Zahl. “It takes a lot of experience in scanning probe microscopy, as well as patience.”

A unique capability and expertise

The nc-AFM at the CFN represents one of a few in this country. Over the past several years, Zahl has upgraded and customized the instrument, most notably with the open-source software and hardware, GXSM (for Gnome X Scanning Microscopy). Zahl has been developing GXSM for more than two decades. A real-time signal processing control system and software continuously records operating conditions and automatically adjusts the tip position as necessary to avoid unwanted collisions when the instrument is operated in an AFM-specific scanning mode to record forces over molecules. Because Zahl wrote the software himself, he can program and implement new imaging or operating modes for novel measurements and add features to help operators better explore the atomic world.

DBT (left column) is one of the sulfur-containing compounds in petroleum; CBZ and ACR (right and middle columns, respectively) are nitrogen-containing compounds. Illustrations and ball-and-stick models of their chemical structures are shown at the top of each column (black indicates carbon atoms; yellow indicates sulfur, and blue indicates nitrogen). The simulated atomic force microscopy images (a, b, d, e, g, and h) well match the ones obtained experimentally (c, f, and i).

For example, recently Zahl applied a custom “slicing” mode to determine the 3-D geometrical configuration in which a single molecule of dibenzothiopene (DBT)—a sulfur-containing aromatic molecule commonly found in petroleum—adsorbs on a gold surface. The DBT molecule is not entirely planar but rather tilted at an angle, so he combined a series of force images (slices) to create a topographic-like representation of the molecule’s entire structure.

“In this mode, obstacles such as protruding atoms are automatically avoided,” said Zahl. “This capability is important, as the force measurements are ideally taken in one fixed plane, with the need to be very close to the atoms to feel the repulsive forces and ultimately to achieve detailed image contrast. When parts stick out of the molecule plane, they will likely negatively impact image quality.”

This imaging of DBT was part of a collaboration with Yunlong Zhang, a physical organic chemist at ExxonMobil Research and Engineering Corporate Strategic Research in New Jersey. Zhang met Zahl at a conference two years ago and realized that the capabilities and expertise in nc-AFM at the CFN would have great potential for his research on petroleum chemistry.

Zahl and Zhang used nc-AFM to image the chemical structure of not only DBT but also of two nitrogen-containing aromatic molecules—carbazole (CBZ) and acridine (ACR)—that are widely observed in petroleum. In analyzing the images, they developed a set of templates of common features in the ring-shaped molecules that can be used to find sulfur and nitrogen atoms and distinguish them from carbon atoms.

Petroleum: a complex mixture

The chemical composition of petroleum widely varies depending on where and how it formed, but in general it contains mostly carbon and hydrogen (hydrocarbons) and smaller amounts of other elements, including sulfur and nitrogen. During combustion, when the fuel is burned, these “heteroatoms” produce sulfur and nitrogen oxides, which contribute to the formation of acid rain and smog, both air pollutants that are harmful to human health and the environment. Heteroatoms can also reduce fuel stability and corrode engine components. Though refining processes exist, not all of the sulfur and nitrogen is removed. Identifying the most common structures of impure molecules containing nitrogen and sulfur atoms could lead to optimized refining processes for producing cleaner and more efficient fuels.

“Our previous research with the IBM group at Zurich on petroleum asphaltenes and heavy oil mixtures provided the first “peek” into numerous structures in petroleum,” said Zhang. “However, more systemic studies are needed, especially on the presence of heteroatoms and their precise locations within aromatic hydrocarbon frameworks in order to broaden the application of this new technique to identify complex molecular structures in petroleum.”

To image the atoms and bonds in DBT, CBZ, and ACR, the scientists prepared the tip of the nc-AFM with a single crystal of gold at the apex and a single molecule of carbon monoxide (CO) at the termination point (the same kind of molecule used in the original IBM experiment). The metal crystal provides an atomically clean and flat support from which the CO molecule can be picked up.

After “functionalizing” the tip, they deposited a few of each of the molecules (dusting amount) on a gold surface inside the nc-AFM under ultrahigh vacuum at room temperature via sublimation. During sublimation, the molecules go directly from a solid to gas phase.

Though the images they obtained strikingly resemble chemical structure drawings, you cannot directly tell from these images whether there is a nitrogen, sulfur, or carbon atom present in a particular site. It takes some input knowledge to deduct this information.

“As a starting point, we imaged small well-known molecules with typical building blocks that are found in larger polycyclic aromatic hydrocarbons—in this case, in petroleum,” explained Zahl. “Our idea was to see what the basic building blocks of these chemical structures look like and use them to create a set of templates for finding them in larger unknown molecular mixtures.”

An illustration showing how nc-AFM can distinguish sulfur- and nitrogen-containing molecules commonly found in petroleum. A tuning fork (grey arm) with a highly sensitive tip containing a single carbon monoxide molecule (black is carbon and red is oxygen) is brought very close to the surface (outlined in white), with the oxygen molecule lying flat on the surface without making contact. As the tip scans across the surface, it “feels” the forces from the bonds between atoms to generate an image of the molecule’s chemical structure. One image feature that can be used to discriminate between the different types of atoms is the relative “size” of the elements (indicated by the size of the boxes in the overlaid periodic table).

For example, for sulfur- and nitrogen-containing molecules in petroleum, sulfur is only found in ring structures with five atoms (pentagon ring structure), while nitrogen can be present in rings with either five or six (hexagonal ring structure) atoms. In addition to this bonding geometry, the relative “size,” or atomic radius, of the elements can help distinguish them. Sulfur is relatively larger than nitrogen and carbon, and nitrogen is slightly smaller than carbon. It is this size, or “height,” that AFM is extremely sensitive to.

“Simply speaking, the force that the AFM records in very close proximity to an atom relates to the distance and thus to the size of that atom; as the AFM scans over a molecule at a fixed elevation, bigger atoms protrude more out of the plane,” explained Zahl. “Therefore, the larger the atom in a molecule, the bigger the force that the AFM records as it gets closer to its atomic shell, and the repulsion increases dramatically. That is why in the images sulfur appears as a bright dot, while nitrogen looks a hint fainter.”

Zahl and Zhang then compared their experimental images to computer-simulated ones they obtained using the mechanical probe particle simulation method. This method simulates the actual forces acting on the CO molecule on the tip end as it scans over molecules and bends in response. They also performed theoretical calculations to determine how the electrostatic potential (charge distribution) of the molecules affects the measured force and relates to their appearance in the nc-AFM images.

“We used density functional theory to study how the forces felt by the CO probe molecule behave in the presence of the charge environment surrounding the molecules,” said Zahl. “We need to know how the electrons are distributed in order to understand the atomic force and bond contrast mechanism. These insights even allow us to assign single or double bonds between atoms by analyzing image details.”

Going forward, Zahl will continue developing and enhancing nc-AFM imaging modes and related technologies to explore many kinds of interesting, unknown, or novel molecules in collaboration with various users. Top candidate molecules of interest include those with large magnetic moments and special spin properties for quantum applications and novel graphene-like (graphene is a one-atom-thick sheet of carbon atoms arranged in a hexagonal lattice) materials with extraordinary electronic properties.

“The CFN has unique capabilities and expertise in nc-AFM that can be applied to a wide range of molecules,” said Zahl. “In the coming years, I believe that artificial intelligence will make a big impact on the field by helping us operate the microscope autonomously to perform the most time-consuming, tedious, and error-prone parts of experiments. With this special power, our chances of winning the “game” will be much improved.”

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One of ten national laboratories overseen and primarily funded by the Office of Science of the U.S. Department of Energy (DOE), Brookhaven National Laboratory conducts research in the physical, biomedical, and environmental sciences, as well as in energy technologies and national security. Brookhaven Lab also builds and operates major scientific facilities available to university, industry and government researchers. The Laboratory’s almost 3,000 scientists, engineers, and support staff are joined each year by more than 5,000 visiting researchers from around the world. Brookhaven is operated and managed for DOE’s Office of Science by Brookhaven Science Associates, a limited-liability company founded by Stony Brook University, the largest academic user of Laboratory facilities, and Battelle, a nonprofit, applied science and technology organization.

From Brookhaven National Lab: “New Approach for Solving Protein Structures from Tiny Crystals”

From Brookhaven National Lab

May 3, 2019
Karen McNulty Walsh
kmcnulty@bnl.gov
(631) 344-8350

Peter Genzer,
genzer@bnl.gov
(631) 344-3174

Technique opens door for studies of countless hard-to-crystallize proteins involved in health and disease.

Wuxian Shi, Martin Fuchs, Sean McSweeney, Babak Andi, and Qun Liu at the FMX beamline at Brookhaven Lab’s National Synchrotron Light Source II [see below], which was used to determine a protein structure from thousands of tiny crystals.

Using x-rays to reveal the atomic-scale 3-D structures of proteins has led to countless advances in understanding how these molecules work in bacteria, viruses, plants, and humans—and has guided the development of precision drugs to combat diseases such as cancer and AIDS. But many proteins can’t be grown into crystals large enough for their atomic arrangements to be deciphered. To tackle this challenge, scientists at the U.S. Department of Energy’s (DOE) Brookhaven National Laboratory and colleagues at Columbia University have developed a new approach for solving protein structures from tiny crystals.

The method relies on unique sample-handling, signal-extraction, and data-assembly approaches, and a beamline capable of focusing intense x-rays at Brookhaven’s National Synchrotron Light Source II (NSLS-II)—a DOE Office of Science user facility—to a millionth-of-a-meter spot, about one-fiftieth the width of a human hair.

“Our technique really opens the door to dealing with microcrystals that have been previously inaccessible, including difficult-to-crystallize cell-surface receptors and other membrane proteins, flexible proteins, and many complex human proteins,” said Brookhaven Lab scientist Qun Liu, the corresponding author on the study, which was published on May 3 in IUCrJ, a journal of the International Union of Crystallography.

Deciphering protein structures

Protein crystallography has been a dominant method for solving protein structures since 1958, improving over time as x-ray sources have grown more powerful, allowing more precise structure determinations. To determine a protein structure, scientists measure how x-rays like those generated at NSLS-II diffract, or bounce off, the atoms in an ordered crystalline lattice consisting of many copies of the same protein molecule all arrayed the same way. The diffraction pattern conveys information about where the atoms are located. But it’s not sufficient.

A cartoon representing the structure of a well-studied plant protein that served as a test case for the newly developed microcrystallography technique. Magenta mesh patterns surrounding sulfur atoms intrinsic to the protein (yellow spheres) indicate the anomalous signals that were extracted using low-energy x-ray diffraction of thousands of crystals measuring less than 10 millionths of a meter, the size of a bacterium.

“Only the amplitudes of diffracted x-ray ‘waves’ are recorded on the detector, but not their phases (the timing between waves),” said Liu. “Both are required to reconstruct a 3-D structure. This is the so-called crystallographic phase problem.”

Crystallographers have solved this problem by collecting phase data from a different kind of scattering, known as anomalous scattering. Anomalous scattering occurs when atoms heavier than a protein’s main components of carbon, hydrogen, and nitrogen absorb and re-emit some of the x-rays. This happens when the x-ray energy is close to the energy those heavy atoms like to absorb. Scientists sometimes artificially insert heavy atoms such as selenium or platinum into the protein for this purpose. But sulfur atoms, which appear naturally throughout protein molecules, can also produce such signals, albeit weaker. Even though these anomalous signals are weak, a big crystal usually has enough copies of the protein with enough sulfur atoms to make them measurable. That gives scientists the phase information needed to pinpoint the location of the sulfur atoms and translate the diffraction patterns into a full 3-D structure.

“Once you know the sulfur positions, you can calculate the phases for the other protein atoms because the relationship between the sulfur and the other atoms is fixed,” said Liu.

But tiny crystals, by definition, don’t have that many copies of the protein of interest. So instead of looking for diffraction and phase information from repeat copies of a protein in a single large crystal, the Brookhaven/Columbia team developed a way to take measurements from many tiny crystals, and then assemble the collective data.

Tiny crystals, big results

To handle the tiny crystals, the team developed sample grids patterned with micro-sized wells. After pouring solvent containing the microcrystals over these well-mount grids, the scientists removed the solvent and froze the crystals that were trapped on the grids.

Micro-patterned sample grids for manipulation of microcrystals.

“We still have a challenge, though, because we can’t see where the tiny crystals are on our grid,” said Liu. “To find out, we used microdiffraction at NSLS-II’s Frontier Microfocusing Macromolecular Crystallography (FMX) beamline to survey the whole grid. Scanning line by line, we can find where those crystals are hidden.”

As Martin Fuchs, the lead beamline scientist at FMX, explained, “The FMX beamline can focus the full intensity of the x-ray beam down to a size of one micron, or millionth of a meter. We can finely control the beam size to match it to the size of the crystals—five microns in the case of the current experiment. These capabilities are crucial to obtain the best signal,” he said.

Wuxian Shi, another FMX beamline scientist, noted that “the data collected in the grid survey contains information about the crystals’ location. In addition, we can also see how well each crystal diffracts, which allows us to pick only the best crystals for data collection.”

The scientists were then able to maneuver the sample holder to place each mapped out microcrystal of interest back in the center of the precision x-ray beam for data collection.

They used the lowest energy available at the beamline—tuned to approach as closely as possible sulfur atoms’ absorption energy—and collected anomalous scattering data.

“Most crystallographic beamlines could not reach the sulfur absorption edge for optimized anomalous signals,” said co-author Wayne Hendrickson of Columbia University. “Fortunately, NSLS-II is a world-leading synchrotron light source providing bright x-rays covering a broad spectrum of x-ray energy. And even though our energy level was slightly above the ideal absorption energy for sulfur, it generated the anomalous signals we needed.”

But the scientists still had some work to do to extract those important signals and assemble the data from many tiny crystals.

“We are actually getting thousands of pieces of data,” said Liu. “We used about 1400 microcrystals, each with its own data set. We have to put all the data from those microcrystals together.”

Scientists used a five-micron x-ray beam at the FMX beamline at NSLS-II to scan the entire grid and locate the tiny invisible crystals. Then a heat map (green) was used to guide the selection of positions for diffraction data acquisition.

They also had to weed out data from crystals that were damaged by the intense x-rays or had slight variations in atomic arrangements.

“A single microcrystal does not diffract x-rays sufficiently for structure solution prior to being damaged by the x-rays,” said Sean McSweeney, deputy photon division director and program manager of the Structural Biology Program at NSLS-II. “This is particularly true with crystals of only a few microns, the size of about a bacterial cell. We needed a way to account for that damage and crystal structure variability so it wouldn’t skew our results.”

They accomplished these goals with a sophisticated multi-step workflow process that sifted through the data, discarded outliers that might have been caused by radiation damage or incompatible crystals, and ultimately extracted the anomalous scattering signals.

“This is a critical step,” said Liu. “We developed a computing procedure to assure that only compatible data were merged in a way to align the individual microcrystals from diffraction patterns. That gave us the required signal-to-noise ratios for structure determination.”

Applying the technique

This technique can be used to determine the structure of any protein that has proven hard to crystallize to a large size. These include cell-surface receptors that allow cells of advanced lifeforms such as animals and plants to sense and respond to the environment around them by releasing hormones, transmitting nerve signals, or secreting compounds associated with cell growth and immunity.

“To adapt to the environment through evolution, these proteins are malleable and have lots of non-uniform modifications,” said Liu. “It’s hard to get a lot of repeat copies in a crystal because they don’t pack well.”

In humans, receptors are common targets for drugs, so having knowledge of their varied structures could help guide the development of new, more targeted pharmaceuticals.

But the technique is not restricted to just small crystals.

“The method we developed can handle small protein crystals, but it can also be used for any size protein crystals, any time you need to combine data from more than one sample,” Liu said.

This research was supported in part by Brookhaven National Laboratory’s “Laboratory Directed Research and Development” program and the National Institutes of Health (NIH) grant GM107462. The NSLS-II at Brookhaven Lab is a DOE Office of Science user facility (supported by DE-SC0012704), with beamline FMX supported primarily by the National Institute of Health, National Institute of General Medical Sciences (NIGMS) through a Biomedical Technology Research Resource P41 grant (P41GM111244), and by the DOE Office of Science.

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One of ten national laboratories overseen and primarily funded by the Office of Science of the U.S. Department of Energy (DOE), Brookhaven National Laboratory conducts research in the physical, biomedical, and environmental sciences, as well as in energy technologies and national security. Brookhaven Lab also builds and operates major scientific facilities available to university, industry and government researchers. The Laboratory’s almost 3,000 scientists, engineers, and support staff are joined each year by more than 5,000 visiting researchers from around the world. Brookhaven is operated and managed for DOE’s Office of Science by Brookhaven Science Associates, a limited-liability company founded by Stony Brook University, the largest academic user of Laboratory facilities, and Battelle, a nonprofit, applied science and technology organization.

From Caltech: “From Beaker to Solved 3-D Structure in Minutes”

From Caltech

11/12/2018

Whitney Clavin
(626) 395-1856
wclavin@caltech.edu

Graduate student Tyler Fulton prepares samples of small molecules in a lab at Caltech. Credit: Caltech

Close-up of a powder containing small molecules like those that gave rise to 3-D structures in the new study. (The copper piece is a sample holder used with microscopes.) Credit: Caltech/Stoltz Lab

Brian Stoltz and Tyler Fulton. Credit: Caltech

UCLA/Caltech team uncovers a new and simple way to learn the structures of small molecules.

In a new study that one scientist called jaw-dropping, a joint UCLA/Caltech team has shown that it is possible to obtain the structures of small molecules, such as certain hormones and medications, in as little as 30 minutes. That’s hours and even days less than was possible before.

The team used a technique called micro-electron diffraction (MicroED), which had been used in the past to learn the 3-D structures of larger molecules, specifically proteins. In this new study, the researchers show that the technique can be applied to small molecules, and that the process requires much less preparation time than expected. Unlike related techniques—some of which involve growing crystals the size of salt grains—this method, as the new study demonstrates, can work with run-of-the-mill starting samples, sometimes even powders scraped from the side of a beaker.

“We took the lowest-brow samples you can get and obtained the highest-quality structures in barely any time,” says Caltech professor of chemistry Brian Stoltz, who is a co-author on the new study, published in the journal ACS Central Science. “When I first saw the results, my jaw hit the floor.” Initially released on the pre-print server Chemrxiv in mid-October, the article has been viewed more than 35,000 times.

The reason the method works so well on small-molecule samples is that while the samples may appear to be simple powders, they actually contain tiny crystals, each roughly a billion times smaller than a speck of dust. Researchers knew about these hidden microcrystals before, but did not realize they could readily reveal the crystals’ molecular structures using MicroED. “I don’t think people realized how common these microcrystals are in the powdery samples,” says Stoltz. “This is like science fiction. I didn’t think this would happen in my lifetime—that you could see structures from powders.”

This movie [animated in the full article] is an example of electron diffraction (MicroED) data collection, in which electrons are fired at a nanocrystal while being continuously rotated. Data from the movie are ultimately converted to a 3-D chemical structure. Credit: UCLA/Caltech

The results have implications for chemists wishing to determine the structures of small molecules, which are defined as those weighing less than about 900 daltons. (A dalton is about the weight of a hydrogen atom.) These tiny compounds include certain chemicals found in nature, some biological substances like hormones, and a number of therapeutic drugs. Possible applications of the MicroED structure-finding methodology include drug discovery, crime lab analysis, medical testing, and more. For instance, Stoltz says, the method might be of use in testing for the latest performance-enhancing drugs in athletes, where only trace amounts of a chemical may be present.

“The slowest step in making new molecules is determining the structure of the product. That may no longer be the case, as this technique promises to revolutionize organic chemistry,” says Robert Grubbs, Caltech’s Victor and Elizabeth Atkins Professor of Chemistry and a winner of the 2005 Nobel Prize in Chemistry, who was not involved in the research. “The last big break in structure determination before this was nuclear magnetic resonance spectroscopy, which was introduced by Jack Roberts at Caltech in the late ’60s.”

Like other synthetic chemists, Stoltz and his team spend their time trying to figure out how to assemble chemicals in the lab from basic starting materials. Their lab focuses on such natural small molecules as the fungus-derived beta-lactam family of compounds, which are related to penicillins. To build these chemicals, they need to determine the structures of the molecules in their reactions—both the intermediate molecules and the final products—to see if they are on the right track.

One technique for doing so is X-ray crystallography, in which a chemical sample is hit with X-rays that diffract off its atoms; the pattern of those diffracting X-rays reveals the 3-D structure of the targeted chemical. Often, this method is used to solve the structures of really big molecules, such as complex membrane proteins, but it can also be applied to small molecules. The challenge is that to perform this method a chemist must create good-sized chunks of crystal from a sample, which isn’t always easy. “I spent months once trying to get the right crystals for one of my samples,” says Stoltz.

Another reliable method is NMR (nuclear magnetic resonance), which doesn’t require crystals but does require a relatively large amount of a sample, which can be hard to amass. Also, NMR provides only indirect structural information.

Before now, MicroED—which is similar to X-ray crystallography but uses electrons instead of X-rays—was mainly used on crystallized proteins and not on small molecules. Co-author Tamir Gonen, an electron crystallography expert at UCLA who began developing the MicroED technique for proteins while at the Howard Hughes Medical Institute in Virginia, said that he only started thinking about using the method on small molecules after moving to UCLA and teaming up with Caltech.

“Tamir had been using this technique on proteins, and just happened to mention that they can sometimes get it to work using only powdery samples of proteins,” says Hosea Nelson (PhD ’13), an assistant professor of chemistry and biochemistry at UCLA. “My mind was blown by this, that you didn’t have to grow crystals, and that’s around the time that the team started to realize that we could apply this method to a whole new class of molecules with wide-reaching implications for all types of chemistry.”

The team tested several samples of varying qualities, without ever attempting to crystallize them, and were able to determine their structures thanks to the samples’ ample microcrystals. They succeeded in getting structures for ground-up samples of the brand-name drugs Tylenol and Advil, and they were able to identify distinct structures from a powdered mixture of four chemicals.

The UCLA/Caltech team says it hopes this method will become routine in chemistry labs in the future.

“In our labs, we have students and postdocs making totally new and unique molecular entities every day,” says Stoltz. “Now we have the power to rapidly figure out what they are. This is going to change synthetic chemistry.”

The study was funded by the National Science Foundation, the National Institutes of Health, the Department of Energy, a Beckman Young Investigators award, a Searle Scholars award, a Pew Scholars award, the Packard Foundation, the Sloan Foundation, the Pew Charitable Trusts, and the Howard Hughes Medical Institute. Other co-authors include Christopher Jones,Michael Martynowycz, Johan Hattne, and Jose Rodriguez of UCLA; and Tyler Fulton of Caltech.

See the full article here .

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The California Institute of Technology (commonly referred to as Caltech) is a private research university located in Pasadena, California, United States. Caltech has six academic divisions with strong emphases on science and engineering. Its 124-acre (50 ha) primary campus is located approximately 11 mi (18 km) northeast of downtown Los Angeles. “The mission of the California Institute of Technology is to expand human knowledge and benefit society through research integrated with education. We investigate the most challenging, fundamental problems in science and technology in a singularly collegial, interdisciplinary atmosphere, while educating outstanding students to become creative members of society.”

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From SLAC National Accelerator Lab: “Researchers create most complete high-res atomic movie of photosynthesis to date”

From SLAC National Accelerator Lab

November 7, 2018

Andrew Gordon
agordon@slac.stanford.edu
(650) 926-2282

In a major step forward, SLAC’s X-ray laser captures all four stable states of the process that produces the oxygen we breathe, as well as fleeting steps in between. The work opens doors to understanding the past and creating a greener future.

Using SLAC’s X-ray laser, researchers have captured the most complete high-res atomic movie to date of Photosystem II, a key protein complex in plants, algae and cyanobacteria responsible for splitting water and producing the oxygen we breathe. (Gregory Stewart, SLAC National Accelerator Laboratory)

Despite its role in shaping life as we know it, many aspects of photosynthesis remain a mystery. An international collaboration between scientists at SLAC National Accelerator Laboratory, Lawrence Berkeley National Laboratory and several other institutions is working to change that. The researchers used SLAC’s Linac Coherent Light Source (LCLS) X-ray laser to capture the most complete and highest-resolution picture to date of Photosystem II, a key protein complex in plants, algae and cyanobacteria responsible for splitting water and producing the oxygen we breathe. The results were published in Nature today.

SLAC/LCLS

Explosion of life

When Earth formed about 4.5 billion years ago, the planet’s landscape was almost nothing like what it is today. Junko Yano, one of the authors of the study and a senior scientist at Berkeley Lab, describes it as “hellish.” Meteors sizzled through a carbon dioxide-rich atmosphere and volcanoes flooded the surface with magmatic seas.

Over the next 2.5 billion years, water vapor accumulating in the air started to rain down and form oceans where the very first life appeared in the form of single-celled organisms. But it wasn’t until one of those specks of life mutated and developed the ability to harness light from the sun and turn it into energy, releasing oxygen molecules from water in the process, that Earth started to evolve into the planet it is today. This process, oxygenic photosynthesis, is considered one of nature’s crown jewels and has remained relatively unchanged in the more than 2 billion years since it emerged.

“This one reaction made us as we are, as the world. Molecule by molecule, the planet was slowly enriched until, about 540 million years ago, it exploded with life,” said co-author Uwe Bergmann, a distinguished staff scientist at SLAC. “When it comes to questions about where we come from, this is one of the biggest.”

A greener future

Photosystem II is the workhorse responsible for using sunlight to break water down into its atomic components, unlocking hydrogen and oxygen. Until recently, it had only been possible to measure pieces of this process at extremely low temperatures. In a previous paper, the researchers used a new method to observe two steps of this water-splitting cycle [Nature]at the temperature at which it occurs in nature.

Now the team has imaged all four intermediate states of the process at natural temperature and the finest level of detail yet. They also captured, for the first time, transitional moments between two of the states, giving them a sequence of six images of the process.

The goal of the project, said co-author Jan Kern, a scientist at Berkeley Lab, is to piece together an atomic movie using many frames from the entire process, including the elusive transient state at the end that bonds oxygen atoms from two water molecules to produce oxygen molecules.

“Studying this system gives us an opportunity to see how metals and proteins work together and how light controls such kinds of reactions,” said Vittal Yachandra, one of the authors of the study and a senior scientist at Berkeley Lab who has been working on Photosystem II for more than 35 years. “In addition to opening a window on the past, a better understanding of Photosystem II could unlock the door to a greener future, providing us with inspiration for artificial photosynthetic systems that produce clean and renewable energy from sunlight and water.”

Sample assembly line

For their experiments, the researchers grow what Kern described as a “thick green slush” of cyanobacteria — the very same ancient organisms that first developed the ability to photosynthesize — in a large vat that is constantly illuminated. They then harvest the cells for their samples.

At LCLS, the samples are zapped with ultrafast pulses of X-rays [Science] to collect both X-ray crystallography and spectroscopy data to map how electrons flow in the oxygen-evolving complex of photosystem II. In crystallography, researchers use the way a crystal sample scatters X-rays to map its structure; in spectroscopy, they excite the atoms in a material to uncover information about its chemistry. This approach, combined with a new assembly-line sample transportation system [Nature Methods], allowed the researchers to narrow down the proposed mechanisms put forward by the research community over the years.

Mapping the process

Previously, the researchers were able to determine the room-temperature structure of two of the states at a resolution of 2.25 angstroms; one angstrom is about the diameter of a hydrogen atom. This allowed them to see the position of the heavy metal atoms, but left some questions about the exact positions of the lighter atoms, like oxygen. In this paper, they were able to improve the resolution even further, to 2 angstroms, which enabled them to start seeing the position of lighter atoms more clearly, as well as draw a more detailed map of the chemical structure of the metal catalytic center in the complex where water is split.

This center, called the oxygen-evolving complex, is a cluster of four manganese atoms and one calcium atom bridged with oxygen atoms. It cycles through the four stable oxidation states, S0-S3, when exposed to sunlight. On a baseball field, S0 would be the start of the game when a player on home base is ready to go to bat. S1-S3 would be players on first, second, and third. Every time a batter connects with a ball, or the complex absorbs a photon of sunlight, the player on the field advances one base. When the fourth ball is hit, the player slides into home, scoring a run or, in the case of Photosystem II, releasing breathable oxygen.

The researchers were able to snap action shots of how the structure of the complex transformed at every base, which would not have been possible without their technique. A second set of data allowed them to map the exact position of the system in each image, confirming that they had in fact imaged the states they were aiming for.

In photosystem II, the water-splitting center cycles through four stable states, S0-S3. On a baseball field, S0 would be the start of the game when a batter on home base is ready to hit. S1-S3 would be players waiting on first, second, and third. The center gets bumped up to the next state every time it absorbs a photon of sunlight, just like how a player on the field advances one base every time a batter connects with a ball. When the fourth ball is hit, the player slides into home, scoring a run or, in the case of Photosystem II, releasing the oxygen we breathe. (Gregory Stewart/SLAC National Accelerator Laboratory)

Sliding into home

But there are many other things going on throughout this process, as well as moments between states when the player is making a break for the next base, that are a bit harder to catch. One of the most significant aspects of this paper, Yano said, is that they were able to image two moments in between S2 and S3. In upcoming experiments, the researchers hope to use the same technique to image more of these in-between states, including the mad dash for home — the transient state, or S4, where two atoms of oxygen bond together — providing information about the chemistry of the reaction that is vital to mimicking this process in artificial systems.

“The entire cycle takes nearly two milliseconds to complete,” Kern said. “Our dream is to capture 50-microsecond steps throughout the full cycle, each of them with the highest resolution possible, to create this atomic movie of the entire process.”

Although they still have a way to go, the researchers said that these results provide a path forward, both in unveiling the mysteries of how photosynthesis works and in offering a blueprint for artificial sources of renewable energy.

“It’s been a learning process,” said SLAC scientist and co-author Roberto Alonso-Mori. “Over the last seven years we’ve worked with our collaborators to reinvent key aspects of our techniques. We’ve been slowly chipping away at this question and these results are a big step forward.”

In addition to SLAC and Berkeley Lab, the collaboration includes researchers from Umeå University, Uppsala University, Humboldt University of Berlin, the University of California, Berkeley, the University of California, San Francisco and the Diamond Light Source.

Key components of this work were carried out at SLAC’s Stanford Synchrotron Radiation Lightsource (SSRL), Berkeley Lab’s Advanced Light Source (ALS) and Argonne National Laboratory’s Advanced Photon Source (APS). LCLS, SSRL, APS, and ALS are DOE Office of Science user facilities. This work was supported by the DOE Office of Science and the National Institutes of Health, among other funding agencies.

SLAC/SSRL

LBNL/ALS

ANL APS

See the full article here .

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SLAC is a multi-program laboratory exploring frontier questions in photon science, astrophysics, particle physics and accelerator research. Located in Menlo Park, California, SLAC is operated by Stanford University for the DOE’s Office of Science.