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Powerful ‘nanopore’ DNA sequencing method tackles proteins too

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Two gloves hands holding a MinION portable and real time device for DNA and RNA sequencing

A nanopore sequencing device is typically used for sequencing DNA and RNA.Credit: Anthony Kwan/Bloomberg/Getty

With its fast analyses and ultra-long reads, nanopore sequencing has transformed genomics, transcriptomics and epigenomics. Now, thanks to advances in nanopore design and protein engineering, protein analysis using the technique might be catching up.

“All the pieces are there to start with to do single-molecule proteomics and identify proteins and their modifications using nanopores,” says chemical biologist Giovanni Maglia at the University of Groningen, the Netherlands. That’s not precisely sequencing, but it could help to work out which proteins are present. “There are many different ways you can identify proteins which doesn’t really require the exact identification of all 20 amino acids,” he says, referring to the usual number found in proteins.

In nanopore DNA sequencing, single-stranded DNA is driven through a protein pore by an electrical current. As a DNA residue traverses the pore, it disrupts the current to produce a characteristic signal that can be decoded into a sequence of DNA bases.

Proteins, however, are harder to crack. They cannot be consistently unfolded and moved by a voltage gradient because, unlike DNA, proteins don’t carry a uniform charge. They might also be adorned with post-translational modifications (PTMs) that alter the amino acids’ size and chemistry — and the signals that they produce. Still, researchers are making progress.

Water power

One way to push proteins through a pore is to make them hitch a ride on flowing water, like logs in a flume. Maglia and his team engineered a nanopore1 with charges positioned so that the pore could create an electro-osmotic flow that was strong enough to unfold a full-length protein and carry it through the pore. The team tested its design with a polypeptide containing negatively charged amino acids, including up to 19 in a row, says Maglia. This concentrated charge created a strong pull against the electric field, but the force of the moving water kept the protein moving in the right direction. “That was really amazing,” he says. “We really did not expect it would work so well.”

Chemists Hagan Bayley and Yujia Qing at the University of Oxford, UK, and their colleagues have also exploited electro-osmotic force, this time to distinguish between PTMs2. The team synthesized a long polypeptide with a central modification site. Addition of any of three distinct PTMs to that site changed how much the current through the pore was altered relative to the unmodified residues. The change was also characteristic of the modifying group. Initially, “we’re going for polypeptide modifications, because we think that’s where the important biology lies”, explains Qing.

And, because nanopore sequencing leaves the peptide chain intact, researchers can use it to determine which PTMs coexist in the same molecule — a detail that can be difficult to establish using proteomics methods, such as ‘bottom up’ mass spectrometry, because proteins are cut into small fragments. Bayley and Qing have used their method to scan artificial polypeptides longer than 1,000 amino acids, identifying and localizing PTMs deep in the sequence. “I think mass spec is fantastic and provides a lot of amazing information that we didn’t have 10 or 20 years ago, but what we’d like to do is make an inventory of the modifications in individual polypeptide chains,” Bayley says — that is, identifying individual protein isoforms, or ‘proteoforms’.

Molecular ratchets

Another approach to nanopore protein analysis uses molecular motors to ratchet a polypeptide through the pore one residue at a time. This can be done by attaching a polypeptide to a leader strand of DNA and using a DNA helicase enzyme to pull the molecule through. But that limits how much of the protein the method can read, says synthetic biologist Jeff Nivala at the University of Washington, Seattle. “As soon as the DNA motor would hit the protein strand, it would fall off.”

Nivala developed a different technique, using an enzyme called ClpX (see ‘Read and repeat’). In the cell, ClpX unfolds proteins for degradation; in Nivala’s method, it pulls proteins back through the pore. The protein to be sequenced is modified at either end. A negatively charged sequence at one end allows the electric field to drive the protein through the pore until it encounters a stably folded ‘blocking’ domain that is too large to pass through. ClpX then grabs that folded end and pulls the protein in the other direction, at which point the sequence is read. “Much like you would pull a rope hand over hand, the enzyme has these little hooks and it’s just dragging the protein back up through the pore,” Nivala says.

Read and repeat. Graphic showing a nanopore protein-sequencing strategy using the push and pull of an electric field through a membrane, enzyme and slip sequence.

Source: Ref. 3

Nivala’s approach has another advantage: when ClpX reaches the end of the protein, a special ‘slip sequence’ causes it to let go so that the current can pull the protein through the pore for a second time. As ClpX reels it back out again and again, the system gets multiple peeks at the same sequence, improving accuracy.

Last October3, Nivala and his colleagues showed that their method can read synthetic protein strands of hundreds of amino acids in length, as well as an 89-amino-acid piece of the protein titin. The read data not only allowed them to distinguish between sequences, but also provided unambiguous identification of amino acids in some contexts. Still, it can be difficult to deduce the amino-acid sequence of a completely unknown protein, because an amino acid’s electrical signature varies on the basis of both its surrounding sequence and its modifications. Nivala predicts that the method will have a ‘fingerprinting’ application, in which an unknown protein is matched to a database of reference nanopore signals. “We just need more data to be able to feed these machine-learning algorithms to make them robust to many different sequences,” he says.

Stefan Howorka, a chemical biologist at University College London, says that nanopore protein sequencing could boost a range of disciplines. But the technology isn’t quite ready for prime time. “A couple of very promising proof-of-concept papers have been published. That’s wonderful, but it’s not the end.” The accuracy of reads needs to improve, he says, and better methods will be needed to handle larger PTMs, such as bulky carbohydrate groups, that can impede the peptide’s movement through the pore.

How easy it will be to extend the technology to the proteome level is also unclear, he says, given the vast number and wide dynamic range of proteins in the cell. But he is optimistic. “Progress in the field is moving extremely fast.”

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These ‘movies’ of proteins in action are revealing the hidden biology of cells

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Since the 1950s, scientists have had a pretty good idea of how muscles work. The protein at the centre of the action is myosin, a molecular motor that ratchets itself along rope-like strands of actin proteins — grasping, pulling, releasing and grasping again — to make muscle cells contract.

The basics were first explained in a pair of landmark papers in Nature1,2, and they have been confirmed and elaborated on by detailed molecular maps of myosin and its partners. Researchers think that myosin generates force by cocking back the long lever-like arm that is attached to the motor portion of the protein.

The only hitch is that scientists had never seen this fleeting pre-stroke state — until now.

In a preprint published in January3, researchers used a cutting-edge structural biology technique to record this moment, which lasts just milliseconds in living cells.

“It’s one of the things in the textbook you sort of gloss over,” says Stephen Muench, a structural biologist at the University of Leeds, UK, who co-led the study. “These are experiments that people wanted to do 40 years ago, but they just never had the technology.”

That technology — called time-resolved cryo-electron microscopy (cryo-EM) — now has structural biologists thinking like cinematographers, turning still snapshots of life’s molecular machinery into motion pictures that reveal how it works.

Muench and his colleagues’ myosin movie isn’t feature-length; it consists of just two frames showing different stages of the molecular motion. Yet it confirmed a decades-old theory and settled debates over the order of the steps in myosin’s choreography. Other researchers are focusing their new-found director’s eye on understanding cell-signalling systems, including those underlying opioid overdoses, the gene-editing juggernaut CRISPR–Cas9 and other molecular machines that have been mostly studied with highly detailed, yet static structural maps.

An animated gif showing a 3D molecular structures of a myosin molecule in two states using a lever arm to pull on an actin fillament

Researchers have been able to capture images of individual myosin proteins as they pull on an actin filament during muscle contraction, confirming key details of the motion. First, myosin becomes cocked or primed, then it attaches to actin and its lever arm swings in a power stroke that slides the filament by about 34 nanometres.Credit: Sean McMillan

“The big picture is to move away, as much as possible, from this single, static snapshot,” says Georgios Skiniotis, a structural biologist at Stanford University in California, whose team used the technique to record the activation of a type of cell-signalling molecule called a G-protein-coupled receptor (GPCR)4. “I want the movie.”

Freeze frame

To underscore the power of cryo-EM, Skiniotis and others like to draw a comparison with one of the first motion pictures ever made. In the 1870s, photographer Eadweard Muybridge used high-speed photography technology, which was cutting edge at the time, to capture a series of still images of a galloping horse. They showed, for the first time, that all four of the animal’s hooves leave the ground at once — something that the human eye could not distinguish.

Similar insights, Skiniotis says, will come from applying the same idea to protein structures. “I want to get a dynamic picture.”

The ability to map proteins and other biomolecules down to the location of individual atoms has transformed biology, underpinning advances in gene editing, drug discovery and revolutionary artificial-intelligence tools such as AlphaFold, which can predict protein structures. But the mostly static images delivered by X-ray crystallography and cryo-EM, the two technologies responsible for the lion’s share of determined protein structures, belie the dynamic nature of life’s molecules.

“Biomolecules are not made up of rocks,” says Sonya Hanson, a computational biophysicist at the Flatiron Institute in New York City. They exist in water and are constantly in motion. “They’re more like jelly,” adds Muench.

Biologists often say that “structure determines function”, but that’s not quite right, says Ulrich Lorenz, a molecular physicist at the Swiss Federal Institute of Technology in Lausanne (EPFL). The protein poses captured by most structural studies are energetically stable ‘equilibrium’ states that provide limited clues to the short-lived, unstable confirmations that are key to chemical reactions and other functions performed by molecular machines. “Structure allows you to infer function, but only incompletely and imperfectly, and you’re missing all of the details,” says Lorenz.

Cryo-EM is a great way to get at the details, but capturing these fleeting states requires careful preparation. Protein samples are pipetted onto a grid and then flash frozen with liquid ethane. They are then imaged using powerful electron beams that record snapshots of individual molecules (sophisticated software classifies and morphs these pictures into structural maps). The samples swim in water before being frozen, so any chemical reaction that can happen in a test tube can, in theory, be frozen in place on a cryo-EM grid — if researchers can catch it quickly enough.

That’s one of the first big challenges says Joachim Frank, a structural biologist at Columbia University in New York City who shared the 2017 Nobel Prize in Chemistry for his work on cryo-EM. “Even for very dexterous people, it takes a few seconds.” In that time, any chemical reactions — and the intermediate structures that mediate the reactions — might be long gone before freezing. “This is the gap we want to fill,” says Frank.

Caught in translation

Frank’s team has attempted to solve this problem using a microfluidic chip. The device quickly mixes two protein solutions, allows them to react for a specified time period and then delivers reaction droplets onto a cryo-EM grid that is instantly frozen.

This year, Frank’s team used their device to study a bacterial enzyme that rescues ribosomes, the cell’s protein-making factories, if they stall in response to antibiotics or other stresses. The enzyme, called HflX, helps to recycle stuck ribosomes by popping their two subunits apart.

Frank’s team captured three images of HflX bound to the ribosome, over a span of 140 milliseconds, which show how it splits the ribosome like someone carefully removing the shell from an oyster. The enzyme breaks a dozen or so molecular bridges that hold a ribosome’s two subunits together, one by one, until just two are left and the ribosome pops open5. “The most surprising thing to me is that it’s a very orderly process,” Frank says. “You would think the ribosome is being split and that’s it.”

Muench and his colleagues, including Charlie Scarff, a structural biologist at the University of Leeds, and Howard White, a kineticist at Eastern Virginia Medical School in Norfolk, Virginia also used a microfluidic chip to make their myosin movie by quickly mixing myosin and actin3.

But the molecular motor is so fast that, to slow things down ever further, they used a mutated version of myosin that operates about ten times slower than normal. This allowed the team to determine two structures, 110 milliseconds apart, that showed the swing of myosin’s lever-like arm. The structures also showed that a by-product of the chemical reaction that powers the motor — the breakdown of a cellular fuel called ATP — exits the protein’s active site before the lever swings and not after. “That is ending decades of conjecture,” says Scarff.

With this new model in mind, Scarff, whose specialty is myosin, and Muench are planning to use time-resolved cryo-EM to study how myosin dynamics are affected by certain drugs and mutations that are known to cause heart disease.

Microfluidic chips aren’t the only way researchers are putting time stamps on protein structures. A team led by Bridget Carragher, a structural biologist and the technical director at the Chan Zuckerberg Imaging Institute in Redwood City, California, developed a ‘spray and mix’ approach that involves shooting tiny volumes of reacting samples onto a grid before flash-freezing them6.

In another set-up — developed by structural physiologist Edward Twomey at Johns Hopkins University School of Medicine in Baltimore, Maryland, and his team — a flash of light triggers light-sensitive chemical reactions, which are stopped by flash-freezing7. Lorenz’s kit, meanwhile, takes already frozen samples and uses laser pulses to reanimate them for a few microseconds before they refreeze, all under the gaze of an electron microscope8.

‘Limitations everywhere’

The different approaches have their pros and cons. Carragher’s spray and mix approach uses minute sample volumes, which should be easy to obtain for most proteins; Twomey says his ‘open-source’ light-triggered device is relatively inexpensive and can be built for a few thousand dollars; and Lorenz says his laser-pulse system has the potential to record many more fleeting events than other time-resolved cryo-EM technologies — down to a tenth of a microsecond.

But these techniques are not yet ready to be rolled out. Currently, there are no commercial suppliers of time-resolved cryo-EM technology, limiting its reach, says Rouslan Efremov, a structural biologist at the VIB-VUB Center for Structural Biology in Brussels. “All these things are fussy and hard to control and they haven’t really caught on,” adds Carragher.

Holger Stark, a structural biologist at the Max Planck Institute for Multidisciplinary Sciences in Göttingen, Germany, says that current forms of time-resolved cryo-EM might be useful for some molecular machines that operate on the basis of large-scale movements — for example, the ribosome. However, the technology is not ready for use on just any biological system. “You have to cherry pick your subject,” he says. “We have limitations everywhere.”

Despite the shortcomings, there are plenty of interesting questions for researchers to start addressing now using these techniques. Twomey is using time-resolved cryo-EM to study Cas9, the DNA-cutting enzyme behind CRISPR gene editing, and says the insights could help to make more efficient gene-editing systems.

Lorenz used his laser-melting method to show how a plant virus swells up after it infects a cell to release its genetic material7 (see ‘Viral blow-up’). He is now studying other viral entry molecules such as HIV’s envelope protein. “We have these static structures, but we don’t know how the system makes it from one state to the other, and how the machinery works,” he says.

VIRAL BLOW UP: infographic showing a viral capsid from contracted to expanded states.

Source: Ref.8

Skiniotis’s team is investigating GPCRs, including one called the β-adrenergic receptor, which has been implicated in asthma. Their work4 shows how activating the receptor triggers it to shed its partner G-protein, a key step in propagating signals in cells.

The researchers are now studying the same process in a GPCR called the µ-opioid receptor, which is activated by morphine and fentanyl among other drugs. In preliminary unpublished results, they have found that the dynamics of the receptor help to explain why some drugs such as fentanyl are so potent in promoting G-protein activation, while others aren’t. Such insights, says Skiniotis, are glimpses of unseen biology that molecular movies promise to reveal. Just don’t forget the popcorn.

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Could AI-designed proteins be weaponized? Scientists lay out safety guidelines

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AlphaFold structure prediction for probable disease resistance protein At1g58602.

The artificial-intelligence tool AlphaFold can design proteins to perform specific functions.Credit: Google DeepMind/EMBL-EBI (CC-BY-4.0)

Could proteins designed by artificial intelligence (AI) ever be used as bioweapons? In the hope of heading off this possibility — as well as the prospect of burdensome government regulation — researchers today launched an initiative calling for the safe and ethical use of protein design.

“The potential benefits of protein design [AI] far exceed the dangers at this point,” says David Baker, a computational biophysicist at the University of Washington in Seattle, who is part of the voluntary initiative. Dozens of other scientists applying AI to biological design have signed the initiative’s list of commitments.

AI tools are designing entirely new proteins that could transform medicine

“It’s a good start. I’ll be signing it,” says Mark Dybul, a global health policy specialist at Georgetown University in Washington DC who led a 2023 report on AI and biosecurity for the think tank Helena in Los Angeles, California. But he also thinks that “we need government action and rules, and not just voluntary guidance”.

The initiative comes on the heels of reports from US Congress, think tanks and other organizations exploring the possibility that AI tools — ranging from protein-structure prediction networks such as AlphaFold to large language models such as the one that powers ChatGPT — could make it easier to develop biological weapons, including new toxins or highly transmissible viruses.

Designer-protein dangers

Researchers, including Baker and his colleagues, have been trying to design and make new proteins for decades. But their capacity to do so has exploded in recent years thanks to advances in AI. Endeavours that once took years or were impossible — such as designing a protein that binds to a specified molecule — can now be achieved in minutes. Most of the AI tools that scientists have developed to enable this are freely available.

To take stock of the potential for malevolent use of designer proteins, Baker’s Institute of Protein Design at the University of Washington hosted an AI safety summit in October 2023. “The question was: how, if in any way, should protein design be regulated and what, if any, are the dangers?” says Baker.

AlphaFold touted as next big thing for drug discovery — but is it?

The initiative that he and dozens of other scientists in the United States, Europe and Asia are rolling out today calls on the biodesign community to police itself. This includes regularly reviewing the capabilities of AI tools and monitoring research practices. Baker would like to see his field establish an expert committee to review software before it is made widely available and to recommend ‘guardrails’ if necessary.

The initiative also calls for improved screening of DNA synthesis, a key step in translating AI-designed proteins into actual molecules. Currently, many companies providing this service are signed up to an industry group, the International Gene Synthesis Consortium (IGSC), that requires them to screen orders to identify harmful molecules such as toxins or pathogens.

“The best way of defending against AI-generated threats is to have AI models that can detect those threats,” says James Diggans, head of biosecurity at Twist Bioscience, a DNA-synthesis company in South San Francisco, California, and chair of the IGSC.

Risk assessment

Governments are also grappling with the biosecurity risks posed by AI. In October 2023, US President Joe Biden signed an executive order calling for an assessment of such risks and raising the possibility of requiring DNA-synthesis screening for federally funded research.

Baker hopes that government regulation isn’t in the field’s future — he says it could limit the development of drugs, vaccines and materials that AI-designed proteins might yield. Diggans adds that it’s unclear how protein-design tools could be regulated, because of the rapid pace of development. “It’s hard to imagine regulation that would be appropriate one week and still be appropriate the next.”

But David Relman, a microbiologist at Stanford University in California, says that scientist-led efforts are not sufficient to ensure the safe use of AI. “Natural scientists alone cannot represent the interests of the larger public.”

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