Tag: Developmental biology

Embryonic development is like an elaborate stage production, in which meticulously choreographed gene-expression programmes organize much of the action. But changes in physical forces such as tension and mechanical properties like elasticity also play a part, coordinating the process’s spatial and temporal footwork.

“Mechanical properties actually become relevant to formation of the first cell lineages before gene expression,” says Jean-Léon Maître, a mechanobiologist at the Curie Institute in Paris. After fertilization, subsequent rounds of cell division produce extensive rearrangements and deformations. The resulting squeezing, stretching, pushing and pulling influences which cells form different embryonic features, and ultimately guides the patterning and development of every tissue in the body. “You first have this difference in mechanics, which then translates to a difference in position, which then translates into fate,” says Maître.

Collection: Mechanobiology

Measuring these properties and how they change over time is therefore essential to understanding embryonic development. But making such observations isn’t easy. Many biophysical and mechanobiology techniques are best suited to artificial systems with cultured cells. Promising techniques are emerging for in vivo studies, but these require further testing to prove their accuracy and quantitative mettle. “All of it is still relatively hard,” says Amy Shyer, a mechanobiologist who co-directs a laboratory at Rockefeller University in New York City with developmental biologist Alan Rodrigues. “There isn’t an off-the-shelf toolkit.” The challenges only mount with live embryos, in which the goal is to measure physical and mechanical phenomena without disrupting development.

But a steady rise in interest in mechanobiology is leading to technologies that offer exciting opportunities to establish a more holistic view of developmental biology. “We’re just at that moment where the field can take off,” says Otger Campàs, a biological physicist at the Technical University Dresden in Germany.

Cells in living tissues have multiple levers that they can pull to reorganize themselves and physically influence their neighbours. The cytoskeleton is a network of proteins that helps to define the shape and organization of cells. By rearranging that internal structure, cells can push and pull and even travel over or between one another, and alter their own mechanical properties, transitioning between being squishy and fluid or firm and viscous.

Under pressure

Furthermore, cell division can contribute to crowding and create changes in embryonic surface tension that are especially important early in development. “To position these cells in a certain place, you need to actually control the way they divide,” says Hervé Turlier, a biophysicist at the College of France in Paris. The early embryo, he notes, starts as a seemingly homogeneous ball of cells; the directionality of cell division helps to disrupt this symmetry and establish the axes that determine an organism’s front, back, top and bottom. One 2016 study, for instance, showed that inward-directed forces associated with certain early cell-division events help to separate cells that form the mouse embryo from those that give rise to extra-embryonic tissues such as the yolk sac¹.

A measure of molecular muscle

Which method is chosen to investigate these processes depends, in part, on the experimental model being studied. Mammalian embryos, such as mice, can be challenging because they generally cannot be maintained outside the mother’s body beyond the stage at which uterine implantation occurs, roughly four days post-fertilization. Moreover, each cell-division event takes place over many hours, requiring long-term experiments. Model organisms such as the roundworm Caenorhabditis elegans offer a simpler alternative. But with their ultrafast development and cycles of cell division that are separated by minutes rather than hours, some methods have trouble keeping up. This frenetic pace also introduces frictional forces that are absent in more slowly dividing embryos, Maître says, making it challenging to model the organism’s mechanical properties.

The good news, says developmental biologist Carl-Philipp Heisenberg at the Institute of Science and Technology Austria in Klosterneuburg, is that many fundamental mechanisms are conserved across species, at least in early development. “The general principles you’re looking at — how cells are interacting, how mechanical forces are being generated, transmitted and sensed — they’re being used again and again in different organisms,” he says.

Full contact

Certain mechanobiology tools are also being used again and again, and two in particular have been popular for decades.

Atomic force microscopy (AFM) was developed in the 1980s as a way to probe the surface texture and stiffness of materials with atomic-scale precision. AFM uses a tiny, cantilevered probe, like a diving board, that is dragged across a sample surface and illuminated with a laser. Changes in the reflected light can be used to calculate tissue stiffness and elasticity, as well as the force with which two cells are attached.

The other approach, micropipette aspiration, dates to the 1950s, when it was used to study sea urchin embryos. Researchers place the tip of an extremely fine glass pipette against a cell surface, then use a pump to draw the membrane part way into the pipette to test membrane surface tension and cell viscosity, or to push air against it to apply a controlled force. With multiple pipettes, researchers can play ‘tug of war’ with adjoining cells to determine how tightly they are coupled. The result is an absolute, rather than relative, measure of force, says Chii Jou Chan, a mechanobiologist at the National University of Singapore.

Maître’s lab has used micropipette aspiration to extensively map surface tension in the earliest stages of mouse development, when embryos are still relatively simple balls of cells. In a study published in May, he and his colleagues applied the technique to study compaction — the process by which early embryos establish tighter couplings between cells — in human embryos². The researchers identified mechanical properties that could inform the selection of viable embryos for reproductive technologies, such as in vitro fertilization.

AFM, in contrast, has limited utility in live embryos. “You need something flat,” explains Shyer. But the technique can deliver important insights when applied to tissues that have been removed from developing embryos and studied in culture. In 2023 study, Shyer and Rodrigues used techniques including AFM and micropipette aspiration to measure how different morphogens — molecules that act as developmental signals —affect the mechanical properties of developing skin and follicles in chicken embryos³. They found that the morphogen-induced biophysical changes that precede follicle formation occur not at the single-cell level as was previously thought, but on the scale of large cell groups. “It was sort of the first time measuring emergent physical properties at this collective cell scale,” says Shyer, noting that most biomechanical studies focus on individual cells.

That said, many aspects of development can be explored only with whole embryos. “If we care about the physics of how embryonic structures are formed, then we need to measure physical fields inside these structures as they’re being formed,” says Campàs.

Inside perspective

The toolbox for such work has been limited, however. Alongside micropipette aspiration, for instance, many researchers evaluated surface tension by using ultraviolet lasers to introduce targeted cuts on the embryo surface. “You observe sort of a recoil of the tissue which you have cut, and this recoil velocity is proportional to the tension in this tissue,” explains Heisenberg. But the embryo is damaged in the process, and the results are difficult to quantify because the strength of recoil is inherently dependent on the — potentially unknown — mechanical properties of the affected cells.

Now, other promising methods are emerging. The Campàs lab has developed oil-encapsulated droplet sensors that can be injected into live embryos and then imaged. In the original iteration of these sensors, the droplets are filled with fluorescent dye, and the stresses they experience at a given injection site are calculated on the basis of how much they deform. This approach has proven especially valuable in fast-developing models such as zebrafish. “You can image an entire organ forming in just a few hours,” says Campàs. But it is also suitable for mammalian development. In 2022, for instance, Campàs and his colleagues used this approach to document how the gradual build-up of internal physical stress governs the initiation and extent of toe growth in embryonic mice⁴. Mutant mice that lack a morphogen that contributes to this stress build-up, developed only short nubs rather than functional digits.

Campàs’s team has also developed more sophisticated microdroplet sensors. One version incorporates an iron-based ‘ferrofluid’ that can be manipulated with a magnetic field⁵. This enables researchers to apply controlled forces at the injection site for extended periods while also measuring the material properties of the surrounding tissue. Another iteration features a water droplet encased in an oil droplet⁶. When this droplet in a droplet is subject to increased osmotic pressure — for example, if a cell or tissue becomes saturated — it takes up more fluid, producing a measurable change in volume. Campàs says that these sensors could be useful for understanding how channels form in hollow organ structures, such as the airways of the lung or in pancreatic ducts.

Optical tweezers, which use focused lasers to physically ‘trap’ and manipulate nanoscale objects, can also be used to push and pull cellular membranes and organelles in vivo. Pierre-François Lenne at Aix Marseille University in France has used this technology to test the strength of cell–cell junctions in live fruit fly embryos. “He traps the junction, and then he would basically just play the guitar with it,” says Maître. Such experiments have historically required complex, home-built apparatus, but commercial platforms are now available, and Maître is using the method in his own lab to test how the material properties of cell and nuclear membranes change as embryos grow. “It’s a very promising tool,” he says.

A hands-off approach

That said, the same sensing mechanisms that allow cells to follow force-induced prompts can also react to being poked with a pipette tip, Heisenberg cautions, and even relatively gentle interventions such as the injection of microdroplets could elicit a response. “You’re putting it into a tissue, and the tissue knows that there is something [there] which doesn’t belong,” says Heisenberg.

Researchers are, therefore, keen to develop contact-free alternatives. One possibility is to computationally deduce the forces that individual cells experience on the basis of how much they deform at junctions with other cells. “The surface is what mostly controls the shape of cells,” explains Turlier. “It’s not things pushing from inside, it’s really the surface that deforms itself.” Given sufficient starting information, researchers could, in principle, map the force landscape of an embryo from image analysis alone.

Many tools for force inference are best suited to flat sheets of epithelial cells, but Turlier’s group has been making headway in 3D force inference using an algorithm called ‘foambryo’⁷. Published in 2023, foambryo models embryos as foams of bubble-like cells, an assumption that seems to be broadly applicable across embryonic systems in terms of the general shape and arrangement of cells. Foambryo has important limitations, Turlier notes — for example, the method provides relative rather than absolute force measurements, and accounts for only a subset of the tensions that embryonic cells are subject to. Still, he says, “I think what we’ve done here is important, because even if it’s not perfect, it will give us a good first guess.”

In another emerging non-invasive method, Brillouin microscopy, samples are scanned with a laser beam in a modified fluorescence microscope. The movement of biomolecules in the scanned tissue influences the extent to which that light is scattered, and measurements of this optical scatter can then be used to calculate mechanical properties associated with cellular stiffness and elasticity.

Brillouin microscopy: an emerging tool for mechanobiology

Jitao Zhang, a biomedical engineer at Wayne State University in Detroit, Michigan, is among the method’s champions and has been using Brillouin microscopy to study essential processes in development that can contribute to life-threatening congenital conditions if derailed, such as spina bifida. “We’ve applied this optical method to chick embryos to monitor how the stiffness changes when the neural tube is closing,” says Zhang. Brillouin allowed his team to document this process over more than 21 hours, measuring a steady increase in tissue stiffness as the neural-tube tissue thickens, bends and closes⁸. Chan has also found Brillouin microscopy to be a valuable tool. “I think it’s a perfect system to study intrafollicular mechanical stiffness,” he says.

Nevertheless, there are open questions about the utility of Brillouin microscopy. The mechanical measurements it enables are influenced by the refractive index and density of the tissue being imaged, and these parameters might not be well defined. “It’s based on assumptions which are sometimes experimentally very difficult to prove,” says Heisenberg. Methods are emerging that make it possible to measure the optical properties of complex biological samples, Chan notes. And although AFM and Brillouin measure different mechanical properties, the two seem to correlate in many biological systems. Cross-validation is therefore important for interpreting Brillouin data, and Zhang says that his group is careful to fact-check its Brillouin data using AFM. “You have to do sample-dependent calibration — that’s the painful part,” he says. His team is setting up a dedicated facility in which both methods can be carried out on the same sample in parallel.

Filling in the gaps

Indeed, when it comes to mechanobiology, even the most tried-and-tested methods present uncertainties. “You’re dealing with a living material,” says Rodrigues. “There’s a little bit of trickiness there because you have to try to build tools about a substance that you don’t fully even understand.”

NatureTech hub

That means any study will require assumptions and models that are based on incomplete information about parameters that might vary considerably between different embryos and over time. For example, Turlier notes that incorrect assumptions about embryonic cell geometry have led to widely divergent results using AFM. It’s not unheard of, he says, for repeated measurements on the same cell to differ by as much as 700%. More in vivo data should prove valuable on this front, he adds, generating values that can be used to build and calibrate more accurate models.

And then there’s the challenge of tying these mechanical phenomena back to the underlying genetic and biomolecular processes that elicit them. Studies like those of Shyer and Rodrigues in avian skin demonstrate the feasibility of drawing explicit connections between specific morphogens or other signalling prompts and large-scale changes in mechanical properties and force maps. Rodrigues is enthusiastic about the opportunity to close the circle here, and perhaps derive more comprehensive explanations for the root causes of poorly understood developmental disorders. “Understanding biophysics at this level”, he says, “could allow us to actually make sense of a lot of genetic information that we haven’t been able to organize.”

Fluorescence microscopy is one of the most powerful tools in the life-sciences toolkit. The unique spectral properties of fluorescent dyes and proteins, collectively called fluorophores, have allowed scientists to capture snapshots — and even movies — of the microscopic universe that makes biology tick. Yet scientists are still perplexed by the fundamental chemistry that keeps the lights on.

All fluorophores work on the same principle: energy absorbed from an incoming photon excites the molecule to a higher energetic state. As the fluorophore ‘relaxes’, it dissipates energy by emitting a new photon of a different (longer) wavelength. The more efficiently a fluorophore absorbs photons and radiates light, the stronger its fluorescent signal.

The problem is that fluorophores can’t perform this light show forever: they dim over time in a phenomenon called photobleaching. Photobleaching occurs more rapidly with high-intensity light, and after repeated rounds of excitation and in oxygen-rich environments. This can end experiments prematurely and sometimes generates toxic by-products that can kill cells. Some of the earliest dyes would fade in seconds under bright light, whereas many of today’s fluorophores can stay bright for minutes or more under the right conditions.

Photobleaching has become an irritating puzzle in the world of biological imaging, but it’s neither glamorous nor easy to solve. Much remains unknown about the chemistry that drives or prevents it, says Luke Lavis, an organic chemist at the Howard Hughes Medical Institute’s Janelia Research Campus in Ashburn, Virginia. Strategies that fortify a fluorophore against bleaching might inadvertently affect other spectral characteristics. And changes that improve one fluorophore can be difficult to apply to others. “It’s the sort of thing that people just live with rather than really trying to study,” says Adam Cohen, a neuroscientist at Harvard University in Cambridge, Massachusetts.

Turning on the lights

Microscopists have a vast palette of dyes and fluorescent proteins at their disposal. Microscope vendor Leica Microsystems, based in Wetzlar, Germany, lists more than 250 fluorescent dyes for life-sciences imaging, and the FPbase fluorescent protein database includes nearly 1,100 entries. Fluorescent dyes are descendants of the nineteenth-century dye industry, when chemists were first learning to make small organic molecules such as fluorescein, rhodamine and cyanine dyes. Fluorescent proteins stem from the discovery in 1962 that certain jellyfish fluoresce because they contain a family of proteins in which a coloured molecule called a chromophore, built from amino acids, is encased in a protective barrel formed from a type of protein structure called β-pleated sheets (corals were subsequently also found to contain the same protein family).

The hunt for red fluorescent proteins

Whatever their chemical nature, these molecules are driven by the same photophysical principles. When a molecule is illuminated by incoming light, the energy from the absorbed photon excites electrons in the material. The fluorophore can’t hold on to that energy forever — the time it spends in the excited state is called the fluorescence lifetime. Eventually it must relax, ideally by offloading that pent-up energy as radiated photons: one photon is released for every photon absorbed.

In practice, excited electrons find myriad non-radiative pathways to return to their ground state — their normal energy level — without emitting a photon. The fluorophore might lose energy through collisions with other molecules or through vibrations in its own bonds, for instance. Off-target photoreactions might generate toxic reactive oxygen species or even destroy the fluorophore. In both cases, fluorescence is temporarily or permanently snuffed out, and quantum yield — the ratio of fluorescence out to excitation light in — suffers.

But it’s rarely clear why that is. Scientists have struggled to isolate bleached fluorophores to do a chemical post-mortem, and it’s nearly impossible to predict all of the non-radiative pathways with which fluorescence competes. But the inevitability of photobleaching means that microscopists have only a limited ‘photon budget’ with which to work, and they must spend it carefully before the fluorophore — and their experiment — goes dark.

One way to protect that return on investment is by shutting down the competition. Researchers have long noted a correlation between a fluorophore’s brightness and the flexibility of its chemical backbone, says Ralph Jimenez, a physical chemist at the University of Colorado Boulder. Stiff molecules tend to have longer fluorescence lifetimes, which corresponds to higher quantum yield, he says.

Smart microscopes spot fleeting biology

“A lot of synthetic chemists tried to make fluorophores brighter by making them very rigid so that basically nothing could wiggle any more,” says chemical biologist Michelle Frei at the Swiss Federal Institute of Technology in Zurich. The drawback was that these ‘rigidified’ molecules were more likely to get trapped in cell membranes before reaching their target, making them less useful in cellular assays, Frei says.

Rigidification has benefits for fluorescent proteins, too, says Dorus Gadella, a molecular scientist at the University of Amsterdam. The chromophore — which is highly conserved across fluorescent proteins — consists of two rings bridged by a methylene group. Yet this molecule does not fluoresce when excited in isolation, Gadella says, because it undergoes a molecular rearrangement known as cis–trans isomerization — a non-radiative pathway that diverts energy from emitting photons. The protein’s β-barrel rigidifies the chromophore and prevents that rearrangement from happening, he says.

Better fluorescent proteins

Working with biochemist Amy Palmer, also at the University of Colorado Boulder, Jimenez and his colleagues developed a bright red fluorescent protein called mCherry-XL, a variant of the popular mCherry. They did so by rigidifying the protein barrel and locking down a squiggly amino-acid side chain that was hampering fluorescence¹; this helped to increase the fluorescence lifetime from 1.6 nanoseconds for mCherry to 3.9 nanoseconds for mCherry-XL. The trade-off is that the more time a fluorophore spends in its excited state, the more likely it is to become damaged, Jimenez says. As a result, mCherry-XL is three times brighter than mCherry, but less photostable.

Too much rigidity, however, can interfere with the protein’s ability to build its protective barrel and perform the multi-step, oxygen-dependent reaction that forms its chromophore, which can slow down the protein’s maturation or quash its fluorescence. A red fluorescent protein developed by Gadella and his colleagues, called mScarlet3, nicely illustrates this balancing act.

According to Gadella, their protein represents a sweet spot in stiffening the barrel so that the chromophore can’t isomerize, but still leaves enough wiggle room for the protein to fold and mature. Thanks in part to its fast maturation rate, mScarlet3 is the brightest red fluorescent protein so far — more than five times brighter than mCherry — and it has a fluorescence lifetime of 4 nanoseconds². But, much like mCherry-XL, mScarlet3 photobleaches faster than its precursors do under intense illumination.

Theoretical chemist Anna Krylov at the University of Southern California in Los Angeles, is developing software that can simulate excited-state processes in fluorophores to enhance their fluorescent properties. By identifying and shutting down non-radiative pathways, the thinking goes, fluorophores can be improved so they are fit for experimental purposes.

Researchers have observed, for instance, that photo-oxidation of the green fluorescent protein EGFP can turn its output from green to red. This behaviour is not observed in the closely related yellow fluorescent protein EYFP, despite both proteins having the same chromophore structure. Using their simulations, in 2016 Krylov and her team uncovered a non-radiative pathway in EGFP through which an electron can hop from the chromophore to a tyrosine residue in the protein barrel and thence to an electron acceptor outside the protein molecule³. That electron-transfer mechanism alters the photophysical properties of the EGFP chromophore, turning it red. In EYFP, however, the comparable tyrosine is chemically out of reach from the chromophore, preventing that electron transfer.

Powerful microscope captures motor proteins in unprecedented detail

The calculations suggested that the team could suppress the non-radiative pathway in EGFP by swapping the tyrosine for leucine, which lacks tyrosine’s electron-hungry oxygen atom. “We were able to propose specific mutations to our experimental collaborators and were able to get a more photostable variant of this protein,” Krylov says. In fact, the resulting mutant was 80 times more photostable than EGFP, but was also dimmer. Although this mutant protein is not suitable for use in biological imaging, it demonstrates how a better understanding of fluorophore photophysics can lead to rational engineering of bespoke materials, Krylov says.

That said, serendipity and random mutagenesis still play an outsized part in the development of fluorescent proteins, Jimenez says. Atsushi Miyawaki, a biochemist at the RIKEN Center for Brain Science near Tokyo, and his team developed what could be the most photostable fluorescent protein so far. They discovered a remarkably photostable, but dim, green fluorescent protein in Cytaeis uchidae, a small jellyfish found along the Japanese coast.

Through random mutagenesis, the researchers found that they could improve the protein’s brightness without compromising photostability by changing a single amino acid from valine to alanine, which increased the protein’s folding and maturation efficiency⁴. Called StayGold, this mutant has a nearly identical structure to the original protein, yet emits ten times more photons before photobleaching than any previous green fluorescent protein, allowing the team to record the dynamic rearrangement of intricate organelle networks in live cells for up to six minutes⁵.

New chemistry

Fluorescent proteins and dyes each have their advantages. Proteins can be expressed in specific cell types, for instance, whereas dyes, as small molecules, are brighter and more easily fine-tuned through synthesis. Now, researchers have developed a fluorophore system that combines the best of both, according to Frei. Self-labelling proteins, such as the commercially available SNAP-tag and HaloTag reagents, work by embedding an enzymatic protein tag in a target of interest. This tagged protein can then be coupled to any of several small fluorescent molecules supplied by the researcher. For Frei, this design provides a platform for fluorescence lifetime imaging (FLIM), a photon-intensive method that differentiates fluorophores by how long they spend in the excited state rather than by the wavelength they emit.

As a doctoral student at the Max Planck Institute for Medical Research in Heidelberg, Germany, Frei created a panel of HaloTag variants that included mutations in the dye docking site. Mutations that helped to hold the fluorophore in place, she saw, extended fluorescence lifetimes — even though the fluorescent dye molecule itself was unchanged. Exploiting this finding, Frei engineered cells to express different self-labelling HaloTag proteins, so they could be treated with the same rhodamine dye but would elicit measurably different signals for each target⁶. Whereas conventional fluorescence microscopy experiments can handle only one dye per colour channel, Frei and her team were able to visualize three targets in a single-channel experiment by adding FLIM to the mix.

To take full advantage of such self-labelling systems, however, fluorescent dyes would need an upgrade. Dye chemistry has changed little since the nineteenth century and requires harsh conditions that are ill-suited to building highly tailored molecules. In a 2023 review, Lavis and Martin Schnermann at the US National Cancer Institute in Frederick, Maryland, argued that chemists could rejuvenate classic dye scaffolds for next-generation microscopy by applying modern synthetic techniques⁷.

Take the classic red dye tetramethylrhodamine (TMR), for example. Lavis and his colleagues reasoned that rotation of the bond between its carbon skeleton and a structure called dimethylamine in the excited state might drive electron transfer to a non-radiative pathway. Chemists have conventionally suppressed that pathway by making rhodamine dyes in which these nitrogen-containing groups are locked in place with clunky fused-ring systems, which improved brightness but hindered permeability when the dye was applied to cells. Then, in 2015, Lavis and his collaborators demonstrated⁸ that a palladium-catalysed reaction could be used to install nitrogen-containing rings of different sizes in place of the original dimethylamine. Lavis’s team used these comparatively sleek functional groups to block rotation of the troublesome bond without the permeability challenges of previous analogues. One such molecule, a TMR analogue with a four-membered ring, had double the quantum yield of its parent, the researchers found.

But that’s in isolation. The team incorporated the molecule, named Janelia Fluor 549, into a self-labelling HaloTag system for cell imaging. When the researchers expressed the chromosomal proteins called histones in live cultured cells labelled with the Janelia Fluor 549 HaloTag ligand, they emitted nearly twice as many photons per second as the commercially available TMR-based HaloTag ligand — and for twice as long⁸. Furthermore, swapping out the dimethylamine groups on other scaffolds — including rhodamines and coumarins — yielded similar improvements to brightness without noticeable side effects.

Photostability in action

Lavis and his colleagues have now expanded the Janelia Fluor series to include a variety of functional groups, such as fluorinated and deuterated substituents⁹, and the dyes are already enabling discoveries. For example, Cohen’s lab at Harvard University uses high-resolution fluorescence microscopy to watch voltage changes as individual neurons fire in the brains of living mice. “It’s a very noisy environment,” Cohen says. “You need to be able to pick out these tiny fluctuations in fluorescence that only last a millisecond.” That means Cohen needs to illuminate his cells at a high frame rate to capture signal transduction in action — a photon-intensive experiment and a classic formula for photobleaching. “If you’re trying to get a lot of photons out of your molecules in a very short time, you need to drive them hard — you need to put a lot of light onto them — and that means they have to be photostable,” Cohen says.

NatureTech hub

In the past, researchers have studied signal transduction in neurons mostly on the basis of voltage changes at the cell body. Neuronal cell bodies receive inputs from dendrites that are ten times thinner than the bodies and are long enough for the voltage difference at the tip of a dendrite to be different from that at its base. It has not been possible to observe these differences using existing fluorescence systems, Cohen says. By better illuminating how electrical signals propagate through dendrites, photostable fluorophores could reveal how neurons process information and learn.

Cohen, Lavis and their colleagues have now combined the red-shifted dye Janelia Fluor JFX608 with a self-labelling, genetically encoded voltage-indicator system to record sub-millisecond voltage changes across dendritic trees in individual neurons in slices of mouse brain¹⁰. In a second study using the same fluorophore system, the researchers observed voltage changes in the neurons of live mice as they responded to external stimuli¹¹. Both papers were first posted on the bioRxiv preprint server in 2023 and are undergoing peer review.

Still, what works for one dye doesn’t necessarily apply to others, and it’s notoriously difficult to compare one fluorophore to another. The key, Gadella suggests, is to get into the lab: users need to test different fluorophores to make sure their molecule is fit for purpose. Like many researchers, Gadella and his team make their materials available through the non-profit organization Addgene, which has already processed more than 700 orders for mScarlet3. Lavis says his lab has shared some 20,000 aliquots of Janelia Fluor dyes through Janelia’s Open Science program.

More dyes and more photons — that’s a recipe that Lavis, Gadella and others hope will allow researchers to observe complex biochemical phenomena that have so far escaped our view.