Piezo1 is a mechanosensitive channel that converts applied force into electrical signals. Partial molecular structures show a bowl-shaped trimer with extended arms. Here we use cryo-electron microscopy (cryo-EM) to show that Piezo1 adopts different degrees of curvature in lipid vesicles of different size. We also use high-speed atomic force microscopy (HS-AFM) imaging to analyze the deformability of Piezo1 under force in membranes on a mica surface: Piezo1 can be flattened reversibly into the membrane plane. By approximating the absolute force applied, we estimate a range of values for a mechanical spring constant for Piezo1. Both methods demonstrate that Piezo1 can deform its shape towards a planar structure. This deformation could explain how lateral membrane tension can be converted into a conformation-dependent free energy change to gate the Piezo1 channel in response to mechanical perturbations.

Living cells are viscoelastic materials, with the elastic response dominating at long timescales (≳1 ms)1. At shorter timescales, the dynamics of individual cytoskeleton filaments are expected to emerge, but active microrheology measurements on cells accessing this regime are scarce2. Here, we develop high-frequency microrheology (HF-MR) to probe the viscoelastic response of living cells from 1Hz to 100 kHz. We report the viscoelasticity of different cell types and upon cytoskeletal drug treatments. At previously inaccessible short timescales, cells exhibit rich viscoelastic responses that depend on the state of the cytoskeleton. Benign and malignant cancer cells revealed remarkably different scaling laws at high frequency, providing a univocal mechanical fingerprint. Microrheology over a wide dynamic range up to the frequency of action of the molecular components provides a mechanistic understanding of cell mechanics.

Conventional atomic force microscopes (AFMs) take at least 30-60 s to capture an image, while dynamic biomolecular processes occur on a millisecond timescale or less. To narrow this large difference in timescale, various studies have been carried out in the past decade. These efforts have led to a maximum imaging rate of 30-60 ms/ frame for a scan range of~250 nm, with a weak tip-sample interaction force being maintained. Recent imaging studies using high-speed AFM with this capacity have shown that this new microscope can provide straightforward and prompt answers to how and what structural changes progress while individual biomolecules are at work. This article first compares high-speed AFM with its competitor (single-molecule fluorescence microscopy) on various aspects and then describes high-speed AFM instrumentation and imaging studies on biomolecular processes. The article concludes by discussing the future prospects of this cutting-edge microscopy.

A fundamental challenge of gene regulation is the accessibility of DNA within nucleosomes. Recent studies performed by various techniques, including single-molecule approaches, led to the realization that nucleosomes are quite dynamic rather than static systems, as they were once considered. Direct data are needed to characterize the dynamics of nucleosomes. Specifically, if nucleosomes are dynamic, the following questions need to be answered. What is the range of nucleosome dynamics? Is a non-ATP-dependent unwrapping of nucleosomes possible? What are the factors facilitating the large-scale opening and unwrapping of nucleosomes? In previous studies using time-lapse atomic force microscopy (AFM) imaging, we were able, for the first time, to observe spontaneous, ATP-independent unwrapping of nucleosomes. However, low temporal resolution did not allow visualization of various pathways of nucleosome dynamics. In the studies described here, we applied high-speed time-lapse AFM (HS-AFM) capable of visualizing molecular dynamics on the millisecond time scale to study the nucleosome dynamics. The mononucleosomes were assembled on a 353 bp DNA substrate containing nucleosome-specific 601 sequence. With HS-AFM, we were able to observe the dynamics of nucleosome on a subsecond time scale and visualize various pathways of nucleosome dynamics, such as sliding and unwrapping to various extents, including complete dissociation. These studies highlight an important role of electrostatic interactions in chromatin dynamics. Overall, our findings shed new light on nucleosome dynamics and provide a novel hypothesis for the mechanisms controlling the spontaneous dynamics of chromatin.

Glutamate transporters are essential for recovery of the neurotransmitter glutamate from the synaptic cleft. Crystal structures in the outward-and inward-facing conformations of a glutamate transporter homolog from archaebacterium Pyrococcus horikoshii, sodium/aspartate symporter Glt Ph , suggested the molecular basis of the transporter cycle. However, dynamic studies of the transport mechanism have been sparse and indirect. Here we present highspeed atomic force microscopy (HS-AFM) observations of membranereconstituted Glt Ph at work. HS-AFM movies provide unprecedented real-space and real-time visualization of the transport dynamics. Our results show transport mediated by large amplitude 1.85-nm "elevator" movements of the transport domains consistent with previous crystallographic and spectroscopic studies. Elevator dynamics occur in the absence and presence of sodium ions and aspartate, but stall in sodium alone, providing a direct visualization of the ion and substrate symport mechanism. We show unambiguously that individual protomers within the trimeric transporter function fully independently.Glt Ph | HS-AFM | transporter | elevator mechanism | dynamics

Annexins are abundant cytoplasmic proteins that can bind to negatively charged phospholipids in a Ca(2+)-dependent manner, and are known to play a role in the storage of Ca(2+) and membrane healing. Little is known, however, about the dynamic processes of protein-Ca(2+)-membrane assembly and disassembly. Here we show that high-speed atomic force microscopy (HS-AFM) can be used to repeatedly induce and disrupt annexin assemblies and study their structure, dynamics and interactions. Our HS-AFM set-up is adapted for such biological applications through the integration of a pumping system for buffer exchange and a pulsed laser system for uncaging caged compounds. We find that biochemically identical annexins (annexin V) display different effective Ca(2+) and membrane affinities depending on the assembly location, providing a wide Ca(2+) buffering regime while maintaining membrane stabilization. We also show that annexin is membrane-recruited and forms stable supramolecular assemblies within ∼5 s in conditions that are comparable to a membrane lesion in a cell. Molecular dynamics simulations provide atomic detail of the role played by Ca(2+) in the reversible binding of annexin to the membrane surface.

Intrinsically disordered (ID) regions of proteins are recognized to be involved in biological processes such as transcription, translation, and cellular signal transduction. Despite the important roles of ID regions, effective methods to observe these thin and flexible structures directly were not available. Herein, we use high-speed atomic force microscopy (AFM) to observe the heterodimeric FACT (facilitates chromatin transcription) protein, which is predicted to have large ID regions in each subunit. Successive AFM images of FACT on a mica surface, captured at rates of 5-17 frames per second, clearly reveal two distinct tail-like segments that protrude from the main body of FACT and fluctuate in position. Using deletion mutants of FACT, we identify these tail segments as the two major ID regions predicted from the amino acid sequences. Their mechanical properties estimated from the AFM images suggest that they have more relaxed structures than random coils. These observations demonstrate that this state-of-the-art microscopy method can be used to characterize unstructured protein segments that are difficult to visualize with other experimental techniques.

The DNA cytosine deaminase APOBEC3G (A3G) is capable of blocking retrovirus replication by editing viral cDNA and impairing reverse transcription. However, the biophysical details of this host-pathogen interaction are unclear. Here we applied atomic force microscopy (AFM) and hybrid DNA substrates to investigate properties of A3G bound to single-stranded DNA (ssDNA). Hybrid DNA substrates included ssDNA with 5′ or 3′ ends attached to DNA duplexes (tail-DNA) and gap-DNA substrates, in which ssDNA is flanked by two double-stranded fragments. We found that A3G binds with similar efficiency to the 5′ and 3′ substrates, suggesting that ssDNA polarity is not an important factor. Additionally, we observed that A3G binds the single-stranded region of the gap-DNA substrates with the same efficiency as tail-DNA. These results demonstrate that single-stranded DNA ends are not needed for A3G binding. The protein stoichiometry does not depend on the ssDNA substrate type, but the ssDNA length modulates the stoichiometry of A3G in the complex. We applied single molecule high-speed AFM to directly visualize the dynamics of A3G in the complexes. We were able to visualize A3G sliding and protein association-dissociation events. During sliding, A3G translocated over a 69 nucleotide ssDNA segment in less than 1 second. Association-dissociation events were more complex, as dimeric A3G could dissociate from the template as a whole, or undergo a two-step process with monomers capable of sequential dissociation. We conclude that A3G monomers, dimers, and higher order oligomers can bind ssDNA substrates independent of strand polarity and availability of free ssDNA ends.