生物物理学

Traditional methods for studying protein–protein interactions often lack the resolution to quantitatively distinguish distinct oligomeric states, particularly for membrane proteins within their native lipid environments. To address this limitation, we developed SiMPull-POP (single-molecule pull-down polymeric nanodisc photobleaching), a single-molecule technique designed to quantify membrane protein oligomerization with high sensitivity and in a near-native context. The goal of SiMPull-POP is to enable precise, quantitative analysis of membrane protein assembly by preserving native lipid interactions using diisobutylene maleic acid (DIBMA) to form nanodiscs. Unlike ensemble methods such as co-immunoprecipitation or FRET, which average out heterogeneous populations, SiMPull-POP uses photobleaching to resolve monomeric, dimeric, and higher-order oligomeric states at the single-molecule level. We validated SiMPull-POP using several model systems. A truncated, single-pass transmembrane protein (Omp25) appeared primarily monomeric, while a membrane-tethered FKBP protein exhibited ligand-dependent dimerization upon addition of the AP ligand. Applying SiMPull-POP to EphA2, a receptor tyrosine kinase, we found it to be mostly monomeric in the absence of its ligand, Ephrin-A1, and shifting toward higher-order oligomers upon ligand binding. To explore factors influencing ligand-independent assembly, we modulated membrane cholesterol content. Reducing cholesterol induced spontaneous EphA2 oligomerization, indicating that cholesterol suppresses receptor self-association. Overall, SiMPull-POP offers significant advantages over conventional techniques by enabling quantitative, single-molecule resolution of membrane protein complexes in a native-like environment. This approach provides critical insights into how membrane properties and external stimuli regulate protein assembly, supporting broader efforts to understand membrane protein function in both normal and disease states.

Most viruses extensively remodel their host cells to establish productive infection. Visualization of virus-induced cellular remodeling by electron microscopy (EM) has been revolutionized in recent years by advances in cryo-focused ion beam (cryo-FIB) milling paired with cryo-electron tomography (cryo-ET). As cryo-FIB/ET becomes more widely available, there is a need for beginner-friendly guides to optimize the preparation of virus-infected mammalian cells on EM grids. Here, we provide an in-house protocol for new users for preparing samples of cells infected with herpes simplex virus 1 (HSV-1) for cryo-FIB/ET. This protocol guides users in how to seed infected cells onto grids, blot, and plunge-freeze grids using basic, manual equipment. It also provides tips on how to screen and prioritize grids for efficient milling and data collection.

Conventional Schlieren optics equipment typically operates on a large optical table, which is inconvenient for imaging small samples or thin layers of transparent materials. We describe an imaging device based on Schlieren optics, aided by a slight shift in light reflected from two surfaces. The device is designed to place the sample between a thick concave mirror and a camera next to a point-light source located at the spherical origin of the concave mirror. The compact device is portable and convenient. It is similarly capable of sensitively detecting patterns in gaseous or liquid media created by a density gradient when the optical effect is too subtle to be detectable by regular cameras and scanners. The new device is particularly suitable for detecting translucent samples, including thin fluid films on the order of micrometers, tissue slices, and other biological samples. We show two examples of how our device can be applied to imaging biological samples. The first compares images acquired using several techniques of a bacterial swarm spread over an agar plate; the second is a set of images of human cells grown on a tissue culture plate.

Autonomic regulation of heart and respiratory rates is essential for understanding brain–body interactions in health and disease. Preclinical cardiovascular recordings are often performed under anesthesia or via telemetry, both of which introduce physiological confounds such as stress or impaired recovery due to the need for acute or chronic implantation of sensors. Here, we present a minimally invasive protocol for simultaneous acquisition of high-quality electrocardiography and respiratory signals in awake mice. Using an in-house-modified physiological monitor in awake, head-fixed mice that were briefly habituated to experimental conditions, we ultimately enable stable, long-term physiological recordings alongside in vivo microscopy. This protocol provides a robust, low-stress method for acquiring physiological signals, enabling the simultaneous study of cardiovascular–cerebral dynamics in awake head-fixed mice, thereby enhancing the translational relevance of preclinical measurements.

Understanding how lipids interact with lipid transfer proteins (LTPs) is essential for uncovering their molecular mechanisms. Yet, many available LTP structures, particularly those thought to function as membrane bridges, lack detailed information on where their native lipid ligands are located. Computational strategies, such as docking or AI-methods, offer a valuable alternative to overcome this gap, but their effectiveness is often restricted by the inherent flexibility of lipid molecules and the lack of large training sets with structures of proteins bound to lipids. To tackle this issue, we introduce a reproducible computational pipeline that uses unbiased coarse-grained molecular dynamics (CG-MD) simulations on a free and open-source software (GROMACS) with the Martini 3 force-field. Starting from a configuration of a lipid in bulk solvent, we run CG-MD simulations and observe spontaneous binding of the lipid to the protein. We show that this protocol reliably identifies lipid-binding pockets in LTPs and, unlike docking methods, suggests potential entry routes for lipid molecules with no a priori knowledge other than the protein’s structure. We demonstrate the utility of this approach in investigating bridge LTPs whose internal lipid-binding positions remain unresolved. Altogether, our study provides a cost-effective, efficient, and accurate framework for mapping binding sites and entry pathways in diverse LTPs.

Characterizing the morphology of amyloid proteins is an integral part of studying neurodegenerative diseases. Such morphological characterization can be performed using atomic force microscopy (AFM), which provides high-resolution images of the amyloid protein fibrils. AFM is widely employed for visualizing mechanical and physical properties of amyloid fibrils, not only from a biological and medical perspective but also in relation to their nanotechnological applications. A crucial step in AFM imaging is coating the protein of interest onto a substrate such as mica. However, existing protocols for this process vary considerably. The conventional sample preparation method often introduces artifacts, particularly due to deposition of excess salt. Hence, an optimized protocol is essential to minimize salt aggregation on the mica surface. Here, we present an optimized protocol for coating amyloid proteins onto mica using the dip-washing method to eliminate background noise. This approach improves the adherence of protein to the mica surface while effectively removing residual salts.

In the field of osteoarthritis (OA), the identification of reliable diagnostic and prognostic biomarkers in patients with hip lesions such as femoroacetabular impingement (FAI) could have an immeasurable value. Calcium crystal detection in synovial fluids (SFs) is one tool currently available to diagnose patients with rheumatologic disorders. Crystals, such as monosodium urate (MSU) and calcium pyrophosphate (CPP), are identified qualitatively by compensated polarized light, whereas basic calcium phosphate (BCP) crystals are visualized under conventional light microscopy by Alizarin red S (ARS) staining. Here, we present an efficient and straightforward protocol to quantify calcium crystals by spectrophotometric analysis in human osteoarthritic SFs after staining with ARS. The type and size of the different crystal species are confirmed by environmental scanning electron microscopy (ESEM).

Understanding the nanoscale organization and molecular rearrangement of synaptic components is critical for elucidating the mechanisms of synaptic transmission and plasticity. Traditional synaptosome isolation protocols involve multiple centrifugation and resuspension steps, which may cause structural damage or alter the synaptosomal fraction, compromising their suitability for cryo-electron tomography (cryo-ET). Here, we present an ultrafast isolation method optimized for cryo-ET that yields two types of synaptosomal fractions: synaptosomes and synaptoneurosomes. This streamlined protocol preserves intact postsynaptic membranes apposed to presynaptic active zones and produces thin, high-quality samples suitable for in situ structural studies. The entire procedure, from tissue homogenization to vitrification, takes less than 15 min, offering a significant advantage for high-resolution cryo-ET analysis of synaptic architecture.

Proper genome organization is essential for genome function and stability. Disruptions to this organization can lead to detrimental effects and the transformation of cells into diseased states. Individual chromosomes and their subregions can move or rearrange during transcriptional activation, in response to DNA damage, and during terminal differentiation. Techniques such as fluorescence in situ hybridization (FISH) and chromosome conformation capture (e.g., 3C and Hi-C) have provided valuable insights into genome architecture. However, these techniques require cell fixation, limiting studies of the temporal evolution of chromatin organization in detail. Our understanding of the heterogeneity and dynamics of chromatin organization at the single-cell level is still emerging. To address this, clustered regularly interspaced short palindromic repeats (CRISPR)/dead Cas9 (dCas9) systems have been repurposed for precise live-cell imaging of genome dynamics. This protocol uses a system called CRISPRainbow, a powerful tool that allows simultaneous targeting of up to seven genomic loci and tracks their locations over time using spectrally distinct fluorescent markers to study real-time chromatin organization. Multiple single-guide RNA (sgRNA), carrying specific RNA aptamers for labeling, can be cloned into a single vector to improve transfection efficiency in human cells. The precise targeting of CRISPRainbow offers distinct advantages over previous techniques while also complementing them by validating findings in live cells.

PIEZO1 is a mechanically activated ion channel essential for mechanotransduction and downstream signaling in almost all organ systems. Western blotting is commonly used to study the expression, stability, and post-translational modifications of proteins. However, as a large transmembrane protein, PIEZO1 contains extensive hydrophobic regions and undergoes post-translational modifications that increase its propensity for nonspecific protein–protein interactions. As a result, conventional sample preparation methods seem unsuitable for PIEZO1. For example, heating and sonicating transmembrane proteins exposes hydrophobic regions, leading to aggregation, improper detergent interactions, and loss of solubility, ultimately compromising their detection in western blots. To address these challenges, we developed a western blot protocol optimized for human PIEZO1 by preparing lysates consistently at lower temperatures and incorporating strong reducing and alkylation reagents into the western blot lysis buffer to ensure proper protein solubilization and minimal cross-linking. Using the same antibody, we also developed an immunoprecipitation protocol with optimized detergents to maintain the solubilization of native human PIEZO1, enabling the discovery of a new family of auxiliary subunits.