生物物理学

Cryo-electron microscopy (cryo-EM) is a powerful technique capable of investigating samples in a hydrated state, compared to conventional high-vacuum electron microscopy that requires samples to be completely dry. During the drying process, numerous features and details may be lost due to damage caused by dehydration. Cryo-EM circumvents these problems by cryo-fixing the samples, thereby retaining the intact and original features of hydrated samples. This protocol describes a step-by-step cryo-scanning electron microscopy (cryo-SEM) experimental procedure with Chlorella sorokiniana as the subject. By employing filter paper as the sample substrate, we propose a simple and reliable method for cryo-fixation and freeze-fracture of Chlorella sorokiniana in water suspension. The advantage of using filter paper as a substrate lies in its ability to support a thin film of sample, enabling a cold knife to make a cut effortlessly and produce a clean freeze-fractured surface for SEM investigation. By following the approach described in this protocol, both the internal structure and surface morphology of Chlorella sorokiniana can be easily resolved with high quality. This protocol is highly versatile and can be applied to samples dispersed in water or solvents, including cyanobacterial cells, algal cells, and any kind of sample that can be adsorbed onto filter paper.

The motile parameters of kinesin superfamily proteins are fundamental to intracellular transport. Single-molecule motility assays using total internal reflection fluorescence (TIRF) microscopy are a gold standard technique for measuring the motile parameters of kinesin motors. With this technique, one can evaluate the velocity, run length, and binding frequency of kinesins on microtubules by directly observing their motility. This protocol provides a comprehensive procedure for single molecule assays of kinesins, including the preparation of labeled microtubules, the measurement of kinesin motility via TIRF microscopy, and the quantification of kinesin motor parameters.

The planar lipid bilayer (PLB) technique represents a highly effective method for the study of membrane protein properties in a controlled environment. The PLB method was employed to investigate the role of mitochondrial inner membrane protein 17 (MPV17), whose mutations are associated with a hepatocerebral form of mitochondrial DNA depletion syndrome (MDS). This protocol presents a comprehensive, step-by-step guide to the assembly and utilization of a PLB system. The procedure comprises the formation of a lipid bilayer over an aperture, the reconstitution of the target protein, and the utilization of electrophysiological recording techniques to monitor channel activity. Furthermore, recommendations are provided for optimizing experimental conditions and overcoming common challenges encountered in PLB experiments. Overall, this protocol highlights the versatility of the PLB technique in advancing our understanding of membrane protein function and its broad application in various fields of research.

Protein carbonylation has been known as the major form of irreversible protein modifications and is also widely used as an indicator of oxidative stress in the biological environment. In the presence of oxidative stress, biological systems tend to produce large amounts of carbonyl moieties; these carbonyl groups do not have particular UV-Vis and fluorescence spectroscopic characteristics that we can differentiate, observe, and detect. Thus, their detection and quantification can only be performed using specific chemical probes. Commercially available fluorescent probes to detect specific carbonylation in biological systems have been used, but their chemical portfolio is still very limited. This protocol outlines the methods and procedures employed to synthesize a probe, (E,Z)-2-(2-(2-hydroxybenzylidene)hydrazonyl)-5-nitrophenol (2Hzin5NP), and assess its impact on carbonylation in human cells. The synthesis involves several steps, including the preparation of its hydrazone compounds mimicking cell carbonyls, 2-Hydrazinyl 5-nitrophenol, (E,Z)-2-(2-ethylidenehydrazonyl)-5-nitrophenol, and the final product (E,Z)-2-(2-(2-hydroxybenzylidene)hydrazonyl)-5-nitrophenol. The evaluation of fluorescence quantum yield and subsequent cell culture experiments are detailed for the investigation of 2Hzin5NP effects on cell proliferation and carbonylation.

Membrane protein structures offer a more accurate basis for understanding their functional correlates when derived from full-length proteins in their native lipid environment. Producing such samples has been a primary challenge in the field. Here, we present robust, step-by-step biochemical and biophysical protocols for generating monodisperse assemblies of full-length transmembrane proteins within lipidic environments. These protocols are particularly tailored for cases where the size and molecular weight of the proteins align closely with those of the lipid islands (nanodiscs). While designed for single-span bitopic membrane proteins, these protocols can be easily extended to proteins with multiple transmembrane domains. The insights presented have broad implications across diverse fields, including biophysics, structural biology, and cryogenic electron microscopy (cryo-EM) studies.

Single-stranded RNA bacteriophages (ssRNA phages) infect their hosts by binding to the host receptor pili. Purification of pili usually involves mechanical shearing of pili from cells followed by precipitation. However, previous methods often result in low efficiency or unstable results due to pili retraction. This protocol presents an optimized method for purifying receptor type IV pili from Acinetobacter genomospecies 16 (A. gp16), incorporating enhancements in shearing and collection steps to achieve high yields. We found that repeated passage through syringe needles increases yield, and temperature control is crucial during purification. Additionally, the CsCl density gradient was optimized specifically for this specific strain. The purified type IV pili are suitable for cryogenic electron microscopy (cryo-EM) and various biochemical experiments.

Protein misfolding fuels multiple neurodegenerative diseases, but existing techniques lack the resolution to pinpoint the location and physical properties of aggregates within living cells. Our protocol describes high-resolution confocal and fluorescent lifetime microscopy (Fast 3D FLIM) of an aggregation probing system. This system involves a metastable HaloTag protein (HT-aggr) labeled with P1 solvatochromic fluorophore, which can be targeted to subcellular compartments. This strategy allows to distinguish between aggregated and folded probe species, since P1 fluorophore changes its lifetime depending on the hydrophobicity of its microenvironment. The probe is not fluorescence intensity-dependent, overcoming issues related to intensity-based measurements of labeled proteins, such as control of probe quantity due to differences in expression or photobleaching of a proportion of the fluorophore population. Our approach reports on the performance of the machinery dealing with aggregation-prone substrates and thus opens doors to studying proteostasis and its role in neurodegenerative diseases.

With the growth of the quantum biology field, the study of magnetic field (MF) effects on biological processes and their potential therapeutic applications has attracted much attention. However, most biologists lack the experience needed to construct an MF exposure apparatus on their own, no consensus standard exists for exposure methods, and protocols for model organisms are sorely lacking. We aim to provide those interested in entering the field with the ability to investigate static MF effects in their own research. This protocol covers how to design, build, calibrate, and operate a static MF exposure chamber (MagShield apparatus), with instructions on how to modify parameters to other specific needs. The MagShield apparatus is constructed of mu-metal (which blocks external MFs), allowing for the generation of experimentally controlled MFs via 3-axial Helmholtz coils. Precise manipulation of static field strengths across a physiologically relevant range is possible: nT hypomagnetic fields, μT to < 1 mT weak MFs, and moderate MFs of several mT. An integrated mu-metal partition enables different control and experimental field strengths to run simultaneously. We demonstrate (with example results) how to use the MagShield apparatus with Xenopus, planarians, and fibroblast/fibrosarcoma cell lines, discussing the modifications needed for cell culture systems; however, the apparatus is easily adaptable to zebrafish, C. elegans, and 3D organoids. The operational methodology provided ensures uniform and reproducible results, affording the means for rigorous examination of static MF effects. Thus, this protocol is a valuable resource for investigators seeking to explore the intricate interplay between MFs and living organisms.

Expansion microscopy (ExM) has significantly reformed the field of super-resolution imaging, emerging as a powerful tool for visualizing complex cellular structures with nanoscale precision. Despite its capabilities, the epitope accessibility, labeling density, and precision of individual molecule detection pose challenges. We recently developed an iterative indirect immunofluorescence (IT-IF) method to improve the epitope labeling density, improving the signal and total intensity. In our protocol, we iteratively apply immunostaining steps before the expansion and exploit signal processing through noise estimation, denoising, and deblurring (NEDD) to aid in quantitative image analyses. Herein, we describe the steps of the iterative staining procedure and provide instructions on how to perform NEDD-based signal processing. Overall, IT-IF in ExM–laser scanning confocal microscopy (LSCM) represents a significant advancement in the field of cellular imaging, offering researchers a versatile tool for unraveling the structural complexity of biological systems at the molecular level with an increased signal-to-noise ratio and fluorescence intensity.

Fluorescence microscopy has been widely accessible and indispensable in cell biology research. This technique enables researchers to label targets, ranging from individual entities to multiple groups, with fluorescent markers. It offers precise determinations of localization, size, and shape, along with accurate quantifications of fluorescence signal intensities. Furthermore, an ideal fluorescence microscope can achieve approximately 250 nm in lateral and 600 nm in axial resolution. Despite its integral role in these measurements, the calibration of fluorescence microscopes is often overlooked. This protocol introduces the use of 3D-Speckler (3D fluorescence speckle analyzer), a semi-automated software tool we have recently developed, for calibrating fluorescence microscopy. Calibration of fluorescence microscopy includes determining resolution limits, validating accuracy in size measurements, evaluating illumination flatness, and determining chromatic aberrations. 3D-Speckler is user-friendly and enables precise quantification of fluorescence puncta, including nanoscale 2D/3D particle size, precise locations, and intensity information. By utilizing multispectral fluorescence beads of known sizes alongside 3D-Speckler, the software can effectively calibrate imaging systems. We emphasize the importance of routine calibration for imaging systems to maintain their integrity and reproducibility, ensuring accurate quantification. This protocol provides a detailed step-by-step guide on using 3D-Speckler to calibrate imaging systems.