生物物理学

The physiological role of a-synuclein (a-syn), an intrinsically disordered presynaptic neuronal protein, is believed to impact the release of neurotransmitters through interactions with the SNARE complex. However, under certain cellular conditions that are not well understood, a-syn will self-assemble into β-sheet-rich fibrils that accumulate and form insoluble neuronal inclusions. Studies of patient-derived brain tissues have concluded that these inclusions are associated with Parkinson’s disease, the second most common neurodegenerative disorder, and other synuclein-related diseases called synucleinopathies. In addition, repetitions of specific mutations to the SNCA gene, the gene that encodes a-syn, result in an increased disposition for synucleinopathies. The latest advances in cryo-EM structure determination and real-space helical reconstruction methods have resulted in over 60 in vitro structures of a-syn fibrils solved to date, with a handful of these reaching a resolution below 2.5 Å. Here, we provide a protocol for a-syn protein expression, purification, and fibrilization. We detail how sample quality is assessed by negative stain transmission electron microscopy (NS-TEM) analysis and followed by sample vitrification using the Vitrobot Mark IV vitrification robot. We provide a detailed step-by-step protocol for high-resolution cryo-EM structure determination of a-syn fibrils using RELION and a series of specialized helical reconstruction tools that can be run within RELION. Finally, we detail how ChimeraX, Coot, and Phenix are used to build and refine a molecular model into the high-resolution cryo-EM map. This workflow resulted in a 2.04 Å structure of a-syn fibrils with excellent resolution of residues 36–97 and an additional island of density for residues 15–22 that had not been previously reported. This workflow should serve as a starting point for individuals new to the neurodegeneration and structural biology fields. Together, this procedure lays the foundation for advanced structural studies of a-syn and other amyloid fibrils.

Interstitial fluid (ISF) is a promising diagnostic sample due to its extensive biomolecular content while being safer and less invasive to collect than blood. However, existing ISF sampling methods are time-consuming, require specialized equipment, and yield small amounts of fluid (<5 μL). We have recently reported a simple and minimally invasive technique for rapidly sampling larger quantities of dermal ISF using a microneedle (MN) array to generate micropores in the skin from which ISF is extracted using a vacuum-assisted skin patch. Here, we present step-by-step protocols for fabricating the MN array and skin patch, as well as for using them to sample ISF from human skin. Using this technique, an average of 20.8 μL of dermal ISF can be collected within 25 min, which is a ∼6-fold improvement over existing ISF sampling methods. Furthermore, the technique is well-tolerated and does not require the use of expensive or specialized equipment. The ability to collect ample volumes of ISF in a quick and minimally invasive manner will facilitate the analysis of ISF for biomarker discovery and its use for diagnostic testing.

During neuronal synaptic transmission, the exocytotic release of neurotransmitters from synaptic vesicles in the presynaptic neuron evokes a change in conductance for one or more types of ligand-gated ion channels in the postsynaptic neuron. The standard method of investigation uses electrophysiological recordings of the postsynaptic response. However, electrophysiological recordings can directly quantify the presynaptic release of neurotransmitters with high temporal resolution by measuring the membrane capacitance before and after exocytosis, as fusion of the membrane of presynaptic vesicles with the plasma membrane increases the total capacitance. While the standard technique for capacitance measurement assumes that the presynaptic cell is unbranched and can be represented as a simple resistance-capacitance (RC) circuit, neuronal exocytosis typically occurs at a distance from the soma. Even in such cases, however, it can be possible to detect a depolarization-evoked increase in capacitance. Here, we provide a detailed, step-by-step protocol that describes how "Sine + DC" (direct current) capacitance measurements can quantify the exocytotic release of neurotransmitters from AII amacrine cells in rat retinal slices. The AII is an important inhibitory interneuron of the mammalian retina that plays an important role in integrating rod and cone pathway signals. AII amacrines release glycine from their presynaptic dendrites, and capacitance measurements have been important for understanding the release properties of these dendrites. When the goal is to directly quantify the presynaptic release, there is currently no other competing method available. This protocol includes procedures for measuring depolarization-evoked exocytosis, using both standard square-wave pulses, arbitrary stimulus waveforms, and synaptic input.

Magnetic resonance imaging (MRI) is an invaluable method of choice for anatomical and functional in vivo imaging of the brain. Still, accurate delineation of the brain structures remains a crucial task of MR image evaluation. This study presents a novel analytical algorithm developed in MATLAB for the automatic segmentation of cerebrospinal fluid (CSF) spaces in preclinical non-contrast MR images of the mouse brain. The algorithm employs adaptive thresholding and region growing to accurately and repeatably delineate CSF space regions in 3D constructive interference steady-state (3D-CISS) images acquired using a 9.4 Tesla MR system and a cryogenically cooled transmit/receive resonator. Key steps include computing a bounding box enclosing the brain parenchyma in three dimensions, applying an adaptive intensity threshold, and refining CSF regions independently in sagittal, axial, and coronal planes. In its original application, the algorithm provided objective and repeatable delineation of CSF regions in 3D-CISS images of sub-optimal signal-to-noise ratio, acquired with (33 μm)³ isometric voxel dimensions. It allowed revealing subtle differences in CSF volumes between aquaporin-4-null and wild-type littermate mice, showing robustness and reliability. Despite the increasing use of artificial neural networks in image analysis, this analytical approach provides robustness, especially when the dataset is insufficiently small and limited for training the network. By adjusting parameters, the algorithm is flexible for application in segmenting other types of anatomical structures or other types of 3D images. This automated method significantly reduces the time and effort compared to manual segmentation and offers higher repeatability, making it a valuable tool for preclinical and potentially clinical MRI applications.

Mitochondrial cristae, formed by folding the mitochondrial inner membrane (IM), are essential for cellular energy supply. However, the observation of the IM is challenging due to the limitations in spatiotemporal resolution offered by conventional microscopy and the absence of suitable in vitro probes specifically targeting the IM. Here, we describe a detailed imaging protocol for the mitochondrial inner membrane using the Si-rhodamine dye HBmito Crimson, which has excellent photophysical properties, to label live cells for imaging via stimulated emission depletion (STED) microscopy. This allows for STED imaging over more than 500 frames (approximately one hour), with a spatial resolution of 40 nm, enabling the observation of cristae dynamics during various mitochondrial processes. The protocol includes detailed steps for cell staining, image acquisition, image processing, and resolution analysis. Utilizing the superior resolution of STED microscopy, the structure and complex dynamic changes of cristae can be visualized.

Cell-generated forces play a critical role in driving and regulating complex biological processes, such as cell migration and division and cell and tissue morphogenesis in development and disease. Traction force microscopy (TFM) is an established technique developed in the field of mechanobiology used to quantify cellular forces exerted on soft substrates and internal mechanical tissue stresses. TFM measures cell-generated traction forces in 2D or 3D environments with varying mechanical and biochemical properties. This technique involves embedding fiducial markers in the substrate, imaging substrate deformations caused by the cells, and using mathematical models to infer forces. This protocol compiles procedures from various previously published studies and software packages and describes how to perform TFM on 2D micropatterned substrates. Although not the focus of this protocol, the methods and software packages shown here also allow to perform monolayer stress microscopy (MSM), a method to calculate internal mechanical stress within the cells by modeling them as a thin plate with linear and homogeneous material properties. TFM and MSM are non-invasive methods capable of yielding spatially and temporally resolved force and stress maps with high throughput. As such, they enable the generation of rich datasets, which can provide valuable insights into the roles of cell-generated forces in various physiological and pathological processes.

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.

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.

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.

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.