Biophysics

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.

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.

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.

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.

A number of extracellular helical protein polymers are crucial for supporting bacterial motility. The bacterial flagellum is a polymeric appendage used to support cellular motility. Historically, structural studies of flagellar and other filaments were limited to those present as or locked into straightened states. Here, we present a robust workflow that produces biologically relevant high-resolution cryo-electron microscopy (cryo-EM) structures of bacterial flagellar filaments. We highlight how a simple purification method, centered around several centrifugation steps, exploits the process of filament ejection in Caulobacter crescentus and results in isolated filaments amenable to transmission electron microscopy (TEM) studies. The quality of the sample is validated by SDS-PAGE and negative stain TEM analysis before a sample is vitrified for cryogenic electron microscopy (cryo-EM) data collection. We provide a detailed protocol for reconstructing either straight or curved flagellar filaments by cryo-EM helical reconstruction methods, followed by an overview of model building and validation. In our hands, this workflow resulted in several flagellar structures below 3 Å resolution, with one data set reaching a global resolution of 2.1 Å. The application of this workflow supports structure-function studies to better understand the molecular interactions that regulate filament architecture in biologically relevant states. Future work will not only examine interactions that regulate bacterial flagellar and other filament organization but also provide a foundation for developing new helical biopolymers for biotech applications.

Tension and force propagation play a central role in tissue morphogenesis, as they enable sub- and supra-cellular shape changes required for the generation of new structures. Force is often generated by the cytoskeleton, which forms complex meshworks that reach cell–cell or cell–extracellular matrix junctions to induce cellular rearrangements. These mechanical properties can be measured through laser microdissection, which concentrates energy in the tissue of interest, disrupting its cytoskeleton. If the tissue is undergoing tension, this cut will induce a recoil in the surrounding regions of the cut. This protocol describes how one can perform laser microdissection experiments and subsequently measure the recoil speed of the sample of interest. While we explain how to carry out these experiments in Drosophila embryos, the recoil calibration and downstream analyses can be applied to other types of preparations.

Key features

• Allows measuring tension in live Drosophila embryos with a relatively simple approach.

• Describes a quick way to mount a high number of embryos.

• Includes a segmentation-free recoil quantification that reduces bias and speeds up analysis.

Graphical overview

Determining the oligomeric state of membrane proteins is critical for understanding their function. However, traditional ex situ methods like clear native gel electrophoresis can disrupt protein subunit interactions during sample preparation. In situ methods such as stepwise photobleaching have limitations due to high expression levels and limitations of optical resolution in microscopy. Super-resolution microscopy techniques such as single-molecule localization microscopy (SMLM) have the potential to overcome these limitations, but the stochastic nature of signals can lead to miscounting due to over-expression, background noise, and temporal separation of signals. Additionally, this technique has limited application due to the limited selection of fluorescent labels and the demanding control of laser power. To address these issues, we developed a dual color colocalization (DCC) strategy that offers higher tolerance to background noise and simplifies data acquisition and processing for high-throughput and reliable counting. The DCC strategy was used to determine the oligomeric states of membrane proteins of the SLC17 and SLC26 family with SMLM, providing a robust and efficient method for studying protein interactions.

Graphical overview

(A) Illustration of the principle for determining the oligomeric state of protein complexes with dual color colocalization–single-molecule localization microscopy (DCC-SMLM). In the inset, as an example, a dimeric protein (brown) is labeled with a marker (M) and an indicator fluorescent protein (F) on each of its two subunits. The overall probability of detecting the dimer with SMLM, as denoted by R, the colocalization ratio, is equal to the ratio of the number of colocalized marker and indicator clusters (NMF) to that of the marker clusters (NM). The plot shows the linear relationship of the oligomeric state (n) vs. the natural logarithm of 1 subtracted by the colocalization ratio, supplemented by the equation of the fitting curve, in which p denotes the recall rate of the indicator fluorescent protein (F). (B) The workflow diagram shows the procedures of DCC-SMLM (Locs: localizations; COM: coefficient of mismatch; LCA: lateral chromatic aberration).

Lipid-conjugated pH sensors based on fluorophores coupled to lipids are a powerful tool for monitoring pH gradients in biological microcompartments and reconstituted membrane systems. This protocol describes the synthesis of pH sensors based on amine-reactive pHrodo esters and the amino phospholipid phosphatidylethanolamine. The major features of this sensor include efficient partitioning into membranes and strong fluorescence under acidic conditions. The protocol described here can be used as a template to couple other amine-reactive fluorophores to phosphatidylethanolamines.

Graphical overview

Synthesis of lipid-conjugated pH sensors based on amine-reactive fluorophore esters and the aminophospholipid phosphoethanolamine (PE)