生物化学

Disruptions and perturbations of the cellular plasma membrane by peptides have garnered significant interest in the elucidation of biological phenomena. Typically, these complex processes are studied using liposomes as model membranes—either by encapsulating a fluorescent dye or by other spectroscopic approaches, such as nuclear magnetic resonance. Despite incorporating physiologically relevant lipids, no synthetic model truly recapitulates the full complexity and molecular diversity of the plasma membrane. Here, biologically representative membrane models, giant plasma membrane vesicles (GPMVs), are prepared from eukaryotic cells by inducing a budding event with a chemical stressor. The GPMVs are then isolated, and bilayers are labelled with fluorescent lipophilic tracers and incubated in a microplate with a membrane-active peptide. As the membranes become damaged and/or aggregate, the resulting fluorescence resonance energy transfer (FRET) between the two tracers increases and is measured periodically in a microplate. This approach offers a particularly useful way to detect perturbations when the membrane complexity is an important variable to consider. Additionally, it provides a way to kinetically detect damage to the plasma membrane, which can be correlated with the kinetics of peptide self-assembly or structural rearrangements.

Key features

• Allows testing of various peptide–membrane interaction conditions (peptide:phospholipid ratio, ionic strength, buffer, etc.) at once.

• Uses intact plasma membrane vesicles that can be prepared from a variety of cell lines.

• Can offer comparable throughput as with traditional synthetic lipid models (e.g., dye-encapsulated liposomes).

Graphical overview

Endosomal recycling is essential for the appropriate function of the endosome. During this process, endosomal coat complexes (i.e., retromer, and Mvp1) are recruited to the endosome, and deform its membrane to form recycling vesicles. To further analyze this, we developed a protocol for the immunoisolation of recycling vesicles from budding yeast. This method is a powerful way to characterize endosomal recycling pathways.

Transbilayer movement of phospholipids in biological membranes is mediated by a diverse set of lipid transporters. Among them are scramblases that facilitate rapid bi-directional movement of lipids without metabolic energy input. In this protocol, we describe the incorporation of phospholipid scramblases into giant unilamellar vesicles (GUVs) formed from scramblase-containing large unilamellar vesicles by electroformation. We also describe how to analyze their activity using membrane-impermeant sodium dithionite, to bleach symmetrically incorporated fluorescent ATTO488-conjugated phospholipids. The fluorescence-based readout allows single vesicle tracking for a large number of settled/immobilized GUVs, and provides a well-defined experimental setup to directly characterize these lipid transporters at the molecular level.

Graphic abstract:

Giant unilamellar vesicles (GUVs) are formed by electroformation from large unilamellar vesicles (LUVs) containing phospholipid scramblases (purple) and trace amounts of a fluorescent lipid reporter (green). The scramblase activity is analyzed by a fluorescence-based assay of single GUVs, using the membrane-impermeant quencher dithionite. Sizes not to scale. Modified from Mathiassen et al. (2021).

Various methods have been developed to generate phosphoglyceride liposomes. Approaches resulting in homogeneous populations of unilamellar bilayer vesicles are generally preferred to mimic various cell membrane situations, as well as to optimize aqueous solute trapping efficiency using the least amount of lipid for biotechnological purposes. Most are time-consuming, often tedious, or require specialized equipment, and produce vesicles with limited shelf-life at room temperature or in cold storage. Herein, we describe a straightforward approach that avoids the preceding complications and streamlines the construction of unilamellar bilayer vesicles from 1-palmitoyl-2-oleoyl phosphatidylcholine (POPC)/dihexanoyl phosphatidylcholine (DHPC) bicelle mixtures at room temperature. The resulting vesicles are small (32-36 nm diameter), unilamellar, bilayer vesicles that are homogeneous, stable, and resistant to freeze-thaw alterations.

Graphic abstract:

Cryo-EM of POPC vesicles formed by dilution of 0.5 q-value POPC/DHPC bicelle mix.

Biolayer interferometry (BLI) is an emerging analytical tool that allows the study of protein complexes in real time to determine protein complex kinetic parameters. This article describes a protocol to determine the K_D of a protein complex using a 6×His tagged fusion protein as bait immobilized on the NTA sensor chip of the FortéBio^® Octet K2 System (Sartorius). We also describe how to determine the half maximal effective concentration (EC₅₀, also known as IC₅₀ for inhibiting effectors) of a metabolite. The complete protocol allows the determination of protein complex K_D and small molecular effector EC₅₀ within 8 h, measured in triplicates.

Graphic abstract:

Principle of the Biolayer interferometry measurement. (Middle, top) Exemplary result of the BLI measurement using Octet^® (Raw Data). Wavelength shift (Δλ) against time. (A) Baseline 1. Baseline measurement. When the sensor is equilibrated in the kinetics buffer. The light is reflected with no difference. (B) Load. The his-tagged proteins (ligand) are loaded onto the sensor surface. The light is reflected with a shift of the wavelength. (C) Baseline 2. The loaded sensor is equilibrated in the kinetics buffer. No further wavelength shift appears. (D) Association. The loaded sensor is dipped into the analyte solution. The analyte binds to the immobilized ligand along with an increased wavelength shift. (E) Dissociation. Afterward, the sensor is dipped again into the kinetics buffer without the analyte. Some analyte molecules dissociate. The wavelength shift decreases. (Subfigures A-E) The left side shows the position of the sensor during the measurement seen in the representative BLI measurement, marked with the figure label. The right side shows the light path in the sensor. Black waves represent the light emitted to the sensor surface. The red waves show the light reflected from the sensor surface back to the detector.

Sphingolipids are major structural components of endomembranes and have also been described as an intracellular second messenger involved in various biological functions in all eukaryotes and a few prokaryotes. Ceramides (Cer), the central molecules of sphingolipids, have been depicted in cell growth arrest, cell differentiation, and apoptosis. With the development of lipidomics, the identification of ceramides has been analyzed in many species, mostly in model insects. However, there is still a lack of research in non-model organisms. Here we describe a relatively simple and sensitive method for the extraction, identification, and quantification of ceramides in Hemiptera Insects (brown planthooper), followed by Ultra-Performance Liquid Chromatography coupled to tandem mass spectrometry (UPLC-MS/MS). C18 is used as the separation column for quantitative detection and analysis on the triple quadruple liquid mass spectrometer. In this protocol, the standard curve method is adopted to confirm the more accurate quantification of ceramides based on the optional detection conditions.

Giant unilamellar vesicles (GUVs) are a widely used model system for a range of applications including membrane biophysics, drug delivery, and the study of actin dynamics. While several protocols have been developed for their generation in recent years, the use of these techniques involving charged lipid types and buffers of physiological ionic strength has not been widely adopted. This protocol describes the generation of large numbers of free-floating GUVs, even for charged lipid types and buffers of higher ionic strength, using a simple approach involving soft polyacrylamide (PAA) gels. This method entails glass cover slip functionalization with (3-Aminopropyl)trimethoxysilane (APTES) and glutaraldehyde to allow for covalent bonding of PAA onto the glass surface. After polymerization of the PAA, the gels are dried in vacuo. Subsequently, a lipid of choice is evenly dispersed on the dried gel surface, and buffers of varying ionic strength can be used to rehydrate the gels and form GUVs. This protocol is robust for the production of large numbers of free-floating GUVs composed of different lipid compositions under physiological conditions. It can conveniently be performed with commonly utilized laboratory reagents.

Lipid membranes are involved in regulating biochemical and biological processes and in modulating the selective permeability of cells, organelles, and vesicles. Membrane composition, charge, curvature, and fluidity all have concerted effects on cellular signaling and homeostasis. The ability to prepare artificial lipid assemblies that mimic biological membranes has enabled investigators to obtain considerable insight into biomolecule-membrane interactions. Lipid nanoscale assemblies can vary greatly in size and composition and can consist of a single lipid monolayer, a bilayer, or other more complex assemblies. This structural diversity makes liposomes suitable for a wide variety of biochemical and clinical applications. Here, we describe a calcein dye leakage assay that we have developed to monitor phospholipid vesicle disruption by alpha-synuclein (αSyn), a presynaptic protein that plays a central role in Parkinson’s disease (PD). We present data showing the effect of adenylylation of αSyn on αSyn-mediated vesicle disruption as an example. This assay can be used to study the effect of mutations or post-translational modifications on αSyn-membrane interactions, to identify protein binding partners or chemical entities that perturb these interactions, and to study the effects of different lipids on the permeabilization activity of αSyn or any other protein.

The emergence and rapid spread of multidrug resistance in bacteria have led to the urgent need for novel antibacterial agents. Membrane permeabilization is the mechanism for many antibacterial molecules that are being developed against gram-negative bacteria. Thus, to determine the efficacy of a potential antibacterial molecule, it is important to assess the change in bacterial membrane permeability after treatment. This study describes the protocol for the assays of outer and inner membrane permeability using the fluorescent probes N-phenyl-1-naphthylamine and propidium iodide. Compared with other experiments, such as electron microscopy and the assay of minimal bactericidal concentration, this methodology provides a simpler, faster, and cost-effective way of estimating the membrane-permeabilizing effect and bactericidal efficacy of antibacterial molecules. This study presents an optimized protocol with respect to the classical protocols by incubating bacteria with antibacterial molecules in the culture condition identical to that of antibacterial assays and then detecting the signal of the fluorescent probe in the buffer without broth and antibacterial molecules. This protocol avoids the effect of nutrient deficiency on the physiological status of bacteria and the interference of antibacterial molecules towards the fluorescent probe. Thus, this method can effectively and precisely evaluate the membrane permeability and match the results obtained from other antibacterial assays, such as minimum inhibitory concentration and time–kill curve assays.

Lipid mixing (redistribution of lipid probes between fusing membranes) has been widely used to study early stages of relatively fast viral and intracellular fusion processes that take seconds to minutes. Lipid mixing assays are especially important for identification of hemifusion intermediates operationally defined as lipid mixing without content mixing. Due to unsynchronized character and the slow rate of the differentiation processes that prime the cells for cell-cell fusion processes in myogenesis, osteoclastogenesis and placentogenesis, these fusions take days. Application of lipid mixing assays to detect early fusion intermediates in these very slow fusion processes must consider the continuous turnover of plasma membrane components and potential fusion-unrelated exchange of the lipid probes between the membranes. Here we describe the application of lipid mixing assay in our work on myoblast fusion stage in development and regeneration of skeletal muscle cells. Our approach utilizes conventional in vitro model of myogenic differentiation and fusion based on murine C2C12 cells. When we observe the appearance of first multinucleated cells, we lift the cells and label them with either fluorescent lipid DiI as a membrane probe or CellTracker^TM Green as a content probe. Redistribution of the probes between the cells is scored by fluorescence microscopy. Hemifused cells are identified as mononucleated cells labeled with both content- and membrane probes. The interpretation must be supported by a system of negative controls with fusion-incompetent cells to account for and minimize contributions of fusion-unrelated exchange of the lipid probes. This approach with minor modifications has been used for investigating fusion of primary murine myoblasts, osteoclast precursors and fusion mediated by a gamete fusogen HAP2, and likely can be adopted for other slow cell-cell fusion processes.