Biochemistry

Membrane transporters mediate the selective movement of ions and molecules across biological membranes and are essential for cellular homeostasis. However, their functional characterization in living cells is often complicated by the complexity of the native membrane environment. Reconstitution into model membrane systems provides a powerful alternative by enabling precise control over lipid composition and experimental conditions. Giant unilamellar vesicles (GUVs) are particularly well suited for transporter studies, as their cell-sized dimensions allow direct microscopic observation and fluorescence-based measurements of protein activity. Here, we describe a two-step reconstitution protocol in which transport proteins are first incorporated into large unilamellar vesicles and then used to generate protein-containing giant unilamellar vesicles (proteo-GUVs) via the poly(vinyl alcohol) swelling method. This two-step approach enhances protein incorporation efficiency and preserves transporter functionality. The method is exemplified using the P3-type ATPase Arabidopsis thaliana plasma membrane H⁺-ATPase isoform 2 (AHA2). We further describe a fluorescence-based assay to assess proton transport activity in proteo-GUVs. Our approach provides a versatile and controlled platform for biochemical, biophysical, and single-molecule analysis of membrane transporters.

Unsaturated fatty acids (UFAs) play key roles in essential cellular functions such as membrane dynamics, metabolism, and animal development. Disruptions in UFA metabolism are linked to metabolic, cardiovascular, and neurodegenerative disorders. Cellular UFAs composition and quantification are normally determined using methods such as gas chromatography and/or mass spectrometry, which require extraction procedures and prevent analysis of live specimens. Here, we describe a protocol that employs uniform ¹³C isotope labeling and high-resolution 2D solution-state nuclear magnetic resonance (NMR) spectroscopy to analyze lipid composition and fatty acid unsaturation directly in the model organism Caenorhabditis elegans. The approach enables in vivo assessment of lipid storage compositions with sufficient resolution and sensitivity to distinguish wild-type animals from those with altered fatty acid desaturation. Complementary analysis of total lipid extracts provides information regarding lipid molecules that are not detected in vivo, such as phospholipid molecules organized in biological membranes. Overall, this non-destructive NMR-based method offers a powerful tool for investigating lipid metabolism in C. elegans and other small model systems that can be isotopically enriched.

Divalent metal ion transporters are conserved across all domains of life and play essential roles in diverse processes such as manganese acquisition during nutritional immunity in bacteria and iron homeostasis in higher eukaryotes [1–3]. Traditional techniques, such as electrophysiological assays, are often unsuitable due to the slow kinetics of many membrane transporters, electroneutral nature of certain transporter types, and the influence of other proteins with similar activity. To overcome these limitations and to investigate both the activity and ion selectivity of transporters, also including those normally expressed intracellularly, we have developed a fluorescence-based transport assay using purified proteins. This in vitro assay uses encapsulated fluorophores to monitor the movement of divalent metal ions (e.g., Mn²⁺, Ca²⁺, Mg²⁺) or protons across liposomal membranes reconstituted with purified transporter proteins. This approach provides detailed functional insight that complements structural and cellular data.

Cellular phenomena such as signal integration and transmission are based on the correct spatiotemporal organization of biomolecules within the cell. Therefore, the targeted manipulation of such processes requires tools that can precisely induce the localizations and interactions of the key players relevant to these processes with high temporal resolution. Chemically induced dimerization (CID) techniques offer a powerful means to manipulate protein function with high temporal resolution and subcellular specificity, enabling direct control over cellular behavior. Here, we present the detailed synthesis and application of dual SLIPT and dual SLIPT^NVOC, which expand the SLIPT (self-localizing ligand-induced protein translocation) platform by incorporating a dual-ligand CID system. Dual SLIPT and dual SLIPT^NVOC independently sort into the inner leaflet of the plasma membrane via a lipid-like anchoring motif, where they present the two headgroup moieties trimethoprim (TMP) and HaloTag ligand (HTL), which recruit and dimerize any two ^iK6eDHFR- and HOB-tagged proteins of interest (POIs). Dual-SLIPT^NVOC furthermore enables this protein dimerization of POIs at the inner leaflet of the plasma membrane in a pre-determined order and light-controlled manner. In this protocol, we detail the synthetic strategy to access dual SLIPT and dual SLIPT^NVOC, while also providing the underlying rationale for key design and synthetic decisions, with the aim of offering a streamlined, accessible, and broadly implementable methodology. In addition to the detailed synthesis, we present representative applications and typical experimental outcomes and recommend strategies for data analysis to support effective use of the system. Notably, dual SLIPT and dual SLIPT^NVOC represent the first CID systems to emulate endogenous lipidation-driven membrane targeting, while retaining hallmark advantages of CID technology—the precision over POI identity, recruitment sequence, high spatiotemporal control, and “plug-and-play” flexibility.

This protocol describes a standardized and economically accessible method for synthesizing mRNA-encapsulated lipid nanoparticles using routine laboratory equipment, including precision syringe pumps, Y-shaped glass microfluidic chips, and silicone tubing. Designed to address the cost and accessibility limitations of commercial microfluidic platforms, the system achieves performance metrics comparable to high-end devices while reducing equipment costs by 90%. By systematically optimizing hydrodynamic parameters (total flow rate: 12 mL/min; lipid-to-aqueous phase ratio: 3:1), the protocol enables consistent production of lipid nanoparticles with key quality attributes: high mRNA encapsulation efficiency (≥ 80%), narrow particle size distribution (100–120 nm, polydispersity index ≤ 0.2), and excellent storage performance (≥ 7 days at 4 °C ).

Phospholipids are major structural and regulatory elements of biological membranes and are involved in many different cellular and physiological processes. In this protocol, we provide an easy, cost-effective, and efficient method to obtain an overview of the phospholipid composition using high-performance thin layer chromatography (HPTLC). While the currently known phospholipid separation methods based on HPTLC display co-migration of certain lipid classes, the method we describe here allows the separation of all phospholipid classes, including anionic phospholipids in plant samples. This protocol combines elements of the classical Vitiello and Touchstone solvent systems to optimize phospholipid separation in a scaled pattern. Here, we provide a full characterization of this method, including statistical analyses of the retention factor of each phospholipid to show the robustness of the method and its efficiency in separating all phospholipid classes of a biological sample.

Phosphatidylinositol 4,5-bisphosphate [PI(4,5)P₂] is a phospholipid enriched on the cytoplasmic leaflet of the plasma membrane, where it plays important roles in membrane trafficking and cytoskeletal dynamics through proteins that directly bind to it. PI(4,5)P₂ can be metabolized to other phosphorylated forms of phosphatidylinositol to regulate numerous processes such as cell growth and development. PI(4,5)P₂ can also be hydrolyzed to generate the second messengers diacylglycerol (DAG) and inositol triphosphate (IP₃). Altered metabolism or mislocalization of PI(4,5)P₂ can perturb one or more of its functions and contribute to disease states. Here, we present a protocol to visualize and quantify the localization of PI(4,5)P₂ in live cells. The protocol uses a highly specific PI(4,5)P₂ protein binding domain coupled to enhanced green fluorescence protein (PH-PLCD1-GFP), enabling localization and quantification of cytosol-facing PI(4,5)P₂ to be determined. Localization and quantification of the PH-PLCD1-GFP, PI(4,5)P₂ specific probe, is enabled by fluorescence imaging and confocal microscopy. This approach can be used to study the dynamics of PI(4,5)P₂ localization temporally in live cells under both physiological and pathological conditions.

Stable isotopes have frequently been used to study metabolic processes in live cells both in vitro and in vivo. Glutamine, the most abundant amino acid in human blood, plays multiple roles in cellular metabolism by contributing to the production of nucleotides, lipids, glutathione, and other amino acids. It also supports energy production via anaplerosis of tricarboxylic acid cycle intermediates. While ¹³C-glutamine has been extensively employed to study glutamine metabolism in various cell types, detailed analyses of specific lipids derived from ¹³C-glutamine via the reductive carboxylation pathway are limited. In this protocol, we present a detailed procedure to investigate glutamine metabolism in human glioblastoma (GBM) cells by conducting ¹³C-glutamine tracing coupled with untargeted metabolomics analysis using liquid chromatography–mass spectrometry (LC–MS/MS). The method includes step-by-step instructions for the extraction and detection of polar metabolites and long-chain fatty acids (LCFAs) derived from ¹³C-glutamine in GBM cells. Notably, this approach enables the distinction between isomers of two monounsaturated FAs with identical masses: palmitoleic acid (16:1n-7) (cis-9-hexadecenoic acid) and palmitelaidic acid (16:1n-7) (trans-9-hexadecenoic acid) derived from ¹³C-glutamine through the reductive carboxylation process. In addition, using this protocol, we also unveil previously unknown metabolic alterations in GBM cells following lysosome inhibition by the antipsychotic drug pimozide.

Membranes are very complex and dynamic structures that are essential for plant cellular functions and whose lipidic composition can be influenced by numerous factors. Anionic phospholipids, which include phosphatidylserine, phosphatidic acid, phosphatidylinositol, and phosphoinositides are key components of these membranes as they are involved in plant cell signaling and as even slight modifications in their quantities may largely impact the cell metabolism. However, the presence of these compounds in low amounts, as well as their poor stability during analysis by mass spectrometry, make their study very complicated. In addition, the precise quantification of all anionic phospholipid species is not possible by lipid separation using thin-layer chromatography followed by the analysis of their fatty acyl chains by gas chromatography. Here, we describe a straightforward strategy for the extraction and semi-quantification of all anionic phospholipid species from plant samples. Our method is based on the derivatization of the anionic phospholipids, and more especially on their methylation using trimethylsilyldiazomethane, followed by analysis by high-performance liquid chromatography coupled with a triple quadrupole mass spectrometer. This approach allows largely improving the sensitivity of the analysis of anionic phospholipids from plant samples, which will help to gain deeper insights into the functions and dynamics of these key parts of plant cellular signaling.

With the advancement of liquid chromatography–mass spectrometry (LC–MS/MS), the quantification of glycerophospholipid (PL) molecules has become more accessible, leading to the discovery of numerous enzymes responsible for determining the acyl groups attached to these molecules. Metabolic tracer experiments using radioisotopes and stable isotopes are powerful tools for defining the function of metabolic enzymes and metabolic flux. We have established an ex vivo muscle experimental system using stable isotope–labeled fatty acids to evaluate fatty acid incorporation into PL molecules. Here, we describe a method to incorporate fatty acids with stable isotope labels into excised skeletal muscle and detect the PL molecules containing labeled acyl chains by LC–MS/MS.