The biophysical drivers of membrane lateral heterogeneity, often termed lipid rafts, have been largely explored using synthetic liposomes or mammalian plasma membrane-derived giant vesicles. Yeast vacuoles, an organelle comparable to mammalian lysosomes, is the only in vivo system that shows stable micrometer scale phase separation in unperturbed cells. The ease of manipulating lipid metabolism in yeast makes this a powerful system for identifying lipids involved in the onset of vacuole membrane heterogeneity. Vacuole domains are induced by stationary stage growth and nutritional starvation, during which they serve as a docking and internalization site for lipid droplet energy stores. Here we describe methods for characterizing vacuole phase separation, its physiological function, and its lipidic drivers. First, we detail methodologies for robustly inducing vacuole domain formation and quantitatively characterizing during live cell imaging experiments. Second, we detail a new protocol for biochemical isolation of stationary stage vacuoles, which allows for lipidomic dissection of membrane phase separation. Third, we describe biochemical techniques for analyzing lipid droplet internalization in vacuole domains. When combined with genetic or chemical perturbations to lipid metabolism, these methods allow for systematic dissection of lipid composition in the structure and function of ordered membrane domains in living cells.

Chaperone-mediated autophagy (CMA) is the most selective form of lysosomal proteolysis, in which proteins are individually selected for lysosomal degradation. CMA degradation targets bear a pentapeptide consensus motif that is recognized by the cytosolic chaperone HSPA8 (Hsc70), which participates in the trafficking of the target to the lysosomal surface. From there, it is translocated into the lysosomal lumen, independent of vesicle fusion, in a process dependent upon the lysosomal transmembrane protein LAMP2A. There are limited tools for studying CMA in whole cells and tissues, and many of the best techniques for studying CMA rely on the preparation of lysosome enriched fractions. Such experiments include (1) the in vitro evaluation of CMA substrate uptake activity, (2) the characterization of changes to lysosomal resident and CMA regulatory proteins, and (3) lysosomal targetomics, i.e., the use of quantitative proteomics to characterize lysosomal degradation targets. Previous studies using discontinuous metrizamide gradients have shown that a subpopulation of liver lysosomes is responsible for the majority of CMA activity (\"CMA+ \"). These CMA+ lysosomes are low density and have higher levels of MTORC2 relative to the \"CMA- \" lysosomes, which are high density and have higher levels of MTORC1. Because of safety concerns surrounding metrizamide, however, this compound is difficult to obtain, and it is impractically expensive. Here, we have provided protocols for isolation of lysosomal subpopulations for CMA-related analyses from mouse liver using Histodenz, a safe and affordable alternative to metrizamide. Supplementary protocols show how to perform CMA activity assays with appropriate statistical analysis, and how to analyze for lysosomal breakage/membrane integrity. © 2024 The Authors. Current Protocols published by Wiley Periodicals LLC. Basic Protocol: Isolation of lysosomal subpopulations from mouse liver using discontinuous Histodenz gradients Alternate Protocol: Isolation of lysosomes from cultured cells using discontinuous Histodenz gradients Support Protocol 1: Verifying enrichment of lysosomal markers in lysosome-enriched fractions Support Protocol 2: Measuring in vitro uptake of CMA substrates Support Protocol 3: Measuring lysosomal membrane integrity by hexosaminidase assay.

The subfractionation of the endoplasmic reticulum (ER) is a widely used technique in cell biology. However, current protocols present limitations such as low yield, the use of large number of dishes, and contamination with other organelles. Here, we describe an improved method for ER subfractionation that solves other reported methods\' main limitations of being time consuming and requiring less starting material. Our protocol involves a combination of different centrifugations and special buffer incubations as well as a fine-tuned method for homogenization followed by western blotting to confirm the purity of the fractions. This protocol contains a method to extract clean ER samples from cells using only five (150 mm) dishes instead of over 50 plates needed in other protocols. In addition, in this article we not only propose a new cell fractionation approach but also an optimized method to isolate pure ER fractions from one mouse liver instead of three, which are commonly used in other protocols. The protocols described here are optimized for time efficiency and designed for seamless execution in any laboratory, eliminating the need for special/patented reagents. Key features • Subcellular fractionation from cells and mouse liver. • Uses only five dishes (150 mm) or one mouse liver to extract highly enriched endoplasmic reticulum without mitochondrial-associated membrane contamination. • These protocols require the use of ultracentrifuges, dounce homogenizers, and/or Teflon Potter Elvehjem. As a result, highly enriched/clean samples are obtained. Graphical overview.

Detailed analysis of mitochondrial function cannot be achieved without good quality preparations of isolated mitochondria. Ideally, the isolation protocol should be quick, while producing a reasonably pure pool of mitochondria that are still intact and coupled. Here, we describe a fast and simple method for the purification of mammalian mitochondria relying on isopycnic density gradient centrifugation. We describe specific steps that should be taken into consideration when functional mitochondria from different tissues should be isolated. This protocol is suitable for the analysis of many aspects of the organelle\'s structure and function.

Mitochondria play a key role in cellular metabolism. Analyses of the genome, the proteome, metabolic, physiological, and biochemical functions of mitochondria frequently require the isolation of intact and functional mitochondria from various plant tissues with sufficient yield. For this purpose, we generated a transgenic Arabidopsis thaliana (Arabidopsis) line which presents a triple hemagglutinin tag on the surface of the outer mitochondrial membrane. The affinity tag enables immunocapture of the organelles in a single step. This chapter gives detailed instructions on how to generate transgenic Arabidopsis lines harboring a ubiquitously expressed 3xHA-sGFP-TOM5 mitochondrial fusion protein that is targeted to the outer mitochondrial membrane and enables purification of the organelles in a single step.

Many biochemical and biophysical related questions require the isolation of functional synaptic vesicles. Isolated synaptic vesicles can be used for transporter kinetics studies, synaptic vesicle content analysis and immuno-labeling of specific synaptic vesicle proteins, etc. Here I describe a fast and reliable isolation procedure to allow researchers to isolate a large amount, as well as physiologically functional synaptic vesicles, by following the subsequent order of cryogrinding, gradient ultracentrifugation, and size exclusion liquid chromatography. This process enriches over 90% of the synaptic vesicle population, with low contamination of Golgi or endoplasmic reticulum vesicles.

Mitochondria are highly dynamic organelles that are vital for eukaryotic cellular metabolism. Mitochondria account only for a small fraction of the total cell volume. As such, intact organelle isolation is a basic requirement to study mitochondrial biochemistry and physiology. However, established isolation procedures are tedious, and require a substantial amount of tissue material to start with. We addressed this matter and developed a single step purification for Arabidopsis thaliana (Arabidopsis) mitochondria. Ubiquitous expression of an 3xHA-sGFP-TOM5 fusion protein in the outer mitochondrial membrane allows affinity purification by immunocapture in a single step. We here provide a detailed step-by-step protocol for rapid immunocapture of intact and physiologically active mitochondria from Arabidopsis plants.

Endosomal microautophagy (eMI) is a type of autophagy that allows for the selective uptake and degradation of cytosolic proteins in late endosome/multi-vesicular bodies (LE/MVB). This process starts with the recognition of a pentapeptide amino acid KFERQ-like targeting motif in the substrate protein by the hsc70 chaperone, which then enables binding and subsequent uptake of the protein into the LE/MVB compartment. The recognition of a KFERQ-like motif by hsc70 is the same initial step in chaperone-mediated autophagy (CMA), a form of selective autophagy that degrades the hsc70-targeted proteins in lysosomes in a LAMP-2A dependent manner. The shared step of substrate recognition by hsc70, originally identified for CMA, makes it now necessary to differentiate between the two pathways. Here, we detail biochemical and imaging-based methods to track eMI activity in vitro with isolated LE/MVBs and in cells in culture using fluorescent reporters and highlight approaches to distinguish whether a protein is a substrate of eMI or CMA.

Intracellular Ca2+ is strictly regulated to maintain optimal levels for function of cellular organelles as well as mitochondrial respiratory signaling at the tricarboxylic acid cycle and electron transport chain level. Optimal Ca2+ concentration for these processes vary between cell types. Furthermore, exposure of mitochondria to sustained, elevated levels of Ca2+ induces mitochondrial Ca2+ overload and damage to mitochondrial oxidative phosphorylation and ATP production. Isolated mitochondria are widely used to study mitochondrial physiology and drug effects on mitochondrial metabolism and respiratory function. However, isolated mitochondria are easily damaged during the mitochondrial isolation process. The present article describes a mitochondrial isolation method using Ca2+-chelation to minimize mitochondrial damage. We follow up the isolation process with an application that requires an optimized buffer Ca2+ concentration: the characterization of their respiratory function using a high-resolution respirometric assay.

Diatoms are unicellular algae and evolved by secondary endosymbiosis, a process in which a red alga-like eukaryote was engulfed by a heterotrophic eukaryotic cell. This gave rise to plastids of remarkable complex architecture and ultrastructure that require elaborate protein importing, trafficking, signaling and intracellular cross-talk pathways. Studying both plastids and mitochondria and their distinctive physiological pathways in organello may greatly contribute to our understanding of photosynthesis, mitochondrial respiration and diatom evolution. The isolation of such complex organelles, however, is still demanding, and existing protocols are either limited to a few species (for plastids) or have not been reported for diatoms so far (for mitochondria). In this work, we present the first isolation protocol for mitochondria from the model diatom Thalassiosira pseudonana. Apart from that, we extended the protocol so that it is also applicable for the purification of a high-quality plastids fraction, and provide detailed structural and physiological characterizations of the resulting organelles. Isolated mitochondria were structurally intact, showed clear evidence of mitochondrial respiration, but the fractions still contained residual cell fragments. In contrast, plastid isolates were virtually free of cellular contaminants, featured structurally preserved thylakoids performing electron transport, but lost most of their stromal components as concluded from Western blots and mass spectrometry. Liquid chromatography electrospray-ionization mass spectrometry studies on mitochondria and thylakoids, moreover, allowed detailed proteome analyses which resulted in extensive proteome maps for both plastids and mitochondria thus helping us to broaden our understanding of organelle metabolism and functionality in diatoms.