Conformational diseases, such as Alzheimer\'s, Parkinson\'s and Huntington\'s diseases as well as ataxias and fronto-temporal disorders, are part of common class of neurological disorders characterised by the aggregation and progressive accumulation of mutant proteins which display aberrant conformation. In particular, Huntington\'s disease (HD) is caused by mutations leading to an abnormal expansion in the polyglutamine (poly-Q) tract of the huntingtin protein (HTT), leading to the formation of inclusion bodies in neurons of affected patients. Furthermore, recent experimental evidence is challenging the conventional view of the disease by revealing the ability of mutant HTT to be transferred between cells by means of extracellular vesicles (EVs), allowing the mutant protein to seed oligomers involving both the mutant and wild type forms of the protein. There is still no successful strategy to treat HD. In addition, the current understanding of the biological processes leading to the oligomerization and aggregation of proteins bearing the poly-Q tract has been derived from studies conducted on isolated poly-Q monomers and oligomers, whose structural properties are still unclear and often inconsistent. Here we describe a standardised biochemical approach to analyse by isopycnic ultracentrifugation the oligomerization of the N-terminal fragment of mutant HTT. The dynamic range of our method allows one to detect large and heterogeneous HTT complexes. Hence, it could be harnessed for the identification of novel molecular determinants responsible for the aggregation and the prion-like spreading properties of HTT in the context of HD. Equally, it provides a tool to test novel small molecules or bioactive compounds designed to inhibit the aggregation of mutant HTT.

Fractionating and characterizing target samples are fundamental to the analysis of biomolecules. Extracellular vesicles (EVs), containing information regarding the cellular birthplace, are promising targets for biology and medicine. However, the requirement for multiple-step purification in conventional methods hinders analysis of small samples. Here, we apply a DNA origami tripod with a defined aperture of binders (e.g., antibodies against EV biomarkers), which allows us to capture the target molecule. Using exosomes as a model, we show that our tripod nanodevice can capture a specific size range of EVs with cognate biomarkers from a broad distribution of crude EV mixtures. We further demonstrate that the size of captured EVs can be controlled by changing the aperture of the tripods. This simultaneous selection with the size and biomarker approach should simplify the EV purification process and contribute to the precise analysis of target biomolecules from small samples.

Plant viruses depend on host cellular factors for their replication and movement. There are cellular proteins that change their localization and/or expression and have a proviral role or antiviral activity and interact with or target viral proteins. Identification of those proteins and their roles during infection is crucial for understanding plant-virus interactions and to design antiviral resistance in crops. Important host proteins have been identified using approaches such as tag-dependent immunoprecipitation or yeast two hybridization that require cloning individual proteins or the entire virus. However, the number of possible interactions between host and viral proteins is immense. Therefore, an alternative method is needed for proteome-wide identification of host proteins involved in host-virus interactions. Here, we present cell fractionation coupled with mass spectrometry as an option to identify protein-protein interactions between viruses and their hosts. This approach involves separating subcellular organelles using differential and/or gradient centrifugation from virus-free and virus-infected cells (1) followed by comparative analysis of the proteomic profiles obtained for each subcellular organelle via mass spectrometry (2). After biological validation, prospect host proteins with proviral or antiviral roles can be subject to fundamental studies in the context of basic biology to shed light on both virus replication and cellular processes. They can also be targeted via gene editing to develop virus-resistant crops.

While RNAs are soluble in vitro, their solubility may be altered when incorporated into some protein complexes inside the cell. The solubility phase transition of RNAs is thus indicative of changes in the function and activity of RNAs. Here, we present a protocol for the assessment of RNA solubility phase transition during Xenopus oocyte maturation. We describe steps for sample preparation, cell fractionation, RNA extraction, real-time PCR, and analysis of the obtained results. For complete details on the use and execution of this protocol, please refer to Hwang et al. (2023).1.

REAP+ is an enhanced version of the rapid, efficient, and practical (REAP) method designed for the isolation of nuclear fractions. This improved version, REAP+, enables fast and effective extraction of mitochondria, cytoplasm, and nuclei. The mechanical cell disruption process has been optimized to cerebral tissues, snap-frozen liver, and HT22 cells with remarkable fraction enrichment. REAP+ is well-suited for samples containing minimal protein quantities, such as mouse hippocampal slices. The method was validated by Western blot and marker enzyme activities, such as LDH and G6PDH for the cytoplasmic fraction and succinate dehydrogenase and cytochrome c oxidase for the mitochondrial fraction. One of the outstanding features of this method is its rapid execution, yielding fractions within 15 min, allowing for simultaneous preparation of multiple samples. In essence, REAP+ emerges as a swift, efficient, and practical technique for the concurrent isolation of nuclei, cytoplasm, and mitochondria from various cell types and tissues. The method would be suitable to study the multicompartment translocation of proteins, such as metabolic enzymes and transcription factors migrating from cytosol to the mitochondria and nuclei. Moreover, its compatibility with small samples, such as hippocampal slices, and its potential applicability to human biopsies, highlights the potential application in medical research.

Information on RNA localisation is essential for understanding physiological and pathological processes, such as gene expression, cell reprogramming, host-pathogen interactions, and signalling pathways involving RNA transactions at the level of membrane-less or membrane-bounded organelles and extracellular vesicles. In many cases, it is important to assess the topology of RNA localisation, i.e., to distinguish the transcripts encapsulated within an organelle of interest from those merely attached to its surface. This allows establishing which RNAs can, in principle, engage in local molecular interactions and which are prevented from interacting by membranes or other physical barriers. The most widely used techniques interrogating RNA localisation topology are based on the treatment of isolated organelles with RNases with subsequent identification of the surviving transcripts by northern blotting, qRT-PCR, or RNA-seq. However, this approach produces incoherent results and many false positives. Here, we describe Controlled Level of Contamination coupled to deep sequencing (CoLoC-seq), a more refined subcellular transcriptomics approach that overcomes these pitfalls. CoLoC-seq starts by the purification of organelles of interest. They are then either left intact or lysed and subjected to a gradient of RNase concentrations to produce unique RNA degradation dynamics profiles, which can be monitored by northern blotting or RNA-seq. Through straightforward mathematical modelling, CoLoC-seq distinguishes true membrane-enveloped transcripts from degradable and non-degradable contaminants of any abundance. The method has been implemented in the mitochondria of HEK293 cells, where it outperformed alternative subcellular transcriptomics approaches. It is applicable to other membrane-bounded organelles, e.g., plastids, single-membrane organelles of the vesicular system, extracellular vesicles, or viral particles. Key features • Tested on human mitochondria; potentially applicable to cell cultures, non-model organisms, extracellular vesicles, enveloped viruses, tissues; does not require genetic manipulations or highly pure organelles. • In the case of human cells, the required amount of starting material is ~2,500 cm2 of 80% confluent cells (or ~3 × 108 HEK293 cells). • CoLoC-seq implements a special RNA-seq strategy to selectively capture intact transcripts, which requires RNases generating 5\'-hydroxyl and 2\'/3\'-phosphate termini (e.g., RNase A, RNase I). • Relies on nonlinear regression software with customisable exponential functions.

The Dynamic Organellar Maps (DOMs) approach combines cell fractionation and shotgun-proteomics for global profiling analysis of protein subcellular localization. Here, we enhance the performance of DOMs through data-independent acquisition (DIA) mass spectrometry. DIA-DOMs achieve twice the depth of our previous workflow in the same mass spectrometry runtime, and substantially improve profiling precision and reproducibility. We leverage this gain to establish flexible map formats scaling from high-throughput analyses to extra-deep coverage. Furthermore, we introduce DOM-ABC, a powerful and user-friendly open-source software tool for analyzing profiling data. We apply DIA-DOMs to capture subcellular localization changes in response to starvation and disruption of lysosomal pH in HeLa cells, which identifies a subset of Golgi proteins that cycle through endosomes. An imaging time-course reveals different cycling patterns and confirms the quantitative predictive power of our translocation analysis. DIA-DOMs offer a superior workflow for label-free spatial proteomics as a systematic phenotype discovery tool.

Peroxisomes are ubiquitous organelles with essential functions in numerous cellular processes such as lipid metabolism, detoxification of reactive oxygen species, and signaling. Knowledge of the peroxisomal proteome including multi-localized proteins and, most importantly, changes of its composition induced by altering cellular conditions or impaired peroxisome biogenesis and function is of paramount importance for a holistic view on peroxisomes and their diverse functions in a cellular context. In this chapter, we provide a spatial proteomics protocol specifically tailored to the analysis of the peroxisomal proteome of baker\'s yeast that enables the definition of the peroxisomal proteome under distinct conditions and to monitor dynamic changes of the proteome including the relocation of individual proteins to a different cellular compartment. The protocol comprises subcellular fractionation by differential centrifugation followed by Nycodenz density gradient centrifugation of a crude peroxisomal fraction, quantitative mass spectrometric measurements of subcellular and density gradient fractions, and advanced computational data analysis, resulting in the establishment of organellar maps on a global scale.

Sophisticated organelle fractionation strategies were the workhorse of early peroxisome research and led to the characterization of the principal functions of the organelle. However, even in the era of molecular biology and \"omics\" technologies, they are still of importance to unravel peroxisome-specific proteomes, confirm the localization of still uncharacterized proteins, analyze peroxisome metabolism or lipid composition, or study their protein import mechanism. To isolate and analyze peroxisomes for these purposes, density gradient centrifugation still represents a highly reliable and reproducible technique. This article describes two protocols to purify peroxisomes from either liver tissue or the HepG2 hepatoma cell line. The protocol for liver enables purification of peroxisome fractions with high purity (95%) and is therefore suitable to study low-abundant peroxisomal proteins or analyze their lipid composition, for example. The protocol presented for HepG2 cells is not suitable to gain highly pure peroxisomal fractions but is intended to be used for gradient profiling experiments and allows easier manipulation of the peroxisomal compartment, e.g., by gene knockdown or protein overexpression for functional studies. Both purification methods therefore represent complementary tools to be used to analyze different aspects of peroxisome physiology. Please note that this is an updated version of a protocol, which has been published in a former volume of Methods in Molecular Biology.

Subcellular fractionation in combination with mass spectrometry-based proteomics is a powerful tool to study localization of key proteins in health and disease. Here we offered a reliable and rapid method for mammalian cell fractionation, tuned for such proteomic analyses. This method proves readily applicable to different cell lines in which all the cellular contents are accounted for, while maintaining nuclear and nuclear envelope integrity. We demonstrated the method\'s utility by quantifying the effects of a nuclear export inhibitor on nucleoplasmic and cytoplasmic proteomes.