Methods and means for determining proteins and derivatives thereof in cells

Abstract

A method for determining at least one structural and / or physicochemical property, or a property thereof, of at least one protein or peptide comprised in a complex mixture of additional proteins and / or other biomolecules in at least one cell, the method comprises the steps of: 1. Delivering a protease into the cells and subjecting the complex mixture to limited proteolysis followed by cell extraction and / or cell lysis to produce a fragment sample; 2. Denaturing the fragment sample to obtain a denatured fragment sample; 3. Optionally, completely fragmenting the denatured fragment sample in a digestion step to obtain a completely fragmented sample; and 4, analytically analyzing the fragmented sample or the completely fragmented sample to determine the at least one protein or peptide or the characteristics thereof.

Description

Technical Field

[0001] This invention relates to methods for determining the properties of proteins and their derivatives in their native biological context, particularly at least one structural and / or physicochemical property, especially conformation and conformational changes, particularly using limited protein hydrolysis, for example in combination with: selected reaction monitoring, data-dependent acquisition (DDA), data-independent acquisition (DIA), including the Sequential Windowed Acquisition of All Theoretical Fragment Ion Mass Spectra (SWATH) method, etc. Existing technology

[0002] Protein function is determined by its structure, which dynamically changes to adapt to cellular perturbations. Protein structure changes due to post-translational modifications, interactions with other proteins or molecules, or in response to physical properties such as temperature.

[0003] Proteins perform a variety of functions in cells, regulating cell growth, differentiation, and survival. A protein's function is determined by its structure, which can change in response to the cellular environment and perturbations. Changes in protein structure can be minor, such as in the case of enzymes binding small regulatory molecules. Other proteins undergo complete remodeling and may even form aggregates, as seen in Alzheimer's disease or Parkinson's disease. To understand the function of proteins in health and disease, it is important to study protein structure and detect structural changes under various perturbations (such as stress, disease, or drug treatment). Protein structure is highly dynamic and adapts to its environment and interacting partners.

[0004] Available nuclear magnetic resonance (NMR) spectroscopy and Förster resonance energy transfer (FRET) techniques provide information on the dynamics of complete intracellular protein structures, but can only study one or a few proteins at a time. On the other hand, structural proteomics techniques, such as limited proteolysis coupled mass spectrometry (LiP-MS), are used to study the dynamics of thousands of protein structures simultaneously. Cryo-electron microscopy (cryoEM) can resolve protein structures in cells and tissues, but can only analyze proteins that meet specific size limitations. It allows only a limited degree of multiplexing and therefore cannot be applied globally in an unbiased manner. Finally, resolution and low throughput issues hinder the detection of protein conformational changes.

[0005] WO-A-2014082733 discloses the aforementioned limited proteolysis (LiP-MS) scheme, namely, a method for detecting the conformational state of proteins contained in complex mixtures of other proteins and / or other biomolecules, particularly in complex natural biological matrices, and the determination for such a method. The method comprises the following steps (if necessary, after extraction and / or lysis steps): 1. performing limited proteolysis on the complex mixture under conditions where the protein is in the conformational state to be detected, producing a first fragment sample; 2. denaturing the first fragment sample to obtain a denatured first fragment sample; 3. completely fragmenting the denatured first fragment sample in a digestion step to obtain a fully fragmented sample; 4. performing analytical analysis on the fully fragmented sample to identify characteristic fragments produced by both the limited proteolysis in step 1 and the complete fragmentation in step 3, thereby determining the conformational state.

[0006] In classic LiP-MS, native proteins are cleaved by nonspecific proteases within a short time, producing structure-specific peptide patterns. Mass spectrometry is used to identify these structural fingerprints. LiP-MS has been successful in capturing structural alterations at the proteome-wide scale in numerous applications. For example, it has enabled the identification of novel structural biomarkers for diseases, global detection of targets for drugs and small molecule metabolites, and the development of new analytical (“omics”) workflows for detecting altered pathways. Standard LiP-MS workflows capture the structural state of proteins in cell lysates.

[0007] Schopper et al., “Measuring protein structural changes on a proteomewidescale using limited proteolysis-coupled mass spectrometry”, *Nature Protocols*, Vol. 12, No. 11, October 26, 2017, pp. 2391-2410, reported protein structural changes induced by external perturbations or internal factors that can significantly affect protein activity and thus regulate cellular physiological states. Limited proteolysis-coupled mass spectrometry (LiP-MS) has been reported as a method to identify protein structural changes directly within the complex biological context of proteins at the whole proteome level. Following a targeted perturbation, the proteome extract undergoes a dual protease digestion step, in which a non-specific protease is applied under native conditions, followed by complete digestion with the sequence-specific protease trypsin under denaturing conditions. This sequential processing yields structure-specific peptides suitable for bottom-up MS analysis. Subsequently, a proteomics workflow involving shotgun or targeted MS and label-free quantification is applied to measure structure-dependent proteolysis patterns directly in the proteome extract. This paper reports potential applications of LiP-MS, including the direct detection of perturbation-induced protein structural changes in biological samples, identification of drug targets, detection of the structural state of disease-related proteins, and analysis of protein aggregates. This method can also identify specific protein regions involved in structural transitions or affected by binding events. Sample preparation takes approximately two days, followed by one to several days of MS and data analysis, depending on the number of samples analyzed.

[0008] Ma R. et al., “Chemo-selection strategy for limited proteolysis experiments on the proteomic scale”, Anal. Chem., Vol. 90, No. 23, November 7, 2018, pp. 14039-14047, describe a chemoselective enrichment strategy, called the semitryptic peptide enrichment strategy for proteolysis procedures (STEPP), for the separation of semitryptic peptides generated in mass spectrometry-based whole proteomics applications of limited proteolysis methods. This strategy involves reacting the ε-amino group of the lysine side chain and any N-terminus generated in the limited proteolysis reaction with an isobaric mass tag. Subsequent sample digestion with trypsin and a chemoselective reaction of the newly exposed N-terminus of the trypsin-cleaved peptides with N-hydroxysuccinimide (NHS)-activated agarose resin resulted in the removal of the trypsin-cleaved peptides from solution, leaving only hemitrypsin-cleaved peptides with a non-trypsin cleavage site generated in the limited proteolysis reaction for subsequent LC-MS / MS analysis. As part of this work, the STEPP technique combines two different proteolysis methods: pulse proteolysis (PP) and limited proteolysis (LiP). The STEPP-PP workflow was evaluated in two proof-of-concept experiments involving proteins from yeast cell lysates and two well-investigated drugs, cyclosporine A and geldanamycin. The STEPP-LiP workflow was evaluated in a proof-of-concept experiment involving proteins from two human breast cancer cell culture models, MCF-7 and MCF-10A cell lines. The STEPP protocol increases the number of hemitrypsin-cleaved peptides detected in LiP and PP experiments by 5 to 10 times. The STEPP protocol not only improves proteomic coverage but also increases the amount of structural information obtainable from limited proteolysis experiments. Furthermore, this protocol enables the quantitative determination of ligand binding affinity.

[0009] Heusel M. et al.: “Complex-centric proteome profiling by SEC-SWATHMS”, Mol. Syst. Biol., Vol. 15, No. 1, Article No. e8438, January 14, 2019, pp. 1-22, describes an integrated experimental and computational technique for quantifying hundreds of protein complexes in a single operation. The method comprises: size exclusion chromatography (SEC) to fractionate native protein complexes; SWATH / DIA mass spectrometry to precisely quantify proteins in each SEC fraction; and the computational framework CCprofiler to detect and quantify protein complexes using prior information from a universal protein interaction map through error-controlled, complex-centric analysis. The inventors' analysis of the HEK293 cell line proteome described 462 complexes composed of 2,127 protein subunits. This technique identified novel subcomplexes and assembly intermediates that centrally regulate complexes, while simultaneously assessing the quantitative subunit distribution within them. They made the CCprofiler toolset freely available and provided the web platform SECexplorer for custom exploration of the modularity of the HEK293 proteome.

[0010] WO-A-2023 / 099341 discloses a method for detecting the conformational state of a protein in a complex mixture of other proteins and other biomolecules, wherein the protein in the complex mixture has undergone conditions that induce structural changes in the protein. The method comprises the following steps in sequence: 1. performing limited proteolysis on an extract mixture under conditions where the protein is in the original conformational state to be detected, producing a first fragment sample; then directly proceeding to 2. removing large peptides and proteins or other biomolecules from the first fragment sample to form an enriched fragment sample; 3. performing analytical analysis on the enriched fragment sample to identify characteristic fragments produced by the limited proteolysis in step 1 and retained after the removal step 2, for determining the conformational state of the at least one protein.

[0011] Van Kelly et al. reported in Mol Cell Proteomics, 2022 Jan; 21(1); 100169. doi:10.1016 / j.mcpro.2021.100169; Epub November 4, 2021 that comprehensive proteomic analysis of rare cell phenotypes remains a significant challenge. They proposed a method for MS-based proteomics for low cell numbers, utilizing proteomic digestion of cells with mild formaldehyde fixation within the cells, which they termed “in-cell digestion.” They combined this with averaged MS1 precursor library matching to quantitatively characterize the proteome from low cell number human lymphoblasts. Approximately 4500 proteins were detected from 2000 cells, and 2500 proteins were quantified from 200 lymphoblasts. The simplicity of sample processing and high sensitivity make this method suitable for proteomic analysis of rare cell states, including immune cell subsets and cell cycle subphases. To demonstrate this method, they characterized proteomic changes across 16 cell cycle states (CCS) isolated from asynchronous TK6 cells, avoiding synchronization. These states included late-stage mitotic cells, which were present at very low frequencies. They identified 119 pseudo-periodic proteins that changed throughout the cell cycle. Clustering of these pseudo-periodic proteins showed abundance patterns consistent with protein degradation “waves” at the late S, G2&M boundary, metaphase, and telogen effluvium. These clusters were distinguished by predicted nuclear localization and significant differences in their interactions with the anaphase-promoting complex / cycle body. The dataset also identified putative anaphase-promoting complex / cycle body substrates during mitosis and the temporal order in which they were targeted for degradation. They showed that a protein signature comprised of these 119 high-confidence cell cycle regulatory proteins could be used to unbiasedly classify the proteome into CCSs. They applied this signature to 296 proteomes covering a range of quantitative methods, cell types, and experimental conditions. The analysis assigned CCSs to 49 proteomes, including the correct classification of proteomes from synchronized cells.

[0012] Schopper et al. reported in Nat Protoc, 2017 Nov; 12(11): 2391-2410; doi: 10.1038 / nprot.2017.100; Epub 26 October 2017 that protein structural changes induced by external perturbations or internal factors can significantly affect protein activity and thus regulate cellular physiological states. Many biophysical methods are available for probing protein structural changes, but these are not applicable to the whole proteome in biological extracts. Limited proteolysis-mass spectrometry (LiP-MS) is a proteomics method that can directly identify protein structural changes in the complex biological environment of proteins at the whole proteome level. Following the target perturbation, the proteome extract undergoes a dual protease digestion step, in which a non-specific protease is applied under native conditions, followed by complete digestion with the sequence-specific protease trypsin under denaturing conditions. This sequential processing yields structure-specific peptides suitable for bottom-up MS analysis. Next, proteomics workflows involving shotgun or targeted MS and label-free quantification are applied to directly measure structure-dependent proteolytic patterns in proteomic extracts. Potential applications of LiP-MS include directly identifying perturbation-induced protein structural changes in biological samples, identifying drug targets, detecting the structural state of disease-related proteins, and analyzing protein aggregates. This method can also identify specific protein regions involved in structural transitions or affected by binding events. Sample preparation takes approximately two days, followed by one to several days for MS and data analysis, depending on the number of samples analyzed. Sample preparation can be performed by personnel with basic biochemistry training. MS measurements and data analysis require a proteomics background.

[0013] In his 2013 NMR paper, "In-cell NMR Spectroscopy in Mammalian Cells," Bekei Beata reported that in vivo protein behavior within cells is influenced by many unique physical parameters, such as macromolecular crowding and intracellular viscosity, as well as by a variety of biological activities that often vary considerably across cell types. Intracellular NMR spectroscopy is a valuable biophysical tool for studying the residue specificity, structure, and function of isotopically labeled proteins in their native cellular environment, making it a preferred tool for investigating intracellular protein behavior. The main objective of this paper was to extend the general applicability of eukaryotic intracellular NMR measurements to include mammalian cells and to develop a universal protocol for the efficient delivery of isotopically labeled proteins into mammalian cells. The results showed that low transduction efficiency, high cell line dependence, and vesicle-like intracellular distribution significantly limited the applicability of CPP-mediated protein delivery attempts. While the SLO- and EP- procedures produce comparable transduction efficiencies at low applied protein concentrations, EP significantly outperforms the SLO method at higher protein concentrations because it enables linear delivery of increased concentrations of exogenous protein with a high correlation to intracellular protein levels. In the second part of the paper, the structural, dynamic, and functional details of human amyloid α-synuclein and its isoforms β and γ-synuclein in different mammalian cells are described for the first time, as determined by high-resolution intracellular NMR experiments. α-synuclein is a small (14.5 kDa) intrinsically disordered neuronal protein known to accumulate in cytoplasmic filamentous inclusions in dopaminergic neurons during neurodegenerative diseases known as synucleinopathy. High-resolution intracellular NMR studies were performed to provide new insights into the native in vivo properties of this protein, particularly regarding its physiological and pathological characteristics, such as intracellular aggregation, membrane association, and post-translational modifications. The selection of mammalian cell models included disease-independent (i.e., HeLa and A2780 cells) and disease-associated dopaminergic neuronal cell lines (i.e., B65 and SK-N-SH cells). High-resolution 2D 1H-15N SOFAST-HMQC spectra of the synuclein protein family revealed its monomeric behavior in five different mammalian cell lines, without significant conformational changes such as amphiphilic helical inversion or aggregation into β-sheet-rich high-molecular-weight oligomers or amyloid fibrils. Intracellular NMR samples of the synuclein protein family obtained under healthy physiological conditions have been reported to possess excellent spectral quality, offering broad prospects for intracellular NMR studies under Parkinson's disease-associated pathogenic conditions and for developing cell-based intracellular NMR models to investigate the intracellular aggregation of these proteins and other proteins. Summary of the Invention

[0014] Limited proteolysis-mass spectrometry (LiP-MS) captures structural alterations adapted to cellular perturbations at the whole-proteome level. In LiP-MS, proteins in native lysates are cleaved by nonspecific or low-specific proteases over a short period, producing structure-specific peptide patterns that are analyzed by mass spectrometry. However, cellular lysis can expose proteins to unnatural conditions due to subcellular compartmentalization and loss of intracellular gradients, and dilution can disrupt unstable interactions. This is, for example, detrimental to structures held together by weak interactions, such as phase-separated protein assemblies.

[0015] Protein structures are highly dynamic and adapt to their environment and interacting pairs. Techniques for determining high-resolution protein structures typically require the purification of proteins or protein complexes. However, using purified proteins is insufficient to capture the perturbation-specific protein dynamics. Cells provide a highly crowded environment where proteins vary with physical parameters such as temperature or pH. In vitro environments cannot simulate the complex intracellular molecular gradients. Protein structures may vary according to their subcellular localization, and low-affinity interacting pairs are lost during cell lysis and purification. One example of this is stress granules, unstable macromolecular assemblages of proteins and RNA that form under stress conditions and are not yet fully characterized at the structural level. Therefore, it is important to study changes in protein structure in their native environment.

[0016] Cellular perturbations typically involve a cascade of multiple proteins over time, ranging from seconds to minutes. These changes involve the binding of small regulatory molecules, post-translational modifications, and alterations in interacting partner molecules. All of these events affect protein structure, while changes in protein abundance occur on longer timescales. The complex interactions of protein structural alterations within pathways significantly influence cellular physiological status.

[0017] To avoid potential protein structural changes during cell lysis and to study protein structures in their cellular environment, the inventors developed a novel method they call Intracellular LiP-MS.

[0018] Until now, LiP-MS has not been able to be applied to intact cells, thus preventing the conduct of limited proteolytic reactions under physiological conditions that maintain molecular crowding, intracellular interactors, intracellular gradients, and compartmentalization within the cell. This is a significant limitation of the current LiP-MS workflow, as cell lysis is likely to lead to changes in at least some protein structures and will inevitably expose proteins to non-physiological conditions (e.g., dilution of cellular contents, loss of interactors, loss of the subcellular environment, etc.). Intracellular LiP-MS overcomes this limitation.

[0019] Currently, other techniques enable the study of intracellular protein structures. Protein surface painting techniques, such as hydroxyl radical foot printing (HRF)-MS or lysine residue demethylation, detect changes in protein surface accessibility by oxidizing with hydroxyl radicals. This method has been applied to intact cells or organisms (such as *C. elegans*) and offers residue-level resolution. It utilizes microfluidic systems and laser devices, while intracellular LiP-MS uses commercially available electroporation equipment. Furthermore, compared to LiP-MS, this method currently exhibits lower reproducibility and significantly lower coverage.

[0020] Thermal proteome profiling (TPP) provides information about proteins that have undergone changes in intracellular stability, but unlike intracellular LiP, it does not provide structural details, i.e., it cannot identify regions of protein sequence / structure that have changed. Furthermore, to collect soluble fractions, cells are lysed and then centrifuged, leading to the loss of information under physiological conditions.

[0021] Nuclear magnetic resonance (NMR) and Foster resonance energy transfer (FRET) techniques provide information on the dynamics of intracellular protein structure, but typically only one protein can be studied at a time. Intracellular LiP-MS provides peptide-level resolution and can simultaneously target thousands of proteins.

[0022] Compared to protein mapping techniques, intracellular LiP-MS offers significantly higher proteomic and protein sequence coverage, as well as superior reproducibility. Compared to TPP, it enables the detection of structural features at peptide-level resolution. Compared to standard LiP-MS, it overcomes the need for cell lysis and the resulting alterations in protein structure and loss of physiological condition. Compared to FRET or NMR, it can simultaneously detect the structure of thousands of proteins, rather than just a single target protein, and does not require labeling (although it cannot determine the 3D coordinates of protein structures like NMR).

[0023] To preserve the natural environment and study protein structure within its cellular context, this paper presents an intracellular LiP-MS protocol. Low-specificity protease proteinase K is introduced into the cell using, for example, electroporation.

[0024] It has been demonstrated that intracellular LiP-MS not only captures known specific structural changes, such as rapamycin binding of FKBP1A, but also captures downstream effects of pathway activation. Furthermore, intracellular LiP-MS captures the formation of phase-separated protein assemblies held together by weak interactions. Using intracellular LiP-MS, the inventors have generated the first in-situ dataset on global protein structural changes during stress granule formation following sodium arsenite treatment. Structural changes of known stress granule components (such as G3BP1) were observed, and a resource was generated on peptide-level structural changes in response to sodium arsenite treatment for hundreds of other proteins. Additionally, it was shown that SERBP1 and PPP1R12A are involved in the early stages of the cellular response to arsenite stress. Furthermore, intracellular LiP provides evidence that phase-separated compartments of nuclear spots change after sodium arsenite treatment, which the inventors verified using fluorescence microscopy.

[0025] A novel approach using intracellular LiP-MS was proposed to study a variety of biological processes in their natural environment, offering peptide-level resolution and enabling studies across the entire proteome.

[0026] Therefore, to study protein structures in their natural environment, a step is used to introduce low-specificity proteases (such as proteinase K) into living cells (e.g., by increasing cell membrane permeability via electroporation). This is preferably performed under optimized conditions (see below). If incubated briefly within the cell, proteinase K cleaves the protein in a solvent-accessible and flexible region. This produces protein fragments that reflect the specific structural state of the protein. The cells are then lysed, and optionally the extracted fragments are further processed with a protease (such as trypsin) to make them suitable for mass spectrometry analysis while preserving structural information, and subsequently identified and quantified using a mass spectrometry-based proteomics workflow. The new combination of increasing cell membrane permeability (e.g., electroporation, a method commonly used to study single proteins using nuclear magnetic resonance (NMR) spectroscopy) with established limited proteolysis-mass spectrometry (LiP-MS) allows for the simultaneous study of thousands of protein structures in their natural environment and how they adapt to a variety of biological perturbations.

[0027] In a typical exemplary experiment, cells are treated with an active compound, exposed to stress conditions, or subjected to any other purposeful perturbation. Cells grown on cell culture dishes are detached and resuspended in physiological buffer (e.g., phosphate-buffered saline, PBS). Next, a protease (e.g., a low-specificity protease such as proteinase K) is added to the cell suspension at a high concentration (e.g., 100 μM). Immediately thereafter, the cells are electroporated to permeate the cell membrane. This allows the protease to diffuse into the cells. After electroporation, the cells are incubated briefly at physiological temperature (typically 1 to 5 minutes at 37°C). Optionally, the cells are then washed with physiological buffer by centrifugation to remove extracellular molecules. A high concentration of a dissociative agent (e.g., 7 M guanidine chloride) is then added. Cells were lysed with a lysis buffer containing guanidinium chloride and heated to quench the protease reaction. Cells were lysed, and fragments were optionally trypsinized to reduce their size, and peptides were analyzed by mass spectrometry. Data from treated and control cells were quantitatively compared, and fragments with varying amounts under different conditions were identified and mapped to protein sequences and (if available) protein structures. This analysis revealed which proteins altered their structure in response to the desired perturbation and precisely identified the altered regions.

[0028] The pulse number and voltage of electroporation can be optimized to allow maximum protease diffusion without impairing cellular physiology and viability. The concentration of the protease and the incubation time after electroporation can be optimized to obtain the maximum number of reproducible structure-specific cleavages. Following electroporation, structure-specific cleavage of thousands of proteins significantly increases in the presence of proteinase K, whereas cleavage occurs primarily at the cell surface without cell electroporation. Electroporation-dependent uptake of proteinase K in target cells can be confirmed using fluorescence Foster resonance energy transfer (FRET)-based protein sensors known to be cleaved by proteinase K. For example, the fluorescent proteins ECFP and YPet are linked via flexible peptide linkers. Upon irradiation of ECFP, energy is internally transferred to YPet, and YPet fluorescence is detected. When cells containing this sensor are incubated with proteinase K and then electroporated, allowing proteinase K to diffuse into the cells, the fluorescent substrate is cleaved, and this cleavage is monitored by the loss of YPet fluorescence and the increase in ECFP fluorescence. However, if proteinase K is added but not electroporated into the cells, the substrate remains intact.

[0029] The novel intracellular LiP-MS method not only captures known protein structural changes but also provides additional biological information compared to established LiP-MS methods using cell extracts. For example, in HEK293 cells treated with 100 nM or 10 μM rapamycin (the drug rapamycin) for 5 or 10 minutes, intracellular LiP-MS detected known structural changes at FKBP1A (the protein target of rapamycin). Furthermore, proteins involved in downstream biological processes also exhibited structural changes that were not observed when LiP-MS was applied to natural lysates. Therefore, intracellular LiP-MS allows for the capture of biological events occurring under physiological conditions. In the case of drugs, it allows for the capture of intracellular target engagement and the pathways underlying the mechanism of action of compounds.

[0030] Furthermore, intracellular LiP-MS can detect the structural features and structural transitions of unstable protein complexes (which are typically disrupted during cell lysis). For example, intracellular LiP-MS has been used to study stress granule formation in mammalian cells. Stress granules are unstable assemblages of proteins and RNA that form under stress conditions. HEK293 cells were treated with 500 μM sodium arsenite for 10, 20, and 90 minutes and compared with untreated cells. Using intracellular LiP-MS, structural changes in a variety of known stress granule proteins were observed after inducing stress granule formation, indicating that this method detects stress granule assembly. Gene ontology enrichment analysis showed that after 20 and 90 minutes of sodium arsenite treatment, structural changes were concentrated in stress granule-related proteins. At 10 minutes of arsenite treatment, intracellular LiP-MS had already identified structural changes in a few specific stress granule proteins before stress granules were visible by fluorescence microscopy.

[0031] Essential for intracellular LiP-MS is the delivery of the protease into the cell, followed by quenching of protease activity and / or cell lysis, and then the remaining standard components of the LiP-MS or Dark-LiP-MS workflow. Delivery via electroporation is specifically illustrated. Possible variations of the protocol enabling limited proteolytic steps within the cell include: delivery of the protease via a cell permeation agent (e.g., detergent or toxin); conjugation of the protease to a cell-penetrating agent (e.g., peptide); delivery within a caging system and vesicles; intracellular expression of the active state of the protease under the control of a tight promoter; and expression of an inactive form of the protease that can be activated upon request (e.g., light-cageed protease, protease expressed in the presence of an inhibitor, or protease conjugated to a degron system). Quenching of protease activity can be achieved through a variety of conditions that denature the protein or inhibit its enzymatic activity, such as increasing temperature, transferring cells to a medium that simultaneously lyses the cells and denatures the proteins (e.g., a mixture of detergent and denaturant compatible with mass spectrometry, a high-concentration acid solution such as trichloroacetic acid), or performing cell lysis in the presence of protease inhibitors. Variations of the remaining components of the LiP-MS workflow have been described in previous patents describing the LiP-MS invention.

[0032] To date, intracellular LiP-MS has been tested on HEK293 cells, but this method also allows for the study of any other cell types, including mammalian cell lines, primary cells, tissues, bacteria, yeast, etc. The setup has been optimized for batch electroporation of 3 million cells, but the electroporation setup can be adjusted to achieve scale-down, even down to single-cell electroporation. Proteinase K was used because of its high activity at physiological temperatures, but any other protease commonly used for structure-specific proteolytic cleavage (e.g., subtilisin, thermophilic protease, papain) can also be used. Optionally, a targeting sequence for specific subcellular localization can be coupled to the protease to be introduced into the cell to enable the probing of structural features of specific subcellular compartments (e.g., nuclear-targeted sequencing).

[0033] LiP-MS allows for the identification of drug or other compound targets within cells and under specific conditions. This method simultaneously provides information downstream of protein-target binding events and can be used to identify regulated pathways and off-target effects. For example, cells grown under hypoxic conditions in a tumor environment may exhibit different cellular responses to drugs. Intracellular LiPs can be used to detect target binding, downstream events, and off-target events under these conditions. Furthermore, intracellular LiP-MS can be used to determine optimal drug concentrations and report the time required for drug uptake and target binding. It can also help elucidate the impact of mutations in pathways affected by drug binding. Additionally, intracellular LiP-MS can be used to study how cells respond when two or more drugs are added simultaneously or sequentially.

[0034] In addition to drug binding, intracellular LiP-MS can also identify protein structural changes induced by specific cellular conditions, such as changes in temperature or pH, which also occur in the tumor microenvironment. LiP-MS can also provide information on global cellular responses to other environmental factors, toxins, and radiation.

[0035] Intracellular LiP-MS can provide mechanistic information about any cellular process involving structural changes, including viral entry, binding of modified T cells during immunotherapy, or nutrient conversion.

[0036] Generally, the proposed invention relates to a method for determining at least one protein or peptide or its properties (preferably simultaneously determining more than one protein or peptide, such as more than 100 or more than 1000 proteins and / or peptides, their structural and / or physicochemical properties), said at least one protein or peptide being contained in a complex mixture of other proteins and / or other biomolecules in at least one cell, said method comprising the following steps:

[0037] 1. Delivering a protease into the (preferably live) cells and performing limited proteolysis on a complex mixture, followed by cell extraction and / or cell lysis to produce a fragment sample (preferably this step is performed by applying an electric field (only) to the cells to increase cell membrane permeability, particularly by using (pulsed) electroporation).

[0038] 2. Denature the fragment sample to obtain a denatured fragment sample;

[0039] 3. Optionally, the denatured fragment sample is completely fragmented during the digestion step to obtain a fully fragmented sample;

[0040] 4. Perform analytical analysis on the fragmented or fully fragmented sample to determine the at least one protein or peptide or its characteristics.

[0041] Step 4, the resolution analysis of fragmented or fully fragmented samples, provides in-cell readouts of the structure and / or physicochemical properties of proteins or peptides. Preferably, the resolution analysis is based on mass spectrometry.

[0042] This method is specifically used to determine at least one structural and / or physicochemical property of the at least one protein or peptide.

[0043] As mentioned, preferably, step 1 is performed by applying an electric field to the cells to increase cell membrane permeability, i.e., applying an electrostatic field in the cell solution. Pulsed electroporation is preferred, for example, applying at least one or more pulses, such as pulses exceeding 300 V / cm or 500 V / cm, each preferably with a length in the range of 20 to 100 ms (generally, the voltage used is in the range of 500 to 1500 V, typically in the range of 800 to 1200 V).

[0044] It should be noted that denaturation in step 2 is not necessary and can be omitted, as is step 3. Denaturation in step 2 is particularly important when a second digestion step 3 is performed. If direct analysis of the large fragment from step 1 by mass spectrometry is desired, denaturation and digestion, i.e., steps 2 and 3, can be completely skipped.

[0045] Therefore, this method can also be used to determine at least one protein or peptide or its properties (particularly to determine at least one structural and / or physicochemical property of said at least one protein or peptide or multiple said proteins or peptides), said at least one protein or peptide being contained in a complex mixture of other proteins and / or other biomolecules in at least one cell, the method comprising the following steps:

[0046] 1. Delivering a protease into the (preferably live) cells and performing limited proteolysis on a complex mixture, followed by cell extraction and / or cell lysis to produce a fragment sample (preferably this step is performed by applying an electric field (only) to the cells to increase cell membrane permeability, particularly by using (pulsed) electroporation).

[0047] 4. Perform analytical analysis on the fragment sample to determine the at least one protein or peptide or its characteristics. Preferably, the analytical analysis is based on mass spectrometry.

[0048] It is noteworthy that this method is fundamentally different from classical proteomics methods that measure protein abundance (rather than structure), such as those reported in the aforementioned paper by Van Kelly et al.: because their method aims to measure protein abundance, it is not necessary to preserve the native protein structure. Therefore, they permeabilized cells with 90% methanol, which partially dissolved the cell membrane and affected protein structure. This step thus hindered protein structure analysis. Additionally, Kelly et al. performed benzonase treatment, which disrupted all interactions between the protein and DNA / RNA, further impacting the possibility of analyzing the native protein structure. Furthermore, the cells used by Kelly et al. were dead. They were fixed in formalin and then permanently permeabilized. In the method presented here, the cells are preferably alive. After, for example, short-pulse electroporation, the pores in the cell membrane close again, and the cells remain viable. Importantly, Kelly et al. performed classic complete trypsin digestion (a very long time, 16 hours). This means that Kelly et al., like Van Kelly, did not digest based on protein structure, but rather on the presence of specific amino acids in the sequence. Trypsin is a sequence-specific protease, while proteinase K is not. Therefore, within 16 hours, trypsin will cleave at any location (approximately every 10 residues) in the protein, regardless of structure, just as in classical proteomics analyses measuring abundance. In the setup proposed here, time-limited digestion steps (typically within minutes) using proteinase K, for example, make the cleavage structure-specific (proteinase K will only cleave at the most accessible regions on each protein structure). Furthermore, Kelly et al. performed only one digestion step with trypsin, which is also consistent with classical proteomics analyses measuring protein abundance. The proposed workflow involves double digestion, for example with proteinase K, followed by digestion with trypsin, for example. The first digestion with proteinase K, for example, is structure-based, and the second digestion is only to reduce the fragment size to make it compatible with mass spectrometry analysis.

[0049] Furthermore, the proposed method is fundamentally different from NMR methods that only allow structural tracking of one protein / peptide, because the proposed method here allows and is preferably used to screen for the structure of more than one, preferably dozens, hundreds or thousands of peptides / proteins simultaneously, and to determine which of these peptides / proteins (if any) undergo structural changes under target conditions, or simply to determine their structural features.

[0050] Regarding the particular suitability of this method for determining at least one structural and / or physicochemical property of the at least one protein or peptide or multiple thereof, the following aspects are relevant: Structural properties also include conformational properties, which highlight the potential for structural dynamics. By determining such structural properties, changes in the "function" of a protein can typically be detected. In other words, structural changes can be detected by the proposed method, and from this information, it can be deduced that the protein involved has also undergone a functional change (e.g., an activity change, if referring to an enzyme). Therefore, the structural and / or physicochemical properties include the corresponding relevant functional properties of the at least one protein or peptide or multiple thereof.

[0051] Therefore, the proposed method allows for the determination / derivation of structure and / or biophysical properties, the detection of structural differences at specific protein regions, and thereby the detection of proteins with altered functions. If the obtained information is combined with modeling tools (such as AlphaFold), even the 3D coordinates of atoms within the structure can be derived.

[0052] According to a first aspect of the invention, it relates to a method for determining the intracellular properties of a protein or peptide by using intracellular limited proteolysis via delivery of a protease into the cell. Using this method, the structure and / or physicochemical properties of a protein or peptide can be determined in situ within the cell, i.e., in a natural biological environment. Therefore, this proposed method, for the first time, allows for the direct derivation of intracellular structural information using mass spectrometric analysis, particularly of fragmented samples. This can be used for a variety of purposes, such as assessing whether the intracellular structure resembles an in vitro structure obtained from a purified protein, or whether the intracellular protein binds to a drug in the same manner as the corresponding purified protein.

[0053] However, if the method relates to comparing the properties of intracellular proteins and / or peptides as a function of or response to external or internal perturbations applied to or to a cell or cell population, then the method may also be, and is particularly applicable.

[0054] According to a second aspect of the invention, it therefore relates to a method for detecting a state change of a protein or peptide in response to a perturbation, wherein the protein or peptide is contained in a complex mixture of other proteins and / or other biomolecules in at least two cells, the method comprising the steps of:

[0055] 1. Perturbing at least one cell in the said cells, followed by or accompanied by delivery of a protease into the cells, and limited proteolysis of a complex mixture, followed by cell extraction and / or cell lysis, to produce a fragment sample;

[0056] 2. Denature the first fragment sample to obtain a denatured fragment sample;

[0057] 3. In the digestion step, the denatured first fragment sample is completely fragmented to obtain a fully fragmented sample (this step is optional).

[0058] Steps 1 to 3 are performed on at least one other cell among the said cells, but in step 1, the at least one other cell is not subjected to the perturbation, to produce a fully fragmented control sample or a fragmented control sample.

[0059] 4. Perform analytical analysis on the fragmented sample or the completely fragmented sample, as well as the completely fragmented control sample or the fragmented control sample, to determine the state changes of the protein.

[0060] The term "change of state" for at least one protein or peptide should be broadly understood as commonly accepted in the art, referring to a change in the sequence and / or spatial arrangement of the constituent atoms of a protein that determine the overall shape of the molecule, or a change in the physicochemical characteristics of a protein or one of its regions (e.g., changes in hydrophobicity or solvent accessibility due to the binding of small molecules to the protein). In other words, the expression "change of state" is not limited to conformational changes, but includes any type of change information, including changes in primary structure as well as information beyond primary structure (i.e., changes in the linear sequence of amino acids in a peptide or protein and their potential chemical modifications). Thus, the expression "change of state" includes changes in primary structure as well as changes in secondary structure (the three-dimensional arrangement of local segments of a protein; the two most common secondary structure elements are α-helices and β-sheets, but also β-turns and Ω-loops), supersecondary structures (motifs, compact three-dimensional protein structures of several adjacent secondary structure elements, smaller than protein domains or subunits), tertiary structures (domains, the three-dimensional shape of a protein), and quaternary structures (protein structures composed of two or more smaller protein chains, also called subunits). When a drug binds to a protein, it does not always induce a conformational change in the protein. It only alters protein properties, such as solvent accessibility, and this is sufficient to change the degree of fragmentation induced by proteinase K. This is why the proposed method can be used to detect protein-drug interactions, which is one of the important applications of the proposed intracellular LiP, and therefore state changes also broadly include such property changes.

[0061] The conformational changes of proteins that can be identified using the proposed method are made possible by their inherent flexibility. These changes can occur with relatively little energy expenditure. At the molecular structural level, conformational changes in a single polypeptide result from changes in the main chain twist angle and side chain orientation. The overall effect of such changes can be limited to the reorientation of a few residues and small twisting changes in the regional main chain. On the other hand, twisting changes limited to a very few critical residues can lead to large tertiary structural changes. The latter type of conformational change is described as domain movement. Conformational changes associated with protein-protein interactions range from local changes in the state of side chain spinomeric states to global structural changes (e.g., collective domain movement).

[0062] When perturbation is mentioned in step 1, followed by or accompanied by the delivery of the protease into the cells, this includes cases where cell perturbation occurs early (e.g., in patient biopsies or cells with mutations), and limited proteolysis occurs only significantly later. It also includes cases where perturbation is applied slightly before or substantially immediately before protease delivery, i.e., the protease is delivered directly into the cells after perturbation. However, it also includes cases where perturbation is applied substantially at the same time as or simultaneously with the delivery of the protease into the cells, initiating limited proteolysis. This is, for example, if the applied perturbation and the delivery of the protease into the set cells are actually applied simultaneously using the same mechanism; thus, for example, if electroporation is used to introduce the protease, and also to introduce the perturbation into the cells. Therefore, the timing between the perturbation and the onset of limited proteolysis can also be used to determine the time dependence of the induced changes. For cases where a perturbation is applied first, followed by the initiation of limited proteolysis, this time dependence can be monitored as a function of the time elapsed between the initiation of the perturbation and the delivery of the protease to the cell and the limited proteolysis. Alternatively, particularly if the timescale of the perturbation is shorter than that of the limited proteolysis, changes under limited proteolysis conditions can be monitored as a function of the time elapsed after the initiation of the perturbation in different experiments. Note that, as detailed below, if the protease is delivered to the cell in an inactivated form, it can be activated in situ within the cell by applying a corresponding stimulus; this also includes cases where the delivery of the inactivated protease occurs long before the application of the perturbation, followed by the application of the perturbation, and subsequently, or accompanied by the initiation of limited proteolysis by applying a stimulus to activate the protease.

[0063] In the context of this invention, the term "cell" includes mammalian cells, but also any other type of animal, bacterial, or plant cell, and therefore includes cells with or without cell walls, eukaryotic cells, and prokaryotic cells. Furthermore, the term includes mature, differentiated cells, but also stem cells or progenitor cells. In particular, the method is of interest if applied to differentiated mammalian cells, mammalian progenitor cells, or mammalian stem cells. Cells can be, for example, fibroblasts, neurons, epithelial cells, hepatocytes, lymphocytes, and macrophages.

[0064] The analysis in step 4 is typically based on a quantitative comparison of the analytical analysis of the fully fragmented sample or the fragmented sample with the analytical analysis of the fully fragmented control sample or the fragmented control sample.

[0065] For the analytical analysis in step 4, mass spectrometry techniques are preferred, such as selected / multiple reaction monitoring (SRM / MRM), data-independent acquisition (DIA) of product ion spectra (including SWATH-MS), and / or data-dependent acquisition (DDA or shotgun method).

[0066] The proteolytic system used in step 1 can be selected from the group of (low-specificity) proteases, such as proteinase K, thermophilic protease, subtilisin, pepsin, papain, α-chymotrypsin, elastase, figase, streptomycin, and mixtures thereof. Note that these proteolytic systems can be supplied and delivered to the cell in either an active or inactivated form. In the latter case, limited proteolysis can be initiated in step 1 by applying a stimulus that activates the corresponding protease in situ within the cell. Furthermore, the proteolytic system can also be expressed intracellularly; therefore, in the context of this invention, "delivery" includes cases where the protease system is not introduced into the cell as a whole, but is expressed in the cell or formed in the cell using another mechanism (e.g., the introduction of a fragment, which is subsequently assembled in the cell).

[0067] The protease used is preferably a low-specificity protease, and is therefore typically characterized as an exopeptidase and an endopeptidase, and / or as a cleavage site on more than two or more three, or more four or five different types of amino acids.

[0068] The delivery of the protease in step 1 can be carried out by increasing the permeability of the cell membrane, preferably by using electroporation.

[0069] After step 1, the steps of removing large peptides and proteins or other biomolecules from the fragment sample can be performed directly to form an enriched fragment sample.

[0070] In the step of removing large peptides and proteins or other biomolecules, the peptides and proteins or other biomolecules are preferably removed in a filtration, separation or additional enrichment step, including: size filtration; chromatography, including size exclusion chromatography, hydrophobic chromatography or anion exchange chromatography; physical removal, including phase separation, adsorption, precipitation; filtration, separation or enrichment based on hydrophilic / hydrophobic properties; filtration, separation or enrichment based on electric / magnetic fields; or combinations thereof.

[0071] In the step of removing large peptides and proteins or other biomolecules, peptides, proteins and / or other biomolecules with a molar weight greater than 20 kDa, preferably greater than 15 kDa, and most preferably greater than 10 kDa can be removed from the first fragment sample.

[0072] The perturbation is preferably selected from the following: temperature changes; pressure changes; ionic strength changes; pH changes; metabolic stimuli changes; ligand addition, including drug / small molecule addition, metabolite addition, protein addition, peptide addition, lipid addition, DNA addition, RNA addition, vitamin addition, viral entry, molecular internalization, irradiation, disease / health status or condition and genetic variation (including mutation), or combinations thereof; addition of ionizing agents; induction of chemical modifications, including post-translational modifications, particularly phosphorylation, disulfide bridge formation, ADP-ribosylation, ubiquitination, SUMOylation, acetylation, methylation, oxidation, glycosylation, or combinations thereof.

[0073] Step 4. This typically includes a proteomics workflow prior to actual analysis, particularly involving denaturation, C18 clearance, phase separation, triggers or disruptors of intermolecular interactions, triggers of protein degradation, or combinations thereof.

[0074] In step 1, the protein hydrolysis system, given as the ratio of enzyme to biomolecule content relative to the total biomolecule content in the sample, is preferably used at a concentration of 1 / 50 to 1 / 10000, more preferably 1 / 100 to 1 / 1000 by weight.

[0075] In terms of molar concentration, proteolysis systems are preferably used at concentrations of 5 to 1000 μM, more preferably 20 to 500 μM, where this is the concentration in the LiP sample containing cells. For example, in the case of proteinase K, a concentration of 100 μM is very suitable, and it works well in the range of 50 to 200 μM, and even up to 500 μM. Under these conditions, reproducibility and coverage can be affected at higher concentrations. These values ​​are suitable for an electroporation volume of approximately 3 million HEK293 cells in 100 μl, using a 25 ms pulse at 1000 V. If higher voltages or more pulses are used, the optimal concentration can be lower. Other cell types may require different conditions.

[0076] Preferably, the protein hydrolysis system used in step 1 is added to the cells in the form of an aqueous solution or suspension (which may be provided as a cell suspension or cell arrangement (including tissues, etc.)). Preferably, the solution or suspension is added at a temperature below 30°C, more preferably below 10°C, and more preferably below 5°C. Typically, the protein hydrolysis system solution or suspension is stored on ice until editing. The temperature value refers to the temperature of the protein hydrolysis system solution or suspension, and not the temperature at which the cell suspension or arrangement is received.

[0077] Furthermore, it is preferable to add a proteolytic system and initiate delivery shortly after its addition, i.e., the mechanism for introducing or activating the proteolytic system into the cell, particularly electroporation. Typically, the introduction of the proteolytic system into the cell (e.g., via electroporation) is initiated or even completed within 20 seconds after the addition of the proteolytic system, preferably within 10 seconds.

[0078] Electroporation parameters can vary depending on cell type and local settings. To determine optimal conditions for the chosen system, electroporation parameters can be adjusted if cell viability and physiological state are important for the experiment in question, aiming to achieve sufficient intracellular cleavage while maintaining cell viability and / or minimizing impact on cell physiology. Cell viability can be assessed, for example, by staining cells with trypan blue after electroporation; trypan blue is a dye that cannot penetrate the intact cell membrane of living cells. For some experiments, the percentage of viable cells can be controlled to prevent a decrease after electroporation. Changes in trypan blue viability measurements can also indicate cell leakage. Furthermore, electroporation can lead to changes in cell morphology, which can indicate physiological alterations. For example, cell clumps can be observed under an optical microscope under excessively high intensity / voltage electroporation (3 pulses of 1400 V). Finally, changes in cell physiology induced by electroporation can also be detected by mass spectrometry or mass spectrometry-based proteomics. This also means that differences in the electroporation procedure or during electroporation can be used as perturbations as described above. On the other hand, to rule out significant changes introduced by electroporation (e.g., pathway alterations, or cleavage within the protein sequence), mass spectrometry can be used to analyze electroporated cells and control cells without electroporation after protein extraction and trypsin digestion, and peptide profiles can be quantitatively compared to detect changes in peptide or protein levels.

[0079] The limited protein hydrolysis stage in step 1 is preferably carried out for a duration of 1 to 60 minutes, preferably 2 to 30 minutes, or 2 to 10 minutes, or 2 to 5 minutes, and is also preferably carried out at a temperature in the range of 20 to 40°C.

[0080] In step 1, the limited proteolysis phase can preferably be terminated by quenching protease activity, particularly by inducing conditions that denature the protein or inhibit its enzyme activity, including increasing the temperature, transferring the cell to a medium that simultaneously lyses the cell and denatures the protein.

[0081] Similarly, in step 1, a washing step can follow the limited protein hydrolysis stage, preferably using a saline solution.

[0082] At least one protein is preferably a protein based solely on protein amino acids, or based on protein amino acids and carrying post-translational modifications. Furthermore, the present invention relates to the use of methods for determining, in particular conformational changes or changes in physicochemical properties of said at least one protein in an assumption-free manner, following a perturbation induced in a complex mixture under study, or for determining, in particular conformational changes, of a protein in an assumption-free manner; for identifying protein-based pharmaceuticals; for the effect of pharmaceuticals or other ligands on proteins; or for the quality control of protein-based pharmaceutical formulations.

[0083] Preferably, this can be combined with peptide fragment enrichment techniques such as TAILS (terminal amine isotopic labeling of substrates) for the peptides produced in step 1.

[0084] Other embodiments of the invention are set forth in the dependent claims. Attached Figure Description

[0085] Some preferred embodiments of the invention are described below with reference to the accompanying drawings, which are for illustrative purposes only and not for limiting the invention. In the drawings,

[0086] Figure 1 The intracellular LiP-MS workflow is illustrated. (a) Low-specificity protease proteinase K was introduced into cells via electroporation. Structure-specific peptides were generated by short-term cell incubation. The peptides were analyzed by mass spectrometry after protease inactivation. (b) Electroporation with 200 μM proteinase K resulted in a significant increase in hemitrypsin-cleaved peptides (bright) and a decrease in whole-trypsin-cleaved peptides (dark) (5 replicates, adjusted p < 0.01, 2-fold change). (c) Adding 200 μM proteinase K without electroporation did not result in a significant increase in hemitrypsin-cleaved peptides (bright) and a decrease in whole-trypsin-cleaved peptides (dark) (5 replicates, adjusted p < 0.01, 2-fold change). (d) Electroporation without proteinase K did not result in a significant increase in hemitrypsin-cleaved peptides (bright) and a decrease in whole-trypsin-cleaved peptides (dark) (5 replicates, adjusted p < 0.01, 2-fold change). (e) Proteinase K-induced cleavage showed a significant increase in hemitrypsin-cleaved peptides and a decrease in whole trypsin-cleaved peptides, primarily occurring on cytoplasmic and nucleoproteins. (f) The intensity of hemitrypsin-cleaved peptides increased with the addition of proteinase K and was even higher after electroporation. (g) Cleavage of each protein under optimized settings, which included the addition of 100 μM proteinase K, one 25 ms pulse electroporation at 1000 V, followed by 2 min of limited proteolysis at 37 °C. A total of 4263 distinct PK cleavage sites were detected on 1656 proteins (adjusted p-values ​​< 0.01, 2-fold change).

[0087] Figure 2Validation of proteinase K delivery and target detection is shown. (a) Electroporation of proteinase K into HEK293 cells expressing the protease-active ECFP / YPET FRET sensor resulted in intracellular cleavage. Adding proteinase K without electroporation, and the electroporation process itself, was insufficient. (b) Structural changes in cells treated with 100 nM rapamycin dissolved in DMSO compared to cells treated with DMSO alone (6 replicates). Cells were electroporated with 100 μM proteinase K 5 min after treatment. The peptide of the rapamycin target FKBP1A is highlighted in medium shade. (c) Structure of rapamycin (bright) and the interacting protein FKBP1A (PDB 2DG3). Dark areas were detected by mass spectrometry, and medium shaded areas showed significant changes after rapamycin treatment. (d) Structural changes in native lysates treated with 10 nM rapamycin dissolved in DMSO compared to cells treated with DMSO alone (6 replicates). Five minutes after treatment, proteinase K was added to the cells at a ratio of 1:100 (relative to total protein concentration). The peptide of rapamycin target FKBP1A is highlighted in medium shade.

[0088] Figure 3 Analysis of structural changes in HEK cells after arsenite treatment is shown. (a) HEK293 cells treated with sodium arsenite. (b) HEK293 cells stained with anti-G3BP1 antibody and Hoechst stain 90 min after sodium arsenite treatment. (c) Changes in peptide levels in HEK cells treated with sodium arsenite compared to untreated cells (6 replicates, q < 0.05, 1.5-fold change). (d) Gene ontology enrichment analysis of proteins with significantly changed peptides.

[0089] Figure 4 Structural changes in HEK293 cells after sodium arsenite treatment are shown. (a) Structural changes on selected proteins after arsenite treatment. The bar graph shows the protein sequence, with the medium shaded region detected by mass spectrometry. The light shaded region changed significantly at the time shown (6 replicates, q value < 0.05, 1.5-fold change). (b) Alphafold structure of G3BP1. The medium shaded region was detected by mass spectrometry. The light shaded region changed significantly at the time shown (6 replicates, q value < 0.05, 1.5-fold change).

[0090] Figure 5This illustrates how nuclear specks become more rounded after treatment with sodium arsenite in HEK293 cells. (a) HEK293 cells were stained with anti-SC35 antibody and Hoechst stain 90 minutes after treatment with sodium arsenite. (b) The roundness of nuclear specks treated with and without sodium arsenite was measured for three biological replicates. The images correspond to immunostaining images from A. Colors correspond to replicates;

[0091] Figure 6 The target detection and reproducibility of intracellular LiP-MS are shown; (a) HEK293 cells treated with 20 μM rapamycin dissolved in DMSO for 10 min, compared with the intensity in cells treated with DMSO alone, in the intracellular LiP setting. Left, middle, and right figures show three independent experiments. Peptide intensity, each data point represents a single peptide; FKBP1A peptides are shown in black. Lines indicate significance levels (FC ≥ 1.5, p < 0.01, n = 6 replicates). (b) Peptide intensity in native lysates of HEK293 cells treated with 10 nM rapamycin dissolved in DMSO for 5 min, compared with the intensity in cells treated with DMSO alone (6 replicates). Each data point represents a single peptide; FKBP1A peptides are shown in red. Gray shaded areas indicate significance levels (FC ≥ 1.5, p < 0.01, n = 4 replicates). The four FKBP1A peptides with the lowest p values ​​are highlighted. (c) Number of peptides missing from quantification for each treatment in (a, b). (d) Median coefficient of variation (CV) for peptide quantification for each treatment in (a, b). (e) Intensity of trypsin-cleaved peptides relative to total peptide intensity in intracellular LiP samples from HEK293 cells treated with rapamycin. (f) Overall sequence coverage of peptides detected in at least 3 replicates under each condition;

[0092] Figure 7A comparison of methods for detecting changes in mammalian cells after glucose starvation is shown. (a) Metabolite intensities in HEK293 cells treated with glucose-free medium for 2 hours compared to untreated cells. Each data point represents a single metabolite. Gray shading indicates significance levels (log2(FC) ≥ 0.5, adjusted p < 0.05, n = 3 replicates). (b, c, d) Peptide intensities in HEK293 cells treated with glucose-free medium for 2 hours compared to untreated cells using the methods shown. Each data point represents a single peptide. Gray shading indicates significance levels (FC ≥ 1.5, q < 0.05, n = 6 replicates). Mitochondrial peptides are highlighted in black. (b) Standard LiP-MS in native lysates. (c) Peptide quantification in (b) corrected for changes in protein abundance. (d) Intracellular LiP-MS in intact cells. Gene ontology enrichment analysis of proteins with significant changes in peptides in (e, f) and (c, d) (p < 0.01; biological process).

[0093] Figure 8 Structural changes after 2 hours of glucose starvation are shown. (a) Standard LiP-MS: Phosphoglycerate mutase dimer (PDB: 2JKV). Dark gray areas are covered by mass spectrometry analysis, and light gray areas (dashed arrows) show significant changes (FC ≥ 1.5, q < 0.01, n = 6 replicates). The second subunit is highlighted in light gray, and the phosphoglycerate binding site is highlighted in dark gray. (b) Intracellular LiP-MS: Pyruvate kinase tetramer (PDB: 4GPZ). Black areas are covered by mass spectrometry analysis, and light gray areas (dashed arrows) show significant changes (FC ≥ 1.5, q < 0.01, n = 6 replicates). The second subunit is highlighted in white, and the fructose-1,6-bisphosphate binding site is highlighted in medium gray. (c) Intracellular LiP-MS: Fructose-bisphosphate aldolase (PDB: 5KY6). (d) Intracellular LiP-MS: Phosphoglucate dehydrogenase dimer (PDB: 2JKV). Black areas are covered by mass spectrometry analysis, and light gray areas (dashed arrows) show significant changes (FC ≥ 1.5, q value < 0.01, n = 6 replicates). The second subunit and areas not covered by mass spectrometry analysis are highlighted in light gray, and NADP+ binding sites are highlighted in medium gray.

[0094] Figure 9Changes following glucose starvation are shown. (a) Gene ontology enrichment analysis of metabolic pathways with significant changes in metabolites after 2 hours of glucose starvation (adjusted p < 0.01). (b) Gene ontology enrichment analysis of proteins with significantly changed peptides after glucose starvation using standard LiP-MS (p < 0.01; biological process). (c to h) Comparison of HEK293 cells treated with glucose-free medium for 2 hours with untreated cells, analyzed using the methods shown. (c) Each data point represents a single peptide. Mitochondrial-related proteins are highlighted in gray. Gray shaded areas indicate significance levels (FC ≥ 1.5, q < 0.05, n = 6 replicates). (d) Overall sequence coverage of peptides detected in at least 3 replicates for each condition. (e) Hemitrypsin-cleaved peptide intensity relative to total peptide intensity. (f) Median coefficient of variation (CV) for peptide quantification for each treatment. (g) Number of peptides for deletion quantification for each treatment. (g) Each data point represents a single protein. The gray shaded areas indicate the significance level (FC ≥ 1.5, q value < 0.05, n = 6 replicates).

[0095] Description of preferred implementation scheme

[0096] To extend the highly reproducible and specific overall protein structure analysis of LiP-MS to living cells, the inventors delivered the low-specificity protease proteinase K into mammalian cells. Various protein delivery techniques have been applied to intracellular NMR and Cas9 delivery for gene editing, including protein-covalently linked cell-penetrating peptides, pore-forming toxins, osmocytosis, extracellular vesicles, and electroporation. Comparisons of delivery methods in intracellular NMR show that electroporation is more efficient than pore-forming toxins and cell-penetrating peptides for intracellular NMR applications. Extracellular vesicles and osmocytosis operate on timescales of hours, while electroporation occurs within milliseconds. This makes electroporation a suitable method for LiP-MS, where precise timing is crucial. Electroporation permeates the cell membrane in a short time and therefore also allows for reproducible protein uptake efficiency compared to other methods with less adequate characterization of protein internalization pathways.

[0097] Electroporation has previously been used to deliver aSyn for intracellular NMR, kinetocytes for fluorescence studies of functional proteins, and cre recombinases for genome editing across the cell wall of Arabidopsis.

[0098] The inventors demonstrate that by combining LiP-MS with electroporation, they can study protein structural changes in living cells at the proteomic level.

[0099] Validation: FRET and rapamycin

[0100] Using a Foster resonance energy transfer (FRET)-based sensor, the inventors determined that active proteinase K is delivered into cells via electroporation. The inventors verified that intracellular LiP-MS captures rapamycin that specifically binds to protein FKBP1A. This interaction was initially reported in yeast (Heitman 2001) and is readily detectable by standard LiP-MS in native lysates of human cells (Piazza 2020). The inventors demonstrate that this interaction is also captured by intracellular LiP-MS.

[0101] New application: Stress particles

[0102] Finally, the inventors applied intracellular LiP-MS to study the formation of stress granules. Stress granules are dynamic assemblages of proteins and RNA that form under stress conditions (e.g., sodium arsenite treatment, oxidative stress, or heat shock) to prevent mRNA degradation. Using intracellular LiP-MS, the inventors obtained a dataset of overall structural changes at different time points during sodium arsenite treatment. The inventors detected known structural transitions in the core protein SGBP1 of stress granules. Furthermore, intracellular LiP-MS showed that SERBP1 and PPP1R12A are involved in the early stages of the cellular response to arsenite stress. In addition, the inventors provided evidence that the shape of nuclear spots became more rounded during sodium arsenite treatment.

[0103] result

[0104] Method Development

[0105] Workflow: To investigate the structural state of proteins in their native environment, the inventors tested whether electroporation introduced proteinase K into mammalian cells. HEK293 cells were isolated from culture dishes, washed, and resuspended in PBS (…). Figure 1 a) Immediately after the addition of proteinase K (PK), cells were permeabilized via electroporation. Cells were then incubated at physiological temperature (37°C) for 1 minute for structure-specific cleavage of intracellular proteins. Extracellular proteinase K was removed by washing with PBS and by adding a dissociative agent (7 M guanidine chloride). The proteinase K activity was quenched by heating to 98°C for 5 minutes. The samples were then flash-frozen. Cells were thawed on ice and lysed by acoustic treatment. After disulfide bond reduction and alkylation, trypsin was added to the lysate, which was measured by quantitative, label-free MS analysis.

[0106] Increased PK cleavage after electroporation: The inventors used proteolytic cleavage as a readout indicator of PK delivery into cells. PK has low sequence specificity compared to trypsin, which is preferably cleaved after lysine and arginine residues. Therefore, peptides with one or two mismatched trypsin recognition motifs at their ends are more likely to be generated by PK cleavage. Thus, an increase in hemitrypsin-cleaved peptides and a decrease in fully trypsin-corresponding peptides report PK activity. The inventors compared proteolytic cleavage in cells electroporated with PK with that in control cells that were PK-added but not electroporated. Figure 1 b). Following electroporation, hemitrypsin-cleaved peptides increased, while whole trypsin-cleaved peptides decreased, corresponding to PK cleavage on 1257 proteins. This indicates that PK did indeed enter the cell after electroporation. This was supported by gene ontology enrichment analysis: PK-cleaved proteins showed significant enrichment in cytoplasmic and nuclear related terms (b). Figure 1 e), indicating that proteinase K diffuses into the cells after electroporation. In contrast, adding PK to cells without electroporation does not lead to increased proteolytic cleavage compared to cells without PK. Figure 1 c). Therefore, adding PK to cells without electroporation is insufficient to significantly improve protein hydrolysis cleavage.

[0107] Minimizing the effects of electroporation on cell physiology and viability: Next, the inventors investigated whether the electroporation process introduces changes in peptide levels. The inventors compared electroporated HEK293 cells with un-electroplated control cells. A 25-millisecond pulse at 1000 V ( Figure 1 d) No significant electroporation-induced changes were observed at 800 V for 1 to 3 25 ms pulses. Whole- and half-trypsin-cleaved peptides were symmetrically distributed in all tested electroporation settings, indicating that electroporation does not induce protein cleavage. The observed peptide level changes at high voltages and pulse numbers corresponded to a significant decrease in protein abundance, indicating that high voltage induces protein aggregation. The inventors assessed the effect of electroporation on cell viability by staining cells with trypan blue within 5 minutes after electroporation. Cell viability was not significantly reduced after 25 ms of electroporation with a single pulse at 1000 V (ANOVA adjusted p < 0.05). Electroporation with a single 25 ms pulse at 1000 V for PK was selected for subsequent experiments to rule out electroporation-induced changes.

[0108] No cell leakage was observed after electroporation: Since the pores created by electroporation allow PK to diffuse into the cells, the inventors tested whether intracellular proteins also diffused into the supernatant. The inventors measured the protein content in the cell supernatant after electroporation using BCA. The protein concentration in the supernatant did not increase after electroporation, indicating that cell leakage was negligible.

[0109] Optimizing Proteome Coverage: To maximize proteome coverage, the inventors investigated how PK concentration affects the number of intracellular cleavages. Following electroporation with a 25-ms pulse at 1000 V, an extracellular PK concentration of 100 μM resulted in the highest number of repeatable intracellular cleavages. The intensity of the cleaved peptides increased with increasing PK concentration. Figure 1 f). At 1000 μM PK, the percentage of trypsin-cleaved peptides was as high as 17% of the total peptide strength (compared to 30% trypsin-cleaved peptides in standard LiP-MS experiments). However, the reproducibility of PK cleavage decreased compared to 100 μM PK, possibly due to overdigestion. Therefore, 100 μM PK was used for electroporation in subsequent experiments. Under optimized conditions, 4263 distinct PK cleavage sites were detected on 1656 proteins. Figure 1 g).

[0110] Increasing the incubation time after electroporation from 1 minute to 2 minutes, and using six replicates instead of five, further improved the number of reproducible electroporation-dependent PK cleavages. Using six replicates under each condition, significant variations were observed in 8045 peptides across 2463 of the 4760 detected proteomes, corresponding to 4263 distinct PK cleavage sites across 1656 proteins. Extending the limited proteolysis to 5 minutes did not further improve PK cleavage. In summary, electroporating 3 million HEK cells with 100 μM PK followed by 2 minutes of incubation at 37°C before washing resulted in the highest number of reproducible PK-induced cleavages, covering over 1600 major intracellular proteins. These settings were used in all subsequent experiments. Since the inventors did not impute missing values, trypsin-cleaved peptides generated only intracellularly were not considered in this analysis. Actual coverage could have been even higher in experiments comparing the two perturbed cell states.

[0111] Method Validation

[0112] Validation of Intracellular Delivery Using FRET: The inventors used HEK cells expressing an ECFP-YPet sensor based on Foster resonance energy transfer (FRET) to determine the diffusion of proteinase K into cells during electroporation. Yellow YPet fluorescence indicates an intact sensor due to the internal energy transfer from CFP to YPet, and an increase in YPet fluorescence relative to ECFP fluorescence indicates cleavage at the junction region. After adding PK to the cells, the sensor remained intact, indicating that PK did not enter the cells. Following electroporation, an increase in cyan fluorescence of the cleavage product indicated that active PK had entered the cells. Figure 2 a) Electroporation without the addition of proteinase K does not result in sensor cleavage.

[0113] Validation of Target Detection: Rapamycin Binding: To validate that limited intracellular proteolysis captures known binding events, the inventors treated HEK293 cells with rapamycin that specifically binds to the protein FKBP1A. After treatment with 100 nM rapamycin for 5 minutes, cells were electroporated in the presence of proteinase K and processed according to a previously optimized workflow. Peptides mapping to FKBP1A were found among the most prominent peptides (6 replicates). Figure 2 b, mapping to a site close to the known rapamycin binding site ( Figure 2 c) Similar results were obtained by treating cells with 10 μM rapamycin for 10 min prior to electroporation with proteinase K. The inventors found significant changes in 61 peptides across 60 proteins (6 replicates, adjusted p < 0.05, 1.5-fold change). One of these peptides mapped to FKBP1A. Thus, limited intracellular proteolysis captures known binding events in a highly complex native context.

[0114] Next, the inventors compared intracellular LiP with standard LiP-MS experiments. Natural lysates of HEK293 cells were treated with 10 μM rapamycin for 5 minutes, followed by the addition of PK. Multiple peptides of FKBP1A (4 replicates) were found among the most prominent peptides. Figure 2 d). In intracellular LiP, the percentage of trypsin-cleaved peptides was lower than in native lysates, indicating less proteinase K cleavage. Proteinase K was one of the most abundant proteins in both methods. Although intracellular LiP-MS replicates were performed one at a time for electroporation and processing, and all lysate samples were processed simultaneously, the variability was similar in both methods.

[0115] Absolute Quantification of Proteinase K: Next, the inventors determined the number of proteinase K molecules that entered the cells after electroporation. Three million HEK293 cells were mixed with 100 μM proteinase K in 120 μl, electroporated with a 25 ms pulse at 1000 V, and incubated at 37°C for 2 min. The sample was then washed twice with PBS to inactivate the proteinase K. The same workflow was repeated for control cells without electroporation. Prior to mass spectrometry analysis, a proteinase K-specific peptide labeled with a heavy amino acid was added to the sample. By comparing with the calibration curve and considering that only a portion of the sample was digested, the inventors calculated that the electroporated sample contained 52.5 pmol of proteinase K, and the control sample contained 16.0 pmol of proteinase K. The inventors assumed that the difference of 36.6 pmol of proteinase K was intracellular (12 mmol of proteinase K added to the cells). For 3 million cells, this corresponds to 12.2 amol of proteinase K / cell, or 7.3 million molecules. For HeLa cells, the protein number was estimated to be 2.0. 10^9 molecules / cell. If the inventors assume that the number of proteins is similar in HEK cells, then this equates to approximately 274 substrate proteins per intracellular proteinase K molecule. In a cell volume of approximately 2000 μm³, this corresponds to 6.1 μM of intracellular proteinase K and 1.7 mM of substrate protein.

[0116] Other applications: stress particles

[0117] Treatment of HEK293 cells with sodium arsenite: After observing the high-affinity interaction between rapamycin and FKBP1A captured in situ by intracellular LiP-MS (In-cell LiP-MS), the inventors inquired whether it also detected structural changes during the formation of low-affinity stress granules. To test this, HEK cells were treated with sodium arsenite, which induced the formation of stress granules. Cells were analyzed in six replicates at 0, 10, 20, and 90 minutes after the addition of arsenite. Cells isolated from culture dishes were pre-warmed to 37°C in PBS, centrifuged at 200 × g for 30 seconds, resuspended in PBS, and immediately electroporated with proteinase K. Samples were then processed according to a previously optimized intracellular LiP protocol. Figure 3 a). Immunofluorescence staining of G3PB1 confirmed the formation of stress granules after 10 to 20 minutes of arsenite treatment. Figure 3 b).

[0118] Intracellular LiP-MS capture of structural changes during stress granule formation: Structural changes in three peptides were detected 10 minutes after treatment, prior to the appearance of stress granules via microscopy, plotted relative to three proteins (q value < 0.05, 1.5-fold change). Figure 3 c). After 20 minutes of arsenite treatment, 2325 peptides out of 1149 proteins showed significant changes. After 90 minutes, 20841 peptides out of 3146 proteins showed significant changes. With a few exceptions, protein levels did not change significantly, and therefore structural changes did not adjust for changes in protein abundance.

[0119] Although PK activity was higher at later time points, gene ontology enrichment of proteins exhibiting structural changes after 20 and 90 minutes of arsenite treatment indicates that proteins showing structural changes are indeed enriched in stress granules. Figure 3 d). Furthermore, RNA binding and RNA-related biological processes were significantly enriched. This confirmed that intracellular LiP-MS reliably identifies unstable macromolecular assemblies that undergo structural changes upon perturbation with sodium arsenite.

[0120] Intracellular LiPs capture known structural changes on G3BP1 at peptide-level resolution: The next step was to inquire whether intracellular LiPs provide reliable information at the protein or even peptide levels. Therefore, structural changes detected for known stress granule-related proteins were observed. Many showed structural changes after arsenite treatment for 20 and 90 minutes, with G3BP1 being a core component of the stress granule assembly. Figure 4 a).

[0121] The formation of stress particles is triggered by structural changes in the G3BP1 region of free RNA. G3BP1 consists of an NTF2-like domain required for dimerization, an RNA recognition motif, and three intrinsically disordered regions (IDRs). Both dimerization and RNA binding are essential for stress particle assembly. It has been shown that G3BP1 normally exists in a closed conformation, held together by electrostatic interactions between IDR1 and IDR3. During arsenite stress, as free RNA levels increase, RNA competitively binds to the RNA recognition motif and IDR3. RNA-bound G3BP1 transitions to an open state, leading to liquid-liquid phase separation and providing the core of the stress particle.

[0122] Interestingly, the inventors detected structural changes in the RNA recognition motif after 20 minutes of arsenite treatment, consistent with the proposed G3BP1 model. At 90 minutes, all domains of G3BP1 underwent structural changes. Figure 4 b).

[0123] Intracellular LiPs SERBP1 and PPP1R12A were newly identified due to their early structural changes: stress granules appeared 20 minutes after arsenite treatment under fluorescence microscopy. Interestingly, significant changes in LiP peptides on three proteins—SERBP1, PPP1R12A, and CKB—were detected after 10 minutes. CKB had never been previously associated with stress granules. PPP1R12A was found to be associated with stress granules. SERBP1 has been reported to act as an inhibitor of ribosomes during mTOR phosphorylation. After arsenite treatment, SERBP1 localizes to both stress granules and the nucleolus. Previous studies reported that SERBP1 is essential for clearing stress granules after arsenite stress by regulating the degradation of G3BP1. Knockout of SERBP1 did not alter stress granules 60 minutes after arsenite treatment, but changes may occur earlier. Intracellular LiPs showed changes in nuclear spots after arsenite treatment.

[0124] Interestingly, gene ontology enrichment analysis revealed significant changes in nuclear spots and spliceosomes located within them after 20 minutes of arsenite treatment. To investigate whether sodium arsenite indeed affected nuclear spots, HEK293 cells were stained with anti-SC35 antibody and Hoechst staining 90 minutes after sodium arsenite treatment. Figure 5 a). It was observed that the nuclear spots of cells treated with sodium arsenite became rounder than those of control cells. Figure 5 b). This indicates that intracellular LiPs can be used to identify new macromolecular rearrangements within cells.

[0125] discuss

[0126] Comparison with other methods: Compared to LiP-MS established in natural lysates, the use of electroporation to introduce the low-specificity protease proteinase K into cells preserves organelle and intracellular molecular concentrations. This eliminates the need for lysis and allows for the study of cell physiology with minimal intrusion. Intracellular LiP-MS is a proteomic screening method that provides a foundation for subsequent functional studies.

[0127] In contrast, HRF-MS reports intracellular structural changes at residue-level resolution but requires more complex microfluidic settings. On the other hand, TPP requires a simple temperature gradient to identify proteins undergoing structural changes but lacks information about the regions of change.

[0128] Compared to intracellular NMR, which also uses electroporation, the inventors' method does not require labeling and is not limited to one or several proteins at a time. However, intracellular LiP reports structural dynamics with peptide-level resolution rather than atomic-level resolution and may suffer from more false positives.

[0129] Workflow: The electroporation settings for intracellular LiP were optimized to maintain cell viability, allowing for the study of cell physiology with minimal intrusion. Proteinase K concentration and incubation time were selected to maximize the number of reproducible cleavages. Under the current settings, thousands of proteins were covered; however, sequence coverage for most proteins remained low. It has been shown that altering proteinase K concentration and incubation time does not significantly improve coverage. However, in applications where cell viability is negligible, higher voltage and pulse count during electroporation can result in a more uniform influx of proteinase K and potentially better sequence coverage by improving reproducibility. However, proteinase K concentration and incubation time may need to be adjusted, taking into account the potential loss of cellular material.

[0130] Intracellular LiPs allow for the study of treatments of cells isolated in solution (as shown with rapamycin) and adherent cells isolated after treatment (as in stress particle studies). The inventors used HEK293 cells, but other mammalian cells can also be used, although viability and proteomic coverage may need to be examined. Intracellular LiPs can even be applied to tissues. Low-specificity proteases can also be introduced into bacteria or yeast using strong electroporation to permeate the cell wall, or pretreatment with cell wall-degrading enzymes.

[0131] If cell material is limited, fewer cells can be used; the limit is the mass spectrometry detection limit. Cell leakage after electroporation was not a major issue. Therefore, washing can be reduced or even eliminated entirely in future experiments. Intracellular LiP-MS was optimized for a physiological temperature of 37°C, but it is applicable to other temperatures, pH values, or different low-specificity proteases. It is even feasible to introduce inactivated proteases via electroporation and initiate limited proteolysis at a later time.

[0132] Intracellular delivery validation: Intracellular delivery of proteinase K was validated using a fluorescence-based FRET sensor expressed in HEK cells. Following electroporation, proteinase K cleaves the linker region. The fluorophore showed resistance to proteinase K cleavage, likely due to their stable β-barrel folds. In the future, this analysis could be used to further quantify proteinase K activity in cells and examine whether cellular or incubation conditions affect proteinase K activity. Furthermore, the fluorescence assay can be used to efficiently screen electroporation conditions and proteinase K concentrations for other cell lines. Additionally, the method was validated by detecting the interaction between the drug rapamycin and its known protein targets.

[0133] Intracellular LiP-MS conditions were optimized to preserve the cells' native state as much as possible. However, cells tended to clump after incubation with extracellular proteinase K, for example, during centrifugation. Clumping during limited proteolysis was reduced if the added proteinase K was ice-cold and electroporation was performed within seconds of mixing the proteinase K with the cells.

[0134] The intracellular concentration of proteinase K after electroporation was determined to be approximately 6.1 μM, which corresponds to one proteinase K molecule per approximately 274 substrate molecules. This is a rough estimate based on a single replicate and assumes that HEK cells are similar in size and protein number to HeLa cells, whose protein number is derived from the literature. Furthermore, this value represents the average of all electroporated cells. Determining the distribution of intracellular proteinase K concentration after single-cell electroporation would be of interest. However, flow cytometry is difficult to apply due to the aforementioned clumping that occurs after incubation with proteinase K. Interestingly, the estimated intracellular proteinase K concentration of 6.1 μM is noted to be significantly lower than the extracellular concentration of 100 μM, which indicates that the cells largely remained intact and that electroporation-induced pore closure occurred.

[0135] In classic LiP-MS analysis of lysates, optimal coverage was found for the addition of 1 μg proteinase K to 100 μg of substrate protein. Direct comparisons of the proteinase K / substrate ratio are limited due to the crowded intracellular environment. Intracellularly, certain regions of proteins accessible in native lysates are protected from proteolytic cleavage. Gene ontology enrichment indicates that cleavage occurs primarily on cytoplasmic and nucleoproteins. Future applications will likely focus on determining whether other organelles, such as mitochondria or lysosomes, are also covered by the electroporation of proteinase K·rapamycin.

[0136] Materials and methods

[0137] Cell culture conditions

[0138] HEK293 cells (ATCC #CRL-1573) were cultured in a humidified environment at 37°C and 5% CO2 in Dulbecco modified Eagle medium (DMEM, Gibco Life Technologies #41965039) supplemented with 10% heat-inactivated fetal bovine serum and 5% penicillin / streptomycin. HEK293 cells were passaged in phosphate-buffered saline (PBS) pH 7.4 (Gibco #10010015) prior to confluence. To measure cell viability and concentration, cells were mixed 1:1 with trypan blue solution and analyzed using a Countess cell counter (Thermo Fisher Scientific).

[0139] Cell transfection

[0140] HEK239 cells were transfected on 150 mm culture dishes at approximately 70% confluence. The culture medium was changed before transfection. In each plate, 1200 μl of jetPrime buffer was mixed with 25 μg of plasmid DNA (ECFP-TevS-YPET, a gift from Charlie Morgan and Jason Chin; Addgene #100097), and 50 μl of jetPrime reagent (Polyplus #101000046) was added. After incubation for 10 minutes, the mixture was added to the cells.

[0141] Fluorescence-based proteinase K activity assay

[0142] HEK293 cells were isolated from the culture dish 2 to 3 days after transfection, washed with PBS, and then cultured at 33.3 × 10⁻⁶ cells per cell. 6 Resuspend the cells in PBS at a concentration of 10 cells / ml. Prior to fluorescence analysis, prepare 90 μl of the cell suspension (equivalent to 10 cells / ml). 6 100 μl of cells were mixed with 30 μl of water or 30 μl of 400 μM proteinase K solution (dissolved in water). For electroporation, 100 μl of the solution was electroporated at 1000 V with a 25 ms pulse using a Neon transfection system (Thermo Fisher Scientific) and transferred to 96-well plates. Control cells that were not electroporated were transferred directly after the proteinase K and cell mixture. The fluorescence of ECFP (430-10 / 480) and FRET against YPet (430-10 / 530-10) were measured using a CLARIOstar Plus plate reader (BMG Labtech).

[0143] Rapamycin treatment of cells

[0144] HEK293 cells were isolated from the culture dish, washed with PBS, and then subjected to a 40×10⁻⁶ ppm solution. 6 Resuspend the cells in PBS at a concentration of 10 cells / ml. For each treatment, resuspend 75 μl of cell suspension (corresponding to 10 cells / ml) in PBS. 6 Mix 15 μl of rapamycin solution with 10 μM rapamycin (rapamycin dissolved in DMSO and PBS; the DMSO concentration for cells is 1% in the 10 μM rapamycin assay and 0.016% in the 100 μM rapamycin assay). After incubation for 5 minutes (for 100 nM rapamycin) or 10 minutes (for 10 μM rapamycin), process the cells according to the intracellular LiP-MS workflow.

[0145] Sodium arsenite treatment of cells

[0146] For each replicate, HEK293 cells were grown in separate dishes. Sodium arsenite dissolved in water was added to the culture medium at a final concentration of 500 μM. After 10, 20, and 90 minutes, the culture medium was removed, cells were washed in PBS, and pelleted by centrifugation at 200 × g for 1 minute. Cells were resuspended in PBS. A fraction of the sample was flash-frozen in liquid nitrogen for protein abundance analysis. For intracellular LiP, cells containing approximately 10 μg of sodium arsenite were processed according to the intracellular LiP-MS workflow. 6 90 μl of each cell.

[0147] Intracellular LiP-MS Workflow

[0148] For intracellular LiP-MS, 90 μl of treated or untreated cells (approximately 10 6 (Number of cells) were mixed with 30 μl of ice-cold proteinase K solution (100 μM final proteinase K concentration; except in the optimization experiments, proteinase K was dissolved in water). To determine the effect of proteinase K on cells, 30 μl of water was added instead. To determine the effect of proteinase K-independent electroporation, PBS was added instead.

[0149] For electroporation, 100 μl of cell suspension was electroporated using a Neon transfection system (Thermo Fisher Scientific). Electroporation was typically performed at 1000 V with a 25 ms pulse, except for optimized experiments testing 1–3 pulses and 800 V–1400 V. Specifically, 15 seconds after electroporation, the sample was heated to 37°C and incubated for 2 minutes (incubation times of 1 to 5 minutes were tested during optimization). For washing, 1 ml of pre-warmed PBS to 37°C was added, and cells were pelleted by centrifugation at 4000 × g for 1 minute. The supernatant was removed, and the cells were washed a second time with 1 ml of PBS. For protease inactivation, 7 M guanidine hydrochloride dissolved in water was used… Add to cells and heat the sample to 98°C for at least 5 minutes. Flash freeze the sample in liquid nitrogen before cell lysis.

[0150] Cell lysis

[0151] Intracellular LiP-MS: Samples were thawed on ice and subjected to two 30-second sonication treatments using a Hielscher UP200St-G sonicator coupled with a VialTweeter (optimization experiment and 10 μM rapamycin assay: 60% duty cycle, 100% amplitude; stress particle assay, 100 nM rapamycin assay: 170 W power, 60% duty cycle). Protein concentrations were measured using a quinolinic acid assay and homogenized. Samples were then digested with trypsin.

[0152] Measuring protein abundance

[0153] To measure protein abundance after arsenite treatment, cell pellets were thawed on ice and supplemented with 8 M urea buffer containing a protease inhibitor mixture (Roche). Cells were acoustically treated as in intracellular LiP-MS samples for stress particle assays, and protein concentration was measured and homogenized using a diquinoline carboxylate assay. The samples were then digested with trypsin.

[0154] Classic LiP-MS: HEK293 cells were isolated from culture dishes, washed with PBS, and resuspended in a buffer consisting of 1 mM MgCl2, 150 mM KCl, and 100 mM HEPES pH 7.4. Cells were lysed using a tissue homogenizer (DKW LifeSciences Kimble Pellet Pestle) in 10 cycles of 10 seconds homogenization followed by a 1-minute pause on ice. The lysates were clarified by centrifugation at 1000 × g for 15 minutes at 4 °C. The supernatant was diluted to a protein concentration of 2 μg / µL and treated with rapamycin.

[0155] Classic LiP-MS workflow for rapamycin treatment

[0156] For classic LiP-MS, 50 μL of native protein lysate (2 μg / μL) was incubated with 1 μL of rapamycin (dissolved in DMSO) or 1 μL of DMSO at 25°C for 5 min. To initiate limited proteolysis, 5 μL of proteinase K (0.2 μg / μL in water) was added for 5 min. For protease inactivation, the sample was heated to 99°C for 5 min, then cooled to 4°C for 5 min. Finally, an equal volume of 10% sodium deoxycholate solution was added. The sample was then digested with trypsin.

[0157] trypsin digestion

[0158] Disulfide bonds were reduced by incubation with 5 mM tris(2-carboxyethyl)phosphonium hydrochloride (TCEP) at 37 °C with shaking for 40 min. Subsequently, the sample was incubated with 40 mM guanidine chloride at room temperature in the dark for 30 min. The sample was then diluted with 0.1 M ammonium bicarbonate (Sigma-Aldrich) to achieve the final guanidine chloride concentration. The concentration was 0.5 M or 1% sodium deoxycholate or < 2 M urea. Proteins were digested by shaking overnight at 37°C with Lys-C (FUJIFILM Wako Pure Chemical Corporation) and trypsin (Promega) at a 1:100 enzyme-substrate ratio. Trypsin digestion was terminated by adding formic acid to a final pH of approximately 2.

[0159] C18-purification (C18-Cleanup) and MS sample preparation

[0160] Wash a 96-well C18 microplate (The Nest Group, 40 to 400 μg) with 200 μl methanol, followed by 100 μl buffer B (50% acetonitrile (ACN), 0.1% FA) and 2 × 200 μl buffer A (0.1% FA). Load the sample onto the plate and wash with 3 × 200 μl buffer A by centrifugation at 1000 × g for 1 min. Elute the peptide with 3 × 100 μl buffer B and dry in a vacuum concentrator. Resuspend the sample in buffer A supplemented with iRT peptide (Biognosys) and analyze by mass spectrometry.

[0161] Absolute quantification of proteinase K

[0162] Four synthetic peptides (peptides 1 through 4) with proteolytic activity were absolutely quantified using proteinase K (AQUA Ultimate Heavy, lysine and arginine stable isotope labeling, Thermo Scientific). Prior to mass spectrometry analysis, the heavily labeled peptides at a final concentration of 500 fmol / μl were added to intracellular LiP samples previously electroporated with 100 μM proteinase K. Peptides were also added to control samples containing proteinase K but not electroporated. For calibration, the heavy peptides were incorporated into proteinase K-free lysates (peptide concentrations of 500 fmol / μl, 50 fmol / μl, 5 fmol / μl, and 500 amol / μl). Samples were measured using PRM. Peptide intensities were determined by summing the areas of the five strongest fragment ions. Linear models for each peptide were fitted to log 10-converted peptide intensities and calibration sample concentrations, respectively. The linear models were used to quantify the corresponding light proteinase K peptides in the target samples. Finally, the amount of proteinase K was calculated as the average of the four determined peptide amounts.

[0163] Liquid chromatography and mass spectrometry data acquisition

[0164] DIA: Samples were injected into an Orbitrap Fusion Lumos Tribrid mass spectrometer equipped with a Waters nanoAQUITY UPLC system over 120 minutes using a linear gradient of 3% to 35% B (eluent A: 0.1% formic acid; eluent B: 99.9% acetonitrile, 0.1% formic acid) at a flow rate of 300 nL / min. MS1 scans were performed from 350 to 1400 m / z, with an orbital trap resolution of 120,000 and an AGC target injection time of 50% or 100 ms. The DIA method consisted of 41 variable-width windows with 1 m / z overlap. Fragmentation was generated via high-energy collision-induced dissociation (HCD) using a fixed collision energy of 28%. MS2-DIA spectra were obtained from 200 to 1800 m / z, with an orbital trap resolution of 30,000 and an AGC target of 200% or an injection time of 54 ms.

[0165] DIA: Using a linear gradient of 3% to 30% B (eluent A: 0.1% formic acid; eluent B: 95% acetonitrile, 0.1% formic acid), the sample, optimized for electroporation settings, PK concentration, and time, was injected into an Orbitrap Eclipse Tribrid mass spectrometer equipped with a Waters nanoAQUITY UPLC system at a flow rate of 300 nL / min over 120 minutes. MS1 scan ranged from 350 to 1400 m / z with an orbital trap resolution of 120,000 and an AGC target resolution of 800,000. The DIA method consisted of 41 variable-width windows with 1 m / z overlap. Fragmentation was generated via high-energy collision-induced dissociation (HCD). MS2-DIA spectra were obtained over a scan range of 150 to 2000 m / z with an orbital trap resolution of 30,000 and an AGC target resolution of 50,000.

[0166] DIA: Intracellular LiP-MS samples containing 10 μM rapamycin were injected into a Q Exactive Plus hybrid quadrupole-orbit trap mass spectrometer equipped with an EASY-nLC 1000 system (Thermo Fisher Scientific). Peptide separation was performed on a 40 cm × 0.75 mm id column (New Objective, PF360-75-10-N-5) packed with 3 μm beads (Dr. Maisch Reprosil-Pur 120). Separation was performed over a total run time of 120 min using a linear gradient of 3% to 30% buffer B, followed by 5 min of isocratic constant concentration of 90% buffer B (eluent A: 0.1% formic acid; eluent B: 95% acetonitrile, 0.1% formic acid). The flow rate was maintained at 300 nl / min. MS1 scan ranged from 350 to 1500 m / z, with an orbital trap resolution of 70,000 m / z and an AGC target injection time of 3e6 or 120 ms. The DIA method consisted of 20 variable-width windows with 1 m / z overlap. Fragmentation was generated via high-energy collision-induced dissociation (HCD). MS2-DIA spectra were obtained within a scan range of 200 to 1800 m / z, with an orbital trap resolution of 35,000 m / z and a 1e6 AGC target.

[0167] PRM: Samples were injected into an Orbitrap Fusion Lumos Tribrid mass spectrometer equipped with a Waters nanoAQUITY UPLC system over 120 minutes at a flow rate of 300 nL / min using a linear gradient of 3% to 35% B (eluent A: 0.1% formic acid; eluent B: 99.9% acetonitrile, 0.1% formic acid). The separation window was 1.2 m / z. Fragmentation was generated via high-energy collision-induced dissociation (HCD) using a fixed collision energy of 28%. MS2 spectra were obtained over a scan range of 200 to 1800 m / z, at an orbital trap resolution of 60,000 and with a 1000% AGC target or an injection time of 118 ms.

[0168] Search engines

[0169] Data was searched using Spectronaut 17 (Biognosys), where the library was constructed from a DIA file. The digestion type was set to semi-specific, and the minimum peptide length was adjusted to 6. Data was searched against the human-reviewed UniProt database and proteinase K sequences. https: / / www.uniprot.org(Searched on April 16, 2023). Data analysis was performed using Spectronaut via classic DIA analysis, which employs a targeted library-based search with the following modifications: single-hit validation by modified sequences, and exclusion of proteins with single hits. Quant 2.0 was selected for label-free protein quantification, with samples globally normalized relative to the median, and PTM localization was chosen.

[0170] Statistical analysis

[0171] For statistical analysis using R version 4.2.2, only peptides with amounts < 1 were considered. Welch's t-test was used for statistical analysis. The Benjamini Hochberg method was used to adjust p-values ​​relative to multiple tests.

[0172] Gene ontology enrichment analysis

[0173] For gene ontology enrichment analysis, the topGP package (version 2.50.0) is used in R. https: / / git.bioconductor.org / packages / to pGO). Gene ontology annotations were obtained from Uniprot ( https: / / www.uniprot.org (Retrieved on May 17, 2022). Enrichment analysis was performed using the elim algorithm from topGO, with Fisher's exact test. Significance was set at a p-value level of < 0.01.

[0174] Peptide map of AlphaFold2 structure

[0175] Peptides were mapped to structures predicted by Alphafold v4 using PyMOL (version 2.5.4).

[0176] Immunostaining and roundness measurement

[0177] HEK293 cells were seeded at 25,000 cells per well on 8-well μ-plates with an ibiTreat surface (ibidi #80806). After 24 hours, 500 μM sodium arsenite (Pfaltz & Bauer #7784-46-5) was added to the cells. After 90 minutes, the cells were fixed for 10 minutes with 4% PFA (Electron Microscopy Sciences #15714) in PBS. After washing twice with PBS, the cells were permeabilized for 10 minutes with 0.5% Triton X-100 (Sigma-Aldrich #T8787) in PBS. The cells were washed twice with PBS and then incubated for 1 hour in PBS containing 0.1% Tween 20 (Sigma-Aldrich #P1379) with primary antibody, secondary antibody, and 1:1000 Hoechst 33258 (Molecular Probes #H-3569). The cells were then washed twice in PBS containing 0.1% Tween 20 and held in PBS for imaging.

[0178] To determine the circularity of the nuclear spots, representative fields of view were captured for each repeat and thresholded in ImageJ to generate a mask of the nuclear spots that matches the spots observed by the eye. Circularity was then measured by analyzing particle function.

[0179] Antibodies used:

[0180]

[0181] As further proof of the concept, the following series of experiments were conducted:

[0182] Intracellular LiP-MS can reproducibly identify structural changes after drug binding.

[0183] Intracellular LiP-MS, which uses a well-characterized binding of rapamycin to protein FKBP1A (FKBP prolyl isomerase 1A, Uniprot P62942) to deliver the protease into the cells and perform limited proteolysis, can capture known structural alterations associated with high-affinity binding events. Cells were electroporated with PK following treatment with 20 μM rapamycin for 10 min, following an optimized workflow. The experiment was repeated twice to assess the reproducibility of intracellular LiP-MS. Additionally, rapamycin-treated HEK293 cell lysates were subjected to standard LiP-MS to allow for comparison with established methods.

[0184] Using intracellular LiP-MS, in a comparison of peptide intensity between rapamycin-treated cells and control (DMSO-treated) cells, multiple FKPB1A-derived peptides were detected in the strongest hits. Figure 6 a; FC > 1.5, p < 0.01, n = 6 replicates). The same FKBP1A peptide varied in two (fragment 1, fragment 2) or all three independent experiments (fragment 3, fragment 4). Standard LiP-MS also identified multiple FKBP1A-derived peptides in the most significant hits (FC > 1.5, p < 0.01, n = 4 replicates); Figure 6 b), as previously shown. The FKBP1A peptide detected by intracellular LiP-MS overlaps with the peptide of the processed lysate detected by standard LiP-MS, but is longer. This may be due to the presence of 0.5 M guanidine hydrochloride used in intracellular LiP-MS. In comparison, trypsin cleavage using 1% sodium deoxycholate, as used in standard LiP-MS, is more efficient. Therefore, intracellular LiP-MS identified the same peptide in the expected target protein in the best hit in three independent experiments.

[0185] Most peptides were consistently quantified in all replicates under each condition. Figure 6 c). The median coefficient of variation for peptides in intracellular LiP-MS ranged from 14% to 24%, compared to 15% to 17% in standard LiP-MS. Figure 6 d). In all experiments, the percentage of HT peptides was close to 18% ( Figure 6 e). In this comparison, similar protein substrate to PK ratios were used under two conditions: i.e., for intracellular LiP, 3 million cells (800 to 1000 μg protein) and 12 nmol (347 μg) PK, and in lysates, 100 μg protein and 35 pmol PK (1 μg). The mass spectrometric sequence coverage of proteins in the intracellular LiP-MS experiments was higher than in the standard LiP-MS experiments, due to the fact that different mass spectrometers were used to acquire the datasets. Figure 6 f; only peptides detected in at least three replicates under each condition are considered. Power calculations were also performed for the first intracellular LiP-MS assay and the standard LiP-MS assay (intracellular LiP-MS: power = 34% at FC = 1.5 and 99% at FC = 2.0, where n = 6, FDR = 5%; standard LiP-MS: power = 46% at FC = 1.5 and 99% at FC = 2.0, where n = 4, FDR = 5%). Therefore, the data quality measurements for intracellular LiP-MS are generally similar to those for established lysate-based LiP-MS methods.

[0186] Intracellular LiP-MS identified structural changes during glucose starvation.

[0187] Intracellular LiP-MS also detected other types of structural changes, such as oligomerization. To test this, HEK293 cells were incubated in glucose-free medium. After 2 hours of glucose starvation, metabolomics measurements identified 21 significantly altered metabolites ( Figure 7 a; log2(FC) ≥ 0.5, adjusted p-value < 0.05, n = 3 replicates). Compared with untreated control cells, gluconeogenesis, glycolysis, and pentose phosphate pathways were the most significantly enriched pathways. Figure 9 a) Intracellular LiP-MS was performed under identical conditions, and protein abundance was measured using a standard proteomics workflow. In parallel, cells were harvested after 2 hours of glucose starvation, lysed via natural lysis, and analyzed by standard LiP-MS. A trypsin control sample, i.e., natural lysate treated with water instead of PK, was also produced to measure changes in protein abundance during standard LiP-MS.

[0188] Standard LiP-MS identified 375 varied peptides on 244 proteins, primarily related to mitochondria. Figure 9 b, c). On the other hand, intracellular LiP-MS identified only three altered peptides ( Figure 9 c; FC ≥ 1.5, q value < 0.05, n = 6 replicates): one peptide on glycogenin-1 (GYG1) and two overlapping peptides on the enzyme pyruvate kinase (PKM). CV, deletion value, HT peptide share, and sequence coverage were similar to or better than previous rapamycin studies. Figure 9 The efficacy was 66% at FC = 1.5 and 99% at FC = 2.0, where n = 6 and FDR = 5%. Since the changes in PKM observed by intracellular LiP-MS were consistent with metabolomics measurements in which changes were found in its substrate phosphoenolpyruvate, the inventors wanted to know about the hundreds of hits identified by standard LiP-MS. Interestingly, the trypsin control of the standard LiP-MS experiments showed a systematic downregulation of mitochondrial proteins after 2 hours of glucose starvation, which was not observed in abundance measurements performed using standard proteomics workflows. Figure 9d). Previous analysis of organelle coverage showed that mitochondrial coverage in standard LiP-MS was lower than that in intracellular LiP-MS, suggesting that mitochondria were not adequately disrupted by natural lysis. Human mitochondria have been reported to fuse into larger networks during glucose starvation. Prior to standard LiP-MS, the natural lysate was clarified by centrifugation at 10,000 × g; in this step, due to differences in mitochondrial size, mitochondria were likely removed to a greater extent from the starved sample compared to the control sample. The conclusion is that the mitochondrial-specific changes observed in standard LiP-MS after glucose starvation are likely an artefact of natural cell lysis, reporting differential centrifugation behavior but not necessarily changes in intracellular protein abundance.

[0189] To identify actual structural changes in proteins, peptide levels in standard LiP-MS were corrected for changes in protein levels in the trypsin control. Multiple p-tests were performed simultaneously on standard LiP-MS data, abundance-corrected standard LiP-MS data, and intracellular LiP-MS data to obtain the same q-value level, which facilitates comparison (…). Figure 7 b to d). Intracellular LiP-MS identified 11 significantly altered peptides on 7 proteins (3 peptides on pyruvate kinase (PKM, Uniprot P14618) and glycogen-1 (GYG1, Uniprot P46976), while abundance-adjusted standard LiP-MS identified 22 peptides on 19 proteins (2 peptides on uridine-cytidine kinase 2 (UCK2, Uniprot Q9BZX2) and 3 peptides on elongation factor 1-α1 (EEF1A1 Uniprot P68104). GO enrichment analysis identified glycolysis as the most significant biological process in both methods. Figure 7 (e, f) Intracellular LiP-MS also identified the pentose phosphate pathway, consistent with metabolomics measurements. The only enzyme altered in both intracellular and standard LiP-MS was glycogenin-1 (GYG1), which initiates glycogen synthesis. Intracellular LiP-MS also identified structural changes in long-chain fatty acid CoA ligase 4 (ACSL4), which participates in lipid synthesis and degradation via β-oxidation.

[0190] In the glycolytic enzymes, standard LiP-MS detected one glycolytic enzyme—phosphoglycerate mutase-1 (PGAM, Uniprot P18669). Figure 8 a) Structural changes; phosphoglycerate mutase-1 is a catalytic... 3-Phosphoglyceric acid Reversible transformation 2-Phosphoglyceric acidThe homodimer of phosphoglycerate also changed in the metabolomics dataset. This structural change occurred at the substrate-binding region, indicating that it reports substrate occupancy. Different peptides located at the substrate-binding region changed in the intracellular LiP-MS dataset, but slightly above the significance cutoff (q = 0.056).

[0191] Intracellular LiP-MS detected three altered peptides on pyruvate kinase (PKM), the final, irreversible, and rate-limiting step in glycolysis. After binding to the allosteric regulator fructose-1,6-bisphosphate, PKM changed between an active tetrameric form and an inactive monomeric or dimer state. The altered peptides were located at the junction of two dimers, directly adjacent to the fructose-1,6-bisphosphate binding site. Figure 7 (b) Therefore, it reported enzyme regulation through oligomerization.

[0192] Further structural changes were detected by intracellular LiP-MS for aldolase (ALDOA, Uniprot P04075). The altered peptide was neither located at the oligomer junction nor near the substrate binding site, which is expected for an enzyme active in both glycolysis and gluconeogenesis. Figure 7 c). Alternatively, the altered peptides are located on the surface of the multimer, which could indicate changes in interactions with other molecules during glucose starvation. For example, it has been shown that aldolases activate AMPK (AMP-activated protein kinase) in the absence of fructose-1,6-bisphosphate, thereby regulating protein translation during this period. Interestingly, changes were also found by intracellular LiP-MS in eukaryotic translation initiation factor 4E binding protein 1 (EIF4EBP1, UniprotQ13541) (a repressor of translation initiation) and small ribosomal subunit protein uS8 (RPS15A, Uniprot P62244) (a component of the ribosome).

[0193] Intracellular LiP-MS identified structural changes in gluconate dehydrogenase (PGD), a major cellular enzyme involved in the pentose phosphate pathway and the primary NADPH-producing enzyme, among enzymes in the pentose phosphate pathway. PGD exists in both inactive monomeric and active oligomeric forms, regulated by NADP+ binding. Following 2 hours of glucose starvation, intracellular LiP-MS revealed peptide changes linking two homodimers and adjacent to the regulatory NADP+ binding site (…). Figure 8 a). The conclusion is that while standard LiP-MS successfully captures substrate binding in PGM, which is expected to be a high-affinity interaction, intracellular LiP-MS may be more sensitive to oligomerization and complex formation, as demonstrated by PKM, PGD, and possibly ALDOA.

[0194] The method used in the second series of experiments differs from the methods described above.

[0195] Intracellular LiP-MS Workflow

[0196] For classic LiP-MS of glucose-starved cells, 50 μl of native protein lysate (2 μg / μl) was incubated at 25 °C for 5 min. To initiate limited proteolysis, 5 μl of PK (0.2 μg / μl in water) was added and incubated for 5 min. For protease inactivation, the sample was heated to 99 °C for 5 min, then cooled to 4 °C for 5 min. Finally, an equal volume of 10% sodium deoxycholate solution was added. The sample was then digested with trypsin.

[0197] Classic LiP-MS Workflow

[0198] For classic LiP-MS of glucose-starved cells, 50 μl of native protein lysate (2 μg / μl) was incubated at 25 °C for 5 min. To initiate limited proteolysis, 5 μl of PK (0.2 μg / μl in water) was added and incubated for 5 min. For protease inactivation, the sample was heated to 99 °C for 5 min, then cooled to 4 °C for 5 min. Finally, an equal volume of 10% sodium deoxycholate solution was added. The sample was then digested with trypsin.

[0199] Liquid chromatography and mass spectrometry data acquisition

[0200] DIA: Glucose-starved intracellular LiP-MS samples at 20 μM were injected into a Q Exactive Plus hybrid quadrupole-orbit trap mass spectrometer, and the 10 μM rapamycin dataset was analyzed as described above.

[0201] Statistical analysis

[0202] In classic LiP-MS experiments under glucose starvation, peptides were adjusted for changes in protein abundance using the MSstatsLiP package (version 1.9.2).

Claims

1. A method for determining at least one structural and / or physicochemical property of at least one protein or peptide, said at least one protein or peptide being contained in a complex mixture of other proteins and / or other biomolecules in at least one cell, said method comprising the steps of:

1. Delivering a protease into the cells and performing limited proteolysis on the complex mixture, followed by cell extraction and / or cell lysis to produce fragment samples; 2. Denature the fragment sample to obtain a denatured fragment sample; 3. Optionally, the denatured fragment sample is completely fragmented during the digestion step to obtain a fully fragmented sample; 4. Perform analytical analysis on the fragmented sample or the fully fragmented sample to determine the at least one protein or peptide or its characteristics.

2. The method of claim 1, used for detecting state changes of said protein or peptide in response to perturbation in said complex mixture of additional proteins and / or other biomolecules contained in at least two cells, said method comprising the steps of:

1. Subjecting at least one cell in the cells to the perturbation, subsequently or simultaneously delivering a protease into the cells and performing limited proteolysis on the complex mixture, followed by cell extraction and / or cell lysis to produce a first fragment sample; 2. Denature the first fragment sample to obtain a denatured fragment sample; 3. Optionally, the denatured first fragment sample is completely fragmented during the digestion step to obtain a fully fragmented sample; Steps 1 to 3 are performed on at least one other cell among the said cells, but in step 1, the at least one other cell is not subjected to the perturbation, to produce a fully fragmented control sample or a fragmented control sample.

4. Perform analytical analysis on the fragmented sample or the completely fragmented sample, as well as the completely fragmented control sample or the fragmented control sample, to determine the state change of the protein.

3. The method of claim 2, wherein the analysis in step 4 is based on a quantitative comparison of analytical analysis of the fully fragmented sample or the fragmented sample with analytical analysis of the fully fragmented control sample or the fragmented control sample.

4. The method according to any one of the preceding claims, wherein for the analytical analysis in step 4., mass spectrometry techniques are used, particularly selected / multiple reaction monitoring (SRM / MRM) and / or data-independent acquisition (DIA) of product ion spectra, including SWATH-MS and / or optionally data-dependent acquisition (DDA or shotgun).

5. The method according to any one of the preceding claims, wherein in step 1, a protein hydrolysis system is used, said protein hydrolysis system being selected from proteinase K, thermophilic protease, subtilisin, pepsin, papain, α-chymotrypsin, elastase, figase, streptomycin, and mixtures thereof. And / or the protein hydrolysis system used in step 1 is added to the at least one cell, preferably in the form of an aqueous solution or suspension, wherein preferably, the temperature of the added solution or suspension is below 30°C, more preferably below 10°C, and more preferably below 5°C.

6. The method according to any one of the preceding claims, wherein the delivery of the protease in step 1 is carried out by: increasing cell membrane permeability, preferably by using electroporation or by using a permeation reagent, or by coupling the protease with a reagent having cell-penetrating properties, by delivery within a cage system and / or vesicles, by expressing the protease in an active state intracellularly under the control of a stringent promoter, by expressing an inactive form of the protease that can be activated when needed, or a combination thereof.

7. The method according to any one of the preceding claims, wherein the step of removing large peptides and proteins or other biomolecules from the fragment sample to form an enriched fragment sample is performed directly after step 1, wherein, preferably, the removal of large peptides and proteins or other biomolecules is performed in a filtration, separation, or additional enrichment step, comprising: Size filtration; chromatography, including size exclusion chromatography, hydrophobic chromatography, or anion exchange chromatography; physical removal, including phase separation, adsorption, and precipitation; Filtration, separation, or enrichment based on hydrophilic / hydrophobic properties; Filtering, separation, or enrichment based on electric / magnetic fields; or combinations thereof, And / or wherein, in the step of removing large peptides and proteins or other biomolecules, peptides, proteins and / or other biomolecules with a molar mass greater than 20 kDa, preferably greater than 15 kDa, and most preferably greater than 10 kDa, are removed from the first fragment sample.

8. The method according to any one of the preceding claims, wherein the perturbation is selected from: temperature change; pressure change; ionic strength change; pH change; metabolic stimuli change; ligand addition, including drug / small molecule addition, metabolite addition, protein addition, peptide addition, lipid addition, DNA addition, RNA addition, vitamin addition, viral entry, molecular internalization, irradiation, disease / health state or condition, and genetic variation including mutation, or combinations thereof; addition of ionizing agents; chemical modification, including post-translational modification, particularly phosphorylation, disulfide bridge formation, ADP-ribosylation, ubiquitination, SUMOylation, acetylation, methylation, oxidation, glycosylation, or combinations thereof; phase separation; triggers or disruptors of intermolecular interactions; and triggers of protein degradation.

9. The method according to any one of the preceding claims, wherein step 4. comprises a proteomics workflow prior to actual analysis, particularly involving denaturation, C18 purification, or a combination thereof.

10. The method according to any one of the preceding claims, wherein in step 1, the concentration of the protein hydrolysis system used, given by the ratio of enzyme to biomolecule content relative to the total biomolecule content in the sample, is in the range of 1 / 50 to 1 / 10000 by weight, preferably in the range of 1 / 100 to 1 / 1000. And / or where, for quantitative determination, a characteristic relabeled fragment generated by the limited protein hydrolysis of step 1, the fragmentation of step 2, or the complete fragmentation of step 3 is incorporated into the first fragment sample and / or the complete fragmented fragment sample.

11. The method according to any one of the preceding claims, wherein the limited protein hydrolysis phase of step 1 is carried out for a duration of 1 to 60 minutes, preferably 2 to 30 minutes, or 2 to 10 minutes, or 2 to 5 minutes, and preferably at a temperature in the range of 20°C to 40°C.

12. The method according to any one of the preceding claims, wherein in step 1, the limited proteolysis phase is terminated by quenching protease activity, particularly by inducing conditions that denature the protein or inhibit its enzymatic activity, including increasing the temperature, transferring the cell to a medium that simultaneously lyses the cell and denatures the protein. And / or in step 1, a washing step is performed after the limited protein hydrolysis stage, preferably using a saline solution.

13. The method according to any one of the preceding claims, wherein the at least one protein is a protein based solely on protein amino acids, or is based on protein amino acids and carries post-translational modifications.

14. Use of the method according to any one of the preceding claims for determining, in a hypothesis-free manner, at least one structural and / or physicochemical property of a protein or peptide, particularly a change in state of said at least one protein or peptide, particularly a conformational change or a change in local or global physicochemical properties, said at least one protein preferably undergoing a change in state, particularly a conformational change or a change in local or global physicochemical properties, following an induced perturbation in the complex mixture under study; or for determining the medically relevant state of said protein, particularly conformational or local or global physicochemical properties; for determining protein-based pharmaceuticals, for the effect of pharmaceuticals or other ligands on proteins, or for quality control of protein-based pharmaceutical formulations.

15. Use of the method according to any one of claims 1 to 13 in combination with peptide fragment enrichment techniques such as TAILS for the peptides produced in step 1.

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