Nucleotide-sugar donors containing bioorthogonal moieties are important tools to study cellular glycosylation. Typically, the acetamide moiety in N-acetylhexosamines such as GlcNAc and GalNAc is replaced by an acylamide with a clickable tag and converted to the corresponding uridine diphosphate analogue. These probes can then be tested for acceptance by glycosyltransferase enzymes in vitro. Lengthy procedures in synthetic chemistry currently limit the availability of bioorthogonal uridine diphosphate (UDP)-sugar analogues. Chemoenzymatic synthesis has proven to be a powerful and effective alternative, and multiple approaches have been published to date. In this protocol, we describe a streamlined method for the generation of bioorthogonal UDP-GlcNAc and UDP-GalNAc analogues. We describe the chemical modification of D-glucosamine and D-galactosamine to incorporate bioorthogonal acylamides, the subsequent one-pot multienzyme conversion to the corresponding UDP-sugar analogues, and reproducible purification. Our approach features the bacterial kinase NahK and human pyrophosphorylase AGX1 as well as a recombinantly expressed AGX1 variant with an expanded substrate profile. The approach further features an inorganic pyrophosphatase and an alkaline phosphatase to improve enzymatic turnover and aid the purification process, respectively. The use of biosynthetic enzymes with substrate promiscuity extends the scope of bioorthogonal nucleotide-sugar analogue structures to aid efforts in chemical glycobiology. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Chemical synthesis of bioorthogonally tagged acylamide analogues of D-GlcNAc and D-GalNAc Alternate Protocol 1: Chemical synthesis of bioorthogonally tagged acylamide analogues of D-GlcNAc and D-GalNAc from a protected GlcNH2 or GalNH2 precursor Basic Protocol 2: Conversion of D-GlcNAc or D-GalNAc analogues to UDP-sugars using analytical- (reaction scouting) and preparative-scale enzymatic synthesis and purification.
Targeted protein degradation using bifunctional molecules like proteolysis targeting chimeras (PROTACs) has revolutionized drug discovery but remains limited by E3 ligase availability and lack of cell selectivity. N-degron degraders provide a smaller, more drug-like alternative that harnesses endogenous N-terminal degradation signals independent of recruited ligases. However, they are prone to proteolysis and lack mechanisms for tissue-specific activation. To overcome these challenges, we developed a "caged" N-degron degrader selectively activated by the immunoproteasome (iCP), a proteasome isoform induced in cancer and inflammatory settings. By appending an iCP-recognition tetrapeptide (Ala-Thr-Met-Trp) capped with a morpholine group to the N-terminus, we shielded the degron from nonspecific cleavage and enabled selective activation in iCP-expressing cells. This design improved stability, enhanced degradation potency, and minimized activity in healthy cells. Our findings establish a generalizable strategy to spatially control N-degron activity through disease-specific proteolytic environments, advancing safer and more selective protein degradation therapeutics. © 2025 Wiley Periodicals LLC. Basic Protocol 1: Synthesis and characterization of an N-degron degrader for Abl Basic Protocol 2: Synthesis and characterization of an immunoproteasome N-degron prodrug Basic Protocol 3: Assessing protein degradation in cells.
Small molecule microarray (SMM) technology has become a powerful tool used in high-throughput screening for target-based drug discovery. One area in which SMMs have found use is the identification of small molecule ligands for RNA. RNAs with unique secondary or tertiary three-dimensional structures are considered to be attractive targets for small molecules. Complex RNA structures can form hydrophobic pockets suitable for small molecule binding, representing an opportunity for developing novel therapeutics. Our lab has previously taken a target-based approach, screening a single target against many small molecules on an SMM platform. Here, we report a screening protocol for SMMs to investigate multiple RNAs simultaneously using multi-color imaging. By introducing a mixture containing different fluorophore-labeled RNAs, the fluorescence signal of each binding event can be observed simultaneously. Thus, the specificity of a hit compound binding to one RNA target over other highly abundant RNAs (such as tRNA or rRNA) can be easily evaluated. © 2020 Wiley Periodicals LLC. Basic Protocol: RNA screening on SMMs by multi-color imaging Support Protocol 1: Preparation of SMM slides Support Protocol 2: Fluorophore labeling of RNA through maleimide chemistry.
Reversible addition-fragmentation chain-transfer (RAFT) polymerization is a commonly used polymerization methodology to generate synthetic polymers. The products of RAFT polymerization, i.e., RAFT polymers, have been widely employed in several biologically relevant areas, including drug delivery, biomedical imaging, and tissue engineering. In this article, we summarize a synthetic methodology to display an azide group at the chain end of a RAFT polymer, thus presenting a reactive site on the polymer terminus. This platform enables a click reaction between azide-terminated polymers and alkyne-containing molecules, providing a broadly applicable scaffold for chemical and bioconjugation reactions on RAFT polymers. We also highlight applications of these azide-terminated RAFT polymers in fluorophore labeling and for promoting organelle targeting capability. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Synthesis of the azide derivatives of chain transfer agent and radical initiator Basic Protocol 2: Installation of an azide group on the α-end of RAFT polymers Alternate Protocol: Installation of an azide group on the ω-end of RAFT polymers Basic Protocol 3: Click reaction between azide-terminated RAFT polymers and alkyne derivatives.
Drug-induced liver injury is an important cause of non-approval in drug development and the withdrawal of already approved drugs from the market. Screening human hepatic cell lines for toxicity has been used extensively to predict drug-induced liver injury in preclinical drug development. Assessing hepatic-cell health with more diverse markers will increase the value of in vitro assays and help predict the mechanism of toxicity. We describe three live cell-based assays using HepG2 cells to measure cell health parameters indicative of hepatotoxicity. The first assay measures cellular ATP levels using luciferase. The second and third assays are multiparametric high-content screens covering a panel of cell health markers including cell count, mitochondrial membrane potential and structure, nuclear morphology, vacuolar density, and reactive oxygen species and glutathione levels. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Measurement of cellular ATP content Basic Protocol 2: High-content analysis assay to assess cell count, mitochondrial membrane potential and structure, and reactive oxygen species Basic Protocol 3: High-content analysis assay to assess nuclear morphology, vacuoles, and glutathione content Support Protocol 1: Subculturing and maintaining HepG2 cells Support Protocol 2: Plating HepG2 cell line Support Protocol 3: Transferring compounds by pin tool Support Protocol 4: Generating dose-response curves.
The emergence of covalent inhibitors and chemoproteomic probes in translational chemical biology research requires the development of robust biophysical and analytical methods to characterize their complex interactions with target biomolecules. Importantly, these methods must efficiently assess target selectivity and accurately discern noncovalent binding from the formation of resultant covalent adducts. One recently reported covalent chemical tool used in tumor immune oncology, covalent immune recruiters (CIRs), increases the proximity of immune cells and cancer cells, promoting immune recognition and response. Herein we describe biolayer interferometry (BLI) biosensor, flow cytometry, and solution fluorescence-based assay approaches to characterize CIR:antibody binding and CIR-antibody covalent-labeling kinetics. BLI technology, akin to surface plasmon resonance, provides the unique opportunity to investigate molecular binding and labeling kinetics both on a solid surface (Basic Protocol 1) and in solution (Alternate Protocol 1). Here, recruitment of mass-containing proteins to the BLI probe via CIR is measured with high sensitivity and is used as a readout of CIR labeling activity. Further, CIR technology is used to label antibodies with a fluorescent handle. In this system, labeling is monitored via SDS-PAGE with a fluorescence gel imager, where increased fluorescence intensity of a sample reflects increased labeling (Basic Protocol 2). Analysis of CIR:antibody target-specific immune activation is demonstrated with a flow cytometry-based antibody-dependent cellular phagocytosis (ADCP) assay (Basic Protocol 3). This ADCP protocol may be further used to discern CIR:antibody binding from covalent adduct formation (Alternate Protocol 3). For the protocols described, each method may be used to analyze characteristics of any covalent-tagging or antibody-recruiting small molecule or protein-based technology. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Determining "on-probe" reaction kinetics of CIR1/CIR4 via biolayer interferometry with Octet RED96 Alternate Protocol 1: Determining "in-solution" reaction kinetics of prostate-specific membrane antigen targeting CIR (CIR3) via biolayer interferometry with Octet RED96 Basic Protocol 2: Reaction kinetics of covalently labeled antibodies via fluorescence SDS-PAGE Basic Protocol 3: Small molecule-directed antibody-dependent cellular phagocytosis on live human cells measured via flow cytometry Alternate Protocol 2: Kinetic analysis of CIR3:antibody labeling via antibody-dependent cellular phagocytosis on flow cytometry Support Protocol 1: Activation of U937 monocytes with interferon γ Support Protocol 2: Labeling streptavidin beads with biotinylated prostate-specific membrane antigen receptor.
Cytidine acetyltransferases are an emerging class of nucleic-acid-modifying enzymes responsible for the establishment of N4 -acetylcytidine (ac4C) in RNA. In contrast to histone acetyltransferases, whose activity is commonly studied by western blotting, relatively few methods exist for quickly assessing the activity of cytidine acetyltransferases from a biological sample of interest or the distribution of ac4C across different RNA species. In this protocol, we describe a method for analysis of cellular cytidine acetyltransferase activity using dot- and immuno-northern-blotting-based detection. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Detection of N4 -Acetylcytidine in RNA by dot blotting Basic Protocol 2: Visualizing N4 -Acetylcytidine Distribution in RNA by northern blotting.
Reverse-polarity activity-based protein profiling (RP-ABPP) is a chemical proteomics approach that uses nucleophilic probes amenable to "click" chemistry deployed into living cells in culture to capture, immunoprecipitate, and identify protein-bound electrophiles. RP-ABPP is used to characterize the structure and function of reactive electrophilic post-translational modifications (PTMs) and the proteins harboring them, which may uncover unknown or novel functions. RP-ABPP has demonstrated utility as a versatile method to monitor the metabolic regulation of electrophilic cofactors, using a pyruvoyl cofactor in S-adenosyl-L-methionine decarboxylase (AMD1), and to discover novel types of electrophilic modifications on proteins in human cells, such as the glyoxylyl modification on secernin-3 (SCRN3). These cofactors cannot be predicted by sequence, and therefore this area is relatively undeveloped. RP-ABPP is the only global, unbiased approach to discover such electrophiles. Here, we describe the utility of these experiments and provide a detailed protocol for de novo discovery, quantitation, and global profiling of electrophilic functionality of proteins. © 2020 The Authors. Basic Protocol 1: Identification and quantification of probe-reactive proteins Basic Protocol 2: Characterization of the site of probe labeling Basic Protocol 3: Determination and quantitation of electrophile structure.
Metabolomic studies allow a deeper understanding of the processes of a given ecological community than nucleic acid-based surveys alone. In the case of the gut microbiota, a metabolic profile of, for example, a fecal sample provides details about the function and interactions within the distal region of the gastrointestinal tract, and such a profile can be generated in a number of different ways. This unit elaborates on the use of 1D 1 H NMR spectroscopy as a commonly used method to characterize small-molecule metabolites of the fecal metabonome (meta-metabolome). We describe a set of protocols for the preparation of fecal water extraction, storage, scanning, measurement of pH, and spectral processing and analysis. We also compare the effects of various sample storage conditions for processed and unprocessed samples to provide a framework for comprehensive analysis of small molecules from stool-derived samples. © 2020 Wiley Periodicals LLC Basic Protocol 1: Extracting fecal water from crude fecal samples Alternate Protocol 1: Extracting fecal water from small crude fecal samples Basic Protocol 2: Acquiring NMR spectra of metabolite samples Alternate Protocol 2: Acquiring NMR spectra of metabolite samples using Bruker spectrometer running TopSpin 3.x Alternate Protocol 3: Acquiring NMR spectra of metabolite samples by semiautomated process Basic Protocol 3: Measuring sample pH Support Protocol 1: Cleaning NMR tubes Basic Protocol 4: Processing raw spectra data Basic Protocol 5: Profiling spectra Support Protocol 2: Spectral profiling of sugars and other complex metabolites.
The reversible oxidation of protein tyrosine phosphatases (PTPs) impairs their ability to dephosphorylate substrates in vivo. This transient inactivation of PTPs occurs as their conserved catalytic cysteine residue reacts with cellular oxidants thereby abolishing the ability of this reactive cysteine to attack the phosphate of the target substrate. Hence, in vivo, the inhibition of specific PTPs in response to regulated and localized rises in cellular oxidants enables phospho-dependent signaling. We present assays that measure the endogenous activity of specific PTPs that become transiently inactivated in cells exposed to growth factors. Here, we describe the methods and highlight the pitfalls to avoid post-lysis oxidation of PTPs in order to assess the inactivation and the reactivation of PTPs targeted by cellular oxidants in signal transduction. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Cell transfection (optional) Support Protocol: Preparation of degassed lysis buffers Basic Protocol 2: Cellular extraction in anaerobic conditions Basic Protocol 3: Enrichment and activity assay of specific PTPs Alternate Protocol: Measurement of active PTPs via direct cysteinyl labeling.
Adenosine to-inosine (A-to-I) RNA editing is a conserved post-transcriptional modification that is critical for a variety of cellular processes. A-to-I editing is widespread in nearly all types of RNA, directly imparting significant global changes in cellular function and behavior. Dysfunctional RNA editing is also implicated in a number of diseases, and A-to-I editing activity is rapidly becoming an important biomarker for early detection of cancer, immune disorders, and neurodegeneration. While millions of sites have been identified, the biological function of the majority of these sites is unknown, and the regulatory mechanisms for controlling editing activity at individual sites is not well understood. Robust detection and mapping of A-to-I editing activity throughout the transcriptome is vital for understanding these properties and how editing affects cellular behavior. However, accurately identifying A-to-I editing sites is challenging because of inherent sampling errors present in RNA-seq. We recently developed Endonuclease V immunoprecipitation enrichment sequencing (EndoVIPER-seq) to directly address this challenge by enrichment of A-to-I edited RNAs prior to sequencing. This protocol outlines how to process cellular RNA, enrich for A-to-I edited transcripts with EndoVIPER pulldown, and prepare libraries suitable for generating RNA-seq data. © 2020 Wiley Periodicals LLC. Basic Protocol 1: mRNA fragmentation and glyoxalation Basic Protocol 2: EndoVIPER pulldown Basic Protocol 3: RNA-seq library preparation and data analysis.
O-GlcNAcylation is a posttranslational modification involving the addition of the single monosaccharide N-acetylglucosamine (GlcNAc) onto serine and threonine residues of intracellular proteins. Though O-GlcNAc is found on ∼1000 proteins in mammals, its specific function on individual substrates remains largely a mystery. To overcome this shortcoming, work has been put toward developing metabolic chemical reporters (MCRs) to label O-GlcNAcylated proteins for subsequent biochemical analysis. Typically, these MCRs are GlcNAc or GalNAc analogs functionalized with azide or alkyne handles. These unnatural sugar moieties can be metabolically incorporated directly on to protein substrates. The protocols outlined in this article describe how to use MCRs as tools for visualizing and identifying potentially O-GlcNAc modified proteins via in-gel fluorescence, Western blotting, and mass spectrometry. Taken together, MCR labeling provides a powerful tool to discover where and when substrates are O-GlcNAc modified. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Treatment of cells and CuAAC Basic Protocol 2: In-gel fluorescence of labeled cell lysates (1 mg scale) Basic Protocol 3: Enrichment of labeled proteins, trypsinolysis, and collection of peptides for proteomics Basic Protocol 4: Proteomic identification of labeled proteins.
Over the past few decades, numerous examples have demonstrated that intrinsic disorder in proteins lies at the heart of many vital processes, including transcriptional regulation, stress response, cellular signaling, and most recently protein liquid-liquid phase separation. The so-called intrinsically disordered proteins (IDPs) involved in these processes have presented a challenge to the classic protein "structure-function paradigm," as their functions do not necessarily involve well-defined structures. Understanding the mechanisms of IDP function is likewise challenging because traditional structure determination methods often fail with such proteins or provide little information about the diverse array of structures that can be related to different functions of a single IDP. Single-molecule fluorescence methods can overcome this ensemble-average masking, allowing the resolution of subpopulations and dynamics and thus providing invaluable insights into IDPs and their function. In this protocol, we describe a ratiometric single-molecule Förster resonance energy transfer (smFRET) routine that permits the investigation of IDP conformational subpopulations and dynamics. We note that this is a basic protocol, and we provide brief information and references for more complex analysis schemes available for in-depth characterization. This protocol covers optical setup preparation and protein handling and provides insights into experimental design and outcomes, together with background information about theory and a brief discussion of troubleshooting. © 2020 by John Wiley & Sons, Inc. Basic Protocol: Ratiometric smFRET detection and analysis of IDPs Support Protocol 1: Fluorophore labeling of a protein through maleimide chemistry Support Protocol 2: Sample chamber preparation Support Protocol 3: Determination of direct excitation of acceptor by donor excitation and leakage of donor emission to acceptor emission channel.
Small-molecule drug discovery can be hindered by the formation of aggregates that act as non-selective inhibitors of drug targets. Such aggregates appear as false positives in high-throughput screening campaigns and can bedevil structure-activity relationships during compound optimization. Protocols are described for resonant waveguide grating (RWG) and dynamic light scattering (DLS) as microplate-based high-throughput approaches to identify compound aggregation. Resonant waveguide grating and dynamic light scattering give equivalent results for the compound test set, as assessed with Bland-Altman analysis. © 2019 The Authors. Basic Protocol 1: Resonant waveguide grating (RWG) in 384-well or 1536-well plate format to detect compound aggregation Basic Protocol 2: Dynamic light scattering (DLS) in 384-well plate format to detect compound aggregation.
Subtiligase is a powerful enzymatic tool for N-terminal modification of proteins and peptides. In a typical subtiligase-catalyzed N-terminal modification reaction, a peptide ester donor substrate is ligated onto the unblocked N terminus of a protein, resulting in the exchange of the ester bond in the donor substrate for an amide bond between the donor substrate and protein N terminus. Using this strategy, new chemical probes and payloads, such as fluorophores, affinity handles, cytotoxic drugs, and reactive functional groups, can be introduced site-specifically into proteins. While the efficiency of this reaction depends on the sequences to be ligated, a panel of mutants was recently developed that expands the scope of substrate sequences that are suitable for subtiligase modification. This article outlines the steps for applying subtiligase or specificity variants for both site-specific bioconjugation of purified proteins and for global modification of cellular N termini to enable their sequencing by tandem mass spectrometry. © 2020 by John Wiley & Sons, Inc. Basic Protocol 1: Subtiligase-catalyzed site-specific protein bioconjugation Support Protocol 1: Expression and purification of subtiligase-His6 Support Protocol 2: Subtiligase substrate synthesis Basic Protocol 2: Subtiligase N terminomics using a cocktail of subtiligase specificity mutants.
Protein phosphorylation plays a critical role in the regulation of cellular function. Information on protein phosphorylation and the responsible kinases is important for understanding intracellular signaling. A method for in vivo screening of kinase substrates named KIOSS (kinase-oriented substrate screening) has been developed. This protocol provides a method that utilizes phosphoprotein-binding modules such as 14-3-3 protein, the pin1-WW domain, and the chek2-FHA domain as biological filters to successfully enrich phosphorylated proteins related to intracellular signaling rather than housekeeping and/or structural proteins. More than 1000 substrate candidates for PKA, PKC, MAPK, and Rho-kinase in HeLa cells, as well as phosphorylation downstream of D1R, NMDAR, adenosine A2a receptor, PKA, PKC, MAPK, and Rho-kinase in mouse brain slice cultures have been identified by this method. An online database named KANPHOS (Kinase-Associated Neural Phospho-Signaling) provides the phosphorylation signals identified by these studies, as well as those previously reported in the literature. © 2019 by John Wiley & Sons, Inc.
Bacterial cells utilize small carbohydrate building blocks to construct peptidoglycan (PG), a highly conserved mesh-like polymer that serves as a protective coat for the cell. PG production has long been a target for antibiotics, and its breakdown is a source for human immune recognition. A key component of bacterial PG, N-acetyl muramic acid (NAM), is a vital element in many synthetically derived immunostimulatory compounds. However, the exact molecular details of these structures and how they are generated remain unknown due to a lack of chemical probes surrounding the NAM core. A robust synthetic strategy to generate bioorthogonally tagged NAM carbohydrate units is implemented. These molecules serve as precursors for PG biosynthesis and recycling. Escherichia coli cells are metabolically engineered to incorporate the bioorthogonal NAM probes into their PG network. The probes are subsequently modified using copper-catalyzed azide-alkyne cycloaddition to install fluorophores directly into the bacterial PG, as confirmed by super-resolution microscopy and high-resolution mass spectrometry. Here, synthetic notes for key elements of this process to generate the sugar probes as well as streamlined user-friendly metabolic labeling strategies for both microbiology and immunological applications are described. © 2019 by John Wiley & Sons, Inc. Basic Protocol 1: Synthesis of peracetylated 2-azido glucosamine Basic Protocol 2: Synthesis of 2-azido and 2-alkyne NAM Basic Protocol 3: Synthesis of 3-azido NAM methyl ester Basic Protocol 4: Incorporation of NAM probes into bacterial peptidoglycan Basic Protocol 5: Confirmation of bacterial cell wall remodeling by mass spectrometry.
Human kinases are a large family of proteins (500+) that catalyze ATP‐dependent phosphorylation of protein and metabolite substrates to regulate diverse facets of cell biology. Dysregulation and mutations of protein kinases are linked to human disease, providing opportunities for developing pharmacological agents as potential therapy. Assessing the selectivity of pharmacological compounds targeting this enzyme class is critical given that off‐target activity of kinase inhibitor drugs may result in toxicity. This set of protocols outlines use of ATP acyl phosphate activity–based probes to evaluate the potency and selectivity of kinase inhibitors via fluorescent gel– and mass spectrometry–based detection methods. These competitive chemical proteomic assays can evaluate engagement of >200 native kinase targets directly in complex proteomes. © 2019 by John Wiley & Sons, Inc.
T4 DNA ligase in high concentrations of certain crowding agents and cosolutes catalyzes the synthesis of a series of backbone‐modified oligonucleotides that are difficult to obtain chemically. Backbone‐modified nucleic acids are often enzymatically and chemically more stable, making them interesting as potential diagnostic or therapeutic agents, as a biosafety tool, or in nanotechnology. In this article, we describe a small‐scale experiment to probe the efficiency of the ligation reaction of modified oligonucleotides in the presence of 3 M betaine and 10% PEG 8000, followed by large‐scale ligation with subsequent isolation of the ligated oligonucleotide. The correct product formation can be verified using denaturing polyacrylamide gel electrophoresis and mass spectrometry. © 2019 by John Wiley & Sons, Inc.
Small open reading frames (smORFs) encode previously unannotated polypeptides or short proteins that regulate translation in cis (eukaryotes) and/or are independently functional (prokaryotes and eukaryotes). Ongoing efforts for complete annotation and functional characterization of smORF-encoded proteins have yielded novel regulators and therapeutic targets. However, because they are excluded from protein databases, initiate at non-AUG start codons, and produce few unique tryptic peptides, unannotated small proteins cannot be detected with standard proteomic methods. Here,, we outline a procedure for mass spectrometry-based detection of translated smORFs in cultured human cells from protein extraction, digestion, and LC-MS/MS, to database preparation and data analysis. Following proteomic detection, translation from a unique smORF may be validated via siRNA-based silencing or overexpression and epitope tagging. This is necessary to unambiguously assign a peptide to a smORF within a specific transcript isoform or genomic locus. Provided that sufficient starting material is available, this workflow can be applied to any cell type/organism and adjusted to study specific (patho)physiological contexts including, but not limited to, development, stress, and disease. © 2019 by John Wiley & Sons, Inc. Basic Protocol 1: Protein extraction, size selection, and trypsin digestion Alternate Protocol 1: In-solution C8 column size selection Support Protocol 1: Chloroform/methanol precipitation Support Protocol 2: Reduction, alkylation, and in-solution protease digestion Support Protocol 3: Peptide de-salting Basic Protocol 2: Two-dimensional LC-MS/MS with ERLIC fractionation Basic Protocol 3: Transcriptomic database construction Alternate Protocol 2: Transcriptomics database generation with gffread Basic Protocol 4: Non-annotated peptide identification from LC-MS/MS data Basic Protocol 5: Validation using isotopically labeled synthetic peptide standards and siRNA Basic Protocol 6: Transcript validation using transient overexpression.
