生物化学

Flippases, a functionally distinct group of transmembrane proteins that flip lipids from the extracellular or luminal side to the cytosolic side of biological membranes, are key players in many important physiological processes, such as membrane trafficking and cellular signaling. To study the function of these membrane proteins under chemically defined conditions, reconstituting them into artificial vesicles is a crucial and effective approach. There are various methods for protein reconstitution involving different detergents and detergent removal techniques to integrate membrane proteins into artificial vesicles. In this protocol, we describe the reconstitution of the yeast flippase complex Drs2-Cdc50, which translocates phosphatidylserine across membranes of the trans-Golgi network at the expense of ATP hydrolysis. The flippase complex is incorporated into liposomes using a zwitterionic detergent, followed by detergent removal via dialysis—a gentle and effective strategy that helps preserve protein function. To evaluate the activity of the reconstituted flippase complex, two complementary assays are employed: (1) a fluorescence-based quenching assay to measure lipid transport; and (2) an ATPase assay using an ATP-regenerating system to measure ATP hydrolysis. Together, these methods provide a robust platform for analyzing the functional reconstitution of Drs2-Cdc50 in a defined membrane environment.

Cathepsin L (CTSL), a lysosomal cysteine protease belonging to the papain-like protease family, is primarily involved in intracellular protein degradation, antigen processing, and extracellular matrix remodeling. It plays critical roles in pathological conditions, including cancer metastasis, neurodegenerative disorders, and viral infection, due to dysregulated activity or overexpression. Thus, inhibitors targeting CTSL are under investigation for therapeutic applications. Current approaches for identifying CTSL inhibitors predominantly rely on fluorescence-labeled substrates, fluorescence resonance energy transfer (FRET), and cell-based screening assays. Here, we applied the principle of fluorescence polarization (FP) to the detection of substrate cleavage activity by CTSL through changes in millipolarization unit (mp) values and established a cost-effective, quantitative, reagent- and time-saving inhibitor high-throughput screening (HTS) assay. We also provide detailed steps for the expression and purification of highly active CTSL from eukaryotic cells, which lays a solid foundation for the FP-based assay. A key advantage of this assay lies in its reduced susceptibility to fluorescence interference, as the fluorescein isothiocyanate (FITC) fluorophore exhibits high quantum efficiency with an emission peak at 535 nm—a wavelength range distinct from most naturally occurring fluorescent molecules. The assay’s adaptability to reaction time, temperature, and dimethyl sulfoxide (DMSO) concentration minimizes false-positive or false-negative results caused by minor experimental inconsistencies, streamlining the screening process. Furthermore, the protocol requires fewer operational steps, reduced incubation time, and lower quantities of CTSL and substrates compared to conventional methods. This rapid, cost-effective, and scalable approach aligns well with the demands of HTS platforms.

The cAMP-dependent protein kinase (PKA) is one of the most extensively distributed kinases among intracellular signal cascades, with a pivotal role in the regulation of various processes, including the capacitation of sperm cells. Traditional assessments of PKA activity rely on the utilization of [γ-³²P] ATP and the Kemptide peptide as a substrate. This strategy presents several major drawbacks, including high costs and health risks derived from the manipulation of radioactive isotopes. In this work, we introduce an enhanced non-radioactive assay to quantify PKA activity, termed kinase mobility shift assay (KiMSA), based on the use of a fluorescent-labeled Kemptide (Kemptide-FITC). Once the kinase reaction is terminated, the products can be easily resolved through electrophoresis on an agarose gel and quantified by fluorescence densitometry. We show that KiMSA is suitable for isolated PKA as well as for the enzyme in cell extracts. In addition, it enables quantification of PKA activity during the progression of mouse sperm capacitation. Furthermore, the assay enables monitoring the inhibition of PKA with pharmacological inhibitors in live cells. Therefore, the experimental and optimal assay conditions are set so that KiMSA can be used to assess in vitro as well as in vivo PKA activity in sperm cells. Finally, this method allows for measurement of cAMP concentrations, rendering a versatile technique for the study of cAMP/PKA pathways.

X-succinate synthase enzymes (XSSs) are a class of glycyl radical enzymes (GREs) that play a pivotal role in microbial anaerobic hydrocarbon degradation. They catalyze the addition of hydrocarbons to fumarate using a protein-based glycyl radical, which must first be installed by a radical S-adenosylmethionine (rSAM) activating enzyme (AE). Once activated, XSS enzymes can undergo multiple catalytic cycles, forming C(sp³)–C(sp³) bonds with high stereoselectivity—a feature that highlights their potential as asymmetric biocatalysts. Due to the insolubility of XSS-AEs when heterologously expressed in Escherichia coli, studies have relied on in vivo radical installation protocols. Although these methods have illuminated fundamental details of XSS mechanisms, the inability to install a glycyl radical in vitro has limited biochemical studies and biotechnological advances using these enzymes. Here, we describe an in vitro protocol for reconstituting the activity of benzylsuccinate synthase (BSS), an XSS that catalyzes the addition of toluene to fumarate to form R-benzylsuccinate. To enable in vitro glycyl radical installation, we identified a soluble homolog via genome mining: 4-isopropylbenzylsuccinate synthase activating enzyme (IbsAE). IbsAE was expressed in E. coli and anaerobically purified in moderate yields (6–8 mg of protein per liter of culture); herein, we outline the expression and anaerobic purification of both IbsAE and BSS proteins. We describe a reproducible method for in vitro glycyl radical installation using these recombinant proteins and provide guidance on quantifying radical formation. Our optimized protocol consistently achieves 30%–50% radical installation, comparable to other in vitro GRE activations. Lastly, we demonstrate the application of this protocol for in vitro hydroalkylation reactions, achieving high assay yields (89%–97%). This protocol enables biochemical experiments that were previously challenging using cell extracts and accelerated advancements in XSS engineering and use in biocatalysis.

Biomolecular condensates are macromolecular assemblies constituted of proteins that possess intrinsically disordered regions and RNA-binding ability together with nucleic acids. These compartments formed via liquid-liquid phase separation (LLPS) provide spatiotemporal control of crucial cellular processes such as RNA metabolism. The liquid-like state is dynamic and reversible, containing highly diffusible molecules, whereas gel, glass, and solid phases might not be reversible due to the strong intermolecular crosslinks. Neurodegeneration-associated proteins such as the prion protein (PrP) and Tau form liquid-like condensates that transition to gel- or solid-like structures upon genetic mutations and/or persistent cellular stress. Mounting evidence suggests that progression to a less dynamic state underlies the formation of neurotoxic aggregates. Understanding the dynamics of proteins and biomolecules in condensates by measuring their movement in different timescales is indispensable to characterize their material state and assess the kinetics of LLPS. Herein, we describe protein expression in E. coli and purification of full-length mouse recombinant PrP, our in vitro experimental system. Then, we describe a systematic method to analyze the dynamics of protein condensates by X-ray photon correlation spectroscopy (XPCS). We also present fluorescence recovery after photobleaching (FRAP)-optimized protocols to characterize condensates, including in cells. Next, we detail strategies for using fluorescence microscopy to give insights into the folding state of proteins in condensates. Phase-separated systems display non-equilibrium behavior with length scales ranging from nanometers to microns and timescales from microseconds to minutes. XPCS experiments provide unique insights into biomolecular dynamics and condensate fluidity. Using the combination of the three strategies detailed herein enables robust characterization of the biophysical properties and the nature of protein phase-separated states.

Xylan is the main component of hemicellulose and consists of a complex heteropolysaccharide with a heterogeneous structure. This framework, in addition to the crystalline structure of cellulosic fibers and the rigidity of lignin, makes lignocellulosic biomass (LCB) highly recalcitrant to degradation. Xylanases are glycoside hydrolases that cleave the β-1,4-glycoside linkages in the xylan backbone and have attracted increasing attention due to their potential uses in various industrial sectors such as pulp and paper, baking, pharmaceuticals, and lignocellulosic biorefining. For decades, the measurement of xylanase activity was based on reducing sugar quantification methods like DNS or Nelson/Somogyi assays, with numerous limitations in terms of specificity and interference from other enzymatic activities. A better alternative is the colorimetric Azo-Xylan assay, which specifically measures the endo-1,4-β-D-xylanase activity. In this study, the Azo-Xylan protocol was adapted from the company Megazyme to determine the enzymatic activity of thermostable xylanases produced by microbial consortia (i.e., microbiomes), aiming to determine biochemical features such as temperature and pH optima, thermostability, and shelf life. This modified approach offers a rapid, cost-effective, and highly specific method for the determination of xylanase activity in complex mixtures, helping the development of a xylanase-based method for the hydrolysis of hard-degrading substrates in bio-based industries.

The motile parameters of kinesin superfamily proteins are fundamental to intracellular transport. Single-molecule motility assays using total internal reflection fluorescence (TIRF) microscopy are a gold standard technique for measuring the motile parameters of kinesin motors. With this technique, one can evaluate the velocity, run length, and binding frequency of kinesins on microtubules by directly observing their motility. This protocol provides a comprehensive procedure for single molecule assays of kinesins, including the preparation of labeled microtubules, the measurement of kinesin motility via TIRF microscopy, and the quantification of kinesin motor parameters.

Alpha-protein kinase 1 (ALPK1) is normally activated by bacterial ADP-heptose as part of the innate immune response, leading to the initiation of downstream signalling events that culminate in the activation of transcription factors such as NF-κB and AP-1. In contrast, disease-causing mutations in ALPK1 that cause ROSAH syndrome or spiradenoma allow ALPK1 to be activated in cells in the absence of bacterial infection (i.e., without ADP-heptose). This protocol describes a semi-quantitative reporter assay based on ALPK1 knockout HEK-Blue cells that measures the activity of transfected wildtype and disease-causing forms of ALPK1 by virtue of their ability to activate the transcription factors NF-κB and AP-1. These cells express a synthetic gene encoding alkaline phosphatase under the control of an NF-κB/AP-1-dependent promoter, and consequently, the activation of ALPK1 leads to the production of alkaline phosphatase, which is secreted into the culture media and can be measured colorimetrically at 645 nm after the addition of a detection reagent.

The planar lipid bilayer (PLB) technique represents a highly effective method for the study of membrane protein properties in a controlled environment. The PLB method was employed to investigate the role of mitochondrial inner membrane protein 17 (MPV17), whose mutations are associated with a hepatocerebral form of mitochondrial DNA depletion syndrome (MDS). This protocol presents a comprehensive, step-by-step guide to the assembly and utilization of a PLB system. The procedure comprises the formation of a lipid bilayer over an aperture, the reconstitution of the target protein, and the utilization of electrophysiological recording techniques to monitor channel activity. Furthermore, recommendations are provided for optimizing experimental conditions and overcoming common challenges encountered in PLB experiments. Overall, this protocol highlights the versatility of the PLB technique in advancing our understanding of membrane protein function and its broad application in various fields of research.

ALPK1 is an atypical protein kinase that is activated during bacterial infection by ADP-heptose and phosphorylates TIFA to activate a cell signaling pathway. In contrast, specific mutations in ALPK1 allow it to also be activated by endogenous human nucleotide sugars such as UDP-mannose, leading to the phosphorylation of TIFA in the absence of infection. This protocol describes a quantitative, cell-free phosphorylation assay that can directly measure the catalytic activity of wildtype and disease-causing ALPK1 in the presence of different nucleotide sugars. In this method, overexpressed ALPK1 is first immunoprecipitated from the extracts of ALPK1 knockout HEK-Blue cells transfected with plasmids encoding either FLAG-tagged wildtype or mutant ALPK1, and then subjected to a radioactive phosphorylation assay in which the phosphorylation of purified GST-tagged TIFA by ALPK1 is quantified by measuring the incorporation of radioactivity derived from radiolabeled ATP.