生物化学

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.

Protein purification is essential for drug development, antibody production, and structural biology. We propose a cost-effective chromatography method using elastin-like polypeptide (ELP) as an aggregating core. In this approach, a chilled (target protein)-ELP fusion is loaded onto an immobilized metal affinity chromatography (IMAC) column equilibrated with low-salt buffer. Impurities are removed with warm high-salt buffer washes. Warming the column above the ELP’s transition temperature (T_m) triggers ELP aggregation, physically trapping the target protein between beads. Subsequent stringent washing (high salt/imidazole) eliminates residual contaminants. Finally, cooling with cold low-salt buffer reverses aggregation, eluting the purified target protein. This method eliminates the need for advanced chromatography systems while achieving high purity through dual mechanisms: (1) IMAC affinity and (2) temperature-dependent physical capture. The ELP’s reversible phase transition offers a simplified yet efficient purification platform, particularly valuable for lab-scale production of challenging proteins.

The voltage-gated proton channel (H_v1) is a membrane protein that dissipates acute cell proton accumulations. To understand the molecular mechanisms explaining H_v1 function, methods for purifying the protein are needed. Previously, methods were developed for expressing and purifying human H_v1 (hH_v1) in yeast and later in bacteria. However, these methodologies produced low protein yields and had high production costs, considerably limiting their usefulness. The protocol described in this work was developed to overcome those limitations. hH_v1 is overexpressed in bacteria, solubilized with the detergent Anzergent 3–12, and purified by immobilized metal affinity chromatography (IMAC) and size-exclusion chromatography (SEC). Our protocol produced higher protein yields at lower costs than previously published methodologies.

Intermediate states are often populated during the folding and unfolding reactions of a protein, and their detection is very challenging as they form transiently. Structural characterization of these short-lived intermediate species is difficult as it requires high-resolution methodologies. Hydrogen exchange-mass spectrometry (HX-MS) can identify and yield direct structural information on folding and unfolding intermediates, as well as information about the cooperativity of the folding or unfolding processes. The mass distributions of intact protein molecules are obtained first to determine their exchange pattern. Then, segment-specific structural information is obtained by analyzing the fragments of the protein. Enzymatic digestion is widely used with HX to determine the sequence-specific structural changes that occur to the protein during folding or unfolding. However, if a protein is an inhibitor of the protease, then alternative methodologies are required. Using electron transfer dissociation (ETD), it is possible to fragment the protein inside a mass spectrometer, and segment-specific structural changes occurring during the folding and unfolding process can be determined. In the case of HX-ETD-MS, protein molecules are first allowed to undergo HX, followed by their fragmentation. Deuterium retention in each fragment is measured. Very little, if any, scrambling of deuterium across fragments occurs during ETD-enabled fragmentation; hence, there is little scope for misinterpretation of the HX data.

Ubiquitination is a post-translational protein modification that regulates a vast majority of processes during protein homeostasis. The covalent attachment of ubiquitin is a highly regulated process carried out by the sequential action of the three enzymes E1, E2, and E3. E3 ligases share a dual function of 1) transferring covalently attached ubiquitin from the catalytic cysteine of E2 (E2~Ub) to the substrate and 2) providing substrate specificity. Our current knowledge of their individual substrate pools is incomplete due to the difficult capture of these transient substrate–E3 ligase interactions. Here, we present an efficient protocol that enables the selective biotinylation of substrates of a given ubiquitin ligase. In brief, the candidate E3 ligase is fused to the biotin ligase BirA and ubiquitin to a biotin acceptor peptide, an Avi-tag variant (-2) AP. Cells are co-transfected with these fusion constructs and exposed to biotin, resulting in a BirA-E3 ligase-catalyzed biotinylation of (-2) AP-Ub when in complex with E2. As the next step, the biotinylated (-2) AP-Ub is transferred covalently to the substrate lysine, which enables an enrichment via denaturing streptavidin pulldown. Substrate candidates can then be identified via mass spectrometry (MS). Our ubiquitin-specific proximity-dependent labeling (Ub-POD) method allows robust biotinylation of the ubiquitylation substrates of a candidate E3 ligase thanks to the wild-type BirA and biotin acceptor peptide fused to the E3 and Ub, respectively. Because of the highly Ub-specific labeling, Ub-POD is more appropriate for identifying ubiquitination substrates compared to other conventional proximity labeling or immunoprecipitation (IP) approaches.

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.

De novo synthesis of purine nucleotide is essential for the production of genetic materials and cellular chemical energy. PRPP amidotransferase (PPAT) is the rate-limiting enzyme in de novo purine synthesis, thereby playing a crucial regulatory role in this pathway. Recent studies suggest that metabolic enzymes, including PPAT, form condensates through phase separation to regulate cellular metabolism in response to environmental changes. However, due to the lack of methods for purifying eukaryotic PPAT, the biophysical properties of the enzyme have remained unknown. Here, I describe a protocol for purifying budding yeast PPAT tagged with green fluorescent protein from yeast cells, as well as an in vitro assay to examine condensation of the fluorescent PPAT by microscopy. These techniques enabled us to elucidate the mechanism controlling PPAT condensation and may also be applicable to the purification and condensation assay of other enzymes.

We have observed that some proinsulin molecules in pancreatic islets and beta cell lines have incomplete or improper intramolecular disulfide bonds. These misfolded monomers can form intermolecular disulfide-linked complexes. Accurately quantifying the fraction of proinsulin molecules contained in these complexes is challenging. By proinsulin immunoblotting after nonreducing SDS-PAGE, the signal for disulfide-linked complexes can exceed the total proinsulin signal detected after reducing SDS-PAGE (i.e., overestimating the abundance of misfolded species due to antibody affinity differences). However, after modification of the SDS-PAGE and electrotransfer protocol, we have been able to more accurately estimate the fraction of proinsulin monomers folded to the native state, as well as misfolded proinsulin monomers and disulfide-linked complexes. Our improved technique offers the ability to obtain a more precise assessment of proinsulin misfolding under different environmental conditions in beta cells and normal islets and in diabetes.

Protein purification is a critical step in both life sciences and biomanufacturing. Traditional affinity chromatography (AC) methods, including His-tag-based purification, provide high-purity proteins but are limited by the high cost of resins and the need for additional tag-removal steps. In this protocol, we present a reusable SpyDock-modified epoxy resin coupled with a pH-inducible self-cleaving intein for direct purification of proteins with authentic N-termini. This method enables efficient protein purification from cell lysates, achieving high purity (>90%) and yields comparable to the His-tag approach, without requiring tag removal. The SpyDock-modified resin protocol is robust, easy to implement, and cost-effective, making it suitable for both research and large-scale industrial applications.

Expansion microscopy (ExM) is an imaging technique that enables super-resolution imaging of biological specimens using conventional confocal microscopy. This process entails the isotropic physical expansion of a (biomolecular) sample that has been cross-linked to a swellable polymer. The grafting of biomolecules (and the subsequent fluorescent readout) is accomplished by introducing an acryloyl group to the amine groups of lysine residues within the proteins, enabling subsequent imaging. However, visualizing actin filaments with high spatial resolution using ExM remains challenging. Herein, we report the construction of a phalloidin conjugate containing actin stains and their application in ExM. This protocol highlights the efficacy of trifunctional linker (TRITON/Actin-ExM) for F-actin imaging, demonstrating that TRITON-labeled actin allows for efficient anchoring and signal retention, enabling robust visualization of actin filaments in expansion microscopy.