Microbiology

Bio-hydrogen production is an eco-friendly alternative to commercial H₂ production, taking advantage of natural systems. Microbial hydrogenases play a main role in biological mechanisms, catalyzing proton reduction to molecular hydrogen (H₂) formation under ambient conditions. Direct determination is an important approach to screen bacteria with active hydrogenase and accurately quantify the amount of H₂ production. Here, we present a detailed protocol for determining hydrogenase activity based on H₂ production using methyl viologen (MV²⁺) as an artificial reductant, directly monitored by gas chromatography. Recombinant Escherichia coli is used as a hydrogenase-enriched model in this study. Even so, this protocol can be applied to determine hydrogenase activity in all biological samples.

Key features

• This protocol is optimized for a wide variety of biological samples; both purified hydrogenase (in vitro) and intracellular hydrogenase (in vivo) systems.

• Direct, quantitative, and accurate method to detect the amount of H₂ by gas chromatography with reproducibility.

• Requires only 2 h to complete and allows testing various conditions simultaneously.

• Kinetic plot of H₂ production allows to analyze kinetic parameters and estimate the efficiency of hydrogenase from different organisms.

Graphical overview

Ubiquitination is a post-translational modification conserved across eukaryotic species. It contributes to a variety of regulatory pathways, including proteasomal degradation, DNA repair, and cellular differentiation. The ubiquitination of substrate proteins typically requires three ubiquitination enzymes: a ubiquitin-activating E1, a ubiquitin-conjugating E2, and an E3 ubiquitin ligase. Cooperation between E2s and E3s is required for substrate ubiquitination, but some ubiquitin-conjugating E2s are also able to catalyze by themselves the formation of free di-ubiquitin, independently or in cooperation with a ubiquitin E2 variant. Here, we describe a method for assessing (i) di-ubiquitin formation by an E1 together with an E2 and an E2 variant, and (ii) the cooperation of an E3 with an E1 and E2 (with or without the E2 variant). Reaction products are assessed using western blotting with one of two antibodies: the first detects all ubiquitin conjugates, while the second specifically recognizes K63-linked ubiquitin. This allows unambiguous identification of ubiquitinated species and assessment of whether K63 linkages are present. We have developed these methods for studying ubiquitination proteins of Leishmania mexicana, specifically the activities of the E2, UBC2, and the ubiquitin E2 variant UEV1, but we anticipate the assays to be applicable to other ubiquitination systems with UBC2/UEV1 orthologues.

Here, we present the first quantitative method for the activity analysis of protealysin-like protease (PLP) inhibitors. This approach is based on a previously developed method for protealysin activity determination by hydrolysis of internally quenched fluorescent peptide substrate 2-aminobenzoyl-L-arginyl-L-seryl-L-valyl-L-isoleucyl-L-(ϵ-2,4-dinitrophenyl)lysine. In this protocol, we significantly reduced enzyme concentration and introduced some minor modifications to decrease variation between replicates. The protocol was validated using emfourin, a novel proteinaceous metalloprotease inhibitor. Data obtained demonstrates that the developed assay method is an affordable approach for characterizing and screening various PLP inhibitors.

Graphical abstract:

Malaria remains a major public health issue, infecting nearly 220 million people every year. The spread of drug-resistant strains of Plasmodium falciparum around the world threatens the progress made against this disease. Therefore, identifying druggable and essential pathways in P. falciparum parasites remains a major area of research. One poorly understood area of parasite biology is the formation of disulfide bonds, which is an essential requirement for the folding of numerous proteins. Specialized chaperones with thioredoxin (Trx) domains catalyze the redox functions necessary for breaking incorrect and forming correct disulfide bonds in proteins. Defining the substrates of these redox chaperones is difficult and immunoprecipitation based assays cannot distinguish between substrates and interacting partners. Further, the substrate or client interactions with the redox chaperones are usually transient in nature. Activity based crosslinkers that rely on the nucleophilic cysteines on Trx domains and the disulfide bond forming cysteines on clients provide an easily scalable method to trap and identify the substrates of Trx-domain containing chaperones. The cell permeable crosslinker divinyl sulfone (DVSF) is active only in the presence of nucleophilic cysteines in proteins and, therefore, traps Trx domains with their substrates, as they form mixed disulfide bonds during the course of their catalytic activity. This allows the identification of substrates that rely on Trx activity for their folding, as well as discovering small molecules that interfere with Trx domain activity.

Graphic abstract:

Identification of thioredoxin domain substrates via divinylsulfone crosslinking and immunoprecipitation-mass spectrometry.

Ustilago maydis, a basidiomycete that infects Zea mays, is one of the top ten fungal models for studying DNA repair, signal transduction pathways, and dimorphic transitions, among other processes. From a metabolic point of view, U. maydis lacks fermentative capacity, pointing to mitochondria as a key player in central metabolism. Oxidative phosphorylation, synthesis of heme groups, Krebs cycle, β-oxidation of fatty acids, and synthesis of amino acids are some of the processes that take place in mitochondria. Given the importance of this organelle in eukaryotic cells in general, and in fungal cells in particular, we present a protocol for the isolation of U. maydis mitochondria based on the enzymatic disruption of U. maydis cell wall and differential centrifugation. The method can easily be extrapolated to other fungal species, by using appropriate lytic enzymes.

Flavodoxin-like proteins (Fld-LPs) are an important constituent of the oxidative stress defense system in several organisms and highly conserved from bacteria to humans. These proteins possess NAD(P)H:quinone oxidoreductase activity and convert quinones to hydroquinones through two-electron reduction, using NAD(P)H and quinone as electron donor and acceptor, respectively. Purified yeast and bacterial Fld-LPs exhibit NAD(P)H:quinone oxidoreductase activity in vitro. Here, we describe a protocol to measure oxidoreductase activity of Fld-LPs that are present in extracts of whole cells. We have recently shown that the assembly and activity of a Fld-LP, CgPst2, is regulated by an aspartyl protease-mediated cleavage of its C-terminus in the pathogenic yeast Candida glabrata. Mutant yeast where the CgPST2 gene was deleted lacked cellular NAD(P)H:quinone oxidoreductase activity and displayed elevated susceptibility to menadione stress. The protocol described herein is based on the measurement of NADH oxidation (conversion of NADH to NAD⁺) by endogenous Fld-LPs in the presence of quinone menadione. This assay can be performed with whole cell lysates prepared by the mechanical lysis of C. glabrata cells and does not require expression and purification of Fld-LPs from a heterogeneous system, thereby allowing researchers to study the effect of different posttranslational modifications and varied structural states of Fld-LPs on their enzymatic activities. Since many FLP-LPs are known to exist in dimeric and tetrameric states possessing differential activities, our efficient and easy-to-use assay can reliably detect and validate their quinone reductase activities. Although we have used menadione with CgPst2 enzyme in our study, the protocol can easily be modified to examine the presence of Fld-LPs with specificity for other quinones. As this assay does not require many expensive chemicals, it can readily be scaled up and adapted for other medically important fungi and potentially be a useful tool to characterize fungal oxidative stress response systems and screen inhibitors specific for fungal Fld-LPs, thereby contributing to our understanding of fungal pathogenesis mechanisms.

Nitrate is one of the major inorganic nitrogen sources for microorganisms. Many bacterial and archaeal lineages can express cytoplasmic assimilatory nitrate reductase (NAS), which catalyzes the rate-limiting reduction of nitrate to nitrite in the nitrate assimilation pathway. Here, we present a detailed protocol for measuring in vitro nitrate reductase (NaR) activity of NAS enzymes from Mycolicibacterium smegmatis crude extract using both physiological and non-physiological electron donors.

Glutamylation is a posttranslational modification where the amino group of a free glutamate amino acid is conjugated to the carboxyl group of a glutamate side chain within a target protein. SidJ is a Legionella kinase-like protein that has recently been identified to perform protein polyglutamylation of the Legionella SdeA Phosphoribosyl-Ubiquitin (PR-Ub) ligase to inhibit SdeA’s activity. The attachment of multiple glutamate amino acids to the catalytic glutamate residue of SdeA by SidJ inhibits SdeA’s modification of ubiquitin (Ub) and ligation activity. In this protocol, we will discuss a SidJ non-radioactive, in vitro glutamylation assay using its substrate SdeA. This will also include a second reaction to assay the inhibition of SdeA by using both modification of free Ub and ligation of ADP-ribosylated Ubiquitin (ADPR-Ub) to SdeA’s substrate Rab33b. Prior to the identification and publication of SdeA’s activity, no SdeA inhibition assays existed. Our group and others have demonstrated various methods to display inhibition of SdeA’s activity. The alternatives include measurement of ADP-ribosylation of Ub using radioactive NAD, NAD hydrolysis, and Western blot analysis of HA-Ub ligation by SdeA. This protocol will describe the inhibition of both ubiquitin modification and the PR-Ub ligation by SdeA using inexpensive standard gels and Coomassie staining.

Kinases and ATPases perform essential biological functions in metabolism and regulation. Activity of these enzymes is commonly measured by coupling ATP consumption to the synthesis of a detectable product. For most assay systems the ATP concentration during the reaction is unknown, compromising the precision of the assay.

Using the ADP-specific hexokinase (ADP-HK) from the thermophilic archaeon Thermococcus litoralis the protocol outlined here allows real time coupling of ATP consumption to downstream signal change enabling accurate kinetic measurements. ADP-HK phosphorylates glucose that is then used by glucose-6-phosphate dehydrogenase to reduce NAD⁺ to NADH which can be measured at 340 nm. We have shown this assay to be sensitive to the detection of micromole quantities of ADP with no detectable background from ATP.

Programmed cell death (PCD) is an irreversible, genetically-controlled form of cell suicide in which an endogenous biochemical pathway leads to morphological changes and ultimately, cellular demise. PCD is accompanied by de-novo protein synthesis of a family of proteases-"caspases" that are often used as a diagnostic marker of PCD. Although phytoplankton do not contain true caspases, caspase-like activity (hypothetical proteins with analogous activity) has been traditionally used as a diagnostic marker of PCD in marine phytoplankton. Increased caspase-like proteolytic activity was demonstrated when synthetic fluorogenic activity substrates specific for caspases (with an Asp at the P1 position) were applied upon PCD induction. Metacaspases, cysteine proteases, share structural properties with those of caspases, yet they are highly specific for Arg and Lys cleavage site at the P1 position implying that caspase specific substrates are not indicative of metacaspase catalytic activity. This method specifically tests direct metacaspase activity in phytoplankton by the cleavage of the fluorogenic metacaspase substrate Ac-VRPR-AMC. Metacaspase activity was tested by the addition of a metacaspase specific peptide that is conjugated to the fluorescent reporter molecule. The cleavage of the peptide by the metacaspase releases the fluorochrome that, when excited by light, emits fluorescence. The level of metacaspase enzymatic activity in the cell lysate is directly proportional to the fluorescence signal detected. The use of specific standards in this test enables the quantification of the fluorescence results. This assay directly allows monitoring the metacaspase cleavage products and thereby tracing evidence for programmed cell death.