Cell Biology

The genome of the dengue virus codes for a single polypeptide that yields three structural and seven non-structural (NS) proteins upon post-translational modifications. Among them, NS protein-3 (NS3) possesses protease activity, involved in the processing of the self-polypeptide and in the cleavage of host proteins. Identification and analysis of such host proteins as substrates of this protease facilitate the development of specific drugs. In vitro cleavage analysis has been applied, which requires homogeneously purified components. However, the expression and purification of both S3 and erythroid differentiation regulatory factor 1 (EDRF1) are difficult and unsuccessful on many occasions. EDRF1 was identified as an interacting protein of dengue virus protease (NS3). The amino acid sequence analysis indicates the presence of NS3 cleavage sites in this protein. As EDRF1 is a high-molecular-weight (~138 kDa) protein, it is difficult to express and purify the complete protein. In this protocol, we clone the domain of the EDRF1 protein (C-terminal end) containing the cleavage site and the NS3 into two different eukaryotic expression vectors containing different tags. These recombinant vectors are co-transfected into mammalian cells. The cell lysate is subjected to SDS-PAGE followed by western blotting with anti-tag antibodies. Data suggest the disappearance of the EDRF1 band in the lane co-transfected along with NS3 protease but present in the lane transfected with only EDRF1, suggesting EDRF1 as a novel substrate of NS3 protease. This protocol is useful in identifying the substrates of viral-encoded proteases using ex vivo conditions. Further, this protocol can be used to screen anti-protease molecules.

Key features

• This protocol requires the cloning of protease and substrate into two different eukaryotic expression vectors with different tags.

• Involves the transfection and co-transfection of both the above recombinant vectors individually and together.

• Involves western blotting of the same PVDF membrane containing total proteins of the cell lysate with two different antibodies.

• Does not require purified proteins for the analysis of cleavage of any suspected substrate by the protease.

Graphical overview

Secreted reporters have been demonstrated to be simple and useful tools for analyzing transcriptional regulation in mammalian cells. The distinctive feature of these assays is the ability to detect reporter gene expression in the culture supernatant without affecting the cell physiology or leading to cell lysis, which allows repeated experimentation and sampling of the culture medium using the same cell cultures. Secreted embryonic alkaline phosphatase (SEAP) is one of the most widely used reporter, which can be easily detected using colorimetry following incubation with a substrate, such as p-nitrophenol phosphate. In this report, we present detailed procedures for detection and quantification of the SEAP reporter. We believe that this step-by-step protocol can be easily used by researchers to monitor and measure molecular genetic events in a variety of mammalian cells due to its simplicity and ease of handling.

Graphical abstract

Schematic overview of the workflow described in this protocol

As one of the main energy metabolism organs, kidney has been proved to have high energy requirements and are more inclined to fatty acid metabolism as the main energy source. Long-chain acyl-CoA dehydrogenases (LCAD) and beta-hydroxyacyl-CoA dehydrogenase (beta-HAD), key enzymes involved in fatty acid oxidation, has been identified as the substrate of acetyltransferase GCN5L1 and deacetylase Sirt3. Acetylation levels of LCAD and beta-HAD regulate its enzymes activity and thus affect fatty acid oxidation rate. Moreover, immunoprecipitation is a key assay for the detection of LCAD and beta-HAD acetylation levels. Here we describe a protocol of immunoprecipitation of acetyl-lysine and western blotting of LCAD and beta-HAD in palmitic acid treated HK-2 cells (human renal tubular epithelial cells). The scheme provides the readers with clear steps so that this method could be applied to detect the acetylation level of various proteins.

In order to acquire fertilizing potential, mammalian sperm must undergo a process known as capacitation, which relies on the early activation of Protein Kinase A (PKA). Frequently, PKA activity is assessed in whole-cell experiments by analyzing the phosphorylation status of its substrates in a western-blot. This technique faces two main disadvantages: it is not a direct measure of the kinase activity and it is a time-consuming approach. However, since PKA can be readily obtained from sperm extracts, in vitro assays such as the “radioactive assay” can be performed using the native enzyme. Unlike western-blot, the radioactive assay is a straightforward technique to evaluate PKA activity by quantification of incorporated ³²P into a peptidic substrate. This approach easily allows the analysis of different agonists or antagonists of PKA. Since mouse sperm is a rich source of soluble PKA, this assay allows a simple fractionation that renders PKA usable both for in vitro testing of drugs on PKA activity and for following changes of PKA activity during the onset of capacitation.

Transglutaminase (TG2) catalyzes protein crosslinking between glutamyl and lysyl residues. Catalytic activity occurs via a transamidation mechanism resulting in the formation of isopeptide bonds. Since TG2-mediated transamidation is of mechanistic importance for a number of biological processes, assays that enable rapid and efficient identification and characterization of candidate substrates are an important first-step to uncovering the function of crosslinked proteins. Herein we describe an optimized and flexible protocol for in vitro TG2 crosslink reactions and substrate incorporation assays. We have previously employed these techniques in the identification of the protein high mobility group box 1 (HMGB1) as a TG2 substrate. However, the protocol can be adapted for identification of any candidate transamidation substrate.

Serine palmitoyltranferase (SPT) is a pyridoxal 5′ phosphate (PLP)-dependent enzyme that catalyzes the first and rate-limiting step of de novo synthesis of sphingolipids. SPT activity is homeostatically regulated in response to increased levels of sphingolipids. This homeostatic regulation of SPT is mediated through small ER membrane proteins termed the ORMDLs. Here we describe a procedure to assay ORMDL dependent lipid inhibition of SPT activity. The assay of SPT activity using radiolabeled L-serine was developed from the procedure established by the Hornemann laboratory. The activity of SPT can also be measured using deuterated L-serine but it requires mass spectrometry, which consumes money, time and instrumentation. The ORMDL dependent lipid inhibition of SPT activity can be studied in both cells and in a cell free system. This assay procedure is applicable to any type of mammalian cell. Here we provide the detailed protocol to measure SPT activity in the presence of either short chain (C8-ceramide) or long chain ceramide (C24-ceramide). One of the greatest advantages of this protocol is the ability to test insoluble long chain ceramides. We accomplished this by generating long chain ceramide through endogenous ceramide synthase by providing exogenous sphingosine and 24:1 acyl CoA in HeLa cell membranes. This SPT assay procedure is simple and easy to perform and does not require sophisticated instruments.

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.

Kinases function as regulators of many cellular processes such as cell migration. These enzymes typically phosphorylate target motif sequences. Mass spec or phospho-specific antibody detection can be used to determine whether a kinase can phosphorylate proteins of interest, however, mass spec can be expensive and phospho-antibodies for the protein of interest may not exist. In this protocol, we will describe an in vitro kinase assay to provide a preliminary readout on whether a protein of interest may be phosphorylated by PKA. Our protein of interest is purified after expression in bacteria and treated with recombinant PKA from bovine heart. Protein is then extracted and a western blot is performed using a phospho-specific antibody for PKA’s target motif. This will allow us to quickly determine if it is possible for PKA to phosphorylate our protein of interest.

Fibrinolysis is an integral part of the matrix remodeling process that contributes to tissue repair. Fibrin clots are broken down during fibrinolysis in a controlled process. Fibrin degradation products (FDPs) have also been shown to have a role in the regulation of cell growth and are implicated in various vascular diseases. This protocol was designed to quantitatively measure the extent of fibrin breakdown and how this can be adapted as a tool to further investigate the pathway involved in fibrinolysis or fibrin degradation products. Until now, we haven’t found an alternative method to analysis fibrinolysis.

Glucocerebrosidase (GCase) is an important enzyme for the metabolism of glycolipids. GCase enzyme deficiency is implicated in human disease and the efficient measurement of GCase activity is important for evaluating the efficacy of therapeutics targeting this enzyme. Existing approaches to measure GCase activity include whole blood mass spectrometry-based assays, where an internal standard is used to measure the accumulation of ceramide following metabolism of the synthetic substrate C12-glucocerebroside, and the utilisation of fluorescent probes that bind active GCase and/or release fluorescent metabolites upon cleavage by GCase. Here, we describe the application of a fluorescence-activated cell sorter-based assay to efficiently quantitate GCase enzyme activity in the monocyte population of human peripheral blood mononuclear cells. The cell-permeable GCase substrate 5-(Pentafluorobenzoylamino) Fluorescein Di-beta-D-Glucopyranoside (PFB-FDGlu) provides a means to measure GCase activity, whereby enzymatic cleavage yields the green-fluorescent PFB-F dye, detectable in the FL-1 channel of a flow cytometer. An inhibitor of lysosomal GCase activity, conduritol B-epoxide, is employed to ensure specificity. This protocol provides an advantageous approach for measuring GCase activity in living individual cells.