Biochemistry

Enzyme immobilization offers a number of advantages that improve biocatalysis; however, finding a proper way to immobilize enzymes is often a challenging task. Implanting enzymes in metal–organic frameworks (MOFs) via co-crystallization, also known as biomineralization, provides enhanced reusability and stability with minimal perturbation and substrate selectivity to the enzyme. Currently, there are limited metal–ligand combinations with a proper protocol guiding the experimental procedures. We have recently explored 10 combinations that allow custom immobilization of enzymes according to enzyme stability and activity in different metals/ligands. Here, as a follow-up of that work, we present a protocol for how to carry out custom immobilization of enzymes using the available combinations of metal ions and ligands. Detailed procedures to prepare metal ions, ligands, and enzymes for their co-crystallization, together with characterization and assessment, are discussed. Precautions for each experimental step and result analysis are highlighted as well. This protocol is important for enzyme immobilization in various research and industrial fields.

Key features

• A wide selection of metal ions and ligands allows for the immobilization of enzymes in metal–organic frameworks (MOFs) via co-crystallization.

• Step-by-step enzyme immobilization procedure via co-crystallization of metal ions, organic linkers, and enzymes.

• Practical considerations and experimental conditions to synthesize the enzyme@MOF biocomposites are discussed.

• The demonstrated method can be generalized to immobilize other enzymes and find other metal ion/ligand combinations to form MOFs in water and host enzymes.

Graphical overview

Dolichyl phosphates (DolP) are ubiquitous lipids that are present in almost all eukaryotic membranes. They play a key role in several protein glycosylation pathways and the formation of glycosylphosphatidylinositol anchors. These lipids constitute only ~0.1% of total phospholipids, and their analysis by reverse phase (RP) liquid chromatography–high-resolution mass spectrometry (LC–HRMS) is challenging due to their high lipophilicity (log P > 20), poor ionization efficiency, and relatively low abundance. To overcome these challenges, we have introduced a new approach for DolP analysis by combining trimethylsilyldiazomethane (TMSD)-based phosphate methylation and HRMS analysis. The analytical method was validated for its reproducibility, sensitivity, and accuracy. The established workflow was successfully applied for the simultaneous characterization and quantification of DolP species with different isoprene units in lipid extracts of HeLa and Saccharomyces cerevisiae cells.

This protocol describes a method for detecting and quantifying calcium ions in the endoplasmic reticulum (ER) and cytoplasm of cultured cells using fluorescent reporter proteins and ImageJ software. Genetically engineered fluorescent reporter proteins, such as G-CEPIA1er and GCaMP6f, localize to intracellular regions of interest (i.e., ER and cytoplasm) and emit green fluorescence upon binding to calcium ions. In this way, the fluorescence brightness of cells transfected with expression vectors for these reporters reflects the calcium ion concentration in each intracellular region. Here, we describe procedures for observing cultured cells expressing these fluorescent reporters under a fluorescence microscope, analyzing the obtained image using the free image analysis software ImageJ (https://imagej.net/ij/index.html), and determining the average fluorescence brightness of multiple cells present in the image. The current method allows us to quickly and easily quantify calcium ions on an image containing multiple cells and to determine whether there are relative differences in intracellular calcium ion concentration among experiments with different conditions.

Key features

• Detection and quantification of calcium ions in the ER and cytoplasm using fluorescent reporter proteins

• Quick and easy verification of measurement results using ImageJ

• Simultaneous comparison between various experimental conditions (drug treatment, mutants, etc.)

Ants use cuticular hydrocarbon (CHC) as a semiochemical for recognizing their nestmates. For socially parasitic ants, deceiving the CHC is an important survival strategy. Profiling and quantifying CHC is a potent approach to understanding such nestmate discrimination behavior. Thus, a highly efficient, stable, and reproducible extraction method for CHC is essential for this purpose. This paper describes a method for socially parasitic ants to disguise the host species’ CHC profile under laboratory conditions, as well as the extraction and measurement of CHC from ants (from a previous study). First, the artificial isotopic substance is applied to the host worker; then, the socially parasitic ant disguises the host-like CHC profile against the above host worker. Next, the CHC is extracted and fractionated from a socially parasitic ant using hexane and silica gel. After concentrating the fractionated product, this product is then used for measurement by gas chromatographymass spectrometry (GC-MS). The CHC extraction protocol described in this paper may be used for various ant species.

Phosphorus is an essential nutrient for plants. Green algae usually store excess P as polyphosphate (polyP) in the vacuoles. PolyP, a linear chain of three to hundreds of phosphate residues linked by phosphoanhydride bonds, is important for cell growth. Based on the previous method of polyP purification with silica gel columns (Werner et al., 2005; Canadell et al., 2016) in yeast cells, we developed a protocol to purify and determine the total P and polyP in Chlamydomonas reinhardtii by a quick, simplified, and quantitative method. We use hydrochloric acid or nitric acid to digest polyP or total P in dried cells and analyze P content using the malachite green colorimetric method. This method may be applied to other microalgae.

ATPase assays are a common tool for the characterization of purified ATPases. Here, we describe a radioactive [γ-³²P]-ATP-based approach, utilizing complex formation with molybdate for phase separation of the free phosphate from non-hydrolyzed, intact ATP. The high sensitivity of this assay, compared to common assays such as the Malachite green or NADH-coupled assay, enables the examination of proteins with low ATPase activity or low purification yields. This assay can be used on purified proteins for several applications including the identification of substrates, determination of the effect of mutations on ATPase activity, and testing specific ATPase inhibitors. Furthermore, the protocol outlined here can be adapted to measure the activity of reconstituted ATPases.

Graphical overview

Paraquat is a cost-effective herbicide, widely used in many countries, that can induce severe oxidative stress in photosynthetic tissues. Studying plant herbicide resistance or antioxidant stress mechanisms requires determining the cellular paraquat level when plants are treated by paraquat. The traditional isotopic labeling method has the potential risk to cause problems to both human health and the environment. For radioisotope manipulation, special operation spaces and strict environmental inspection are also required. In addition, the radiolabeled paraquat is increasingly hard to buy due to the extended production cycle. Here, we describe a nonradioactive method to determine the paraquat level in a small number of Arabidopsis tissues or protoplasts, using a high resolution ultra-high-performance liquid chromatography (UHPLC)-mass spectrometry (MS)/MS method. This method is highly selective and sensitive, and more environmentally compatible and technically feasible than the isotope detection method.

Ethylene is an important plant hormone that is involved in the regulation of numerous processes in plant development. It also acts as a signaling molecule in response to biotic and abiotic stress conditions. Most studies have investigated ethylene evolution of harvested fruit or small herbaceous plants under controlled conditions, but only a few explored ethylene release in other plant tissues, such as leaves and buds, particularly those of subtropical crops. However, in light of increasing environmental challenges in agriculture (such as temperature extremes, droughts, floods, and high solar radiation), studies on these challenges and on potential chemical treatments for mitigating their effects on plant physiology have become more and more important. Thus, adequate techniques for the sampling and analysis of tree crops are needed to ensure accurate ethylene quantification. As part of a study on ethephon as a mitigating agent to improve litchi flowering under warm winter conditions, a protocol was developed for ethylene quantification in leaf and bud tissue of litchi following ethephon application, taking into account that these plant organs release lower ethylene concentrations than fruit. At sampling, leaves and buds were placed in glass vials of appropriate sizes for the respective plant tissue volumes and allowed to equilibrate for 10 min to release possible wound ethylene before incubating the samples for 3 h at ambient temperature. Thereafter, ethylene samples were aspirated from the vials and analyzed using a gas chromatograph with flame ionization detection, the TG-BOND Q+ column for separation of ethylene, and helium as the carrier gas. Quantification was achieved based on a standard curve derived from an external standard gas calibration with certified ethylene gas. This protocol will also be appropriate for other tree crops with similar plant materials as study foci. It will enable researchers to accurately determine ethylene production in various studies investigating the role of ethylene in general plant physiology or stress-induced plant responses following a range of treatment conditions.

Macrophages are at the center of innate immunity and are the main target cells of the intracellular pathogen Salmonella enterica serovar Typhi. The production of reactive oxygen and nitrogen species (ROS/RNS) is the host’s early response to invading microbes, as oxidative stress is highly toxic for bacteria. Adequate ROS/RNS production in infected macrophages is critical for the clearance of intracellular pathogens; this is achieved by several enzymes, including inducible NADPH phagocyte oxidase (NOX) and nitric oxide synthase (iNOS), respectively. The pro-inflammatory cytokine interferon gamma (IFNγ), primarily produced by activated natural killer cells and T-helper cells type 1, is a potent inducer of iNOS. Therefore, it is crucial for infection control through oxidative microbicidal activity.

To characterize the early oxidative stress response via ROS formation, which is critical for the reduction of Salmonella proliferation within macrophages, we established an in vitro model of murine macrophages infected with Salmonella enterica serovar Typhimurium (S.tm). This serovar induces a systemic infection in mice that is frequently used as a model for typhoid fever, which, in human subjects, is caused by Salmonella Typhi.

We generated bone marrow–derived macrophages (BMDM) from C57BL/6N wildtype mice using macrophage colony-stimulating factor (M-CSF) stimulation for six days. Thereafter, we infected BMDM with S.tm for one hour. Shortly before infection, cells were stained with CellROX^TM Deep Red reagent. In its reduced form, CellROX^TM is non-fluorescent. As a result of oxidation by ROS, this reagent exhibits strong fluorescence and persists within the cells. Subsequently, changes as a result of the oxidative stress response can be measured with a TECAN Spark microplate reader over time.

We designed this protocol to measure oxidative stress in macrophages through the course of an infection with an intracellular bacterium. The protocol has several advantages over established techniques. First, it allows to continuously monitor and quantify ROS production in living cells from the very start of the infection to the final clearance of the intracellular pathogen. Second, this protocol enables efficient ROS detection without stressing the cells by detaching or staining procedures.

Graphical abstract

Ion homeostasis is a fundamental regulator of cellular processes and depends upon lipid membranes, which function as ion permeability barriers. Ionophores facilitate ion transport across cell membranes and offer a way to manipulate cellular ion composition. Here, we describe a calcein quenching assay based on large unilamellar vesicles that we used to evaluate divalent cation transport of the ionophore 4-Br-A23187. This assay can be used to study metal transport by ionophores and membrane proteins, under well-defined conditions.

Graphical abstract: