细胞生物学

The placenta is a metabolically active organ whose mitochondrial activity is tightly linked to fetal growth, oxygenation, and nutrient transport, mediating fetal susceptibility to environmental exposures. Accordingly, aberrant mitochondrial function has been implicated in the progression of placental dysfunction. However, existing respirometry platforms require primarily fresh or cryopreserved placental tissue and offer limited throughput, rendering these platforms impractical in the context of large-scale placental dissections. Here, we describe and validate a Seahorse XF approach for measuring mitochondrial respiration in previously frozen placentae, enabling the functional interrogation of placental mitochondria in prenatal studies. Our protocol fundamentally relies on the restoration of matrix substrates that are depleted due to increased mitochondrial membrane permeability following freeze-thaw cycles. We provide a strategy to assess complex I and II-associated respiration adapted for the Seahorse XFe24 Analyzer and further demonstrate comparable oxygen consumption readouts between fresh and frozen placentae. We further demonstrate distinct differences in the magnitude of oxygen consumption between fresh and frozen placentae in the absence of exogenous NADH. Taken together, we present a simplified and convenient protocol for the assessment of respiratory enzyme complex-associated respiration from archived placental tissue.

Protein–protein interactions facilitate cellular functions through the creation of networks and multi-protein complexes. Mapping the interactions within and between protein networks and elucidating the composition of protein complexes provides critical insight into biological processes. Interactions among soluble cytoplasmic proteins have been extensively investigated through the application of immunoaffinity capture as well as conventional nuclear two-hybrid testing. The integrated membrane yeast two-hybrid provides a method to investigate protein–protein interactions between integral membrane proteins in their native membrane environment. This procedure makes use of the ability of the amino-terminal fragment of ubiquitin (Nub) and the carboxyl-terminal fragment of ubiquitin (Cub) to refold reconstituting functional ubiquitin, which can be recognized by a ubiquitin peptidase. Appending a fusion protein composed of Cub fused to LexA and VP16 (CLV) to a candidate "bait" protein and Nub to candidate "prey" proteins allows a test of their interaction. If the two proteins interact closely, the CLV fragment is cleaved and enters the nucleus to activate the expression of reporter genes, signaling the interaction. When the bait and prey proteins are tagged with CLV and NubG, respectively, at their genomic loci, they are only copies of the bait and prey in the cell and are expressed under the regulation of their native promoters. This avoids overexpression artifacts that can occur if the tagged proteins are expressed from plasmids while the untagged chromosomally encoded copies of the bait and prey continue to be expressed.

Oxidative protein damage is important in various biological processes and age-related diseases. Protein carbonylation is the predominant and most frequently studied form of protein oxidation. It is most frequently detected following its derivatization with 2,4-dinitrophenylhydrazine (DNPH) hapten, followed by its detection with an anti-DNP antibody. However, when used to detect protein carbonylation by western blotting, this method suffers from diminished sensitivity, distortion of protein migration patterns, and unsatisfactory representation of low-abundance proteins. This is due to the poor solubility of DNPH in typical buffer solutions, the acidic protein precipitation due to the use of strong acid for its dissolution, the instability in solution, and the distorted protein migration patterns introduced by an additional salt content generated by the required pH adjustment prior to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). To address the DNPH method limitations, a new Oxime blot technique was developed. This method is based on forming the stable oxime bonds between the protein carbonyl groups and biotin-aminooxy probe in the presence of a p-phenylenediamine (pPDA) catalyst at neutral pH conditions. The derivatization reaction reaches a plateau within 3 h. It ensures efficient and complete derivatization of carbonylated proteins, which are separated by SDS-PAGE without additional manipulation and detected with avidin-HRP and enhanced chemiluminescence (ECL) in western blotting. The Oxime blot protocol allows researchers to reliably and sensitively detect carbonylated proteins and provides a valuable tool for studying oxidative stress in diverse biological settings.

Mitochondria are dynamic organelles with essential roles in energetics and metabolism. Several metabolites are common to both the cytosolic and mitochondrial fractions of the cell. The compartmentalization of metabolites within the mitochondria allows specialized uses for mitochondrial metabolism. Inorganic phosphate (Pi) is one such critical metabolite required for ATP synthesis, via glycolysis and mitochondrial oxidative phosphorylation. Estimating total cellular Pi levels cannot distinguish the distribution of Pi pools across different cellular compartments, such as the cytosol and mitochondria, and therefore separate the contributions made toward glycolysis or other cytosolic metabolic processes vs. mitochondrial outputs. Quantifying Pi pools in mitochondria can therefore be very useful toward understanding mitochondrial metabolism and phosphate homeostasis. Here, we describe a protocol for the fairly rapid, efficient isolation of mitochondria from Saccharomyces cerevisiae by immunoprecipitation for quantitative estimation of mitochondrial and cytosolic Pi pools. This method utilizes magnetic beads to capture FLAG-tagged mitochondria (Tom20-FLAG) from homogenized cell lysates. This method provides a valuable tool to investigate changes in mitochondrial phosphate dynamics. Additionally, this protocol can be coupled with LC–MS approaches to quantitatively estimate mitochondrial metabolites and proteins and can be similarly used to assess other metabolite pools that are partitioned between the cytosol and mitochondria.

The Seahorse 96 XF Analyzer (Agilent Technologies, Santa Clara, CA, USA) has been an effective tool in non-invasively measuring mitochondrial function for the past decade. It is a high-throughput respirometer that is considered the “gold standard” for quantifying mitochondrial function and bioenergetics in cells. Peripheral blood mononuclear cells (PBMCs) play a selective role in immune system responses and are key components of human immunity. Recent studies have suggested that these cell populations provide an overview of systemic changes within the body and therefore provide a source of sensitive biomarkers. Assessing mitochondrial function in PBMCs has been shown to provide an indication of metabolic stress associated with diseases such as diabetes and neurodegenerative conditions such as Alzheimer’s disease. In this protocol, we use two adhesive compounds, Poly-D-Lysine (PDL) and Poly-L-Lysine (PLL), at 50 μg/mL each per well, to immobilize PBMCs to a specialized Seahorse microplate to perform mitochondrial stress assay using the Seahorse Analyzer. We compared six cell densities of PBMCs to identify the optimal cell density for use in Seahorse Mito Stress analysis. This protocol includes the immobilization of freshly isolated PBM cells into a Seahorse microplate, hydration and calibration of the sensor cartridge, cell seeding, running the Seahorse Analyzer for the Mito Stress test, and simple data analysis to compare the effectiveness of PLL and PDL as the coating agent for PBMCs. The data analysis indicates that there is no statistical difference between PLL and PDL.

With the advancement of liquid chromatography–mass spectrometry (LC–MS/MS), the quantification of glycerophospholipid (PL) molecules has become more accessible, leading to the discovery of numerous enzymes responsible for determining the acyl groups attached to these molecules. Metabolic tracer experiments using radioisotopes and stable isotopes are powerful tools for defining the function of metabolic enzymes and metabolic flux. We have established an ex vivo muscle experimental system using stable isotope–labeled fatty acids to evaluate fatty acid incorporation into PL molecules. Here, we describe a method to incorporate fatty acids with stable isotope labels into excised skeletal muscle and detect the PL molecules containing labeled acyl chains by LC–MS/MS.

Analysis of mitochondrial function has broad applicability in many research specialties. Neurodegenerative disorders such as chemotherapy-induced peripheral neuropathy (CIPN) often exhibit damaged mitochondria or reduced mitochondrial respiratory capacity. Isolation of intact mitochondria for protein analysis or respiration measurements has been previously reported in numerous model organisms. Here, we describe an adaptation of previous protocols to isolate intact functional mitochondria from Drosophila melanogaster for use in a model of CIPN. Whole Drosophila are ground in isolation buffer, and mitochondria are purified using differential centrifugation through a sucrose and mannitol solution. The intact mitochondria are plated as a monolayer for measurements of mitochondrial oxygen consumption rates and response to inhibitor compounds on an Agilent Seahorse analyzer. This experimental protocol is quick and yields a purified population of intact mitochondria that may be used for functional assays for several hours after isolation. The isolated mitochondria may be used for respiration measurements, which reflect their health, and stored for protein or genetic analysis. Mitochondrial populations from multiple strains or treatment groups can be easily compared simultaneously. The rapid biochemical assessment of mitochondria, in combination with the utility of Drosophila as an in vivo genetic model system, offers great potential for researchers to probe the impact of genetics and pharmacologic interventions on mitochondrial respiratory capacity.

Two aconitase isoforms are present in mammalian cells: the mitochondrial aconitase (ACO2) that catalyzes the reversible isomerization of citrate to isocitrate in the citric acid cycle, and the bifunctional cytosolic enzyme (ACO1), which also plays a role as an RNA-binding protein in the regulation of intracellular iron metabolism. Aconitase activities in the different subcellular compartments can be selectively inactivated by different genetic defects, iron depletion, and oxidative or nitrative stress. Aconitase contains a [4Fe-4S]²⁺ cluster that is essential for substrate coordination and catalysis. Many Fe-S clusters are sensitive to oxidative stress, nitrative stress, and reduced iron availability, which forms the basis of redox- and iron-mediated regulation of intermediary metabolism via aconitase and other Fe-S cluster-containing metabolic enzymes, such as succinate dehydrogenase. As such, ACO1 and ACO2 activities can serve as compartment-specific surrogate markers of oxygen levels, reactive oxygen species (ROS), reactive nitrogen species (RNS), iron bioavailability, and the status of intermediary and iron metabolism. Here, we provide a protocol describing a non-denaturing polyacrylamide gel electrophoresis (PAGE)-based procedure that has been successfully used to monitor ACO1 and ACO2 aconitase activities simultaneously in human and mouse cells and tissues.

Macrophages are at the center of innate immunity and iron metabolism. In the case of an infection, macrophages adapt their cellular iron metabolism to deprive iron from invading bacteria to combat intracellular bacterial proliferation. A concise evaluation of the cellular iron content upon an infection with bacterial pathogens and diverse cellular stimuli is necessary to identify underlying mechanisms concerning iron homeostasis in macrophages. For the characterization of cellular iron levels during infection, we established an in vitro infection model where the murine macrophage cell line J774A.1 is infected with Salmonella enterica serovar Typhimurium (S.tm), the mouse counterpart to S. enterica serovar Typhi, under normal and iron-overload conditions using ferric chloride (FeCl₃) treatment. To evaluate the effect of infection and iron stimulation on cellular iron levels, the macrophages are stained with FerroOrange. This fluorescent probe specifically detects Fe²⁺ ions and its fluorescence can be quantified photometrically in a plate reader. Importantly, FerroOrange fluorescence does not increase with chelated iron or other bivalent metal ions. In this protocol, we present a simple and reliable method to quantify cellular Fe²⁺ levels in cultured macrophages by applying a highly specific fluorescence probe (FerroOrange) in a TECAN Spark microplate reader. Compared to already established techniques, our protocol allows assessing cellular iron levels in innate immune cells without the use of radioactive iron isotopes or extensive sample preparation, exposing the cells to stress.

Key features

• Easy quantification of Fe²⁺ in cultured macrophages with a fluorescent probe.

• Analysis of iron in living cells without the need for fixation.

• Performed on a plate reader capable of 540 nm excitation and 585 nm emission by trained employees for handling biosafety level 2 bacteria.

Graphical overview

The mitochondrial electron transport chain (ETC) is a multi-component pathway that mediates the transfer of electrons from metabolic reactions that occur in the mitochondrion to molecular oxygen (O₂). The ETC contributes to numerous cellular processes, including the generation of cellular ATP through oxidative phosphorylation, serving as an electron sink for metabolic pathways such as de novo pyrimidine biosynthesis and for maintaining mitochondrial membrane potential. Proper functioning of the mitochondrial ETC is necessary for the growth and survival of apicomplexan parasites including Plasmodium falciparum, a causative agent of malaria. The mitochondrial ETC of P. falciparum is an attractive target for antimalarial drugs, due to its essentiality and its differences from the mammalian ETC. To identify novel P. falciparum ETC inhibitors, we have established a real-time assay to assess ETC function, which we describe here. This approach measures the O₂ consumption rate (OCR) of permeabilized P. falciparum parasites using a Seahorse XFe96 flux analyzer and can be used to screen compound libraries for the identification of ETC inhibitors and, in part, to determine the targets of those inhibitors.

Key features

• With this protocol, the effects of candidate inhibitors on mitochondrial O₂ consumption in permeabilized asexual P. falciparum parasites can be tested in real time.

• Through the sequential injection of inhibitors and substrates into the assay, the molecular targets of candidate inhibitors in the ETC can, in part, be determined.

• The assay is applicable for both drug discovery approaches and enquiries into a fundamental aspect of parasite mitochondrial biology.

Graphical overview

Seahorse assay experimental workflow. Prior to the assay, coat the cell culture microplate with Cell-Tak to help adhere the parasites to the wells; hydrate the cartridge wells to ensure proper sensor functionality and design the assay template using the Agilent Seahorse Wave Desktop software (Analyze Seahorse data files, Seahorse Wave desktop software|Agilent). On the day of the assay, prepare the inhibitors/substrates that are to be injected into the ports. Then, separate 3 × 10⁸ trophozoite-stage parasites from the uninfected red blood cells (RBCs) and ring-stage parasites using a MACS^® magnetic column. Check the purity of the parasites with Giemsa-stained smears. Determine the concentration of infected RBCs in the sample using a hemocytometer and dilute to approximately 5 × 10⁷ parasites per milliliter. Treat infected RBCs with saponin to permeabilize the host cell membrane and seed approximately 5 × 10⁶ parasites (100 μL) per well in mitochondria assay solution (MAS) buffer. Supplement MAS buffer with digitonin to permeabilize the parasite plasma membrane. Load the ports with the prepared inhibitors/substrates and run the assay using a Seahorse XFe96 analyzer. Once the assay is completed, analyze the data using the Wave desktop software. Further data processing can be done using statistical analysis software.