Neuroscience

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.

Mitochondrial dysfunction is associated with perturbations in the cellular oxidative status, changes in energy production and metabolic rate, and the onset of pathological processes. Classic methods of assessing mitochondrial dysfunction rely on indirect measures, such as evaluating mitochondrial DNA copy numbers, or direct but more costly and skilled techniques, such as electron microscopy. The protocol presented here was recently implemented to evaluate mitochondrial dysfunction in response to insecticide exposure in Drosophila melanogaster larvae, and it relies on the use of a previously established MitoTimer mutant strain. MitoTimer is a genetically engineered mitochondrial protein that shows green fluorescence when newly synthetized, irreversibly turning into red as mitochondria age. The protocol described here allows for the easy and direct assessment of shifts in mitochondrial turnover, with tissue-specific accuracy. This protocol can be adapted to assess changes in mitochondrial turnover in response to drugs, rearing conditions, and/or mutations in larva, pupa, or adult fruit flies.

In this protocol, we describe the analysis of protein stability over time, using synthesis shutoff. As an example, we express HA-tagged yeast mitofusin Fzo1 in Saccharomyces cerevisiae and inhibit translation via cycloheximide (CHX). Proteasomal inhibition with MG132 is performed, as an optional step, before the addition of CHX. Proteins are extracted via trichloroacetic acid (TCA) precipitation and subsequently separated via SDS-PAGE. Immunoblotting and antibody-decoration are performed to detect Fzo1 using HA-specific antibodies. We have adapted the method of blocking protein translation with cycloheximide to analyze the stability of high molecular weight proteins, including post-translational modifications and their impact on protein turnover.

The efficient ATP production in mitochondria relies on the highly specific organization of its double membrane. Notably, the inner mitochondrial membrane (IMM) displays a massive surface extension through its folding into cristae, along which concentrate respiratory complexes and oligomers of the ATP synthase. Evidence has accumulated to highlight the importance of a specific phospholipid composition of the IMM to support mitochondrial oxidative phosphorylation. Contribution of specific phospholipids to mitochondrial ATP production is classically studied by modulating the activity of enzymes involved in their synthesis, but the interconnection of phospholipid synthesis pathways often impedes the determination of the precise role of each phospholipid. Here, we describe a protocol to specifically enrich mitochondrial membranes with cardiolipin or phosphatidylcholine, as well as a fluorescence-based method to quantify phospholipid enrichment. This method, based on the fusion of lipid vesicles with isolated mitochondria, may further allow a precise evaluation of phospholipid contribution to mitochondrial functions.

Mitochondrial reactive oxygen species (mROS) are naturally produced signalling molecules extremely relevant for understanding both health- and disease-associated biological processes. The study of mROS in the brain is currently underway to decipher their physiopathological roles and contributions in neurological diseases. Recent advances in this field have highlighted the importance of studying mROS signalling and redox biology at the cellular level. Neurons are especially sensitive to the harmful effects of excess mROS while astrocytic mROS have been shown to play a relevant physiological role in cerebral homeostasis and behaviour. However, given the complexity of the brain, investigating mROS formation in a specific cell-type in adult animals is methodologically challenging. Here we propose an approach to specifically assess mROS abundance in astrocytes that combines i) a targeting strategy based on the use of adeno-associated virus (AAV) vectors expressing the green fluorescent protein (GFP) under an astrocyte (glial fibrillary acidic protein or GFAP) promoter, along with, ii) a robust and widely extended protocol for the measurement of mROS by flow cytometry using commercial probes. The significance of this work is that it allows the selective study of astrocytic mROS abundance by means of easily accessible technology.

In recent years, fluorescent dyes have been frequently used for monitoring mitochondrial membrane potential to evaluate mitochondrial viability and function. However, the reproducibility of the results across laboratories strongly depends upon following well validated and reliable protocols along with the appropriate controls. Herein, we provide a practical user guide for monitoring mitochondrial membrane potential in whole cells using a fluorescent cationic probe. The data analysis of mitochondrial membrane potential we provide is one associated with the impact of xenobiotics such as tobacco smoking on blood-brain barrier endothelial cells including both mouse primary (mBMEC) and a mouse-based endothelial cell line (bEnd3) in a side by side comparison.

Mitochondria form dynamic cytoplasmic networks which undergo morphological changes in order to adapt to cellular stresses and signals. These changes can include alterations in size and number within a given cell. Analysis of the whole network can be a useful metric to assess overall mitochondrial health, particularly in neurons, which are highly sensitive to mitochondrial dysfunction. Here we describe a method which combines immunofluorescence and computerized image analysis to measure mitochondrial morphology (quantification of number, density, and area) in dopaminergic neurites of mice expressing mitochondrially-targeted eYFP.

This protocol describes the in vitro phosphorylation of ubiquitin and Parkin by the kinase PINK1 using recombinant proteins. Both substrates, ubiquitin and Parkin, are phosphorylated at the conserved serine 65 residue (pS65-ubiquitin and pS65-Parkin). The protocol also includes the use of monomeric and K48- and K63-linked poly-ubiquitin chains as alternative substrates. Although there are commercially available antibodies, we have not tested their performance in this assay since, but used validated antibodies from our laboratory. An alternative antibody-independent method, the use of phos-tag gels to detect pS65-ubiquitin and pS65-Parkin, is described in addition.

In addition to methods aimed at the study of mitochondrial function in-situ, a full understanding of mitochondrial function requires their purification from cells or tissues under specific physiological or pathological conditions. This protocol illustrates a sequential procedure to obtain functional mitochondria with high yield from mice brain tissue. Mitochondria obtained with this method can be used to assess different mitochondrial parameters, including oxygen consumption, membrane potential and calcium retention capacity.

Mitochondria are eukaryotic organelles that play a crucial role in several cellular processes, including energy production, β-oxidation of fatty acids and regulation of calcium homeostasis. In the last 20 years there has been a hightened interest in the study of mitochondria following the discoveries that mitochondria are central to the process of programmed cell death and that mitochondrial dysfunctions are implicated in numerous diseases including a wide range of neurological disorders such as Parkinson’s disease, Alzheimer’s disease, Huntington’s disease and amyotrophic lateral sclerosis. In order to identify and study changes in mitochondrial function related to specific neurological conditions the mitochondria are often isolated from the compartment of the central nervous system most affected during disease. Here, we describe a protocol for the isolation of mitochondria from mouse spinal cord, a compartment of the central nervous system that is significantly affected in neuromuscular diseases such as amyotrophic lateral sclerosis. This method relies on differential centrifugation to separate the mitochondria from the other subcellular compartments.