Name: analyses of mitochondrial calcium influx in isolated mitochondria and cultured cells
Uploaded: 2018-04-27T13:30:00
Description: Watch this Scientific Journal Video about Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells at JoVE.com

This work was supported by grant funding from the American Heart Association (J.Q.K.).

All methods described in this protocol have been approved by the Institutional Animal Care and Use Committee of Emory University.
NOTE: The first part is the experimental procedure for measuring mitochondrial Ca2+ influx in isolated cardiac mitochondria using a plate reader.
1. Reagents and Solutions
<ol>
	<li>Make 500 mL of MS-EGTA buffer for mitochondrial isolation: 225 mM mannitol, 75 mM sucrose, 5 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic aci...

<ol>
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Here, we describe two different approaches to measure mitochondrial Ca2+ influx. The plate reader-based calcium green-5N method monitors extramitochondrial Ca2+ levels and is a Ca2+ uptake assay that is well suited for measurements in isolated mitochondria. While we have shown representative results from isolated murine cardiac mitochondria, this assay can be readily adapted for mitochondria isolated from tissues with high mitochondrial abundance including the liver, skeletal muscle, and ...

Mitochondria are critical sites of intracellular Ca2+ storage and signaling. Decades of research have demonstrated that mitochondria have the ability to import and sequester Ca2+ 1,2. Mitochondria, however, are not merely passive sites of Ca2+ storage. Ca2+ at the mitochondrial compartment performs fundamental signaling functions including regulation of metabolic output and activation of mitochondrial-mediated cell death pathways, which has been reviewed previously3. For metabolic regulation, Ca2+ enhances the activity of three matrix-localized dehydrogenases of the tricarboxylic acid cycle as well as respiratory chain complexes, to increase mitochondrial energy production4,5. With mitochondrial Ca2+ overload and dysregulated mitochondrial Ca2+ handling, Ca2+ triggers mitochondrial permeability transition pore (MPTP) opening, leading to mitochondrial inner membrane permeabilization, membrane potential loss, mitochondrial dysfunction, swelling, rupture and ultimately, cell death6,7,8,9. Thus, mitochondrial Ca2+ signaling directly impacts both cellular life and death pathways through metabolic control and MPTP-death axis.
In recent years, there has been rapidly expanding interest in the study of mitochondrial Ca2+ dynamics due in large part to the identification of the molecular constituents of the mitochondrial Ca2+ uniporter complex, a mitochondrial inner membrane transporter that is a primary mode of Ca2+ import into the mitochondrial matrix 10,11,12. Identification of these structural and regulatory subunits of the uniporter has brought forth the possibility of genetically targeting mitochondrial Ca2+ influx to modulate mitochondrial function and dysfunction and facilitated the study of the contribution of the uniporter complex and mitochondrial Ca2+ influx to disease13,14,15. Indeed, mitochondrial Ca2+ signaling has been implicated in the pathologies of a diverse array of diseases ranging from cardiac disease to neurodegeneration, and cancer16,17,18,19,20.
Given the fundamental importance of mitochondrial Ca2+ signaling in metabolism and cell death, and combined with the broad reach of biological systems that mitochondrial Ca2+ signaling impacts, methods to assess mitochondrial Ca2+ influx are of great interest. Not surprisingly, a variety of techniques and tools to measure mitochondrial Ca2+ have been developed. These include methods that utilize tools such as fluorescent Ca2+-sensitive dyes21,22 and genetically-encoded Ca2+ sensors targeted to the mitochondria, such as cameleon and aequorin23,24. The goal of this article is to highlight different methods and model systems in which mitochondrial Ca2+ uptake can be measured. We present two experimental methods to assess mitochondrial Ca2+ influx capacity. Using cardiac mitochondria as an example, we detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N that is ideally suited for isolated tissue mitochondria14. Using cultured NIH 3T3 cells, we also describe a confocal microscopy imaging-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM25.

<table>
	<tbody>
		<tr>
			<td>Olympus FV1000 Laser Scanning confocal microscope</td>
			<td>Olympus</td>
			<td>FV1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Synergy Neo2 Multimode microplate reader with injectors</td>
			<td>Biotek</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Homogenizer</td>
			<td>Kimble</td>
			<td>886000-0022</td>
			<td></td>
		</tr>
		<tr>
			<td>22 x 22 mm coverslips</td>
			<td>Corning</td>
			<td>2850-22</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well plate</td>
			<td>Corning</td>
			<td>3628</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well plate</td>
			<td>Corning</td>
			<td>3506</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Green-5N</td>
			<td>Invitrogen</td>
			<td>C3737</td>
			<td></td>
		</tr>
		<tr>
			<td>MitoTracker green FM</td>
			<td>Invitrogen</td>
			<td>M7514</td>
			<td></td>
		</tr>
		<tr>
			<td>Rhod-2, AM</td>
			<td>Invitrogen</td>
			<td>R1244</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Invitrogen</td>
			<td>D12345</td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic F-127</td>
			<td>Invitrogen</td>
			<td>P3000MP</td>
			<td></td>
		</tr>
		<tr>
			<td>D-Mannitol</td>
			<td>Sigma</td>
			<td>M9546</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>EMD Millipore</td>
			<td>8510</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Sigma</td>
			<td>E8145</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Fisher</td>
			<td>BP366-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic</td>
			<td>Sigma</td>
			<td>P0662</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride</td>
			<td>Sigma</td>
			<td>M2670</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium pyruvate</td>
			<td>Sigma</td>
			<td>P2256</td>
			<td></td>
		</tr>
		<tr>
			<td>L-malic acid</td>
			<td>Sigma</td>
			<td>M1125</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride</td>
			<td>Sigma</td>
			<td>C4901</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium acetate</td>
			<td>Fisher</td>
			<td>BP364-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine 5&#8242;-triphosphate magnesium salt</td>
			<td>Sigma</td>
			<td>A9187</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphocreatine disodium salt</td>
			<td>Sigma</td>
			<td>P7936</td>
			<td></td>
		</tr>
		<tr>
			<td>Saponin</td>
			<td>Sigma</td>
			<td>S7900</td>
			<td></td>
		</tr>
		<tr>
			<td>Ru360</td>
			<td>Calbiochem</td>
			<td>557440</td>
			<td></td>
		</tr>
	</tbody>
</table>

analyses of mitochondrial calcium influx in isolated mitochondria and cultured cells

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ability to accurately measure the influx and efflux of Ca2+ from mitochondria is important for determining the role of mitochondrial Ca2+ handling in these processes. In this report, we present two methods for the measurement of mitochondrial Ca2+ handling in both isolated mitochondria and cultured cells. We first detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N. The plate reader-based format circumvents the need for specialized equipment, and the calcium green-5N dye is ideally suited for measuring Ca2+ from isolated tissue mitochondria. For our application, we describe the measurement of mitochondrial Ca2+ uptake in mitochondria isolated from mouse heart tissue; however, this procedure can be applied to measure mitochondrial Ca2+ uptake in mitochondria isolated from other tissues such as liver, skeletal muscle, and brain. Secondly, we describe a confocal microscopy-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM and imaging using 2-dimensional laser-scanning microscopy. This permeabilization protocol eliminates cytosolic dye contamination, allowing for specific recording of changes in mitochondrial Ca2+. Moreover, laser-scanning microscopy allows for high frame rates to capture rapid changes in mitochondrial Ca2+ in response to various drugs or reagents applied in the external solution. This protocol can be applied to measure mitochondrial Ca2+ uptake in many cell types including primary cells such as cardiac myocytes and neurons, and immortalized cell lines.

Figure 1 shows mitochondrial Ca2+ uptake measurements in isolated cardiac mitochondria using the plate reader-based platform and the Ca2+ dye calcium green-5N. Under control conditions (Figure 1A), cardiac mitochondria were suspended in KCl buffer containing calcium green-5N and then challenged with sequential pulses of CaCl2 (5 &#956;L of a 0.6 mM CaCl2 solution) added at the 30 s, 150...

Watch this Scientific Journal Video about Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells at JoVE.com

Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells

Here, we present two protocols for the measurement of mitochondrial Ca2+ influx in isolated mitochondria and cultured cells. For isolated mitochondria, we detail a plate reader-based Ca2+ import assay using the Ca2+ sensitive dye calcium green-5N. For cultured cells, we describe a confocal microscopy method using the Ca2+ dye Rhod-2/AM.

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ...

The overall goal of this procedure is to measure mitochondrial calcium influx in isolated mitochondria and cultured cells. This method can help answer key questions in the calcium signaling field such as how mitochondria help to shape the cytosolic calcium landscape, and the role of mitochondrial calcium dynamics in disease conditions. The main advantage of this technique is that our protocols can be adapted to a variety of model systems to assess mitochondrial calcium influx capacity.I will be demonstrating the plate-reader-based mitochondrial calcium uptake procedure. And Dr.Joshua Maxwell, an instructor in the Department of Pediatrics will be demonstrating the confocal-microscope-based procedure to measure mitochondrial calcium influx. Start this experiment by thoroughly rinsing previously isolated heart in 25 milliliters of ice-cold one-X PBS making sure that the blood is completely squeezed out of the ventricles.In five milliliters of ice-cold one-X PBS, using a pair of sharp scissors, mince the heart tissue into small pieces. Carefully discard the PBS and transfer the tissue to a pre-chilled seven-milliliter glass Teflon Dounce homogenizer. Then, add five milliliters of ice-cold MSEGTA buffer and homogenize until there are no visible tissue pieces.Transfer the homogenate to a 15-milliliter tube, and then centrifuge at 600 times G at four degrees Celsius for five minutes to pellet unwanted nuclei and unbroken cells. Transfer the supernatant to a fresh 15-milliliter tube, and centrifuge at 10, 000 times G at four degrees Celsius for 10 minutes. After discarding the supernatant, keep the mitochondrial pellet on ice.To wash the pellet, add five milliliters of ice-cold MSEGTA buffer to it, resuspend and then centrifuge at 10, 000 times G at four degrees Celsius for 10 minutes. After discarding the supernatant, repeat the wash and keep the pellet on ice. To start mitochondrial calcium uptake measurement using a plate reader, program the reader to perform a kinetic read of Calcium Green-5N florescence every second for a total time of 1, 000 seconds.Program the reagent injectors to dispense five microliters of calcium chloride solution at various time points. Then, prime the injectors with the calcium chloride solution that will be used. Next, add 200 micrograms of isolated mitochondria to one well of a 96-well plate and add the appropriate volume of potassium chloride buffer to achieve a total volume of 197 microliters.Then, add one microliter of one molar pyruvate, one microliter of 500-millimolar malate and one microliter of one-millimolar Calcium Green-5N stock and mix gently by pipetting. Incubate protected from light at room temperature for two minutes for mitochondria to become energized. Finally, start the pre-programmed kinetic protocol and monitor Calcium Green-5N florescence.For this part of the protocol, prepare Rhod-2 AM MitoTracker Green working solution by mixing 20 microliters of Rhod-2 AM, 0.2 microliters of MitoTracker Green, 2.5 microliters of 20%pluronic F-127 and one milliliter of Tyrode solution. Onto a previously-prepared coverslip, plated with NIH/3T3 cells, add the Rhod-2 AM MitoTracker Green solution drop-wise until covered. For the cells to load with the dyes, incubate the coverslip protected from light for 30 minutes at room temperature.To deesterify Rhod-2 AM, gently remove the Rhod-2 AM MitoTracker Green solution, and replace it with approximately 300 microliters of fresh Tyrode solution to cover the cells. Incubate the coverslip protected from light for 30 minutes at room temperature. Now, transfer the coverslip to the microscope imaging chamber and fill the chamber with wash solution.Adjust the focus to observe the cells and phase contrast at 40X magnification. To permeabilize the plasma membrane of Rhod-2 AM MitoTracker Green loaded cells, remove the wash solution from the coverslip and replace it with approximately 300 microliters of permeabilization solution to cover the cells. Monitor the plasma membrane morphology during this process.When permeabilized, cells will develop a roughened surface. Remove the permeabilization solution immediately after complete permeabilization and replace it with zero-calcium internal solution. Next, focus on permeabilized cells displaying a clear co-localization of Rhod-2 and MitoTracker Green to image Rhod-2 and Mitotracker Green florescence simultaneously.Next, decrease the microscope laser and gain settings to make the mitochondrial Rhod-2 florescence dim and barely visible. To capture the kinetics of mitochondrial calcium changes, select microscope settings to acquire two-dimensional scans at an appropriate frame-rate and time course. Remove the zero-calcium internal solution making sure not to disturb the cells and microscope focus and then start image acquisition.Using a syringe, manually add calcium-replete internal solution at the 10-seconds time point. Finally, in the image acquisition software, select regions of interest to encompass regions of co-localization of Rhod-2 and MitoTracker Green signal. When calcium chloride is added to isolated cardiac mitochondria, florescence from the calcium dye increases as mitochondria uptake calcium from the buffer florescence decreases.However, at the third edition of calcium, florescence suddenly starts increasing after a drop indicating mitochondrial calcium overload and mitochondrial permeability transition pore opening which can lead to mitochondrial inner membrane permeabilization. On the other hand, when mitochondria are pre-treated with an inhibitor of mitochondrial calcium uptake, there is an increase in florescence following each calcium addition. In cultured NIH/3T3 cells, the mitochondrial network is stained with MitoTracker Green dye.Cells are also stained with Rhod-2 dye and after saponin permeabilization, cytosolic Rhod-2 is washed away leaving just mitochondrial staining which co-localizes with the MitoTracker. Since Rhod-2 can accumulate in non-mitochondrial structures, the analysis should focus only on regions where MitoTracker and Rhod-2 co-localize. The addition of calcium causes an increase in Rhod-2 florescence in control cells whereas in cells treated with the inhibitor of mitochondrial calcium uptake, there is no change in florescence.These protocols can be applied to measure mitochondrial calcium uptake in many cell types including primary cells such as cardiac myocytes and neurons in immortalized cell lines as well as isolated mitochondria from many tissue types including heart, liver and brain. Calcium handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in different cells. The ability to accurately measure the influx of calcium from mitochondria is an important part of determining the role of mitochondrial calcium handling in these processes.After watching this video, you should have a good understanding of how to measure mitochondrial calcium influx in isolated mitochondria and cultured cells.

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Research

JoVE Journal

Biochemistry

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Isolation and Respiratory Measurements of Mitochondria from Arabidopsis thaliana

Mitochondria are essential organelles involved in numerous metabolic pathways in plants, most notably the production of adenosine triphosphate (ATP) from the oxidation of reduced compounds such as nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide (FADH2). The complete annotation of the Arabidopsis thaliana genome has established it as the most widely used plant model system, and thus the need to purify mitochondria from a variety of organs (leaf, root, or flower) is necessary to fully utilize the tools that are now available for Arabidopsis to study mitochondrial biology. Mitochondria are isolated by homogenization of the tissue using a variety of approaches, followed by a series of differential centrifugation steps producing a crude mitochondrial pellet that is further purified using continuous colloidal density gradient centrifugation. The colloidal density material is subsequently removed by multiple centrifugation steps. Starting from 100 g of fresh leaf tissue, 2 - 3 mg of mitochondria can be routinely obtained. Respiratory experiments on these mitochondria display typical rates of 100 - 250 nmol O2 min-1 mg total mitochondrial protein-1 (NADH-dependent rate) with the ability to use various substrates and inhibitors to determine which substrates are being oxidized and the capacity of the alternative and cytochrome terminal oxidases. This protocol describes an isolation method of mitochondria from Arabidopsis thaliana leaves using continuous colloidal density gradients and an efficient respiratory measurements of purified plant mitochondria.

Isolation and Respiratory Measurements of Mitochondria from Arabidopsis thaliana

As mitochondria are only a small percentage of the plant cell, they need to be purified for a range of studies. Mitochondria can be isolated from a variety of plant organs by homogenization, followed by differential and density gradient centrifugation to obtain a highly purified mitochondrial fraction.

Mitochondria are essential organelles involved in numerous metabolic pathways in plants, most notably the production of adenosine triphosphate (ATP) from ...

Interactions with and Membrane Permeabilization of Brain Mitochondria by Amyloid Fibrils

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary mechanism of amyloid aggregate-induced toxicity in neurodegenerative diseases. However, most reports describing the mechanisms of membrane disruption are based on phospholipid model systems, and studies directly targeting events occurring at the level of biological membranes are rare. Described here is a model for studying the mechanisms of amyloid toxicity at the membrane level. For mitochondrial isolation, density gradient medium is used to obtain preparations with minimal myelin contamination. After mitochondrial membrane integrity confirmation, the interaction of amyloid fibrils arising from &#945;-synuclein, bovine insulin, and hen egg white lysozyme (HEWL) with rat brain mitochondria, as an in vitro biological model, is investigated. The results demonstrate that treatment of brain mitochondria with fibrillar assemblies can cause different degrees of membrane permeabilization and ROS content enhancement. This indicates structure-dependent interactions between amyloid fibrils and mitochondrial membrane. It is suggested that biophysical properties of amyloid fibrils and their specific binding to mitochondrial membranes may provide explanations for some of these observations.

Provided here is a protocol for investigating the interactions between native form, prefibrillar, and mature amyloid fibrils of different peptides and proteins with mitochondria isolated from different tissues and various areas of the brain.

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary ...

Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore

Eryptosis, erythrocyte programmed cell death, occurs in a number of hematological diseases and during injury to erythrocytes. A hallmark of eryptotic cells is the loss of compositional asymmetry of the cell membrane, leading to the translocation of phosphatidylserine to the membrane outer leaflet. This process is triggered by increased intracellular concentration of Ca2+, which activates scramblase, an enzyme that facilitates bidirectional movement of phospholipids between membrane leaflets. Given the importance of eryptosis in various diseased conditions, there have been efforts to induce eryptosis in vitro. Such efforts have generally relied on the calcium ionophore, ionomycin, to enhance intracellular Ca2+ concentration and induce eryptosis. However, many discrepancies have been reported in the literature regarding the procedure for inducing eryptosis using ionomycin. Herein, we report a step-by-step protocol for ionomycin-induced eryptosis in human erythrocytes. We focus on important steps in the procedure including the ionophore concentration, incubation time, and glucose depletion, and provide representative result. This protocol can be used to reproducibly induce eryptosis in the laboratory.

A protocol for the induction of eryptosis, programmed cell death in erythrocytes, using the calcium ionophore, ionomycin, is provided. Successful eryptosis is evaluated by monitoring the localization phosphatidylserine in the membrane outer leaflet. Factors affecting the success of the protocol have been examined and optimal conditions provided.

Eryptosis, erythrocyte programmed cell death, occurs in a number of hematological diseases and during injury to erythrocytes. A hallmark of eryptotic ...

Measurement of Protein Turnover Rates in Senescent and Non-Dividing Cultured Cells with Metabolic Labeling and Mass Spectrometry

Mounting evidence has shown that the accumulation of senescent cells in the central nervous system contributes to neurodegenerative disorders such as Alzheimer&#39;s and Parkinson&#39;s diseases. Cellular senescence is a state of permanent cell cycle arrest that typically occurs in response to exposure to sub-lethal stresses. However, like other non-dividing cells, senescent cells remain metabolically active and carry out many functions that require unique transcriptional and translational demands and widespread changes in the intracellular and secreted proteomes. Understanding how protein synthesis and decay rates change during senescence can illuminate the underlying mechanisms of cellular senescence and find potential therapeutic avenues for diseases exacerbated by senescent cells. This paper describes a method for proteome-scale evaluation of protein half-lives in non-dividing cells using pulsed stable isotope labeling by amino acids in cell culture (pSILAC) in combination with mass spectrometry. pSILAC involves metabolic labeling of cells with stable heavy isotope-containing versions of amino acids. Coupled with modern mass spectrometry approaches, pSILAC enables the measurement of protein turnover of hundreds or thousands of proteins in complex mixtures. After metabolic labeling, the turnover dynamics of proteins can be determined based on the relative enrichment of heavy isotopes in peptides detected by mass spectrometry. In this protocol, a workflow is described for the generation of senescent fibroblast cultures and similarly arrested quiescent fibroblasts, as well as a simplified, single-time point pSILAC labeling time-course that maximizes coverage of anticipated protein turnover rates. Further, a pipeline is presented for the analysis of pSILAC mass spectrometry data and user-friendly calculation of protein degradation rates using spreadsheets. The application of this protocol can be extended beyond senescent cells to any non-dividing cultured cells such as neurons.

This protocol describes the workflow for metabolic labeling of senescent and non-dividing cells with pulsed SILAC, untargeted mass spectrometry analysis, and a streamlined calculation of protein half-lives.

Mounting evidence has shown that the accumulation of senescent cells in the central nervous system contributes to neurodegenerative disorders such as ...

High-Throughput Quantification of the Synthesis and Distribution of mtDNA

High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells produce most of their ATP in the mitochondria by oxidative phosphorylation. Mitochondria are unique organelles because they have their own genome that is replicated and passed on to the next generation of cells. In contrast to the nuclear genome, there are multiple copies of the mitochondrial genome in the cell. The detailed study of the mechanisms responsible for the replication, repair, and maintenance of the mitochondrial genome is essential for understanding the proper functioning of mitochondria and whole cells under both normal and disease conditions. Here, a method that allows the high-throughput quantification of the synthesis and distribution of mitochondrial DNA (mtDNA) in human cells cultured in vitro is presented. This approach is based on the immunofluorescence detection of actively synthesized DNA molecules labeled by 5-bromo-2'-deoxyuridine (BrdU) incorporation and the concurrent detection of all the mtDNA molecules with anti-DNA antibodies. Additionally, the mitochondria are visualized with specific dyes or antibodies. The culturing of cells in a multi-well format and the utilization of an automated fluorescence microscope make it easier to study the dynamics of mtDNA and the morphology of mitochondria under a variety of experimental conditions in a relatively short time.

Author Spotlight: High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution  

A procedure for studying the dynamics of mitochondrial DNA (mtDNA) metabolism in cells using a multi-well plate format and automated immunofluorescence imaging to detect and quantify mtDNA synthesis and distribution is described. This can be further used to investigate the effects of various inhibitors, cellular stresses, and gene silencing on mtDNA metabolism.

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells ...

Isolation of Mitochondrial Contact Sites

An Improved Method to Isolate Mitochondrial Contact Sites

Mitochondria are present in virtually all eukaryotic cells and perform essential functions that go far beyond energy production, for instance, the synthesis of iron-sulfur clusters, lipids, or proteins, Ca2+ buffering, and the induction of apoptosis. Likewise, mitochondrial dysfunction results in severe human diseases such as cancer, diabetes, and neurodegeneration. In order to perform these functions, mitochondria have to communicate with the rest of the cell across their envelope, which consists of two membranes. Therefore, these two membranes have to interact constantly. Proteinaceous contact sites between the mitochondrial inner and outer membranes are essential in this respect. So far, several contact sites have been identified. In the method described here, Saccharomyces cerevisiae mitochondria are used to isolate contact sites and, thus, identify candidates that qualify for contact site proteins. We used this method to identify the mitochondrial contact site and cristae organizing system (MICOS) complex, one of the major contact site-forming complexes in the mitochondrial inner membrane, which is conserved from yeast to humans. Recently, we further improved this method to identify a novel contact site consisting of Cqd1 and the Por1-Om14 complex.

Author Spotlight: Unveiling Mitochondrial Contact Sites and Architectural Insights 

Mitochondrial contact sites are protein complexes that interact with mitochondrial inner and outer membrane proteins. These sites are essential for the communication between the mitochondrial membranes and, thus, between the cytosol and the mitochondrial matrix. Here, we describe a method to identify candidates qualifying for this specific class of proteins.

Mitochondria are present in virtually all eukaryotic cells and perform essential functions that go far beyond energy production, for instance, the ...

5-Bromo-2&#39;-Deoxyuridine (BrdU) Labeling of Mitochondrial DNA in HeLa Cells

5-Bromo-2'-Deoxyuridine (BrdU) Labeling of Mitochondrial DNA in HeLa Cells  

To begin the mitochondria labeling prepare a 20 micromolar solution of BrdU in DMEM with 10%fetal bovine serum or FBS. Then add the mitochondria tracking dye to adjust the final concentration to 1.1 micromolar. After 15 hours add 10 microliters of the mitochondria tracking dye solution to the HeLa cell suspension containing wells.Incubate the cells for one hour. The conditions used to label newly synthesized DNA molecules with BrdU allowed the detection of BrdU labeled DNA in the mitochondria of HeLa cells. The signal obtained with anti-BrdU antibodies was specific and only observed in cells treated with BrdU.Regardless of BrdU treatment no anti-BrdU or anti-DNA antibody signal was detected in the mitochondria of row zero cells. Quantifying the fluorescent signal from the anti-BrdU antibodies indicated that row zero cells treated with BrdU showed the same low level of fluorescence as the BrdU untreated cells, while the signal for the parental lines A549 and HeLa were 50 fold higher.

Locus-Specific Chromatin Isolation from Yeast Cells

Single-Copy Gene Locus Chromatin Purification in Saccharomyces cerevisiae

The basic organizational unit of eukaryotic chromatin is the nucleosome core particle (NCP), which comprises DNA wrapped ~1.7 times around a histone octamer. Chromatin is defined as the entity of NCPs and numerous other protein complexes, including transcription factors, chromatin remodeling and modifying enzymes. It is still unclear how these protein-DNA interactions are orchestrated at the level of specific genomic loci during different stages of the cell cycle. This is mainly due to the current technical limitations, which make it challenging to obtain precise measurements of such dynamic interactions. Here, we describe an improved method combining site-specific recombination with an efficient single-step affinity purification protocol to isolate a single-copy gene locus of interest in its native chromatin state. The method allows for the robust enrichment of the target locus over genomic chromatin, making this technique an effective strategy for identifying and quantifying protein interactions in an unbiased and systematic manner, for example by mass spectrometry. Further to such compositional analyses, native chromatin purified by this method likely reflects the in vivo situation regarding nucleosome positioning and histone modifications and is, therefore, amenable to further structural and biochemical analyses of chromatin derived from virtually any genomic locus in yeast.

Author Spotlight: Advancing Chromatin Research and Overcoming Limitations with a High-Enrichment Locus-Specific Chromatin Isolation Protocol 

This protocol presents a locus-specific chromatin isolation method based on site-specific recombination to purify a single-copy gene locus of interest in its native chromatin context from budding yeast, Saccharomyces cerevisiae.

The basic organizational unit of eukaryotic chromatin is the nucleosome core particle (NCP), which comprises DNA wrapped ~1.7 times around a histone ...

Generating Submitochondrial Vesicles from Saccharomyces cerevisiae Mitochondria to Isolate Mitochondrial Contact Sites

Generating Submitochondrial Vesicles from  Saccharomyces cerevisiae Mitochondria to Isolate Mitochondrial Contact Sites  

To begin, take 10 milligrams of freshly-isolated crude yeast mitochondria and re-suspend them in 1.6 milliliters of SM buffer at four degrees celsius by pipetting. Then, transfer the mitochondrial suspension to a pre-cooled 100-milliliter Erlenmeyer flask. Slowly, add 16 milliliters of the swelling buffer, using a 20-milliliter glass pipette while applying constant mild stirring on ice.Incubate the samples under constant mild stirring for 30 minutes on ice. Then, slowly add five milliliters of 2.5-molar sucrose solution using a five-milliliter glass pipette, allowing the inner membrane to shrink. Incubate the samples under mild stirring for 15 minutes on ice.To generate sub-mitochondrial membrane vesicles, transfer the mitochondrial suspension to a pre-cool rosette cell, and sonicate the suspension at 10%amplitude for 30 seconds while cooling the rosette cell in an ice bath. Rest the suspension for 30 seconds in an ice bath.