The Italian Ministry of Health supported this study (RC 2013-BIO 15). We thank Barbara Micheli for her excellent technical assistance.

In this protocol, the human hearts are collected during multiorgan explantation (cold ischemia time of 4-12 h, mean 6 &#177; 2 h) from multi-organ donors excluded from organ transplantation for technical or functional reasons, despite normal echocardiographic parameters. They are sent to the Cardiovascular Tissue Bank of Milan, Monzino Cardiologic Center (Milan, Italy) for the banking of the aortic and pulmonary valves. The mitral posterior leaflets are not used for clinical purposes, so they are collected during the aortic and pulmonary valve isolation after informed consent is obtained from the donors&#39; relatives. The tissue for transplantation and research is collected only after parental consent; on the consent sheet, they authorize (or not) the use of the cardiac tissue for research only if it is not suitable for human clinical use (i.e., microbiological, functional, and serological problems), following the guidelines of the ethics committee of Monzino Cardiologic Center.
1. Mitral valve preparation
<ol>
	<li>Harvest the human mitral valve as soon as possible after organ explantation (cold ischemia time of 4-12 h).</li>
	<li>In a clean room, remove the heart from the transport bag containing a cold (4 &#176;C) solution (i.e., saline solution or balanced medium Eurocollins or Wisconsin). Put it into a bucket and place it in a biosafety cabinet (biohazard vertical air flow, class A, Good Manufacturing Practices (GMP) classification) to proceed with the valve preparation.</li>
	<li>Place the heart on a sterile disposable drape in the cabinet. Using a sterile disposable scalpel, cut the heart completely, perpendicularly to its major axis, on the level of the left and right ventricles, about 4 cm away from the apex.</li>
	<li>Move the ascending aorta and pulmonary artery to display the left atrial roof.</li>
	<li>With sterile autoclavable forceps and picks, cut around the left auricle on the left atrial roof, making the mitral valve visible and allowing for the great mitral leaflet (anterior) and the small mitral leaflet (posterior) to be identified. 
	NOTE: Antero-lateral and the posterior medial commissures define the border of the anterior leaflet and the posterior area.</li>
	<li>Using sterile autoclavable scissors and non-traumatic forceps, dissect the left atrium and the ventricle wall thickness around the circumference of the whole mitral valve.</li>
	<li>Identify the mitro-aortic valve continuity. 
	NOTE: The left ventricle contains the whole mitral valve and chords.</li>
	<li>Separate the anterior mitral valve leaflet from the posterior mitral valve leaflet, cutting the posterior leaflet along the insertion with the ventricle (commissure).</li>
	<li>Wash the posterior leaflet in the saline solution. Cut the leaflet into small pieces (&#60;1 cm2) and individually wrap them in aluminum foil. Snap-freeze them with liquid nitrogen. 
	Caution: Follow organizational safety procedures when using liquid nitrogen.
	<ol>
		<li>Sanitize the table of the cabinet with a 70% isopropyl alcohol solution and a 6% hydrogen peroxide solution at the end of the procedure.</li>
	</ol>
	</li>
</ol>
2. Protein extraction
<ol>
	<li>Use forceps to pick up the sample stored in liquid nitrogen and immediately place it on dry ice while still wrapped in the aluminum foil. Do not leave the sample to thaw during any transfers.</li>
	<li>Prior to grinding, chill the porcelain/zirconium mortar and pestles of a grinder system (e.g., CryoGrinder), together with the sample, by putting them in a Dewar flask containing liquid nitrogen (~500 mL). 
	Caution: Follow organizational safety procedures when using liquid nitrogen.</li>
	<li>Put the mortar and pestles in a polystyrene box containing dry ice. Remove the sample from the aluminum foil and put it into the mortar.</li>
	<li>Grind the sample with the big pestle against the mortar 15-20 times, using the screwdriver to rotate the pestle. Mix the sample with the tip of a pre-chilled spatula during the grinding process.
	<ol>
		<li>Repeat with the small pestle.</li>
	</ol>
	</li>
	<li>Transfer the ground sample to a previously weighed tube (e.g., 15 mL centrifuge tube) by inverting the tube, placing it over the mortar, and inverting them together to move the sample to the tube. Use a pre-chilled spatula to recover all material from the mortar.
	<ol>
		<li>Keep the tube with the sample on dry ice to avoid sample thawing during the transfer.</li>
	</ol>
	</li>
	<li>Calculate the net weight of the sample.</li>
	<li>Clean the mortar and the pestles after each sample and decontaminate them by autoclaving or heating them at 200 &#176;C for 2 h.</li>
	<li>Transfer the powdered sample from the centrifuge tube to the glass tube of a homogenizer by inversion.</li>
	<li>Add filtered urea buffer (8 M urea, 2 M thiourea, 4% w/v CHAPS, 20 mM Tris, and 55 mM dithiotreitol) to the glass tube, 200 &#181;L of urea buffer for every 10 mg of powdered tissue. 
	NOTE: Residual powdered sample left in the centrifuge tube can be recovered using part of the calculated volume of urea buffer.</li>
	<li>Homogenize the sample using a stirrer equipped with a borosilicate glass mortar and a polytetrafluoroethylene (PTFE) pestle. Slowly press the pestle onto the sample with a twisting motion (1,500 rpm) 10 times.</li>
	<li>Recover the supernatant and transfer it to a clean 1.7 mL centrifuge tube. Again extract the remaining sample with fresh urea buffer, adding half of the volume used during the first extraction.</li>
	<li>Repeat step 2.10.</li>
	<li>Recover the supernatant and combine it with the supernatant from step 2.11. Place the combined supernatant on a tube rotator for 30 min.</li>
	<li>Centrifuge the tube for 30 min at 13,000 x g and 4 &#176;C.</li>
	<li>Recover the supernatant and measure the protein concentration using the Bradford protein assay, per the manufacturer&#39;s instructions. Store the sample at -80 &#176;C until use.</li>
</ol>

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The authors have nothing to disclose.

One critical step of this protocol is the use of liquid nitrogen to freeze the sample and to chill the grinder system. The use of liquid nitrogen prevents biological degradation and allows for efficient powdering, but it requires specific training for safe handling.
In this protocol features a grinder system for sample grinding because small samples are difficult to recover from standard mortar and pestles. In this case, small samples spread as a fine powder over the mortar surface, rendering ...

Recent evidence has altered the understanding of the roles of the many regulatory mechanisms that occur after mRNA synthesis. Indeed, translational, post-transcriptional, and proteolytic processes can regulate protein abundance and function. The dogma &#8211; which says that mRNA concentrations are proxies to those of the corresponding proteins, assuming that transcript levels are the main determinant of protein abundance &#8211; has been partially revised.Indeed, transcript levels only partially predict protein abundance, suggesting that post-transcriptional events occur to regulate the proteins within cells1,2.
Furthermore, proteins ultimately dictate the function of the cell and therefore dictate its phenotype, which can undergo dynamic changes in response to autocrine, paracrine, and endocrine factors; blood-borne mediators; temperature; drug treatment; and disease development. Thus, an expression analysis focused on the protein level is useful to characterize the proteome and to unravel the critical changes that occur to it as part of disease pathogenesis3.
Therefore, the opportunities that proteomics present to clarify health and disease conditions are formidable, despite the existing technological challenges. The particularly promising areas of research to which proteomics can contributeinclude: the identification of altered protein expression at any level (i.e., whole cells or tissue, subcellular compartments, and biological fluids); the identification, verification, and validation of novel biomarkers useful for the diagnosis and prognosis of disease; and, hopefully, the identification of new protein targets that can be used for therapeutic purposes, as well as for the assessment of drug efficacy and toxicity4.
Capturing the complexity of the proteome represents a technological challenge. The current proteomic tools offer the opportunity to perform large-scale, high-throughput analysis for the identification, quantification, and validation of altered protein levels. In addition, the introduction of fractionation and enrichment techniques, aimed at avoiding the interference caused by the most abundant proteins, has also improved protein identification by including the least abundant proteins. Finally, proteomics has been complemented by the analysis of post-translational modifications, which progressively emerge as important modulators of protein function.
However, the sample preparation and protein recovery in the biological specimens under analysis still remain the limiting steps in the proteomic workflow and increase the potential for possible pitfalls5. Indeed,in most of the molecular biology techniques that must be optimized, the first steps are tissue homogenization and cell lysis, especially during the analysis of low-abundance proteins for which amplification methods do not exist. In addition, the chemical nature of proteins can influence their own recovery. For example, the analysis of highly hydrophobic proteins is very challenging, because they easily precipitate during isoelectric focusing, while trans-membrane proteins are almost insoluble (reviewed in Reference 5). Furthermore, the tissue composition variability creates a significant barrier to developing a universal extraction method. Finally, because almost all of the clinical specimens are of limited quantity, it is essential to enable protein preparation with maximal recovery and reproducibility from minimal sample amounts6.
This work describes an optimized protocol for protein extraction from the normal human cardiac mitral valve, which represents a very challenging sample for proteomic analysis. The normal mitral valve is a complex structure lying between the left atrium and the left ventricle of the heart (Figure 1). It plays an important role in the control of blood flow from the atrium to the ventricle, preventing backflow and ensuring the proper level of oxygen supply to the whole body, thus maintaining an adequate cardiac output. However, it is often considered to be an &#34;inactive&#34; tissue, with a low cellularity and few components, mainly in the extracellular matrix. This is because, in normal conditions, the resident valvular interstitial cells (VICs) present a quiescent phenotype with a low protein biosynthesis rate7.
However, it has been demonstrated that, in a pathological state, the number of VICs in the spongiosa increases and their protein synthesis is activated, together with other functional and phenotypical changes8. Therefore, it is not surprising that the minimal data available in the literature focus on the analysis of pathological mitral valves9,10, in which the increased number of activated VICs might explain the relatively high number of identified proteins.
In conclusion, the present protocol may serve to develop the understanding of the pathogenic mechanisms responsible for mitral valve diseases through the study of mitral valve protein components. Indeed, a greater understanding of the underlying pathological processes could help to improve the clinical management of valve diseases, whose current indications for intervention are largely predicated on hemodynamic considerations.

<table><tbody><tr><td>Saline solution</td><td></td><td></td><td>0.9 % NaCl</td></tr><tr><td>Eurocollins A</td><td>SALF</td><td>30874046</td><td>Balanced organ&#039;s transport medium. Combine 400 mL of Eurocollins A with 100 mL Eurocollins B to obtain balanced medium Eurocollins</td></tr><tr><td>Eurocollins B</td><td>SALF</td><td>30874022</td><td>Balanced organ&#039;s transport medium. Combine 400 mL of Eurocollins A with 100 mL Eurocollins B to obtain balanced medium Eurocollins</td></tr><tr><td>Wisconsin</td><td>Bridge life</td><td>RM/N 4081</td><td>Balanced organ&#039;s transport medium</td></tr><tr><td>Biohazard vertical flow air</td><td>Burdinola</td><td></td><td>Class A GMP classification</td></tr><tr><td>Dewar Flask</td><td>Thermo Scientific</td><td>Nalgene 4150-1000</td><td></td></tr><tr><td>Cryogrinder system</td><td>OPS diagnostics</td><td>CG 08-01</td><td>Grinder system containing mortars, pestles and screwdriver</td></tr><tr><td>Stainless steel forceps</td><td></td><td></td><td></td></tr><tr><td>Stainless steel spatula</td><td></td><td></td><td></td></tr><tr><td>Disposable sterile scalpel</td><td>Medisafe</td><td>MS-10</td><td></td></tr><tr><td>Stainless steel scissors</td><td></td><td></td><td>Autoclavable</td></tr><tr><td>Stainless steel picks</td><td></td><td></td><td>Autoclavable</td></tr><tr><td>Disposable sterile drap</td><td>Mon&amp;amp;Tex</td><td>3.307.08</td><td></td></tr><tr><td>Sterilizing solution with isopropyl alcohol</td><td></td><td></td><td>70% isopropyl alcohol</td></tr><tr><td>Sterilizing solution with hydrogen peroxide</td><td></td><td></td><td>6% hydrogen peroxide</td></tr><tr><td>Micropipette, 1 mL, with tips</td><td></td><td></td><td></td></tr><tr><td>15 mL centrifuge tubes</td><td>VWR international</td><td>9278</td><td></td></tr><tr><td>1.7 mL centrifuge tubes</td><td>VWR international</td><td>PIER90410</td><td></td></tr><tr><td>Urea buffer</td><td></td><td></td><td>8 M urea, 2 M thiourea, 4 % w/v CHAPS, 20 mM Trizma, 55 mM Dithiotreitol</td></tr><tr><td>Urea</td><td>Sigma aldrich</td><td>U6504-1KG</td><td>To be used for Urea buffer</td></tr><tr><td>Thiourea</td><td>Sigma aldrich</td><td>T8656</td><td>To be used for Urea buffer</td></tr><tr><td>CHAPS</td><td>Sigma aldrich</td><td>C3023-5GR</td><td>To be used for Urea buffer</td></tr><tr><td>Dithiotreitol</td><td>Sigma aldrich</td><td>D0632-5G</td><td>To be used for Urea buffer</td></tr><tr><td>Syringe 50 mL</td><td>PIC</td><td></td><td>To be used to filter Urea buffer</td></tr><tr><td>0.22 &amp;micro;m filter</td><td>Millipore</td><td>SLGP033RB</td><td>To be used to filter Urea buffer</td></tr><tr><td>PFTE Pestle, 2 mL</td><td>Kartell</td><td>6302</td><td>Part of Potter-Elvehjem homogenizer</td></tr><tr><td>Borosilicate glass mortar</td><td>Kartell</td><td>6102</td><td>Part of Potter-Elvehjem homogenizer</td></tr><tr><td>Stirrer</td><td>VELP scientifica</td><td>Stirrer DLH</td><td>To be used for homogenization by Potter-Elvehjem</td></tr><tr><td>Bradford Protein assay</td><td>Bio-Rad laboratories</td><td>5000006</td><td></td></tr><tr><td>Tube rotator</td><td>Pbi International</td><td>F205</td><td></td></tr><tr><td>Liquid nitrogen</td><td></td><td></td><td></td></tr><tr><td>Aluminum foil</td><td></td><td></td><td></td></tr><tr><td>Ice</td><td></td><td></td><td></td></tr><tr><td>Polystyrene box</td><td></td><td></td><td></td></tr><tr><td>Dry ice</td><td></td><td></td><td></td></tr><tr><td>Centrifuge</td><td></td><td></td><td>For centrifugation of 1.7 mL centrifuge tubes at 13,000 x g</td></tr><tr><td>Freezer -80&amp;deg;C</td><td></td><td></td><td></td></tr><tr><td>Precision balance</td><td></td><td></td><td></td></tr><tr><td>Autoclave</td><td></td><td></td><td>For sterilization</td></tr><tr><td>Cryogenic gloves for liquid nitrogen</td><td></td><td></td><td></td></tr><tr><td>Gloves</td><td></td><td></td><td></td></tr><tr><td>Professional forced ventilation and natural air convection oven</td><td></td><td></td><td>For sterilization</td></tr><tr><td>Protease inhibitor cocktail</td><td>Sigma aldrich</td><td>P8340-5ML</td><td>100X solution</td></tr><tr><td>ProteoExtract Protein Precipitation Kit</td><td>Calbiochem</td><td>539180</td><td></td></tr><tr><td>RapiGest</td><td>Waters</td><td>186001861</td><td></td></tr><tr><td>Cytoscape</td><td>www.cytoscape.org</td><td>version 2.7</td><td>Software platform for Gene Ontology analysis</td></tr><tr><td>BiNGO</td><td>http://apps.cytoscape.org/apps/bingo</td><td>version 3.0.3</td><td>Plugin for Gene ontology analysis</td></tr><tr><td>AlphaB Crystallin/CRYAB Antibody</td><td>Novus Biologicals</td><td>NBP1-97494</td><td>Mouse monoclonal antibody against CryAB</td></tr><tr><td>Septin-11 Antibody</td><td>Novus Biologicals</td><td>NBP1-83824</td><td>Rabbit polyclonal antibody against septin-11</td></tr><tr><td>FHL1 Antibody</td><td>Novus Biologicals</td><td>NBP-188745</td><td>Rabbit polyclonal antibody against FHL-1</td></tr><tr><td>Dermatopontin Antibody</td><td>Novus Biologicals</td><td>NB110-68135</td><td>Rabbit polyclonal antibody against dermatopontin</td></tr><tr><td>Goat Anti mouse IgG HRP</td><td>Sigma aldrich</td><td>A4416-0.5ML</td><td>Secondary antibody for immunoblotting</td></tr><tr><td>Goat Anti rabbit IgG HRP</td><td>Bio-Rad laboratories</td><td>170-5046</td><td>Secondary antibody for immunoblotting</td></tr></tbody></table>

optimized protocol for the extraction of proteins from the human mitral valve

Analysis of the cellular proteome can help to elucidate the molecular mechanisms underlying diseases due to the development of technologies that permit the large-scale identification and quantification of the proteins present in complex biological systems.The knowledge gained from a proteomic approach can potentially lead to a better understanding of the pathogenic mechanisms underlying diseases, allowing for the identification of novel diagnostic and prognostic disease markers, and, hopefully, of therapeutic targets. However, the cardiac mitral valve represents a very challenging sample for proteomic analysis because of the low cellularity in proteoglycan and collagen-enriched extracellular matrix. This makes it challenging to extract proteins for a global proteomic analysis. This work describes a protocol that is compatible with subsequent protein analysis, such as quantitative proteomics and immunoblotting. This can allow for the correlation of data concerning protein expression with data on quantitative mRNA expression and non-quantitative immunohistochemical analysis. Indeed, these approaches, when performed together, will lead to a more comprehensive understanding of the molecular mechanisms underlying diseases, from mRNA to post-translational protein modification. Thus, this method can be relevant to researchers interested in the study of cardiac valve physiopathology.

The extraction and dissolution of proteins in the urea buffer is directly compatible with proteomic methods based on isoelectrofocusing (two-dimensional electrophoresis (2-DE)11 and liquid-phase isoelectric focusing (IEF)12) and with immunoblotting after dilution in Laemmli buffer13 containing a protease inhibitor cocktail14.
For gel-...

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Optimized Protocol for the Extraction of Proteins from the Human Mitral Valve

The protein composition of the human mitral valve is still partially unknown, because its analysis is complicated by low cellularity and therefore by low protein biosynthesis. This work provides a protocol to efficiently extract protein for the analysis of the mitral valve proteome.

Analysis of the cellular proteome can help to elucidate the molecular mechanisms underlying diseases due to the development of technologies that permit ...

optimized-protocol-for-extraction-proteins-from-human-mitral

Research

JoVE Journal

Biochemistry

12.8K Views.  Centro Cardiologico Monzino IRCCS.  De eiwitsamenstelling van de menselijke mitrale klep is nog steeds gedeeltelijk onbekend, omdat de analyse ervan gecompliceerd is door lage cellulariteit en derhalve door biosynthese met een lage eiwit. Dit werk biedt een protocol om eiwit efficiënt te extraheren voor de analyse van het mitrale klep proteome. 

Video: Optimized Protocol for the Extraction of Proteins from the Human Mitral Valve

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this technique is also frequently referred to as immunoprecipitation. Affinity capture can be applied in a bench-scale and in a high-throughput context. When coupled with protein mass spectrometry, affinity capture has proven to be a workhorse of interactome analysis. Although there are potentially many ways to execute the numerous steps involved, the following protocols implement our favored methods. Two features are distinctive: the use of cryomilled cell powder to produce cell extracts, and antibody-coupled paramagnetic beads as the affinity medium. In many cases, we have obtained superior results to those obtained with more conventional affinity capture practices. Cryomilling avoids numerous problems associated with other forms of cell breakage. It provides efficient breakage of the material, while avoiding denaturation issues associated with heating or foaming. It retains the native protein concentration up to the point of extraction, mitigating macromolecular dissociation. It reduces the time extracted proteins spend in solution, limiting deleterious enzymatic activities, and it may reduce the non-specific adsorption of proteins by the affinity medium. Micron-scale magnetic affinity media have become more commonplace over the last several years, increasingly replacing the traditional agarose- and Sepharose-based media. Primary benefits of magnetic media include typically lower non-specific protein adsorption; no size exclusion limit because protein complex binding occurs on the bead surface rather than within pores; and ease of manipulation and handling using magnets.

Here we describe protocols to disrupt mammalian cells by solid-state milling at a cryogenic temperature, produce a cell extract from the resulting cell powder, and isolate protein complexes of interest by affinity capture upon antibody-coupled micron-scale paramagnetic beads.

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this ...

Rapid Isolation of the Mitoribosome from HEK Cells

The human mitochondria possess a dedicated set of ribosomes (mitoribosomes) that translate 13 essential protein components of the oxidative phosphorylation complexes encoded by the mitochondrial genome. Since all proteins synthesized by human mitoribosomes are integral membrane proteins, human mitoribosomes are tethered to the mitochondrial inner membrane during translation. Compared to the cytosolic ribosome the mitoribosome has a sedimentation coefficient of 55S, half the rRNA content, no 5S rRNA and 36 additional proteins. Therefore, a higher protein-to-RNA ratio and an atypical structure make the human mitoribosome substantially distinct from its cytosolic counterpart.
Despite the central importance of the mitoribosome to life, no protocols were available to purify the intact complex from human cell lines. Traditionally, mitoribosomes were isolated from mitochondria-rich animal tissues that required kilograms of starting material. We reasoned that mitochondria in dividing HEK293-derived human cells grown in nutrient-rich expression medium would have an active mitochondrial translation, and, therefore, could be a suitable source of material for the structural and biochemical studies of the mitoribosome. To investigate its structure, we developed a protocol for large-scale purification of intact mitoribosomes from HEK cells. Herein, we introduce nitrogen cavitation method as a faster, less labor-intensive and more efficient alternative to traditional mechanical shear-based methods for cell lysis. This resulted in preparations of the mitoribosome that allowed for its structural determination to high resolution, revealing the composition of the intact human mitoribosome and its assembly intermediates. Here, we follow up on this work and present an optimized and more cost-effective method requiring only ~1010 cultured HEK cells. The method can be employed to purify human mitoribosomal translating complexes, mutants, quality control assemblies and mitoribosomal subunits intermediates. The purification can be linearly scaled up tenfold if needed, and also applied to other types of cells.

Mitochondria have specialized ribosomes that diverged from their bacterial and cytoplasmic counterparts. Here we show how mitoribosomes can be obtained from their native compartment in HEK cells. The method involves isolation of mitochondria from suspension cells and consequent purification of mitoribosomes.

The human mitochondria possess a dedicated set of ribosomes (mitoribosomes) that translate 13 essential protein components of the oxidative ...

Dynamic Proteomic and miRNA Analysis of Polysomes from Isolated Mouse Heart After Langendorff Perfusion

Studies in dynamic changes in protein translation require specialized methods. Here we examined changes in newly-synthesized proteins in response to ischemia and reperfusion using the isolated perfused mouse heart coupled with polysome profiling. To further understand the dynamic changes in protein translation, we characterized the mRNAs that were loaded with cytosolic ribosomes (polyribosomes or polysomes) and also recovered mitochondrial polysomes and compared mRNA and protein distribution in the high-efficiency fractions (numerous ribosomes attached to mRNA), low-efficiency (fewer ribosomes attached) which also included mitochondrial polysomes, and the non-translating fractions. miRNAs can also associate with mRNAs that are being translated, thereby reducing the efficiency of translation, we examined the distribution of miRNAs across the fractions. The distribution of mRNAs, miRNAs, and proteins was examined under basal perfused conditions, at the end of 30 min of global no-flow ischemia, and after 30 min of reperfusion. Here we present the methods used to accomplish this analysis&#8212;in particular, the approach to optimization of protein extraction from the sucrose gradient, as this has not been described before&#8212;and provide some representative results.

Here we present a protocol to perform polysome profiling on the isolated perfused mouse heart. We describe methods for heart perfusion, polysome profiling, and analysis of the polysome fractions with respect to mRNAs, miRNAs, and the polysome proteome.

Studies in dynamic changes in protein translation require specialized methods. Here we examined changes in newly-synthesized proteins in response to ...

Hybrid Clear/Blue Native Electrophoresis for the Separation and Analysis of Mitochondrial Respiratory Chain Supercomplexes

Complexes of the oxidative phosphorylation machinery form supramolecular protein arrangements named supercomplexes (SCs), which are believed to confer structural and functional advantages to mitochondria. SCs have been identified in many species, from yeast to mammal, and an increasing number of studies report disruption of their organization in genetic and acquired human diseases. As a result, an increasing number of laboratories are interested in analyzing SCs, which can be methodologically challenging. This article presents an optimized protocol that combines the advantages of Blue- and Clear-Native PAGE methods to resolve and analyze SCs in a time-effective manner. With this hybrid CN/BN-PAGE method, mitochondrial SCs extracted with optimal amounts of the mild detergent digitonin are exposed briefly to the anionic dye Coomassie Blue (CB) at the beginning of the electrophoresis, without exposure to other detergents. This short exposure to CB allows to separate and resolve SCs as effectively as with traditional BN-PAGE methods, while avoiding the negative impact of high CB levels on in-gel activity assays, and labile protein-protein interactions within SCs. With this protocol it is thus possible to combine precise and rapid in gel activity measurements with analytical techniques involving 2D electrophoresis, immuno-detection, and/or proteomics for advanced analysis of SCs.

Here we present a protocol to extract, resolve and identify mitochondrial supercomplexes which minimizes exposure to detergents and Coomassie Blue. This protocol offers an optimal balance between resolution, and preservation of enzyme activities, while minimizing the risk of losing labile protein-protein interactions.

Complexes of the oxidative phosphorylation machinery form supramolecular protein arrangements named supercomplexes (SCs), which are believed to confer ...

Measurement of Protein Import Capacity of Skeletal Muscle Mitochondria

Mitochondria are key metabolic and regulatory organelles that determine the energy supply as well as the overall health of the cell. In skeletal muscle, mitochondria exist in a series of complex morphologies, ranging from small oval organelles to a broad, reticulum-like network. Understanding how the mitochondrial reticulum expands and develops in response to diverse stimuli such as alterations in energy demand has long been a topic of research. A key aspect of this growth, or biogenesis, is the import of precursor proteins, originally encoded by the nuclear genome, synthesized in the cytosol, and translocated into various mitochondrial sub-compartments. Mitochondria have developed a sophisticated mechanism for this import process, involving many selective inner and outer membrane channels, known as the protein import machinery (PIM). Import into the mitochondrion is dependent on viable membrane potential and the availability of organelle-derived ATP through oxidative phosphorylation. Therefore its measurement can serve as a measure of organelle health. The PIM also exhibits a high level of adaptive plasticity in skeletal muscle that is tightly coupled to the energy status of the cell. For example, exercise training has been shown to increase import capacity, while muscle disuse reduces it, coincident with changes in markers of mitochondrial content. Although protein import is a critical step in the biogenesis and expansion of mitochondria, the process is not widely studied in skeletal muscle. Thus, this paper outlines how to use isolated and fully functional mitochondria from skeletal muscle to measure protein import capacity in order to promote a greater understanding of the methods involved and an appreciation of the importance of the pathway for organelle turnover in exercise, health, and disease.

Mitochondria are key metabolic organelles that exhibit a high level of phenotypic plasticity in skeletal muscle. The import of proteins from the cytosol is a critical pathway for organelle biogenesis, essential for the expansion of the reticulum and the maintenance of mitochondrial function. Therefore, protein import serves as a barometer of cellular health.

Mitochondria are key metabolic and regulatory organelles that determine the energy supply as well as the overall health of the cell. In skeletal muscle, ...

Mitochondrial Isolation from Rodents for High-Quality Bioenergetic Studies

Robust Mitochondrial Isolation from Rodent Cardiac Tissue

Mitochondrial isolation has been practiced for decades, following procedures established by pioneers in the fields of molecular biology and biochemistry to study metabolic impairments and disease. Consistent mitochondrial quality is necessary to properly investigate mitochondrial physiology and bioenergetics; however, many different published isolation methods are available for researchers. Although different experimental strategies require different isolation methods, the basic principles and procedures are similar. This protocol details a method capable of extracting well-coupled mitochondria from a variety of tissue sources, including small animals and cells. The steps outlined include organ dissection, mitochondrial purification, protein quantification, and various quality control checks. The primary quality control metric used to identify high-quality mitochondria is the respiratory control ratio (RCR). The RCR is the ratio of the respiratory rate during oxidative phosphorylation to the rate in the absence of ADP. Alternative metrics are discussed. While high RCR values relative to their tissue source are obtained using this protocol, several steps can be optimized to suit the individual needs of researchers. This procedure is robust and has consistently resulted in isolated mitochondria with above-average RCR values across animal models and tissue sources.

Author Spotlight: Uncovering the Role of Mitochondrial Calcium Phosphate in Heart Failure and Bioenergetics

Bioenergetic and metabolomic studies on mitochondria have revealed their multifaceted role in many diseases, but the isolation methods for these organelles vary. The method detailed here is capable of purifying high-quality mitochondria from multiple tissue sources. Quality is determined by respiratory control ratios and other metrics assessed with high-resolution respirometry.

Mitochondrial isolation has been practiced for decades, following procedures established by pioneers in the fields of molecular biology and biochemistry ...

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in current research and a plethora of methods for their investigation is available1. Protein-protein interactions often are highly dynamic and may depend on subcellular localization, post-translational modifications and the local protein environment2. Therefore, they should be investigated in their natural environment, for which co-immunoprecipitation approaches are the method of choice3. Co-precipitated interaction partners are identified either by immunoblotting in a targeted approach, or by mass spectrometry (LC-MS/MS) in an untargeted way. The latter strategy often is adversely affected by a large number of false positive discoveries, mainly derived from the high sensitivity of modern mass spectrometers that confidently detect traces of unspecifically precipitating proteins. A recent approach to overcome this problem is based on the idea that reduced amounts of specific interaction partners will co-precipitate with a given target protein whose cellular concentration is reduced by RNAi, while the amounts of unspecifically precipitating proteins should be unaffected. This approach, termed QUICK for QUantitative Immunoprecipitation Combined with Knockdown4, employs Stable Isotope Labeling of Amino acids in Cell culture (SILAC)5 and MS to quantify the amounts of proteins immunoprecipitated from wild-type and knock-down strains. Proteins found in a 1:1 ratio can be considered as contaminants, those enriched in precipitates from the wild type as specific interaction partners of the target protein. Although innovative, QUICK bears some limitations: first, SILAC is cost-intensive and limited to organisms that ideally are auxotrophic for arginine and/or lysine. Moreover, when heavy arginine is fed, arginine-to-proline interconversion results in additional mass shifts for each proline in a peptide and slightly dilutes heavy with light arginine, which makes quantification more tedious and less accurate5,6. Second, QUICK requires that antibodies are titrated such that they do not become saturated with target protein in extracts from knock-down mutants.
Here we introduce a modified QUICK protocol which overcomes the abovementioned limitations of QUICK by replacing SILAC for 15N metabolic labeling and by replacing RNAi-mediated knock-down for affinity modulation of protein-protein interactions. We demonstrate the applicability of this protocol using the unicellular green alga Chlamydomonas reinhardtii as model organism and the chloroplast HSP70B chaperone as target protein7 (Figure 1). HSP70s are known to interact with specific co-chaperones and substrates only in the ADP state8. We exploit this property as a means to verify the specific interaction of HSP70B with its nucleotide exchange factor CGE19.

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

We present a variation of the QUICK (QUantitative Immunoprecipitation Combined with Knockdown) approach that was introduced previously to distinguish between true and false protein-protein interactions. Our approach is based on 15N metabolic labeling, the modulation of affinities of protein-protein interactions by the presence/absence of ATP, immunoprecipitation, and quantitative mass spectrometry.

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in ...

Biology

Protein Isolation from the Developing Embryonic Mouse Heart Valve Region

Western blot analysis is a commonly employed technique for detecting and quantifying protein levels. However, for small tissue samples, this analysis method may not be sufficiently sensitive to detect a protein of interest. To overcome these difficulties, we examined protocols for obtaining protein from adult human cardiac valves and modified these protocols for the developing early embryonic mouse counterparts. In brief, the mouse embryonic aortic valve regions, including the aortic valve and surrounding aortic wall, are collected in the minimal possible volume of a Tris-based lysis buffer with protease inhibitors. If required based on the breeding strategy, embryos are genotyped prior to pooling four embryonic aortic valve regions for homogenization. After homogenization, an SDS-based sample buffer is used to denature the sample for running on an SDS-PAGE gel and subsequent western blot analysis. Although the protein concentration remains too low to quantify using spectrophotometric protein quantification assays and have sample remaining for subsequent analyses, this technique can be used to successfully detect and semi-quantify phosphorylated proteins via western blot from pooled samples of four embryonic day 13.5 mouse aortic valve regions, each of which yields approximately 1 &#956;g of protein. This technique will be of benefit for studying cell signaling pathway activation and protein expression levels during early embryonic mouse valve development.

The analysis of protein expression in young embryonic mouse valves has been hampered by the limited tissue available. This manuscript provides a protocol for preparing protein from developing embryonic mouse valve regions for western blot analysis.

Western blot analysis is a commonly employed technique for detecting and quantifying protein levels. However, for small tissue samples, this analysis ...

Isolation of Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High Throughput Microplate Respiratory Measurements

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial respirometric assays from isolated mitochondrial preparations allow for the assessment of mitochondrial function, as well as determination of the mechanism(s) of action of drugs and proteins that modulate metabolism. Current isolation procedures often require large quantities of tissue to yield high quality mitochondria necessary for respirometric assays. The methods presented herein describe how high quality purified mitochondria (~ 450 &#181;g) can be isolated from minimal quantities (~75-100 mg) of mouse skeletal muscle for use in high throughput respiratory measurements. We determined that our isolation method yields 92.5&#177; 2.0% intact mitochondria by measuring citrate synthase activity spectrophotometrically. In addition, Western blot analysis in isolated mitochondria resulted in the faint expression of the cytosolic protein, GAPDH, and the robust expression of the mitochondrial protein, COXIV. The absence of a prominent GAPDH band in the isolated mitochondria is indicative of little contamination from non-mitochondrial sources during the isolation procedure. Most importantly, the measurement of O2 consumption rate with micro-plate based technology and determining the respiratory control ratio (RCR) for coupled respirometric assays shows highly coupled (RCR; &#62;6 for all assays) and functional mitochondria. In conclusion, the addition of a separate mincing step and significantly reducing motor driven homogenization speed of a previously reported method has allowed the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle that results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Here, we present a modification of a previously reported method that allows for the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle. This procedure results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial ...

Multifunctional, Micropipette-based Method for Incorporation And Stimulation of Bacterial Mechanosensitive Ion Channels in Droplet Interface Bilayers

MscL, a large conductance mechanosensitive channel (MSC), is a ubiquitous osmolyte release valve that helps bacteria survive abrupt hypo-osmotic shocks. It has been discovered and rigorously studied using the patch-clamp technique for almost three decades. Its basic role of translating tension applied to the cell membrane into permeability response makes it a strong candidate to function as a mechanoelectrical transducer in artificial membrane-based biomolecular devices. Serving as building blocks to such devices, droplet interface bilayers (DIBs) can be used as a new platform for the incorporation and stimulation of MscL channels. Here, we describe a micropipette-based method to form DIBs and measure the activity of the incorporated MscL channels. This method consists of lipid-encased aqueous droplets anchored to the tips of two opposing (coaxially positioned) borosilicate glass micropipettes. When droplets are brought into contact, a lipid bilayer interface is formed. This technique offers control over the chemical composition and the size of each droplet, as well as the dimensions of the bilayer interface. Having one of the micropipettes attached to a harmonic piezoelectric actuator provides the ability to deliver a desired oscillatory stimulus. Through analysis of the shapes of the droplets during deformation, the tension created at the interface can be estimated. Using this technique, the first activity of MscL channels in a DIB system is reported. Besides MS channels, activities of other types of channels can be studied using this method, proving the multi-functionality of this platform. The method presented here enables the measurement of fundamental membrane properties, provides a greater control over the formation of symmetric and asymmetric membranes, and is an alternative way to stimulate and study mechanosensitive channels.

Bacterial mechanosensitive channels can be used as mechanoelectrical transducers in biomolecular devices. Droplet interface bilayers (DIBs), cell-inspired building blocks to such devices, represent new platforms to incorporate and stimulate mechanosensitive channels. Here, we demonstrate a new micropipette-based method of forming DIBs, allowing the study of mechanosensitive channels under mechanical stimulation.

MscL, a large conductance mechanosensitive channel (MSC), is a ubiquitous osmolyte release valve that helps bacteria survive abrupt hypo-osmotic shocks. ...

Optimized Protocol for the Extraction of Proteins from the Human Mitral Valve

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