We would like to acknowledge Daniel A. Beard and Kalyan C. Vinnakota for their foundational contributions to this protocol. This work was funded by NSF CAREER grant MCB-2237117.

The use and treatment of all vertebrate animals were performed in accordance with approved protocols reviewed and accepted by the Institutional Animal Care and Use Committee (IACUC) at Michigan State University. This protocol was designed using both male and female Hartley albino guinea pigs and Sprague Dawley (SD) rats. For the isolation of cardiac mitochondria from guinea pigs, animals were sacrificed at ages between 4-6 weeks (300-450 g). Cardiac mitochondria from SD rats of both sexes were obtained between the ages of 10-13 weeks (250-400 g). Recipes for buffers are described in Table 1 and should be prepared in advance. Details of the reagents and equipment used in this study are listed in the Table of Materials.
1. Experimental preparation
<ol>
	<li>Before starting the isolation, label two 50 mL centrifugation tubes as &#34;Spins 1 and 2&#34; and &#34;Spin 3&#34;.</li>
	<li>Place tubes, freshly thawed isolation buffer (IB), sharp mincing scissors, assembled homogenization probe, and a 5 mL beaker or equivalent-sized container on ice.</li>
	<li>Place freshly thawed respiration buffer (RB) in an incubator to warm for subsequent respiratory assays.</li>
	<li>Pre-chill a refrigerated centrifuge to 4 &#176;C.</li>
	<li>Ensure that all equipment is arranged and 20 &#181;L of 7-15 U/mg protease from Bacillus licheniformis has been added to the tube labeled &#34;Spins 1 and 2&#34;.</li>
	<li>Set up a gravity-dependent pressure system for perfusion via cannulation using Cardioplegia buffer (CB, Table 1), glass cannula, and plastic tubing with a stopcock valve attached.</li>
	<li>Arrange surgical tools and include proper scissors and forceps for dissection and cannulation of the heart. 
	NOTE: All water should be of pure quality (18.2 M&#937;&#183;cm)</li>
</ol>
2. Tissue dissection
<ol>
	<li>Inject animals with sterile heparin sulfate intraperitoneally at a dose of 500 U/kg.</li>
	<li>Allow the animal to sit in the induction chamber for 15 min after heparin administration. During this time, supply 2 L/min of pure O2 gas to fully oxygenate the animals, calm them, and minimize any stress that may have adverse consequences on tissue of interest.</li>
	<li>Start anesthesia induction by a continuous flow of isoflurane at 0.5%. After 1 min, increase to 1%. Continue toincrease by 1% every minute until 5% is reached. Once at 5%, wait for 1 min and monitor the animal&#39;s breathing pattern.</li>
	<li>Once breathing has slowed and becomes labored (approximately at the 6.5 min mark) turn off isoflurane and oxygen flow.</li>
	<li>Quickly remove the animal from the induction chamber and check for appropriate anesthetic depth by squeezing a paw and checking for the corneal reflex. If the animal responds to either stimulus, then place it back in the induction chamber, readminister anesthesia, and repeat the anesthetic depth check.</li>
	<li>Once the proper depth of anesthesia is reached, quickly decapitate with a guillotine to severe the cervical spine and place the prone body on ice.</li>
	<li>Make two parallel vertical incisions from the clavicles proceeding along the lateral rib cage down the length of the thorax. Ensure that the incisions are deep enough to cut through the ribs on the lateral thorax, but avoid damaging intrathoracic structures such as the heart or great vessels. 
	NOTE: Vertical incision sizes depend on the animal being used. If using guinea pigs and rats, cut to the diaphragm (approximately 6.35 cm for rats and 11 cm for guinea pigs). If using mice, perform a standard thoracotomy27.</li>
	<li>Expose the heart using a hemostat to displace the anterior thorax and pack the exposed thoracic cavity with ice. This step minimizes warm ischemia time and enhances the viability of the isolated organelles.</li>
	<li>Use tweezers to bluntly dissect the thymus and pericardium and fully expose the heart.</li>
	<li>Using forceps, apply gentle inferior traction on the heart to expose and identify the aorta. The aorta is the thickest great vessel branching out from the base of the heart. Other large vessels, such as the pulmonary vein, are noticeably more translucent than the aorta.</li>
	<li>Cut the aorta approximately 4-6 mm above the aortic root but below the carotid branching points.</li>
	<li>Cannulate the aorta and perfuse the heart28,29 with ice-cold cardioplegia (CB) solution using a gravity-dependent pressure head until the coronary arteries are cleared of blood and the organ appears blanched. 
	NOTE: For large rodents, a cannula diameter of 1.8-2.2 mm works well, while for smaller rodents, a diameter range of 1.4-1.8 mm is recommended. Retro-perfusion should take no more than 15-30 s for the coronary vessels to clear of blood.</li>
	<li>Isolate the ventricular myocardium by dissecting away the atria, cartilaginous valvular tissue, and fatty tissues.</li>
	<li>Place ventricles in a pre-chilled 10 mL beaker containing 0.1-0.2 mL ice-cold IB.</li>
	<li>Mince tissue with sharp surgical scissors until pieces are approximately 1 mm3.</li>
</ol>
3. Mitochondrial purification and protein quantification
<ol>
	<li>Transfer the minced tissue into the pre-chilled centrifuge tube labeled &#34;Spins 1 and 2&#34; containing the protease solution.</li>
	<li>Add ice-cold IB to a final volume of 25 mL.</li>
	<li>Using a motorized handheld rotor-stator homogenizer, disperse the tissue at 18,000 rpm on ice for 20-25 s.</li>
	<li>Centrifuge homogenized tissue in a tube labeled &#34;Spins 1 and 2&#34; at 8,000 x g for 10 min at 4 &#176;C.</li>
	<li>Discard the supernatant (which contains protease) by pouring it into the waste carboy and gently rinse the pellet with 5 mL of IB to remove residual protease.</li>
	<li>After discarding the rinse, resuspend the pellet with fresh ice-cold IB to a final volume of 25 mL by gentle vortex.</li>
	<li>Centrifuge at 800 x g for 10 min at 4 &#176;C.</li>
	<li>Remove the supernatant (contains mitochondria) by gently pouring it into a pre-chilled 50 mL tube labeled &#34;Spin 3&#34;. While pouring, take care to avoid dislodging the loose upper portion of the pellet. As an alternative, a transfer pipette or stripette can be used to collect the supernatant.</li>
	<li>Centrifuge the supernatant at 8000 x g for 10 min at 4 &#176;C.</li>
	<li>Discard the resulting supernatant and retain the mitochondria-containing pellet.</li>
	<li>Use a lint-free wipe to absorb excess supernatant from the inside wall of the tube, taking care to avoid disturbing the pellet. Keep the pellet at 4 &#176;C on ice as much as possible.</li>
	<li>To resuspend the mitochondria, add 80 &#181;L of ice-cold IB to the bottom of the tube. Gently resuspend the pellet by repeatedly washing IB over the pellet.</li>
	<li>To avoid creating bubbles, set a micropipette to aspirate and deliver between 40-60 &#181;L of volume.</li>
	<li>As the mitochondrial pellet disperses, increase the micropipette volume to efficiently resuspend the pelleted mitochondria. Avoid touching the pellet with the pipette tip, and avoid making bubbles.</li>
	<li>Once resuspended, transfer the mitochondria to a pre-chilled microcentrifuge tube and label it as stock mitochondria. Make a note of the total resuspension volume.</li>
	<li>To determine the mitochondrial protein concentration in the sample, conduct a total protein assay using the well-known BCA or Bradford protein assays as defined by the manufacturer&#39;s instructions. 
	NOTE: An alternative or complementary strategy to assess yield is to determine the citrate synthase activity. For reference, mitochondrial content can be quantified by following the protocol described in Vinnakota et al.30.</li>
</ol>
4. Quality control checks
<ol>
	<li>In a pre-chilled microcentrifuge tube, dilute the mitochondrial stock to the desired working concentration with IB. 
	NOTE: Mitochondrial stocks are diluted to 40 mg/mL to work at 0.1 mg/mL final concentration for respirometry assays when using isolates from cardiac tissue.</li>
	<li>Rinse oxygraph chambers, stoppers, and microliter syringes ten times with distilled water to clean them before use in respirometry assays.</li>
	<li>To test the viability and quality of mitochondrial isolates, load 2.3 mL of respiration buffer (RB) into oxygraph chambers and allow the oxygen consumption signal to equilibrate at 37 &#176;C for about 10 min or when the rate is near 0.</li>
	<li>Once the signal is equilibrated, push down the stoppers and aspirate the excess buffer that emerges from the capillary of the stopper.</li>
	<li>Add 1 mM EGTA using a microliter syringe to chelate any residual calcium in the buffers or mitochondrial sample.</li>
	<li>To fuel respiration, add 5 mM sodium pyruvate and 1 mM L-malate.</li>
	<li>Following the addition of substrates, add a bolus of diluted mitochondria to achieve working concentration and allow respiration to occur for 5 min. This period is termed LEAK or State 2 respiration.</li>
	<li>At the 5 min mark, add a bolus of 500 &#181;M ADP to initiate State 3 respiration, also termed OXHPOS, and allow the mitochondria to respire until anoxia.</li>
	<li>Calculate the respiratory control ratio (RCR) by dividing the maximal rate of oxygen consumption during State 3 by the rate of oxygen consumption just before the addition of ADP in State 2 (see Figure 1). 
	NOTE: An alternative RCR expression of 1 - 1/RCR can also be used as a metric of quality, which is bounded between 0 and 1; however, it makes it difficult to differentiate quality using this metric when the RCR &#62; 10 (see Figure 2).</li>
	<li>Rinse chambers and stoppers 10 times with pure water to clean the oxygraph for subsequent assays. If respirometry is complete, fill chambers with 70% ethanol and place stoppers in chambers until the next use.</li>
</ol>

Mitochondrial Isolation from Rodents for High-Quality Bioenergetic Studies

The authors declare that there is nothing to disclose.

Adhering to the methods concisely described in this protocol will ensure the isolation of well-coupled mitochondria from the cardiac tissue of small rodents, in addition to other tissue types and sources. Overall, the process should take a total of 3-3.5 h, during which all animal tissue, samples, and isolates should remain on ice at 4 &#176;C as much as possible to limit degradation. This procedure is robust and can be altered in several ways to better fit experimental goals and models utilized. One modulation that can ...

Mitochondria are subcellular organelles that establish cytoplasmic energetic conditions optimized for specific cell functions. While cellular, tissue, and organism-level studies can provide insights into mitochondrial function, isolating the organelles offers a level of experimental control not possible otherwise. Mitochondrial isolations have been performed since the 1940s, allowing mechanistic studies of metabolism and respiration across a variety of cells and tissues1,2. The historical relevance of mitochondria is also well-documented3. As the main producers of ATP, mitochondria play many key roles that are vital for optimal cellular and organ function4. Within the mitochondrial matrix, substrates are oxidized by the TCA cycle, producing reducing equivalents and mobile electron carriers such as NADH and UQH25,6. Cytochrome C is the third main mobile electron carrier in the mitochondrial biochemical reaction network7. These molecules are then oxidized by the transmembrane complexes of the electron transport system (ETS) embedded in the inner mitochondrial membrane8. Redox reactions of the ETS are coupled with proton translocation from the matrix to the intermembrane space. These processes establish an electrochemical proton gradient that is used to phosphorylate ADP with Pi by F1F0 ATP synthase to produce ATP9,10. The individual processes that occur at each complex can be explored with high-resolution respirometry using Clark-type electrodes or microplate oxygen consumption assays11,12. Additionally, disease models and treatments using isolated mitochondria can determine the impact or importance of mitochondrial function in the progression of certain pathologies. This has proven fruitful in the field of cardiology, where alterations in fuel and substrate delivery have been used to elucidate how mitochondrial dysfunction influences heart failure13,14,15,16. Mitochondria are also known to impact the development of other disease states such as diabetes, cancer, obesity, neurological disorders, and myopathies17,18. Therefore, the use of isolated or purified mitochondria enables mechanistic investigations of oxidative metabolism and ATP production in the source tissue.
There is no shortage of mitochondrial isolation protocols due to their importance in bioenergetic research. Additionally, highly specific methods tailored to subpopulations of mitochondria within tissues and cells can be found19,20. The basic procedural steps are similar between isolation methods, but variations can be made to buffer composition, homogenization steps, and centrifugation spins to improve the amount and quality of mitochondria. Changes to these aspects are based on the metabolic demand of the tissue, overall organ function, mitochondrial density, and other factors. In tissues such as the liver and skeletal muscle, handheld homogenizers are used to preserve mitochondrial integrity and limit damage to the mitochondrial membranes21. However, when isolating from kidneys, some protocols suggest using manually driven homogenization or commercial kits to yield better results22. Although both methods yield functional mitochondria, the quality of the organelles can become compromised by the additional time it takes to complete isolations using these protocols. Centrifugation is also vital to the extraction of mitochondrial protein, as it separates cellular components such as nuclei and other organelles from mitochondria23. During the isolation process, it is debated whether differential or density-based centrifugation should be implemented to obtain purer isolates24. While density centrifugation can separate mitochondria from organelles of similar specific gravity, such as peroxisomes, it is not well-established if mitochondria from these methods better represent in situ organelle function compared to mitochondria isolated using differential centrifugation. In the field of mitochondrial physiology, density-based centrifugation is preferred and can be easily altered to increase isolate purity. Whether changes to g-forces, centrifugation duration, and the number of centrifugation spins are incorporated should be considered before experimentation due to their influence on mitochondrial purification. Furthermore, mitochondrial resuspension, arguably the most crucial step during isolation, differs greatly between studies, with the use of scraping, vortex-based mixing, or homogenization23,25,26. Mechanical resuspension of these types can be too abrasive and impair the membrane integrity of mitochondria. For this reason, gentle washing should be performed to correct this. Despite the plethora of modulations and suggested methodologies, there are fewer comprehensive protocols with high reproducibility and adaptability for rodent models.
The methods described herein outline a detailed, robust, and highly reproducible protocol that yields purified, well-coupled mitochondria from small animal cardiac tissue. As demonstrated, this method can be easily adapted to accommodate the specific needs of each experiment and/or laboratory environment for isolating mitochondria from kidneys, liver, and cultured cells. Further modifications can be made to isolate mitochondria from tissues and other animals not listed here. Buffer recipes used for all isolations are provided and can be modified if needed. Similar to other published protocols, motorized homogenization, and differential centrifugation are implemented; however, adjustments are made to both the shearing time and the force at which the samples are centrifuged to consistently deliver high-quality mitochondrial isolates, depending on the isolation source. Notably, this protocol differs from others by using gentle washing via pipetting to resuspend pelleted mitochondria, which helps preserve mitochondrial membrane integrity and maintain the overall functionality of the organelles. Mitochondrial protein is quantified either by total protein determination or by measuring citrate synthase activity. The utility and broad applicability of this isolation method are further supported by the values of respiratory control ratios (RCR) achieved across various organisms and tissue sources.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.7 mL microcentrifuge tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10 mL glass beaker</td>
			<td></td>
			<td></td>
			<td>For organ disection and mincing</td>
		</tr>
		<tr>
			<td>50 mL centrifuge tubes</td>
			<td></td>
			<td></td>
			<td>Centrifugation</td>
		</tr>
		<tr>
			<td>Adenosine 5&#39;-diphosphate monopotassium salt dihydrate</td>
			<td>Sigma&#160;</td>
			<td>A5285</td>
			<td>Respirometry assays</td>
		</tr>
		<tr>
			<td>BSA Protein Assay Kit</td>
			<td>Thermo Scentific</td>
			<td>PI23225</td>
			<td>Mitochondrial protein quantification</td>
		</tr>
		<tr>
			<td>Dextrose</td>
			<td>Sigma&#160;</td>
			<td>DX0145</td>
			<td>For buffer (CB)</td>
		</tr>
		<tr>
			<td>Ethylene glycol-bis(2-amino-ethylether)-N,N,N&#39;,N&#39;-tetraacetic acid&#160;</td>
			<td>Sigma&#160;</td>
			<td>E4378</td>
			<td>For buffers (CB and IB) and respirometry assays</td>
		</tr>
		<tr>
			<td>Glass cannula</td>
			<td>Radnoti</td>
			<td></td>
			<td>Guinea pig and rat heart perfusion</td>
		</tr>
		<tr>
			<td>Heparin sodium porcine mucosa</td>
			<td>Sigma&#160;</td>
			<td>SRE0027-250KU</td>
			<td>Animal IP injection</td>
		</tr>
		<tr>
			<td>High-resolution respirometer</td>
			<td></td>
			<td></td>
			<td>Clark-type electrode; oxygraph with 2 mL chambers</td>
		</tr>
		<tr>
			<td>Induction chamber</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Sigma&#160;</td>
			<td>792632</td>
			<td>Anesthetic&#160;</td>
		</tr>
		<tr>
			<td>L-malic acid</td>
			<td>Sigma&#160;</td>
			<td>02288-50G</td>
			<td>Respirometry assays</td>
		</tr>
		<tr>
			<td>Magnesium chloride hexahydrate</td>
			<td>Sigma&#160;</td>
			<td>M9272</td>
			<td>For buffer (RB)</td>
		</tr>
		<tr>
			<td>Mannitol</td>
			<td>Sigma&#160;</td>
			<td>MX0214</td>
			<td>For buffer (IB)</td>
		</tr>
		<tr>
			<td>Microliter syringes</td>
			<td></td>
			<td></td>
			<td>Sizes ranging from 5&#8211;50 &#181;L</td>
		</tr>
		<tr>
			<td>Microplate reader</td>
			<td></td>
			<td></td>
			<td>Must be able to incubate at 37 &#176;C&#160;</td>
		</tr>
		<tr>
			<td>MOPS</td>
			<td>Sigma&#160;</td>
			<td>475898</td>
			<td>For all buffers</td>
		</tr>
		<tr>
			<td>O2 tank</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>OMNI THQ Homogenizer</td>
			<td>OMNI International&#160;</td>
			<td>12-500</td>
			<td>Similar rotor stator homogenizers will work</td>
		</tr>
		<tr>
			<td>pipettes</td>
			<td></td>
			<td></td>
			<td>Volumes of&#160; 2&#8211;20 &#181;L; 20&#8211;200 &#181;L; 200&#8211;1000 &#181;L</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma&#160;</td>
			<td>P3911</td>
			<td>For buffers (RB and CB)</td>
		</tr>
		<tr>
			<td>Potassium phosphate dibasic</td>
			<td>Sigma&#160;</td>
			<td>795496</td>
			<td>For buffers (IB and RB)</td>
		</tr>
		<tr>
			<td>Protease from Bacillus licheniformis</td>
			<td>Sigma&#160;</td>
			<td>P5459</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma&#160;</td>
			<td>S9888</td>
			<td>For buffer (CB)</td>
		</tr>
		<tr>
			<td>Sodium pyruvate</td>
			<td>Fisher bioreagents</td>
			<td>BP356-100</td>
			<td>Respirometry assays</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma&#160;</td>
			<td>8510-OP</td>
			<td>For buffer (IB)</td>
		</tr>
		<tr>
			<td>Surgical dissection kit</td>
			<td></td>
			<td></td>
			<td>Depends on animal and tissue source</td>
		</tr>
		<tr>
			<td>Tabletop centrifuge</td>
			<td></td>
			<td></td>
			<td>Must cool to 4 &#176;C&#160;</td>
		</tr>
	</tbody>
</table>

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.

Upon completion of mitochondrial isolation, the quality and functionality of the isolates should be tested via quantifying rates of oxygen consumption (JO2) using high-resolution respirometry. To do so, mitochondrial stocks were diluted to 40 mg/mL to allow for working concentrations of 0.1 mg/mL in 2 mL of RB for all respirometry assays using isolated cardiac mitochondria. Respiration was fueled by 5 mM sodium pyruvate and 1 mM L-malate in the presence of 1 mM EGTA, a calcium chelator, and was allowe...

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.

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 ...

robust-mitochondrial-isolation-from-rodent-cardiac-tissue

Research

JoVE Journal

Biochemistry

508 Views.  Michigan State University. 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.

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

Isolation of Cognate RNA-protein Complexes from Cells Using Oligonucleotide-directed Elution

Ribonucleoprotein particles direct the biogenesis and post-transcriptional regulation of all mRNAs through distinct combinations of RNA binding proteins. They are composed of position-dependent, cis-acting RNA elements and unique combinations of RNA binding proteins. Defining the composition of a specific RNP is essential to achieving a fundamental understanding of gene regulation. The isolation of a select RNP is akin to finding a needle in a haystack. Here, we demonstrate an approach to isolate RNPs associated at the 5' untranslated region of a select mRNA in asynchronous, transfected cells. This cognate RNP has been demonstrated to be necessary for the translation of select viruses and cellular stress-response genes.
The demonstrated RNA-protein co-precipitation protocol is suitable for the downstream analysis of protein components through proteomic analyses, immunoblots, or suitable biochemical identification assays. This experimental protocol demonstrates that DHX9/RNA helicase A is enriched at the 5' terminus of cognate retroviral RNA and provides preliminary information for the identification of its association with cell stress-associated huR and junD cognate mRNAs.

This manuscript describes an approach to isolate select cognate RNPs formed in eukaryotic cells via a specific oligonucleotide-directed enrichment. We demonstrate the applicability of this approach by isolating a cognate RNP bound to the retroviral 5' untranslated region that is composed of DHX9/RNA helicase A.

Ribonucleoprotein particles direct the biogenesis and post-transcriptional regulation of all mRNAs through distinct combinations of RNA binding proteins. ...

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 ...

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 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 ...

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 ...

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human organs, possess highly conserved metabolic pathways and have a relatively short lifespan that allows the study of different fundamental mechanisms in a short period of time. It is, however, surprising that one of the mechanisms essential for cellular metabolism, the mitochondrial respiration, has not been thoroughly investigated in this model. It is likely because the measure of the mitochondrial respiration in Drosophila usually requires a very large number of individuals and the results obtained are not highly reproducible. Here, a method allowing the precise measurement of mitochondrial oxygen consumption using minimal amounts of tissue from Drosophila is described. In this method, the thoraxes are dissected and permeabilized both mechanically with sharp forceps and chemically with saponin, allowing different compounds to cross the cell membrane and modulate the mitochondrial respiration. After permeabilization, a protocol is performed to evaluate the capacity of the different complexes of the electron transport system (ETS) to oxidize different substrates, as well as their response to an uncoupler and to several inhibitors. This method presents many advantages compared to methods using mitochondrial isolations, as it is more physiologically relevant because the mitochondria are still interacting with the other cellular components and the mitochondrial morphology is conserved. Moreover, sample preparations are faster, and the results obtained are highly reproducible. By combining the advantages of Drosophila as a model for the study of metabolism with the evaluation of mitochondrial respiration, important new insights can be unveiled, especially when the flies are experiencing different environmental or pathophysiological conditions.

In this paper, a method to measure oxygen consumption using high-resolution respirometry in permeabilized thoraxes of Drosophila is described. This technique requires a minimal amount of tissue compared to the classic mitochondrial isolation technique and the results obtained are more physiologically relevant.

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human ...

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 ...

Isolation of F1-ATPase from the Parasitic Protist Trypanosoma brucei

F1-ATPase is a membrane-extrinsic catalytic subcomplex of F-type ATP synthase, an enzyme that uses the proton motive force across biological membranes to produce adenosine triphosphate (ATP). The isolation of the intact F1-ATPase from its native source is an essential prerequisite to characterize the enzyme&#39;s protein composition, kinetic parameters, and sensitivity to inhibitors. A highly pure and homogeneous F1-ATPase can be used for structural studies, which provide insight into molecular mechanisms of ATP synthesis and hydrolysis. This article describes a procedure for the purification of the F1-ATPase from Trypanosoma brucei, the causative agent of African trypanosomiases. The F1-ATPase is isolated from mitochondrial vesicles, which are obtained by hypotonic lysis from in vitro cultured trypanosomes. The vesicles are mechanically fragmented by sonication and the F1-ATPase is released from the inner mitochondrial membrane by the chloroform extraction. The enzymatic complex is further purified by consecutive anion exchange and size-exclusion chromatography. Sensitive mass spectrometry techniques showed that the purified complex is devoid of virtually any protein contaminants and, therefore, represents suitable material for structure determination by X-ray crystallography or cryo-electron microscopy. The isolated F1-ATPase exhibits ATP hydrolytic activity, which can be inhibited fully by sodium azide, a potent inhibitor of F-type ATP synthases. The purified complex remains stable and active for at least three days at room temperature. Precipitation by ammonium sulfate is used for long-term storage. Similar procedures have been used for the purification of F1-ATPases from mammalian and plant tissues, yeasts, or bacteria. Thus, the presented protocol can serve as a guideline for the F1-ATPase isolation from other organisms.

Isolation of F1-ATPase from the Parasitic Protist Trypanosoma brucei

This protocol describes the purification of F1-ATPase from the cultured insect stage of Trypanosoma brucei. The procedure yields a highly pure, homogeneous, and active complex suitable for structural and enzymatic studies.

F1-ATPase is a membrane-extrinsic catalytic subcomplex of F-type ATP synthase, an enzyme that uses the proton motive force across biological membranes to ...

Measuring Liver Mitochondrial Oxygen Consumption and Proton Leak Kinetics to Estimate Mitochondrial Respiration in Holstein Dairy Cattle

Oxygen consumption, proton motive force (PMF) and proton leak are measurements of mitochondrial respiration, or how well mitochondria are able to convert NADH and FADH into ATP. Since mitochondria are also the primary site for oxygen use and nutrient oxidation to carbon dioxide and water, how efficiently they use oxygen and produce ATP directly relates to the efficiency of nutrient metabolism, nutrient requirements of the animal, and health of the animal. The purpose of this method is to examine mitochondrial respiration, which can be used to examine the effects of different drugs, diets and environmental effects on mitochondrial metabolism. Results include oxygen consumption measured as proton dependent respiration (State 3) and proton leak dependent respiration (State 4). The ratio of State 3 / State 4 respiration is defined as respiratory control ratio (RCR) and can represent mitochondrial energetic efficiency. Mitochondrial proton leak is a process that allows dissipation of mitochondrial membrane potential (MMP) by uncoupling oxidative phosphorylation from ADP decreasing the efficiency of ATP synthesis. Oxygen and TRMP+ sensitive electrodes with mitochondrial substrates and electron transport chain inhibitors are used to measure State 3 and State 4 respiration, mitochondrial membrane PMF (or the potential to produce ATP) and proton leak. Limitations to this method are that liver tissue must be as fresh as possible and all biopsies and assays must be performed in less than 10 h. This limits the number of samples that can be collected and processed by a single person in a day to approximately 5. However, only 1 g of liver tissue is needed, so in large animals, such as dairy cattle, the amount of sample needed is small relative to liver size and there is little recovery time needed.

Here, we share methods for measuring mitochondrial oxygen consumption, a defining concept of nutritional energetics, and proton leak, the primary cause of inefficiency in mitochondrial generation of ATP. These results can account for 30% of the energy lost in nutrient utilization to help evaluate mitochondrial function.

Oxygen consumption, proton motive force (PMF) and proton leak are measurements of mitochondrial respiration, or how well mitochondria are able to convert ...

Isolation of Tissue Extracellular Vesicles from the Liver

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in cell-to-cell communication by shuttling bioactive molecules such as RNA or protein from one cell to another. Most studies of EVs have been performed in cell culture models or in EVs isolated from body fluids. There is emerging interest in the isolation of EVs from tissues to study their contribution to physiological processes and how they are altered in disease. The isolation of EVs with sufficient yield from tissues is technically challenging because of the need for tissue dissociation without cellular damage. This method describes a procedure for the isolation of EVs from mouse liver tissue. The method involves a two-step process starting with in situ collagenase digestion followed by differential ultra-centrifugation. Tissue perfusion using collagenase provides an advantage over mechanical cutting or homogenization of liver tissue due to its increased yield of obtained EVs. The use of this two-step process to isolate EVs from the liver will be useful for the study of tissue EVs.

This is a protocol to isolate tissue extracellular vesicles (EVs) from the liver. The protocol describes a two-step process involving collagenase perfusion followed by differential ultracentrifugation to isolate liver tissue EVs.

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in ...

Characterization at the Molecular Level using Robust Biochemical Approaches of a New Kinase Protein

Extensive whole genome sequencing has identified many Open Reading Frames (ORFs) providing many potential proteins. These proteins may have important roles for the cell and may unravel new cellular processes. Among proteins, kinases are major actors as they belong to cell signaling pathways and have the ability to switch on or off many processes crucial to the fate of the cell, such as cell growth, division, differentiation, motility, and death.
In this study, we focused on a new potential kinase protein, LIMK2-1. We demonstrated its existence by Western Blot using a specific antibody. We evaluated its interaction with an upstream regulating protein using coimmunoprecipitation experiments. Coimmunoprecipitation is a very powerful technique able to detect the interaction between two target proteins. It may also be used to detect new partners of a bait protein. The bait protein may be purified either via a tag engineered to its sequence or via an antibody specifically targeting it. These protein complexes may then be separated by SDS-PAGE (Sodium Dodecyl Sulfate PolyAcrylamide Gel) and identified using mass spectrometry. Immunoprecipitated LIMK2-1 was also used to test its kinase activity in vitro by &#947;[32P] ATP labeling. This well-established assay may use many different substrates, and mutated versions of the bait may be used to assess the role of specific residues. The effects of pharmacological agents may also be evaluated since this technique is both highly sensitive and quantitative. Nonetheless, radioactivity handling requires particular caution. Kinase activity may also be assessed with specific antibodies targeting the phospho group of the modified amino acid. These kinds of antibodies are not commercially available for all the phospho modified residues.

We characterized a new kinase protein using robust biochemical approaches: Western Blot analysis with a dedicated specific antibody on different cell lines and tissues, interactions by coimmunoprecipitation experiments, kinase activity detected by Western Blot using a phospho-specific antibody and by &#947;[32P] ATP labeling.

Extensive whole genome sequencing has identified many Open Reading Frames (ORFs) providing many potential proteins. These proteins may have important ...