This work was supported by the Swedish Foundation for Strategic Research (Future Leaders Grant FFL15:0325), Ragnar S&#246;derberg Foundation (Fellowship in Medicine M44/16), Swedish Research Council (Starting Grant NT&#215;2015-04107), FEBS Long-Term Fellowship (SA), Horizon 2020 - Marie Sklodowska-Curie Innovative Training Network (721757) (VS).

All mammalian cell culture work must be performed inside a biological safety cabinet. Use sterile equipment if contact with cells. Wear nitrile gloves and a lab coat and follow good tissue culture practice.
1. Cell Culture
<ol>
	<li>Maintain HEK293S suspension cells in Dulbecco&#8217;s Modified Eagle Medium (DMEM) supplemented with 10% tetracycline-free fetal bovine serum (FBS), 5 &#956;g/mL blasticidin and 200 &#956;g/mL Azithromycin, at 37 &#176;C and 5% CO2.</li>
	<li>Scale up to 9 x T175 flasks. At 90% confluency, harvest cells and spin them down at 500 x g for 5 min. Resuspend the pelleted cells in Freestyl supplemented with 5% tetracycline-free FBS. Count cells using an automated cell counter and adjust cell concentration to 1.5 x 106 cells/mL in a vented shaking flask. 
	Note: This will typically correspond to a starting volume of 300 mL in a 1,000 mL vented flask.</li>
	<li>Incubate cells in a shaking incubator at 37 &#176;C and 5% CO2 at 120 rpm (50 mm shaking diameter).</li>
	<li>After 2 days count the cells and proceed to split the cells if the cell density is above 3.0 x 106 cells/mL. Split the cells by spinning down 2 x 150 mL cell culture in 2 x 250 mL conical bottles at 500 g for 5 min. Resuspend cells in 2 x 10 mL fresh media and transfer to a 2 x 1, 000 mL vented flasks containing the appropriate volume for a final cell density of 1.5 x 106 cell/mL, e.g. 2 x 300 mL from a starting cell density of 3.0 x 106 cells/mL.</li>
	<li>Incubate cells in a shaking incubator at 37 &#176;C and 5% CO2 at 120 rpm (50 mm shaking diameter).</li>
	<li>After two days count the cells and proceed to split the cells if the cell density is above 3.0 x 106 cells/mL. Split the cells by spinning down 4 x 150 mL cell culture in 4 x 250 mL conical bottles at 500 g for 5 min. Resuspend cells in 2 x 10 mL fresh media and transfer to 2 x 2,000 mL vented flask containing the appropriate volume for a final cell density of 1.5 x 106 cell/mL, e.g. 2 x 700 mL from a starting cell density of 3.0 x 106 cells/mL.</li>
	<li>Incubate cells in a shaking incubator at 37 &#176;C and 5% CO2 at 120 rpm (50 mm shaking diameter).</li>
	<li>After two days count the cells and proceed to split the cells if the cell density is above 3.0 x 106 cells/mL. Split the cells by pouring the 2 x 700 mL cell culture into 2 x 2800 mL vented flasks and topping up with 2 x 300 mL fresh media to reach a volume of 2 x 1 L.</li>
	<li>Incubate cells in a shaking incubator at 37&#176;C and 5% CO2 at 120 rpm (50 mm shaking diameter).</li>
	<li>After 24 h count the cells and harvest the cells if the cell density is between 3.0 - 4.0 x 106 cells/mL.</li>
</ol>
2. Mitochondria Isolation
<ol>
	<li>Buffers required 
	Note: The quantities are given for a preparation from 2 L cells. Prepare the following&#160;stock solutions&#160;ahead of time for mitochondrial isolation buffer (MIB), sucrose/mannitol buffer (SM4), experimental buffer (MIBSM), resuspension buffer (SEM).
	<ol>
		<li>Make 0.5L MIB buffer: 50mM HEPES-KOH, pH 7.5, 10 mM KCl, 1.5mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1mM DTT, protease inhibitors.</li>
		<li>Make 100 mL SM4 buffer: 280 mM sucrose, 840 mM mannitol, 50 mM HEPES-KOH, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1m M DTT, protease inhibitors.</li>
		<li>Make 160 mL MIBSM buffer: 120 mL MIB buffer + 40 mL SM4 buffer.</li>
		<li>Store 200 mL PBS at 4 &#176;C.</li>
		<li>Make 5 mL SEM buffer: 250 mM sucrose, 20 mM HEPES-KOH, pH 7.5, 1 mM EDTA.</li>
		<li>Make 4 x 10 mL stock solutions for the stepwise sucrose gradient containing 20 mM HEPES-KOH, pH 7.5, 1 mM EDTA and 60% / 32% /23% and 15% sucrose, respectively.</li>
	</ol>
	</li>
	<li>Mitochondria isolation 
	Note: It is important to work quickly and keep everything on ice throughout the procedure.</li>
	<li>Precool the nitrogen cavitation chamber prior to use.
	<ol>
		<li>Harvest 2 x 1 L cells (3-4 x 109 cells per centrifuge tube) by centrifugation at 1,000 x g for 7 min, 4 &#176;C.</li>
		<li>Decant the supernatant carefully and resuspend the pelleted cells quickly in 2 x 100 mL PBS. Pool the cells.</li>
		<li>Centrifuge the resuspended cells at 1,200 x g for 10 min, 4 &#176;C.</li>
		<li>Decant the supernatant carefully and weigh the pellet (~20 g).</li>
		<li>Resuspend the pellet in 120 mL MIB buffer.</li>
		<li>Allow cells to swell by gently stirring in a cold room for 10 min.</li>
		<li>Place the nitrogen cavitation chamber on the ice and transfer the swelled cells to the nitrogen cavitation chamber. Add 45 mL SM4 buffer (1/3 of the final volume of resuspended cells, about 140 mL, yielding a final concentration of 70 mM sucrose and 210 mM mannitol).</li>
		<li>Fasten the nitrogen cavitation chamber and fill with nitrogen until the pressure reaches 500 psi. Close taps and keep isolated on ice for 20 min.</li>
		<li>Slowly release the pressure in the nitrogen cavitation chamber and collect the lysate (approximately 185 mL).</li>
		<li>Centrifuge the lysed material to remove the cell debris and nuclei at 800 x g for 15 min, 4 &#176;C.</li>
		<li>Collect the supernatant by pouring it through the cheesecloth into a beaker kept on ice. Do not discard the pellet.</li>
		<li>Resuspend the pellet in 90 mL MIBSM buffer (1/2 the previous volume). Homogenize using Teflon/glass Dounce homogenizer and repeat the centrifugation at 800 x g for 15 min, 4 &#176;C.</li>
		<li>Collect the supernatant by pouring it through the cheesecloth into a beaker kept on ice. Combine this supernatant with the supernatant from step 2.2.11.</li>
		<li>Centrifuge the combined supernatants at 1,000 x g for 15 min, 4 &#176;C.</li>
		<li>Collect the supernatant by pouring it through the cheesecloth into a beaker kept on ice.</li>
		<li>Discard the pellet and centrifuge the supernatant containing crude mitochondria at 10,000 x g for 15 min, 4 &#176;C. 
		Note: The pellet will typically consist of two parts: loose and tight.</li>
		<li>Carefully wash out the loose pellet without disturbing the tight portion. Resuspend the tight pellet in 10 mL MIBSM buffer.</li>
		<li>Perform a protein concentration assay using a commercial Protein Assay Kit or similar method. Typical concentration yields from 2 L starting culture are ~2 mg/mL.</li>
		<li>Add 200 Units of RNase free DNase and leave to rotate on a roller in the cold room for 20 min to evenly mix the DNase for removal of the genomic DNA.</li>
		<li>Centrifuge at 10,000 x g for 15 min, 4 &#176;C and resuspend the pellet in 2 mL SEM buffer. Homogenize gently using a small Teflon/glass Dounce homogenizer to resuspend any remaining aggregates. Perform not more than five up-and-down passes to avoid breakage. Keep on ice.</li>
		<li>Prepare the sucrose gradient in 14 mL SW40 tubes by carefully pipetting 1.5 mL of 60% sucrose stock buffer into the bottom of the tube. Carefully add 4.5 mL of the 32% sucrose stock buffer on top of the 60% band without disturbing it. Repeat with 1.5 mL of 23% sucrose stock buffer and again with 1.5 mL of 15% sucrose stock buffer.</li>
		<li>Load the entire mitochondrial suspension (approximately 3 mL) on top of the sucrose gradient.</li>
		<li>Centrifuge in SW40 rotor at 139, 065 x g for 60 min.</li>
		<li>Carefully collect the brown band migrating to the interface of 32% and 60% sucrose using a transfer pipette, typically 2-3 mL.</li>
		<li>Snap-freeze the purified mitochondria in liquid nitrogen and store at -80 &#176;C.&#160;&#160;&#160;</li>
	</ol>
	</li>
</ol>
3. Mitoribosome Preparation
<ol>
	<li>Buffers required 
	Note: Prepare the following&#160;stock solutions&#160;ahead of time for the lysis buffer, sucrose cushion/gradient, and resuspension buffer.
	<ol>
		<li>Make 10 mL lysis buffer: 25 mM HEPES-KOH, pH 7.5, 150 mM KCl, 50mM MgOAc, 2% Polyethylene glycol octylphenyl ether, 2 mM DTT, protease inhibitors.</li>
		<li>Make 10 mL sucrose cushion buffer: 1 M sucrose (34% w/v), 20 mM HEPES-KOH, pH 7.5, 100 mM KCl, 20 mM MgOAc, 1% Polyethylene glycol octylphenyl ether, 2 mM DTT.</li>
		<li>Make 5 mL resuspension buffer: 20 mM HEPES-KOH, pH 7.5, 100 mM KCl, 20mM MgOAc, 2 mM DTT</li>
		<li>Make 15%-30% linear sucrose gradients with resuspension buffer in TLS-55 polycarbonate tubes.</li>
	</ol>
	</li>
	<li>&#160;Mitoribosome purification
	<ol>
		<li>Defrost the frozen mitochondria on ice.</li>
		<li>Add 2 volumes of the lysis buffer,e.g. add 6 mL lysis buffer to 3 mL mitochondria. Mix immediately by inverting the tube several times.</li>
		<li>Homogenize with a small Teflon/glass dounce homogenizer to assist the lysis and incubate for 5-10 min on ice to complete the lysis.</li>
		<li>Centrifuge the lysed material (approximately 9 mL) at 30,000 x g for 20 min, 4 &#176;C to remove the insoluble material. Decant the supernatant carefully from the pellet and discard the pellet.</li>
		<li>Repeat the centrifugation at 30,000 x g for 20 min at 4 &#176;C to ensure clarification of the supernatant. Decant the supernatant carefully from the pellet and discard the pellet.</li>
		<li>Prepare the sucrose cushion in TLA 120.2 tubes (ultra-clear): 0.4 mL sucrose cushion per tube. Prepare one tube per mL of lysed material.</li>
		<li>Layer lysed mitochondria on the sucrose cushions: approximately 1 mL per tube, resulting in a lysate:cushion ratio of 2.5:1.</li>
		<li>Centrifuge the sample at 231, 550 x g for 45 min in TLA120.2 rotor at 4 &#176;C.</li>
		<li>Discard the supernatant and rinse the tubes sequentially with 100 &#181;l of the resuspension buffer to remove residual sucrose.</li>
		<li>Resuspend the pellets in total 100 &#181;l resuspension buffer.</li>
		<li>Vortex on slow speed for 30 s to dissolve the remaining aggregates and centrifuge at 17,949 x g for 10 min in microtube centrifuge at 4 &#176;C.</li>
		<li>Carefully collect the supernatant and repeat the centrifugation.</li>
		<li>Measure mitoribosome absorption at A260. 
		Note: The typical yield is 7 A260, with A260: A280 ratio of 1.3.</li>
		<li>Load the entire sample onto a single linear sucrose gradient tube. Centrifuge in TLS-55 rotor at 213, 626 x g for 120 min at 4 &#176;C.</li>
		<li>Fractionate the gradient, determine the optical density at A260 and pool the fraction corresponding to the nucleic acid peak together. 
		Note: The typical A260: A280 ratio of the peak is &#62;1.6.</li>
		<li>Exchange the buffer if necessary, using a method of choice. Calculate the final concentration by using the conversion 1 A260 = 0.1 mg/mL.&#160;&#160;</li>
		<li>Snap freeze the purified mitoribosome sample in the resuspension buffer and store at -80 &#176;C.</li>
	</ol>
	</li>
</ol>

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<li>Brown, A., Amunts, A., Bai, X. C., Sugimoto, Y., Edwards, P. C., Murshudov, G., Ramakrishnan, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Structure+of+the+large+ribosomal+subunit+from+human+mitochondria%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Structure of the large ribosomal subunit from human mitochondria.</a> Science. 346 (6210), 718-722 (2014).</li>
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<li>Rice, J. E., Lindsay, J. G. Subcellular fractionation of mitochondria. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aRice%2C+JE+%22Subcellular+Fractionation%3A+A+Practical+Approach%22">Subcellular Fractionation: A Practical Approach</a>. Grahamand, J. M., Rickwood, D. , IRL Press. Oxford. 107-142 (1997).</li>
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Regarding the source of starting material, although the relatively easy availability of animal tissues that are known to be a rich source of mitochondria, makes it a popular choice for mitoribosomes3,5,12,13, they cannot be genetically modified and reproduced in the lab easily. Hence, there is a clear practical need for developing protocols involving homogeneously cultured human cell lines. The major difference between the protocols dealing with tissues and cultured cells is the mode of lysis and homogenization. For cultured cells grown as a monolayer, a typical method is Teflon/glass dounce homogenization14. The original preparation protocols while efficient were developed for small scales15. A direct scaling up employing a homogenizer with the capacity of 500 mL is possible11, however, it requires ~2 h of manual labor to achieve 80% lysis. This introduced at least three issues: aggregation of organelles due to long waiting times, non-homogeneous lysis due to heavy material sinking to the bottom of a large vessel, heating of the sample due to the necessity of multiple strokes. Therefore, a preferable method of lysis utilizes nitrogen cavitation, which is based on decompression from a pressurized vessel16,17. In this method, cells are first swelled in the cold room in order to soften the cells and make them more susceptible to lysis. As they are placed in the nitrogen cavitation device a buffer containing sucrose and mannitol is added in order to maintain an osmotic pressure that will help keep mitochondria intact. The nitrogen cavitation device is then pressurized with a large volume of oxygen-free nitrogen, which dissolves into the cells. As the pressure is released nitrogen bubbles out of solution resulting in rupturing of the cell membranes. This method offers several advantages over mechanical homogenizing methods involving shear stresses and friction as follows: 1) any external physical stress on the cells is avoided; 2) an adiabatic expansion that cools the sample, ensures no heat damage to organelles; 3) cell components are protected from oxidation by the inert nitrogen gas; 4) no pH alteration of the suspending medium; 5) the process is uniform and reproducible because the same disruptive forces are applied within each cell and throughout the sample; 6) the process is fast and can be completed within 20-30 min.
Isolation of mitochondria from cultured cells and tissues has been described in the literature extensively, and it is based on a gentle cell disruption followed by a series of differential centrifugations. Most of the currently used protocols follow the original procedures developed in the middle of the previous century18. While the basic biochemical approaches are correct, there are several misconceptions that are highlighted in the literature and remained undetected. To optimize the protocol for mitoribosomes, we systematically investigated the reported general principles and conclude that: 1) Inclusion of Mg2+ and K+ in the buffer is not crucial. It has been argued that the KCl helps to prevent cytoplasmic proteins from forming a gel19, however, provided a sufficient dilution as described in our protocol, this phenomenon does not occur. Furthermore, exclusion of Mg2+ is useful to reduce contaminations by cytoplasmic ribosomes20; 2) There is no need to keep maximal possible volume ratio of cells to medium to protect the released organelles from the hypo-osmotic environment. The osmotic support in our protocol is sufficiently provided by sucrose-containing buffers, and the dilution of cells with mitochondria isolation buffer (step 2.II.5) is critical for the efficient separation of organelles in a large-scale preparation.
The pH of mitochondria isolation buffer is close to physiological values, i.e. 7.5, which is also the optimal pH for the following mitoribosome preparation. Binding agents EDTA and EGTA are added to the isolation media to chelate contaminating ions, and they chelate free magnesium and calcium, respectively. It has been discussed that inclusion of EDTA can lead to the damage of the inner mitochondrial membrane14, therefore the concentration is limited to 1 mM as a precaution. We have not observed any difference when using 5 mM EDTA in the final purification step of sucrose density gradient.
The protocol described herein uses HEK293S-derived human cells for purification of the mitoribosome. The quality of the preparation allows the obtained sample to be investigated biochemically and structurally to atomic resolution. This allows one to apply the method to human mitoribosomal translating complexes, quality control assemblies, and mitoribosomal subunits intermediates. Moreover, since cancer cells have amplified OXPHOS capacity and elevated mitochondrial protein translation compared with adjacent stromal tissue21, mitoribosomes are established drug targets for cancer22. Therefore, using this protocol for specific purification of mitoribosomes in presence of inhibitors will have medical applications. Also, mitoribosomal mutations have been linked to hereditary mitochondrial diseases23. Since these mutations have direct effects on the structure, the approach presented here will be useful for their structural characterization. The protocol can be experimentally expanded and applied to a variety of scientific questions tackling the fundamental understanding of translation in human mitochondria and its medical importance.

The process of mitochondrial protein synthesis is based on 13 essential mt-mRNAs that are translated by a specialized membrane-attached mitoribosome to form the catalytic core of the respiratory chain. The altered mitochondrial genomes and the need for the proteins to insert co-translationally into the membrane have substantially shaped the architecture of the mitochondrial ribosomes1. Recent high-resolution structures of the mammalian mitoribosome showed a strikingly different overall appearance to the bacterial counterpart2,3. Particularly, at least 36 mitochondria-specific proteins are added, contributing ~1 MDa extra molecular mass, while mt-rRNA is reduced by twofold and highly diverged. The structural rearrangements alter almost all critical functional features that were previously generally accepted to be universally conserved4.
New principal elements have been acquired by each one of the mitoribosomal subunits, for example, the small subunit has incorporated an intrinsic GTPase protein mS29 into its &#39;head&#39; region. GTPase activity has not been found in other translation systems, andthe structure indicates that the GTPase might have a role in subunit assembly2. The 5S rRNA that was believed to be a landmark of all known ribosomal large subunits, forming the core of the central protuberance, is missing in the mammalian mitoribosome and has adopted mt-tRNA-Val as an integral building block instead2. Ban and colleagues showed that the porcine mitoribosome has mt-tRNA-Phe and not &#8211;Val5. Chrzanowska-Lightowlers, Minczuk, and colleagues followed up on these data and found that mitoribosomes from patients with compromised mt-tRNA-Val stability can in principle accommodate mt-tRNA-Phe6,7. Why the mitoribosomes have incorporated these specific elements and what pathways and trans-factors are required for these unique assemblies remain unknown.
Overall, the high complexity of the human mitoribosome, new protein components, and the unique association of the mt-tRNA as a structural element imply the involvement of as-yet-unknown mitochondria-specific trans-factors. However, because many of the features of this system are unique to mitochondria, which have traditionally been difficult to investigate8, little is known about the molecular and quality control mechanism. With the development of high-resolution single particle analysis by electron cryo-microscopy (cryo-EM)20, opportunities now arise to comprehensively study the molecular mechanisms underlying the assembly, action and quality control of the human mitoribosome. Our report of the first structure of the human mitoribosome assembly intermediate provides the recognition that it is possible to visualize how the human mitoribosome is formed and shows that cryo-EM is instrumental in the identification of new trans-acting assembly factors9.
To expand on this initial effort, we describe a rapid protocol for human mitoribosome purification in detail. In the first part of the protocol, a large-scale isolation of highly pure intact mitochondria from suspension cells is described. This procedure requires 9 h and can be easily modified and adapted to different cell types and scales. A significant step in this protocol is the use of nitrogen cavitation to break open the cells. The second part of the protocol was developed to purify mitoribosomes. This procedure requires 7 h and yields a sufficient amount of mitoribosomes for biochemical and structural studies. Use of pure mitochondria as a starting material offers high-quality final preparations and can be extrapolated to other mitochondrial macromolecules.

<table><tbody><tr><td>Dulbecco&amp;rsquo;s Modified Eagle Medium (DMEM) high glucose, GLUTAMAX supplement, pyruvate</td><td>Thermo Scientific</td><td>31966-021</td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Thermo Scientific</td><td>16000-044</td><td></td></tr><tr><td>Blasticidin S HCl</td><td>Thermo Scientific</td><td>R210-01</td><td></td></tr><tr><td>Zeocin selection reagent</td><td>Thermo Scientific</td><td>R25001</td><td>100 mg/ml</td></tr><tr><td>Freestyle 293 Expression medium</td><td>Thermo Scientific</td><td>12338026</td><td></td></tr><tr><td>T175 tissue culture flask with vented cap</td><td>Sarstedt</td><td>83.3912.002</td><td></td></tr><tr><td>Shaker flasks with vented cap</td><td>Thermo Scientific</td><td>4115-0500, 4115-1000, 4115-2000, 4115-2800</td><td></td></tr><tr><td>250 ml conical bottle tubes, sterile</td><td>Corning</td><td>430776</td><td></td></tr><tr><td>Eve automated cell counter</td><td>NanoEnTek</td><td>E1000</td><td></td></tr><tr><td>Nitrogen cavitation cell disruption vessel</td><td>Parr Instruments</td><td>4635, 4639</td><td>safety volume: 40ml, 600ml respectively</td></tr><tr><td>Dnase (RNA-free)</td><td>HT Biotechnology</td><td>N401a</td><td></td></tr><tr><td>Teflon/glass dounce homogenizers</td><td>Cambridge Glassblowing Limited</td><td></td><td>Size designed upon request</td></tr><tr><td>SW40 tubes for mitochondria gradient</td><td>Beckman</td><td>344060</td><td>Polypropylene, thin wall</td></tr><tr><td>Transfer pipettes</td><td>Sarstedt</td><td>86.1171</td><td></td></tr><tr><td>TLA 120.2 tubes for cushion</td><td>Beckman</td><td>343778</td><td>Polycarbonate, thick wall</td></tr><tr><td>TLS-55 tubes for gradient</td><td>Beckman</td><td>347356</td><td>Ultra-clear</td></tr><tr><td>Gradient Station IP</td><td>BioComp</td><td>153-002</td><td></td></tr></tbody></table>

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.

Dividing and highly viable cells is an essential starting point for purification of active mitoribosomes. This protocol is applicable to any HEK293 suspension cells. We are using in-house cell line T501, which is stably expressing a transporter under tetracycline-inducible control. The parental cell line is HEK293S-GnTI- cells (Table of Materials)10. During the cell growth and expansion in the FreeStyle 293 Expression Medium the minimal density should be kept at 1.5 x 106 cell/mL, in order to ensure a doubling rate of every two days, whereas the maximal cell density should not exceed ~5 x 106 cell/mL. Upon reaching a cell density above 3 x 106 cell/mL the cells are pelleted and resuspended in a fresh, preheated media in an expanded volume to achieve the minimum cell density of 1.5 x 106 cell/mL. This splitting is performed repeatedly every 2-3 days until a desired cell mass for the procedure is achieved. The final cell density can vary in the range of 3-4.5 x 106 cell/mL, and at least 2 L is required as a starting point for the isolation of mitochondria. The cell viability should be generally maintained &#62;90%, and for the final culture that is harvested &#62;95%. Treatment of the large-scale cell culture with antibiotics is not recommended.
After the series of differential centrifugations, the volume of the mitochondrial suspension after step 2.2.20 is typically in the range of 3-5 mL. The mitochondria are then separated on the sucrose gradient (Figure 1) from other organelles. The stepwise sucrose gradient is prepared such that the volume of the gradient and the volume of the mitochondrial suspension together fill up the centrifugation tube to its maximal volume. Here special care is needed to collect the brown band migrating to the 60% / 32% interface with minimal contamination from the surrounding buffer. This is important in order to keep constant protein:detergent ratio in the following step of mitochondria solubilization. It is advised to assess the mitochondrial protein concentration at this stage, and a typical yield of 15-20 mg total mitochondrial protein is expected from ~1010 cultured HEK cells.
Upon successful mitochondria isolation, they are lysed by the addition of polyethylene glycol octylphenyl ether, and mitoribosomes are separated through a sucrose cushion (Figure 2). To separate mitoribosomes from the hydrophobic membranous substance, pellets are resuspended in the buffer not containing detergent, and hydrophobic complexes are pelleted by centrifugation. This procedure is repeated, and the purification degree of mitoribosomes is quantified by A260/A280 ratio, which is expected to be ~1.3. Typical yield is 7 A260 from a 2 L starting culture. This mitoribosomal fraction also contains additional large soluble mitochondrial complexes, such as pyruvate dehydrogenase and glutamate dehydrogenase. To separate mitoribosomes from the soluble mitochondrial complexes, the supernatant is applied to a sucrose density gradient. Fractions containing the mitoribosome are generally located at the bottom third of the tube. The fractionation of the sucrose gradient can then be done either with an automated piston or manually by carefully taking 50 &#181;L fractions with a pipette or by punching the tube bottom with a 21 G needle and collecting the drops.
Two major mitoribosomal populations are identified in the gradient: monosome 55S and large subunit 39S, as illustrated in Figure 3 and Figure 4. The presence of the large subunit fraction suggests that cells are harvested in a highly active dividing state11. The ratio between the monosome and large subunit peaks may change. An additional peak located closer to the bottom may appear in preparations with contaminating cytoplasmic ribosomes 80S. Please note that the small sucrose gradient using the swinging bucket rotor TLS-55 allows for the rapid purification of ~1 mL of mitoribosomes at an optical density of 0.4-1 A260. The separation of the monosome and large subunit peaks may vary slightly depending on the fractionation, but there is usually some overlap of the two peaks. Hence, which fractions to collect, and pool should be taken into consideration in order to ensure the highest proportion of monosome, or alternatively large subunit, in the sample. For high-resolution cryo-EM studies, the separation between the two mitoribosomal populations is not absolutely required (due to additional in silico operations). However, if better separation is necessary, it is recommended to use larger tubes and corresponding running times.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57877/57877fig1.jpg" /> 
Figure 1: Purification of mitochondria on a sucrose gradient. Subcellular organelles from a 2 L starting culture were fractionated through a series of differential centrifugations as described in the protocol, and mitochondria were separated on a discontinuous sucrose gradient. The purified mitochondria are found in the lower band at the 32%/60% interface.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57877/57877fig2.jpg" /> 
Figure 2: Purification of mitoribosomes on a sucrose cushion. Crude mitoribosomes from a 2 L starting culture are sedimented through 1 M sucrose cushion. The pellets are resuspended in a detergent-free buffer, and mitoribosomes are clarified by two centrifugations as described in the protocol. The absorbance is recorded to assess the quality of the preparation and typical A260/A280 ratio of ~1.3 (right panel) certifies a mitoribosome rich fraction. <a href="/files/ftp_upload/57877/57877fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57877/57877fig3.jpg" /> 
Figure 3: Fine purification of mitoribosomes on a sucrose gradient. Absorbance trace from a 2 L starting culture. Fractions are numbered from the top to the bottom of the gradient. Two major mitoribosomal populations are identified: large subunit (peak 1) and monosome 55S (peak 2). The ratio between the populations may change. <a href="/files/ftp_upload/57877/57877fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57877/57877fig4.jpg" /> 
Figure 4: Electron micrograph, 2D classes, and 3D reconstruction. Left panel: a micrograph with the sample from the monosome peak 2 at a calibrated magnification of 1.23 A/pixel. Middle panel: A representative post-processing data (2D classes) revealing intact monosomes. Right panel: 3D reconstruction. <a href="/files/ftp_upload/57877/57877fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Rapid Isolation of the Mitoribosome from HEK Cells at JoVE.com

Rapid Isolation of the Mitoribosome from HEK 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 ...

rapid-isolation-of-the-mitoribosome-from-hek-cells

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JoVE Journal

Biochemistry

10.9K Views.  Stockholm University. Het algemene doel van deze procedure is om ribosomen te zuiveren uit mitochondriën van menselijke cellijnen op grote schaal die voldoende is voor structurele studies. Deze mitochondriale ribosome zuiveringsmethode maakt de karakterisering van het vertalen van complexen, mutanten, kwaliteitscontrole assemblages, en mitoribosomale subunit tussenproducten. Deze kunnen worden gebruikt om belangrijke vragen over eiwitsynthese in mitochondriën te beantwoorden.Het belangrijkste voordeel van...

Video: Rapid Isolation of the Mitoribosome from HEK Cells

An Oligonucleotide-based Tandem RNA Isolation Procedure to Recover Eukaryotic mRNA-Protein Complexes

RNA-binding proteins (RBPs) play key roles in the post-transcriptional control of gene expression. Therefore, biochemical characterization of mRNA-protein complexes is essential to understanding mRNA regulation inferred by interacting proteins or non-coding RNAs. Herein, we describe a tandem RNA isolation procedure (TRIP) that enables the purification of endogenously formed mRNA-protein complexes from cellular extracts. The two-step protocol involves the isolation of polyadenylated mRNAs with antisense oligo(dT) beads and subsequent capture of an mRNA of interest with 3&#39;-biotinylated 2&#39;-O-methylated antisense RNA oligonucleotides, which can then be isolated with streptavidin beads. TRIP was used to recover in vivo crosslinked mRNA-ribonucleoprotein (mRNP) complexes from yeast, nematodes and human cells for further RNA and protein analysis. Thus, TRIP is a versatile approach that can be adapted to all types of polyadenylated RNAs across organisms to study the dynamic re-arrangement of mRNPs imposed by intracellular or environmental cues.

A tandem RNA isolation procedure (TRIP) for recovery of endogenously formed mRNA-protein complexes is described. Specifically, RNA-protein complexes are crosslinked in vivo, polyadenylated RNAs are isolated from extracts with oligo(dT) beads, and particular mRNAs are captured with modified RNA antisense oligonucleotides. Proteins bound to mRNAs are detected by immunoblot analysis.

RNA-binding proteins (RBPs) play key roles in the post-transcriptional control of gene expression. Therefore, biochemical characterization of mRNA-protein ...

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 ...

Isolation of Translating Ribosomes Containing Peptidyl-tRNAs for Functional and Structural Analyses

Recently, structural and biochemical studies have detailed many of the molecular events that occur in the ribosome during inhibition of protein synthesis by antibiotics and during nascent polypeptide synthesis. Some of these antibiotics, and regulatory nascent polypeptides mostly in the form of peptidyl-tRNAs, inhibit either peptide bond formation or translation termination1-7. These inhibitory events can stop the movement of the ribosome, a phenomenon termed "translational arrest". Translation arrest induced by either an antibiotic or a nascent polypeptide has been shown to regulate the expression of genes involved in diverse cellular functions such as cell growth, antibiotic resistance, protein translocation and cell metabolism8-13. Knowledge of how antibiotics and regulatory nascent polypeptides alter ribosome function is essential if we are to understand the complete role of the ribosome in translation, in every organism.
Here, we describe a simple methodology that can be used to purify, exclusively, for analysis, those ribosomes translating a specific mRNA and containing a specific peptidyl-tRNA14. This procedure is based on selective isolation of translating ribosomes bound to a biotin-labeled mRNA. These translational complexes are separated from other ribosomes in the same mixture, using streptavidin paramagnetic beads (SMB) and a magnetic field (MF). Biotin-labeled mRNAs are synthesized by run-off transcription assays using as templates PCR-generated DNA fragments that contain T7 transcriptional promoters. T7 RNA polymerase incorporates biotin-16-UMP from biotin-UTP; under our conditions approximately ten biotin-16-UMP molecules are incorporated in a 600 nt mRNA with a 25% UMP content. These biotin-labeled mRNAs are then isolated, and used in in vitro translation assays performed with release factor 2 (RF2)-depleted cell-free extracts obtained from Escherichia coli strains containing wild type or mutant ribosomes. Ribosomes translating the biotin-labeled mRNA sequences are stalled at the stop codon region, due to the absence of the RF2 protein, which normally accomplishes translation termination. Stalled ribosomes containing the newly synthesized peptidyl-tRNA are isolated and removed from the translation reactions using SMB and an MF. These beads only bind biotin-containing messages.
The isolated, translational complexes, can be used to analyze the structural and functional features of wild type or mutant ribosomal components, or peptidyl-tRNA sequences, as well as determining ribosome interaction with antibiotics or other molecular factors 1,14-16. To examine the function of these isolated ribosome complexes, peptidyl-transferase assays can be performed in the presence of the antibiotic puromycin1. To study structural changes in translational complexes, well established procedures can be used, such as i) crosslinking to specific amino acids14 and/or ii) alkylation protection assays1,14,17.

A major impediment to biochemical analyses of ribosomes containing nascent peptidyl-tRNAs has been the presence of other ribosomes in the same samples, ribosomes not involved in the translation of the specific mRNA sequence being analyzed. We developed a simple methodology to purify, exclusively, the ribosomes containing the nascent peptidyl-tRNA of interest.

Recently, structural and biochemical studies have detailed many of the molecular events that occur in the ribosome during inhibition of protein synthesis ...

Biology

Method for the Isolation and Identification of mRNAs, microRNAs and Protein Components of Ribonucleoprotein Complexes from Cell Extracts using RIP-Chip

As a result of the development of high-throughput sequencing and efficient microarray analysis, global gene expression analysis has become an easy and readily available form of data collection. In many research and disease models however, steady state levels of target gene mRNA does not always directly correlate with steady state protein levels. Post-transcriptional gene regulation is a likely explanation of the divergence between the two. Driven by the binding of RNA Binding Proteins (RBP), post-transcriptional regulation affects mRNA localization, stability and translation by forming a Ribonucleoprotein (RNP) complex with target mRNAs. Identifying these unknown de novo mRNA targets from cellular extracts in the RNP complex is pivotal to understanding mechanisms and functions of the RBP and their resulting effect on protein output. This protocol outlines a method termed RNP immunoprecipitation-microarray (RIP-Chip), which allows for the identification of specific mRNAs associated in the ribonucleoprotein complex, under changing experimental conditions, along with options to further optimize an experiment for the individual researcher. With this important experimental tool, researchers can explore the intricate mechanisms associated with post-transcriptional gene regulation as well as other ribonucleoprotein interactions.

A step by step protocol to isolating and identifying RNA associated complexes through RIP-Chip.

As a result of the development of high-throughput sequencing and efficient microarray analysis, global gene expression analysis has become an easy and ...

Using SecM Arrest Sequence as a Tool to Isolate Ribosome Bound Polypeptides

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway that polypeptide chain follows during co-translational folding to achieve its functional form is still an enigma. In order to understand this process and to determine the exact conformation of the co-translational folding intermediates, it is essential to develop techniques that allow the isolation of RNCs carrying nascent chains of predetermined sizes to allow their further structural analysis.
SecM (secretion monitor) is a 170 amino acid E. coli protein that regulates expression of the downstream SecA (secretion driving) ATPase in the secM-secA operon6. Nakatogawa and Ito originally found that a 17 amino acid long sequence (150-FSTPVWISQAQGIRAGP-166) in the C-terminal region of the SecM protein is sufficient and necessary to cause stalling of SecM elongation at Gly165, thereby producing peptidyl-glycyl-tRNA stably bound to the ribosomal P-site7-9. More importantly, it was found that this 17 amino acid long sequence can be fused to the C-terminus of virtually any full-length and/or truncated protein thus allowing the production of RNCs carrying nascent chains of predetermined sizes7. Thus, when fused or inserted into the target protein, SecM stalling sequence produces arrest of the polypeptide chain elongation and generates stable RNCs both in vivo in E. coli cells and in vitro in a cell-free system. Sucrose gradient centrifugation is further utilized to isolate RNCs.
The isolated RNCs can be used to analyze structural and functional features of the co-translational folding intermediates. Recently, this technique has been successfully used to gain insights into the structure of several ribosome bound nascent chains10,11. Here we describe the isolation of bovine Gamma-B Crystallin RNCs fused to SecM and generated in an in vitro translation system.

We describe here a technique that is now routinely used to isolate stably bound ribosome nascent chain complexes (RNCs). This technique takes advantage of the discovery that a 17 amino acid long SecM "arrest sequence" can halt translation elongation in a prokaryotic (E. coli) system, when inserted into (or fused to the C-terminus) of virtually any protein.

Extensive research has provided ample evidences suggesting that protein folding in the cell is a co-translational process1-5. However, the exact pathway ...

Isolation of mRNAs Associated with Yeast Mitochondria to Study Mechanisms of Localized Translation

Most of mitochondrial proteins are encoded in the nucleus and need to be imported into the organelle. Import may occur while the protein is synthesized near the mitochondria. Support for this possibility is derived from recent studies, in which many mRNAs encoding mitochondrial proteins were shown to be localized to the mitochondria vicinity. Together with earlier demonstrations of ribosomes&#8217; association with the outer membrane, these results suggest a localized translation process. Such localized translation may improve import efficiency, provide unique regulation sites and minimize cases of ectopic expression. Diverse methods have been used to characterize the factors and elements that mediate localized translation. Standard among these is subcellular fractionation by differential centrifugation. This protocol has the advantage of isolation of mRNAs, ribosomes and proteins in a single procedure. These can then be characterized by various molecular and biochemical methods. Furthermore, transcriptomics and proteomics methods can be applied to the resulting material, thereby allow genome-wide insights. The utilization of yeast as a model organism for such studies has the advantages of speed, costs and simplicity. Furthermore, the advanced genetic tools and available deletion strains facilitate verification of candidate factors.

Many mRNAs encoding mitochondrial proteins are associated with the mitochondria outer membrane. We describe a subcellular fractionation procedure aimed at isolation of yeast mitochondria with its associated mRNAs and ribosomes. This protocol can be applied to cells grown under diverse conditions&#160;in order to reveal mechanisms of mRNA localization and localized translation near the mitochondria.

Most of mitochondrial proteins are encoded in the nucleus and need to be imported into the organelle. Import may occur while the protein is synthesized ...

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

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 ...

Rapid In Vivo Fixation and Isolation of Translational Complexes from Eukaryotic Cells

Rapid responses involving fast redistribution of messenger(m)RNA and alterations of mRNA translation are pertinent to ongoing homeostatic adjustments of the cells. These adjustments are critical to eukaryotic cell survivability and &#39;damage control&#39; during fluctuating nutrient and salinity levels, temperature, and various chemical and radiation stresses. Due to the highly dynamic nature of the RNA-level responses, and the instability of many of the RNA:RNA and RNA:protein intermediates, obtaining a meaningful snapshot of the cytoplasmic RNA state is only possible with a limited number of methods. Transcriptome-wide, RNA-seq-based ribosome profiling-type experiments are among the most informative sources of data for the control of translation. However, absence of a uniform RNA and RNA:protein intermediate stabilization can lead to different biases, particularly in the fast-paced cellular response pathways. In this article, we provide a detailed protocol of rapid fixation applicable to eukaryotic cells of different permeability, to aid in RNA and RNA:protein intermediate stabilization. We further provide examples of isolation of the stabilized RNA:protein complexes based on their co-sedimentation with ribosomal and poly(ribo)somal fractions. The separated stabilized material can be subsequently used as part of ribosome profiling-type experiments, such as in Translation Complex Profile sequencing (TCP-seq) approach and its derivatives. Versatility of the TCP-seq-style methods has now been demonstrated by the applications in a variety of organisms and cell types. The stabilized complexes can also be additionally affinity-purified and imaged using electron microscopy, separated into different poly(ribo)somal fractions and subjected to RNA sequencing, owing to the ease of the crosslink reversal. Therefore, methods based on snap-chilling and formaldehyde fixation, followed by the sedimentation-based or other type of RNA:protein complex enrichment, can be of particular interest in investigating finer details of rapid RNA:protein complex dynamics in live cells.

Rapid In Vivo Fixation and Isolation of Translational Complexes from Eukaryotic Cells

We present a technique to rapidly stabilize translational (protein biosynthesis) complexes with formaldehyde crosslinking in live yeast and mammalian cells. The approach enables dissecting transient intermediates and dynamic RNA:protein interactions. The crosslinked complexes can be used in multiple downstream applications such as in deep sequencing-based profiling methods, microscopy, and mass-spectrometry.

Rapid responses involving fast redistribution of messenger(m)RNA and alterations of mRNA translation are pertinent to ongoing homeostatic adjustments of ...

Improved Protein Quantification Using Two-Step Mitochondrial Isolation

Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification

Most physiological and disease processes, from central metabolism to immune response to neurodegeneration, involve mitochondria. The mitochondrial proteome is composed of more than 1,000 proteins, and the abundance of each can vary dynamically in response to external stimuli or during disease progression. Here, we describe a protocol for isolating high-quality mitochondria from primary cells and tissues. The two-step procedure comprises (1) mechanical homogenization and differential centrifugation to isolate crude mitochondria, and (2) tag-free immune capture of mitochondria to isolate pure organelles and eliminate contaminants. Mitochondrial proteins from each purification stage are analyzed by quantitative mass spectrometry, and enrichment yields are calculated, allowing the discovery of novel mitochondrial proteins by subtractive proteomics. Our protocol provides a sensitive and comprehensive approach to studying mitochondrial content in cell lines, primary cells, and tissues.

Author Spotlight: Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification  

We present a two-step protocol for high-quality mitochondria isolation that is compatible with protein discovery and quantification at a proteome scale. Our protocol does not require genetic engineering and is thus suitable for studying mitochondria from any primary cells and tissues.

Rapid Isolation of the Mitoribosome from HEK Cells

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