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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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&#8217;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

Nanyang Technological University

Research

JoVE Journal

Biochemistry

10.8K Views.  Stockholm University. 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.

Video: Rapid Isolation of the Mitoribosome from HEK Cells

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. ...

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

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

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

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

An Efficient Method for the Isolation of Highly Purified RNA from Seeds for Use in Quantitative Transcriptome Analysis

Plant seeds accumulate large amounts of storage reserves comprising biodegradable organic matter. Humans rely on seed storage reserves for food and as industrial materials. Gene expression profiles are powerful tools for investigating metabolic regulation in plant cells. Therefore, detailed, accurate gene expression profiles during seed development are required for crop breeding. Acquiring highly purified RNA is essential for producing these profiles. Efficient methods are needed to isolate highly purified RNA from seeds. Here, we describe a method for isolating RNA from seeds containing large amounts of oils, proteins, and polyphenols, which have inhibitory effects on high-purity RNA isolation. Our method enables highly purified RNA to be obtained from seeds without the use of phenol, chloroform, or additional processes for RNA purification. This method is applicable to Arabidopsis, rapeseed, and soybean seeds. Our method will be useful for monitoring the expression patterns of low level transcripts in developing and mature seeds.

We have succeeded in establishing a method for RNA isolation from plant seeds containing large amounts of oils, proteins, and polyphenols, which have inhibitory effects on high-purity RNA isolation. Our method is suitable for monitoring the expression of genes with low level transcripts in seeds.

Plant seeds accumulate large amounts of storage reserves comprising biodegradable organic matter. Humans rely on seed storage reserves for food and as ...

Tandem Affinity Purification of Protein Complexes from Eukaryotic Cells

The purification of active protein-protein and protein-nucleic acid complexes is crucial for the characterization of enzymatic activities and de novo identification of novel subunits and post-translational modifications. Bacterial systems allow for the expression and purification of a wide variety of single polypeptides and protein complexes. However, this system does not enable the purification of protein subunits that contain post-translational modifications (e.g., phosphorylation and acetylation), and the identification of novel regulatory subunits that are only present/expressed in the eukaryotic system. Here, we provide a detailed description of a novel, robust, and efficient tandem affinity purification (TAP) method using STREP- and FLAG-tagged proteins that facilitates the purification of protein complexes with transiently or stably expressed epitope-tagged proteins from eukaryotic cells. This protocol can be applied to characterize protein complex functionality, to discover post-translational modifications on complex subunits, and to identify novel regulatory complex components by mass spectrometry. Notably, this TAP method can be applied to study protein complexes formed by eukaryotic or pathogenic (viral and bacterial) components, thus yielding a wide array of downstream experimental opportunities. We propose that researchers working with protein complexes could utilize this approach in many different ways.

We describe here a novel, robust, and efficient tandem affinity purification (TAP) method for the expression, isolation, and characterization of protein complexes from eukaryotic cells. This protocol could be utilized for the biochemical characterization of discrete complexes as well as the identification of novel interactors and post-translational modifications that regulate their function.

The purification of active protein-protein and protein-nucleic acid complexes is crucial for the characterization of enzymatic activities and de novo ...

Purification of Biotinylated Cell Surface Proteins from Rhipicephalus microplus Epithelial Gut Cells

Rhipicephalus microplus &#8211; the cattle tick &#8211; is the most significant ectoparasite in terms of economic impact on livestock as a vector of several pathogens. Efforts have been dedicated to the cattle tick control to diminish its deleterious effects, with focus on the discovery of vaccine candidates, such as BM86, located on the surface of the tick gut epithelial cells. Current research focuses upon the utilization of cDNA and genomic libraries, to screen for other vaccine candidates. The isolation of tick gut cells constitutes an important advantage in investigating the composition of surface proteins upon the tick gut cells membrane. This paper constitutes a novel and feasible method for the isolation of epithelial cells, from the tick gut contents of semi-engorged R. microplus. This protocol utilizes TCEP and EDTA to release the epithelial cells from the subepithelial support tissues and a discontinuous density centrifugation gradient to separate epithelial cells from other cell types. Cell surface proteins were biotinylated and isolated from the tick gut epithelial cells, using streptavidin-linked magnetic beads allowing for downstream applications in FACS or LC-MS/MS-analysis.

Purification of Biotinylated Cell Surface Proteins from Rhipicephalus microplus Epithelial Gut Cells

A modified density centrifugation gradient-based methodology was utilized to isolate epithelial cells from Rhipicephalus microplus gut tissue. Surface-bound proteins were biotinylated and purified through streptavidin magnetic beads for utilization in downstream applications.

Rhipicephalus microplus &#8211; the cattle tick &#8211; is the most significant ectoparasite in terms of economic impact on livestock as a vector of ...

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 ...

Leaf Spray Mass Spectrometry: A Rapid Ambient Ionization Technique to Directly Assess Metabolites from Plant Tissues

Plants produce thousands of small molecules that are diverse in their chemical properties. Mass spectrometry (MS) is a powerful technique for analyzing plant metabolites because it provides molecular weights with high sensitivity and specificity. Leaf spray MS is an ambient ionization technique where plant tissue is used for direct chemical analysis via electrospray, eliminating chromatography from the process. This approach to sampling metabolites allows for a wide range of chemical classes to be detected simultaneously from intact plant tissues, minimizing the amount of sample preparation needed. When used with a high-resolution, accurate mass MS, leaf spray MS facilitates the rapid detection of metabolites of interest. It is also possible to collect tandem mass fragmentation data with this technique to facilitate a compound identification. The combination of accurate mass measurements and fragmentation is beneficial in confirming compound identities. The leaf spray MS technique requires only minor modifications to a nanospray ionization source and is a useful tool to further expand the capabilities of a mass spectrometer. Here, fresh leaf tissue from Sceletium tortuosum (Aizoaceae), a traditional medicinal plant from South Africa, is analyzed; numerous mesembrine alkaloids are detected with leaf spray MS.

Leaf spray mass spectrometry is a direct chemical analysis technique that minimizes the sample preparation and eliminates chromatography, allowing for the rapid detection of small molecules from plant tissues.

Plants produce thousands of small molecules that are diverse in their chemical properties. Mass spectrometry (MS) is a powerful technique for analyzing ...

Dissipative Microgravimetry to Study the Binding Dynamics of the Phospholipid Binding Protein Annexin A2 to Solid-supported Lipid Bilayers Using a Quartz Resonator

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic properties of the absorbed/immobilized mass on sensor surfaces, allowing the measurements of the interaction of proteins with solid-supported surfaces, such as lipid bilayers, in real-time and with a high sensitivity. Annexins are a highly conserved group of phospholipid-binding proteins that interact reversibly with the negatively charged headgroups via the coordination of calcium ions. Here, we describe a protocol that was employed to quantitatively analyze the binding of annexin A2 (AnxA2) to planar lipid bilayers prepared on the surface of a quartz sensor. This protocol is optimized to obtain robust and reproducible data and includes a detailed step-by-step description. The method can be applied to other membrane-binding proteins and bilayer compositions.

Here, we present an experimental protocol that can be employed to determine the binding affinities and mode of interaction of label-free phospholipid-binding protein annexin A2 with immobilized solid-supported bilayers (SLB) by simultaneously measuring the mass uptake and the viscoelastic properties of the protein annexin A2.

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic ...

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 ...

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.