This work was supported in part by a Research Support Funds Grant (RSFG) from Indiana University-Purdue University, Indianapolis, and in part by an award from the American Heart Association 14GRNT20390154, to A.R.K.

1. Bacterial Expression.
Note: Expression plasmids used in this study are described in Table 1. Solutions, media, and buffers used in this study are described in Table 2. The cloning of archaeal proteasome subunit genes and the generation of expression plasmids are described elsewhere18,20. In brief, plasmids for recombinant coexpression of subunits employ a bicistronic operon strategy which helps in obtaining comparable expression levels of individual subunits18,20. The expression parameters listed below were empirically determined to be optimal for the proteasome subunits in this study. It may be necessary to optimize expression for other proteasome mutants, for proteasomes from other archaeal species, or for other recombinant protein complexes (see Discussion).
<ol>
	<li>Transform expression plasmid of interest into chemically competent E.coli BL21(DE3) cells.</li>
	<li>Add 1 - 2 &#956;L&#160;of plasmid (typically 50 - 100 ng) to an aliquot of DE3&#160;cells that were freshly thawed on ice, and continue to incubate on ice for 20 min.</li>
	<li>Heat shock the cells at 42 &#176;C for 45 s and return tube to ice for additionl 2 min.</li>
	<li>Add 1 mL of LB medium and incubate at 37 &#176;C with shaking (150 rpm) for 1 h.</li>
	<li>Spread 100 - 200 &#956;L&#160;of the transformation mixture onto LB-kan plates (plates containing solid LB media, supplemented with kanamycin (all plasmids used in this study encode resistance to this antibiotic)). Incubate plates at 37 &#186;C O/N.</li>
	<li>Next morning, inoculate a single colony from the LB-kan plate into 3 mL of liquid LB-kan media in a glass culture tube. Incubate at 37 &#186;C with shaking (150 rpm) for approximately 2.5 - 3 h, or till turbidity is observed.</li>
	<li>Measure the optical density of the culture at 600 nm (OD600) in a spectrophotometer. Dilute the cell culture with an appropriate amount of prewarmed LB-kan media to an OD600 of 0.4 in a final volume of 6 mL. Resume shaking at 37 &#186;C for 40 min.</li>
	<li>Induce protein expression in the liquid cultures by adding IPTG from a stock solution to a final concentration of 1 mM. Incubate at 37 &#186;C with shaking (150 rpm) for approximately 6 - 7 h.</li>
	<li>Harvest bacterial cell cultures in their entirety into 1.5 mL&#160;microcentrifuge tubes.</li>
	<li>Add 1.5 mL&#160;of a culture into microcentrifuge tube. Centrifuge at 10,000 &#215; g for 1 min.</li>
	<li>Discard the supernatant and add another 1.5 mL&#160;of the same culture to the pellet. Centrifuge at 10,000 &#215; g for 1 min.</li>
	<li>Repeat step 1.11 until the entire culture is harvested into the microcentrifuge tube.</li>
	<li>Store the induced pellets at -80 &#176;C until lysis.</li>
</ol>
2. Bacterial Lysis and Lysate Mixing.
Note: For samples studied via coexpression, follow section 2.1 (and its subsections). For samples requiring lysate mixing, follow section 2.2 (and its subsections). The TSP (Total, Soluble, Pellet) analysis that is included in the protocol is useful in optimizing protein expression which can help ensure that approximately equal amounts of &#945; and &#946; subunits are combined during lysate mixing (see Discussion). It also provides an important control if downstream results are not as expected (i.e. it verifies that protein was expressed and soluble). Thus, we highly recommend including TSP analysis initially. Once an optimal protocol has been achieved for a particular subunit combination, TSP analysis is not strictly required which is why it is described as optional below.
<ol>
	<li>Lysis of bacterial cells and preparation of soluble fractions.
	<ol>
		<li>Thaw the induced cell pellet on ice for 5 min. Resuspend the pellet in 600 &#181;L&#160;of lysis buffer.</li>
		<li>Incubate the suspension at 30 &#186;C with shaking (150 rpm) for 30 min to generate total crude lysate. 
		Note: The following step and the subsection that follows are optional.</li>
		<li>Remove two 25 &#956;L&#160;aliquots from the total crude lysate into separate 1.5 mL&#160;microcentrifuge tubes and carry out TSP analysis as follows.
		<ol>
			<li>To one of the two 25 &#956;L&#160;aliquots, add 5X SDS&#160;sample buffer to a final concentration of 1X. Label the tube with &#34;T&#34; for &#34;total crude lysate&#34;.</li>
			<li>Incubate the &#34;T&#34; sample at 100 &#186;C for 5 min to completely denature proteins.</li>
			<li>Meanwhile, take the second 25 &#956;L&#160;aliquot and centrifuge it at 10,000 &#215; g for 10 min. Carefully remove the supernatant without disturbing the tiny pellet, and transfer it to a new 1.5 mL&#160;microcentrifuge tube. Label this new tube &#34;S&#34; for &#34;soluble fraction&#34;.</li>
			<li>To the tiny pellet left behind in the previous step, add 25 &#956;L&#160;of lysis buffer and vortex to resuspend. Label this tube &#34;P&#34; for &#34;pellet fraction&#34;.</li>
			<li>Add 6 &#956;L&#160;of 5X SDS&#160;sample buffer to the &#34;S&#34; and &#34;P&#34; tubes and incubate at 100 &#186;C as described above. 
			Note: If the TSP samples will not be used that day to analyze expression, they may be frozen at -20 &#186;C until needed. Once thawed, they must be reincubated at 100 &#186;C, as described above, immediately prior to loading on a 12% and/or 15% standard SDS-PAGE gel.</li>
		</ol>
		</li>
		<li>While the TSP analysis is carried out, centrifuge the remaining 550 &#956;L&#160;of total crude lysate (or the entire 600 &#956;L&#160;of total crude lysate if TSP analysis is not carried out) at 10,000 &#215; g for 10 min. Collect the supernatant in a fresh 1.5 mL&#160;microcentrifuge tube. This is the soluble lysate that will be used for purification.</li>
	</ol>
	</li>
	<li>Lysate mixing.
	<ol>
		<li>Carry out lysis as described in steps 2.1.1 and 2.1.2 above.</li>
		<li>Mix 600 &#181;L&#160;of total crude lysate from bacteria expressing the desired &#945; subunit with 600 &#181;L&#160;of total crude lysate from bacteria expressing the desired &#946; subunit. Incubate at 37 &#186;C with slow shaking for 30 min. 
		Note: It may be necessary to optimize incubation time to achieve maximum assembly during lysate mixing (see Discussion). The time and temperature presented here are optimal for the recombinant proteins in this study and were determined elsewhere18.</li>
		<li>Centrifuge the mixed lysate at 10,000 &#215; g for 10 min to separate soluble material from insoluble pellet. Transfer the supernatant to a new 1.5 mL&#160;microcentrifuge tube. Use this mixed soluble lysate for protein purification.</li>
	</ol>
	</li>
</ol>
3. Protein Purification via Immobilized&#160;Cobalt Affinity Resin (ICAR).
<ol>
	<li>Equilibrate the resin.
	<ol>
		<li>Thoroughly resuspend the resin by inverting the bottle and transfer 50 &#181;L&#160;of the slurry to a 1.5 mL&#160;microcentrifuge tube. Centrifuge at 700 &#215; g for 2 min to pellet resin. Carefully aspirate supernatant. 
		Note: We carry out pipette aspiration using a blue 1 mL&#160;pipette tip to remove the bulk of the liquid. A white 2 &#956;L&#160;tip is used for fine control of removal of the remaining supernatant, leaving a very thin layer of liquid covering the beads.</li>
		<li>Add 1 mL&#160;of Buffer A. Mix gently to resuspend resin and centrifuge at 700 &#215; g for 2 min to pellet resin. Carefully aspirate the supernatant and repeat this wash step one more time.</li>
	</ol>
	</li>
	<li>Apply soluble lysate obtained previously in section 2.1 or 2.2 to the equilibrated resin. Incubate the tube with gentle rotation at 4 &#186;C for 60 min. Centrifuge the tube at 700 &#215; g for 5 min and carefully aspirate the supernatant.</li>
	<li>Wash the resin as follows to remove nonspecifically bound proteins.
	<ol>
		<li>Resuspend resin with 1 mL&#160;of Buffer A and incubate with gentle rocking at 4 &#186;C for 10 min. Centrifuge the tube at 700 &#215; g for 5 min and carefully aspirate the supernatant. Repeat this wash step one more time.</li>
		<li>Resuspend resin with 1 mL&#160;of Buffer B (Buffer A with 5 mM Imidazole) and incubate with gentle rocking at 4 &#186;C for 5 min. Centrifuge the tube at 700 &#215; g for 5 min and carefully aspirate the supernatant. Repeat this wash step one more time.</li>
		<li>Resuspend resin with 1 mL&#160;of Buffer C (Buffer A with 10 mM Imidazole) and incubate with gentle rocking at 4 &#186;C for 5 min. Centrifuge the tube at 700 &#215; g for 5 min and carefully aspirate the supernatant.</li>
	</ol>
	</li>
	<li>Elute the protein by adding 400 &#181;L of Buffer E (Buffer A with 200 mM Imidazole) to the resin. Incubate with gentle rocking at 4 &#186;C for 5 min. Centrifuge at 700 &#215; g for 5 min.</li>
	<li>Transfer supernatant containing purified protein to a new 1.5 mL&#160;centrifuge tube.</li>
	<li>Desalt the purified protein by serial centrifugation as follows.
	<ol>
		<li>To 400 &#956;L&#160;of purified protein, add 100 &#956;L&#160;Buffer A (this reduces Imidazole concentration from 200 mM to 160 mM). Apply purified protein to 0.5 mL&#160;ultracentrifugal filters with a 10 kDa molecular weight cut-off. Centrifuge at 14,000 &#215; g for 5 min.</li>
		<li>Discard the filtrate and add 400 &#956;L&#160;Buffer A to dilute the retentate and centrifuge again. Continue the cycles of centrifugation/dilution until imidazole concentration falls below 4 mM. As an example, if each centrifugation concentrates 500 &#956;l down to ~70 &#956;l (or approximately 7-fold), then two cycles will reduce the starting 160 mM imidazole concentration (7&#215;7 = 49-fold) to approximately 3.3 mM.</li>
		<li>Measure the protein concentration of the desalted sample using the BCA assay21. 
		Note: Desalting is required to reduce Imidazole levels below the tolerance limit for the BCA assay, as described in the manufacturer&#39;s instructions. Our lab prefers the BCA assay because it is sensitive, has a large dynamic range, and exhibits much less protein-to-protein variation. However, other methods to determine protein concentration can be substituted for the BCA assay. The key is to be aware of each method&#39;s advantages and limitations.</li>
	</ol>
	</li>
	<li>Add 5X native sample buffer to a final concentration of 1X and proceed to electrophoresis. Alternatively, store samples at -20 &#186;C for later analysis.</li>
</ol>
4. Nondenaturing Polyacrylamide Gel Electrophoresis (PAGE).
Caution: Unpolymerized acrylamide is a neurotoxin. Wear appropriate protective gear.
<ol>
	<li>Prepare the nondenaturing PAGE gel as follows.
	<ol>
		<li>Prepare the gel cassette for casting using clean glass plates and casting stand. Place gradient maker on top of a small magnetic stirrer and place a tiny stir bar into each chamber.</li>
		<li>Using 40% (w/v) acrylamide and 2% (w/v) bisacrylamide stock solutions, prepare 5% and 10% (w/v) acrylamide gel solutions in a native resolving buffer. Ensure that the ratio of total acrylamide to bisacrylamide is 37.5:1. Chill on ice prior to pouring. 
		Note: The nondenaturing PAGE system used here is essentially identical to the Laemmli SDS-PAGE system, with SDS omitted22.</li>
		<li>Add Ammonium Persulfate (from a 10% (w/v) stock solution prepared in water) to the acrylamide solutions to initiate polymerization. Final concentration of Ammonium Persulfate is 0.1% (w/v).</li>
		<li>Pour the acrylamide solutions into the two chambers of the gradient maker. With the outlet tubing inserted between the plates of the gel cassette, activate the magnetic stirrer and open the gradient maker valves. Pour the 5 - 10% nondenaturing gradient gel.</li>
		<li>Once poured, overlay the gel with a thin layer of Isopropanol and allow the gel to polymerize for 30 min. During this time, prepare fresh 5% acrylamide gel solution in native resolving buffer. After the gel sets, pour off Isopropanol and rinse the top of the gel with deionized water from a squirt bottle.</li>
		<li>To the 5% gel solution prepared above, add Ammonium Persulfate (exactly as described in 4.1.3) and pour on top of the polymerized gel until glass plates are full. Insert gel comb. Allow the overlaid gel to polymerize for an additional 30 min.</li>
		<li>Once gel is polymerized, assemble gel cassette into electrophoresis apparatus. Alternatively, store the gel at 4 &#186;C, wrapped in moistened paper towels and plastic wrap until ready for use.</li>
	</ol>
	</li>
	<li>Once nondenaturing PAGE gel is assembled into electrophoresis apparatus, fill the tank with 1X native running buffer prepared fresh from a 10X native running buffer stock.</li>
	<li>Load 10 &#956;g of purified protein, obtained at the end of section 3, into each well using a glass syringe.</li>
	<li>Load 2 &#956;L&#160;of high molecular weight native protein standard (diluted in 1X native running buffer) into one of the wells.</li>
	<li>Run gel at 55 V and 4 &#186;C until the dye front runs off the gel (approximately 4 - 4.5 h).</li>
	<li>In addition to the nondenaturing gel, analyze aliquots of the purified protein by standard SDS-PAGE22.
	<ol>
		<li>Mix aliquots (10 &#956;g) of the purified protein samples, obtained at the end of section 3, with 5X SDS-sample buffer to a final concentration of 1X.</li>
		<li>Incubate at 100 &#186;C for 5 min and load onto standard 12% and/or 15% SDS-PAGE gels22.</li>
		<li>Run at 80V for 20 min and then at 120 V until the dye front runs off the gel (this is approximately an additional 75 min for a 12% gel, and 120 min for a 15% gel).</li>
	</ol>
	</li>
</ol>
5. Visualizing Activity and Protein Staining.
<ol>
	<li>Following electrophoresis, carefully separate glass plates and transfer the nondenaturing gel to a gel tray containing 50 mL&#160;of deionized water. Rinse gel with gentle rocking for 5 min. Discard water and repeat this wash step three times.</li>
	<li>Perform substrate overlay assay as follows.
	<ol>
		<li>Add 1 mL&#160;of developing buffer containing the fluorogenic peptide substrate Suc-LLVY-AMC and spread uniformly over the gel. Incubate at 37 &#186;C for 30 min. 
		Note: A glass rod or a gel releaser (small wedge of plastic used to separate glass plates) can be used to spread the liquid over the gel.</li>
		<li>Carefully transfer the gel onto the UV transilluminator of the gel imaging system and observe fluorescence due to the cleaved peptide substrate. Record image. Carefully transfer the gel back to the gel tray.</li>
		<li>Rinse the gel twice with 50 mL of deionized water for 5 min with gentle rocking.</li>
		<li>Stain the gel with 10 mL of a colloidal coomassie stain reagent for 60 min with gentle rocking. Destain gel with 50 mL water until background becomes clear. This staining step applies to the standard SDS-PAGE gels as well.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

We demonstrate the benefit of a combined approach to analyzing proteasome assembly by nondenaturing PAGE using recombinant archaeal proteasomes. The usual method9,11 of bacterial coexpression of proteasome subunits allows for rapid analysis but may not reveal key assembly intermediates. We suggest combining coexpression with lysate mixing to develop a broader picture of assembly events.
The advantage of this combined approach is that despite requiring separate expression of the &#94...

Multiprotein complexes carry out numerous critical cellular activities1. For many of these complexes, much more is known about their structure and function than about their assembly2,3. The proteasome is one such complex and is found in all domains of life. In eukaryotes, this molecular machine is at the core of the Ubiquitin/Proteasome System (UPS) and provides the major route of intracellular protein degradation4. The eukaryotic proteasome (referred to as the 26S proteasome) is comprised of two major sub assemblies: a 20S proteasome, or Core Particle (CP)5, that can be capped on one or both ends by a 19S Regulatory Particle (RP)6.
The 20S proteasome&#160;is a large compartmentalized protease. Its quaternary structure is absolutely conserved across all domains of life and consists of a stack of four seven-membered rings containing two types of structurally related subunits, &#945; and &#946;5,7,8. In eukaryotes, the two outer rings are each comprised of seven distinct &#945; subunits and the two inner rings are each comprised of seven distinct &#946; subunits; proteolytic activity resides within three of the &#946; subunits. By contrast, the CP rings of archaea and bacteria are usually comprised of only one type of &#945; and one type of &#946; subunit. Archaeal proteasomes have provided an important model system to study proteasome assembly due to both their compositional simplicity and their sharing a common assembly mechanism with their eukaryotic counterparts9-13. In brief, &#945; subunits assemble into &#945; rings first, which serve as a scaffold onto which &#946; subunits assemble. The resulting half-proteasomes (&#945;7&#946;7) dimerize, giving rise to fully assembled CP (&#945;7&#946;7&#946;7&#945;7). During dimerization, the propeptides present on &#946; subunits are autocatalytically removed, exposing the catalytic N-terminal threonines. The use of archaeal proteasomes to model assembly frequently takes advantage of the production of recombinant archaeal proteasome proteins in Escherichia coli. This is a worthwhile approach because it enables the subunits to be produced in various combinations, as both WT and mutant versions, in a host organism that does not produce its own proteasomes.
Monitoring the assembly of multi-protein complexes biochemically requires some kind of fractionation method that separates fully assembled complexes from assembly intermediates and precursors. Due to its superior resolving capacity, nondenaturing Polyacrylamide Gel Electrophoresis (PAGE) has proven to be especially useful in the fractionation of various large multiprotein complexes14-17. The combination of recombinant archaeal proteasome production and nondenaturing PAGE has become a powerful approach in dissecting proteasome assembly9,11,12,18. However, the usual method by which this approach is applied (i.e. via the recombinant coexpression of &#945; and &#946; subunits) has an important drawback. Assembly reactions are cooperative and strongly concentration-dependent3. Given that the protein concentration inside cells is very high19, due to excluded volume effects, assembly reactions proceed rapidly in vivo. Hence it is possible to miss important assembly intermediates when &#945; and &#946; subunits are coexpressed.
Here, we argue for a combined approach in the study of proteasome assembly using recombinant archaeal proteasome subunits. In this approach, both coexpression and lysate mixing methods are employed. The former allows for rapid analysis of assembly because coexpression is less labor intensive. The latter depends on separate expression of &#945; and &#946; subunits, followed by mixing. Though this requires a bit more effort than coexpression, it is more than compensated for by the ability to detect intermediates that are missed during coexpression. Together, these two methods can provide a more complete picture of proteasome assembly.

<table><tbody><tr><td>Acrylamide (40%) solution</td><td>Biorad</td><td>1610104</td><td>Unpolymerized acrylamide is a neurotoxin. Wear proper protective gear</td></tr><tr><td>Amicon ultra 0.5 mL centrifugal filters</td><td>EMDMillipore</td><td>UFC501024</td><td></td></tr><tr><td>Ammonium Persulfate</td><td>Sigma</td><td>A3678</td><td></td></tr><tr><td>ATP</td><td>Sigma</td><td>A7699</td><td></td></tr><tr><td>BCA assay kit</td><td>Pierce</td><td>23225</td><td></td></tr><tr><td>Bisacrylamide (2%) solution</td><td>Biorad</td><td>1610142</td><td></td></tr><tr><td>Bromophenol blue</td><td>Sigma</td><td>B8026</td><td></td></tr><tr><td>DNaseI</td><td>Sigma</td><td>DN25</td><td></td></tr><tr><td>Dithiothreitol (DTT)</td><td>Thermo Fisher</td><td>BP172</td><td></td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; BL21 competent cells</td><td>EMD Millipore</td><td>69450</td><td></td></tr><tr><td>GelCode Blue</td><td>Thermo Fisher</td><td>24592</td><td>Colloidal coomassie stain reagent for gels</td></tr><tr><td>Gel doc EZ system</td><td>Biorad</td><td>1708270</td><td>Gel documentation system</td></tr><tr><td>Gel releasers</td><td>Biorad</td><td>1653320</td><td>Wedge shaped plastic used to separate gel plates; useful for spreading liquid.</td></tr><tr><td>Glass rod</td><td>Thermo Fisher</td><td>11-380B</td><td></td></tr><tr><td>Glycerol</td><td>Sigma</td><td>49767</td><td></td></tr><tr><td>Glycine</td><td>Thermo Fisher</td><td>BP3865</td><td></td></tr><tr><td>Hamilton syringe</td><td>Thermo Fisher</td><td>14-813-38</td><td>Glass syringe for loading gels</td></tr><tr><td>HEPES</td><td>US Biologicals</td><td>H2010</td><td></td></tr><tr><td>HMW Native calibration kit</td><td>GE Healthcare</td><td>170445-01</td><td>High molecular weight protein standards</td></tr><tr><td>Hoefer SG30</td><td>Thermo Fisher</td><td>03-500-277</td><td>Gradient maker</td></tr><tr><td>Imidazole</td><td>US Biologicals</td><td>280671</td><td></td></tr><tr><td>IPTG</td><td>US Biologicals</td><td>I8500</td><td>For induction of protein expression</td></tr><tr><td>Isopropanol</td><td>Thermo Fisher</td><td>BP26181</td><td></td></tr><tr><td>Kanamycin sulfate</td><td>US Biologicals</td><td>K0010</td><td></td></tr><tr><td>Lysozyme</td><td>Sigma</td><td>L6876</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Fluka analytical</td><td>630680</td><td></td></tr><tr><td>Mini Protean Tetra Cell</td><td>Biorad</td><td>1658002EDU</td><td>Gel electrophoresis apparatus</td></tr><tr><td>NaCl</td><td>Thermo Fisher</td><td>S640-3</td><td></td></tr><tr><td>NaOH</td><td>Thermo Fisher</td><td>S318-1</td><td></td></tr><tr><td>Pefabloc SC</td><td>Roche</td><td>11429876001</td><td>Protease inhibitor</td></tr><tr><td>pET42</td><td>EMD Millipore</td><td>70562</td><td>Expression plasmid</td></tr><tr><td>Precision plus all blue standard</td><td>Biorad</td><td>1610373</td><td>Molecular protein standard for SDS-PAGE</td></tr><tr><td>Quickchange mutagenesis kit</td><td>Agilent technologies</td><td>200521</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate (SDS)</td><td>Thermo Fisher</td><td>BP166</td><td></td></tr><tr><td>Suc-LLVY-AMC</td><td>Enzo lifesciences</td><td>BML P802-0005</td><td>Fluorogenic substrate</td></tr><tr><td>Talon Metal Affinity Resin</td><td>Clontech</td><td>635502</td><td>Immobilized Cobalt Affinity Resin</td></tr><tr><td>TEMED</td><td>Sigma</td><td>T7024</td><td></td></tr><tr><td>Tris</td><td>US Biologicals</td><td>T8600</td><td></td></tr><tr><td>Triton-X100</td><td>Sigma</td><td>93426</td><td></td></tr><tr><td>Tryptone</td><td>Bacto BD</td><td>211699</td><td></td></tr><tr><td>UV sample tray</td><td>Biorad</td><td>1708271</td><td>For UV imaging of gels</td></tr><tr><td>Yeast extract</td><td>Bacto BD</td><td>212720</td><td></td></tr></tbody></table>

examining proteasome assembly with recombinant archaeal proteasomes and nondenaturing page:  the case for a combined approach

Proteasomes are found in all domains of life. They provide the major route of intracellular protein degradation in eukaryotes, though their assembly is not completely understood. All proteasomes contain a structurally conserved core particle (CP), or 20S proteasome, containing two heptameric&#160;&#946; subunit rings sandwiched between two heptameric&#160;&#945; subunit rings. Archaeal 20S proteasomes are compositionally simpler compared to their eukaryotic counterparts, yet they both share a common assembly mechanism. Consequently, archaeal 20S proteasomes continue to be important models for eukaryotic proteasome assembly. Specifically, recombinant expression of archaeal 20S proteasomes coupled with nondenaturing polyacrylamide gel electrophoresis (PAGE) has yielded many important insights into proteasome biogenesis. Here, we discuss a means to improve upon the usual strategy of coexpression of archaeal proteasome &#945; and &#946; subunits prior to nondenaturing PAGE. We demonstrate that although rapid and efficient, a coexpression approach alone can miss key assembly intermediates. In the case of the proteasome, coexpression may not allow detection of the half-proteasome, an intermediate containing one complete &#945;-ring and one complete &#946;-ring. However, this intermediate is readily detected via lysate mixing. We suggest that combining coexpression with lysate mixing yields an approach that is more thorough in analyzing assembly, yet remains labor nonintensive. This approach may be useful for the study of other recombinant multiprotein complexes.

Proteasome assembly (Figure 1) begins when &#945; subunits combine to form rings9. This can be illustrated when &#945; subunits from the archaeon Methanococcus maripaludis S2 are expressed in E. coli as C-terminally hexahistidine tagged (his-tagged) derivatives (Table 1). When the recombinant &#945;-his protein was purified by ICAR&#160;and analyzed by nondenaturing PAGE, two bands were observed (Figure 2A, lane 1). We have previously demonst...

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Examining Proteasome Assembly with Recombinant Archaeal Proteasomes and Nondenaturing PAGE:  The Case for a Combined Approach

This protocol uses both subunit coexpression and postlysis subunit mixing for a more thorough examination of recombinant proteasome assembly.

Proteasomes are found in all domains of life. They provide the major route of intracellular protein degradation in eukaryotes, though their assembly is ...

examining-proteasome-assembly-with-recombinant-archaeal-proteasomes

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

6.6K Views.  Indiana University-Purdue University, Indianapolis (IUPUI). This protocol uses both subunit coexpression and postlysis subunit mixing for a more thorough examination of recombinant proteasome assembly.

Video: Examining Proteasome Assembly with Recombinant Archaeal Proteasomes and Nondenaturing PAGE:  The Case for a Combined Approach

Multimer-PAGE: A Method for Capturing and Resolving Protein Complexes in Biological Samples

There are many well-developed methods for purifying and studying single proteins and peptides. However, most cellular functions are carried out by networks of interacting protein complexes, which are often difficult to investigate because their binding is non-covalent and easily perturbed by purification techniques. This work describes a method of stabilizing and separating native protein complexes from unmodified tissue using two-dimensional polyacrylamide gel electrophoresis. Tissue lysate is loaded onto a non-denaturing blue-native polyacrylamide gel, then an electric current is applied until the protein migrates a short distance into the gel. The gel strip containing the migrated protein is then excised and incubated with the amine-reactive cross-linking reagent dithiobis(succinimidyl propionate), which covalently stabilizes protein complexes. The gel strip containing cross-linked complexes is then cast into a sodium dodecyl sulfate polyacrylamide gel, and the complexes are separated completely. The method relies on techniques and materials familiar to most molecular biologists, meaning it is inexpensive and easy to learn. While it is limited in its ability to adequately separate extremely large complexes, and has not been universally successful, the method was able to capture a wide variety of well-studied complexes, and is likely applicable to many systems of interest.

A method for stabilizing and separating native protein complexes from unmodified tissue lysate using an amine-reactive protein cross-linker coupled to a novel two-dimensional polyacrylamide gel electrophoresis (PAGE) system is presented.

There are many well-developed methods for purifying and studying single proteins and peptides. However, most cellular functions are carried out by ...

Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological processes. Affinity purification coupled to mass spectrometry (AP-MS) has become the method of choice for identifying protein-protein interactions. However, conventional AP-MS studies provide information on protein interactions, but the organizational information is lost. To address this issue, we developed a strategy to unravel the distinct functional assemblies a protein might be involved in, by resolving affinity-purified protein complexes prior to their characterization by mass spectrometry. Protein complexes isolated through affinity purification of a bait protein using an epitope tag and competitive elution are separated through blue native electrophoresis. Comparison of protein migration profiles through correlation profiling using quantitative mass spectrometry allows assignment of interacting proteins to distinct molecular entities. This method is able to resolve protein complexes of close molecular weights that might not be resolved by traditional chromatographic techniques such as gel filtration. With little more work than conventional AP-geLC-MS/MS, we demonstrate this strategy may in many cases be adequate for obtaining protein complex topological information concomitantly to identifying protein interactions.

Here we present protocols for affinity purification of protein complexes and their separation by blue native PAGE, followed by protein correlation profiling using label free quantitative mass spectrometry. This method is useful to resolve interactomes into distinct protein complexes.

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological ...

Combining X-Ray Crystallography with Small Angle X-Ray Scattering to Model Unstructured Regions of Nsa1 from S. Cerevisiae

Determination of the full-length structure of ribosome assembly factor Nsa1 from Saccharomyces cerevisiae (S. cerevisiae) is challenging because of the disordered and protease labile C-terminus of the protein. This manuscript describes the methods to purify recombinant Nsa1 from S. cerevisiae for structural analysis by both X-ray crystallography and SAXS. X-ray crystallography was utilized to solve the structure of the well-ordered N-terminal WD40 domain of Nsa1, and then SAXS was used to resolve the structure of the C-terminus of Nsa1 in solution. Solution scattering data was collected from full-length Nsa1 in solution. The theoretical scattering amplitudes were calculated from the high-resolution crystal structure of the WD40 domain, and then a combination of rigid body and ab initio modeling revealed the C-terminus of Nsa1. Through this hybrid approach the quaternary structure of the entire protein was reconstructed. The methods presented here should be generally applicable for the hybrid structural determination of other proteins composed of a mix of structured and unstructured domains.

Combining X-Ray Crystallography with Small Angle X-Ray Scattering to Model Unstructured Regions of Nsa1 from S. Cerevisiae

This method describes the cloning, expression, and purification of recombinant Nsa1 for structural determination by X-ray crystallography and small-angle X-ray scattering (SAXS), and is applicable for the hybrid structural analysis of other proteins containing both ordered and disordered domains.

Determination of the full-length structure of ribosome assembly factor Nsa1 from Saccharomyces cerevisiae (S. cerevisiae) is challenging because of the ...

2 in 1: One-step Affinity Purification for the Parallel Analysis of Protein-Protein and Protein-Metabolite Complexes

Cellular processes are regulated by interactions between biological molecules such as proteins, metabolites, and nucleic acids. While the investigation of protein-protein interactions (PPI) is no novelty, experimental approaches aiming to characterize endogenous protein-metabolite interactions (PMI) constitute a rather recent development. Herein, we present a protocol that allows simultaneous characterization of the PPI and PMI of a protein of choice, referred to as bait. Our protocol was optimized for Arabidopsis cell cultures and combines affinity purification (AP) with mass spectrometry (MS)-based protein and metabolite detection. In short, transgenic Arabidopsis lines, expressing bait protein fused to an affinity tag, are first lysed to obtain a native cellular extract. Anti-tag antibodies are used to pull down protein and metabolite partners of the bait protein. The affinity-purified complexes are extracted using a one-step methyl tert-butyl ether (MTBE)/methanol/water method. Whilst metabolites separate into either the polar or the hydrophobic phase, proteins can be found in the pellet. Both metabolites and proteins are then analyzed by ultra-performance liquid chromatography-mass spectrometry (UPLC-MS or UPLC-MS/MS). Empty-vector (EV) control lines are used to exclude false positives. The major advantage of our protocol is that it enables identification of protein and metabolite partners of a target protein in parallel in near-physiological conditions (cellular lysate). The presented method is straightforward, fast, and can be easily adapted to biological systems other than plant cell cultures.

Protein-protein and protein-metabolite interactions are crucial for all cellular functions. Herein, we describe a protocol that allows parallel analysis of these interactions with a protein of choice. Our protocol was optimized for plant cell cultures and combines affinity purification with mass spectrometry-based protein and metabolite detection.

Cellular processes are regulated by interactions between biological molecules such as proteins, metabolites, and nucleic acids. While the investigation of ...

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

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

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

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

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

Biology

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The introduced protocol provides a tool for the analysis of multiprotein complexes in the thylakoid membrane, by revealing insights into complex composition under different conditions. In this protocol the approach is demonstrated by comparing the composition of the protein complex responsible for cyclic electron flow (CEF) in Chlamydomonas reinhardtii, isolated from genetically different strains. The procedure comprises the isolation of thylakoid membranes, followed by their separation into multiprotein complexes by sucrose density gradient centrifugation, SDS-PAGE, immunodetection and comparative, quantitative mass spectrometry (MS) based on differential metabolic labeling (14N/15N) of the analyzed strains. Detergent solubilized thylakoid membranes are loaded on sucrose density gradients at equal chlorophyll concentration. After ultracentrifugation, the gradients are separated into fractions, which are analyzed by mass-spectrometry based on equal volume. This approach allows the investigation of the composition within the gradient fractions and moreover to analyze the migration behavior of different proteins, especially focusing on ANR1, CAS, and PGRL1. Furthermore, this method is demonstrated by confirming the results with immunoblotting and additionally by supporting the findings from previous studies (the identification and PSI-dependent migration of proteins that were previously described to be part of the CEF-supercomplex such as PGRL1, FNR, and cyt f). Notably, this approach is applicable to address a broad range of questions for which this protocol can be adopted and e.g. used for comparative analyses of multiprotein complex composition isolated from distinct environmental conditions.

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on 15N Metabolic Labeling and Quantitative Mass Spectrometry

The described comparative, quantitative proteomic approach aims at obtaining insights into the composition of multiprotein complexes&#160;under different conditions and is demonstrated by comparing genetically different strains. For quantitative analysis equal volumes of different fractions from a sucrose density gradient are mixed and analyzed by mass spectrometry.

The introduced protocol provides a tool for the analysis of multiprotein complexes in the thylakoid membrane, by revealing insights into complex ...

Genome-wide Protein-protein Interaction Screening by Protein-fragment Complementation Assay (PCA) in Living Cells

Proteins are the building blocks, effectors and signal mediators of cellular processes. A protein&#8217;s function, regulation and localization often depend on its interactions with other proteins. Here, we describe a protocol for the yeast protein-fragment complementation assay (PCA), a powerful method to detect direct and proximal associations between proteins in living cells. The interaction between two proteins, each fused to a dihydrofolate reductase (DHFR) protein fragment, translates into growth of yeast strains in presence of the drug methotrexate (MTX). Differential fitness, resulting from different amounts of reconstituted DHFR enzyme, can be quantified on high-density colony arrays, allowing to differentiate interacting from non-interacting bait-prey pairs. The high-throughput protocol presented here is performed using a robotic platform that parallelizes mating of bait and prey strains carrying complementary DHFR-fragment fusion proteins and the survival assay on MTX. This protocol allows to systematically test for thousands of protein-protein interactions (PPIs) involving bait proteins of interest and offers several advantages over other PPI detection assays, including the study of proteins expressed from their endogenous promoters without the need for modifying protein localization and for the assembly of complex reporter constructs.

Genome-wide Protein-protein Interaction Screening by Protein-fragment Complementation Assay PCA in Living Cells

Proteins interact with each other and these interactions determine in a large part their functions. Protein interaction partners can be identified at high-throughput in vivo using a yeast fitness assay based on the dihydrofolate reductase protein-fragment complementation assay (DHFR-PCA).

Proteins are the building blocks, effectors and signal mediators of cellular processes. A protein&#8217;s function, regulation and localization often ...

Combining Chemical Cross-linking and Mass Spectrometry of Intact Protein Complexes to Study the Architecture of Multi-subunit Protein Assemblies

Proteins interact with their ligands to form active and dynamic assemblies which carry out various cellular functions. Elucidating these interactions is therefore fundamental for the understanding of cellular processes. However, many protein complexes are dynamic assemblies and are not accessible by conventional structural techniques. Mass spectrometry contributes to the structural investigation of these assemblies, and particularly the combination of various mass spectrometric techniques delivers valuable insights into their structural arrangement.
In this article, we describe the application and combination of two complementary mass spectrometric techniques, namely chemical cross-linking coupled with mass spectrometry and native mass spectrometry. Chemical cross-linking involves the covalent linkage of amino acids in close proximity by using chemical reagents. After digestion with proteases, cross-linked di-peptides are identified by mass spectrometry and protein interactions sites are uncovered. Native mass spectrometry on the other hand is the analysis of intact protein assemblies in the gas phase of a mass spectrometer. It reveals protein stoichiometries as well as protein and ligand interactions. Both techniques therefore deliver complementary information on the structure of protein-ligand assemblies and their combination proved powerful in previous studies.

The architecture of protein complexes is essential for their function. Combining various mass spectrometric techniques proved powerful to study their assembly. We provide protocols for chemical cross-linking and native mass spectrometry and show how these complementary techniques help to elucidate the architecture of multi-subunit protein assemblies.

Proteins interact with their ligands to form active and dynamic assemblies which carry out various cellular functions. Elucidating these interactions is ...

In Situ Monitoring of Transiently Formed Molecular Chaperone Assemblies in Bacteria, Yeast, and Human Cells

J-domain proteins (JDPs) form the largest and the most diverse co-chaperone family in eukaryotic cells. Recent findings show that specific members of the JDP family could form transient heterocomplexes in eukaryotes to fine-tune substrate selection for the 70 kDa heat shock protein (Hsp70) chaperone-based protein disaggregases. The JDP complexes target acute/chronic stress induced aggregated proteins and presumably help assemble the disaggregases by recruiting multiple Hsp70s to the surface of protein aggregates. The extent of the protein quality control (PQC) network formed by these physically interacting JDPs remains largely uncharacterized in vivo. Here, we describe a microscopy-based in situ protein interaction assay named the proximity ligation assay (PLA), which is able to robustly capture&#160;these transiently formed chaperone complexes in distinct cellular compartments of eukaryotic cells. Our work expands the employment of PLA from human cells to yeast (Saccharomyces cerevisiae) and bacteria (Escherichia coli), thus rendering an important tool to monitor the dynamics of transiently formed protein assemblies in both prokaryotic and eukaryotic cells.

Cognate J-domain proteins cooperate with the Hsp70 chaperone to assist in a myriad of biological processes ranging from protein folding to degradation. Here, we describe an in situ proximity ligation assay, which allows the monitoring of these transiently formed chaperone machineries in bacterial, yeast and human cells.

J-domain proteins (JDPs) form the largest and the most diverse co-chaperone family in eukaryotic cells. Recent findings show that specific members of the ...

Examining Proteasome Assembly with Recombinant Archaeal Proteasomes and Nondenaturing PAGE: The Case for a Combined Approach

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Multimer-PAGE: A Method for Capturing and Resolving Protein Complexes in Biological Samples

Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling

Combining X-Ray Crystallography with Small Angle X-Ray Scattering to Model Unstructured Regions of Nsa1 from S. Cerevisiae

2 in 1: One-step Affinity Purification for the Parallel Analysis of Protein-Protein and Protein-Metabolite Complexes

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

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

A New Approach for the Comparative Analysis of Multiprotein Complexes Based on ¹⁵N Metabolic Labeling and Quantitative Mass Spectrometry

Genome-wide Protein-protein Interaction Screening by Protein-fragment Complementation Assay (PCA) in Living Cells

Combining Chemical Cross-linking and Mass Spectrometry of Intact Protein Complexes to Study the Architecture of Multi-subunit Protein Assemblies

In Situ Monitoring of Transiently Formed Molecular Chaperone Assemblies in Bacteria, Yeast, and Human Cells