Name: analysis of protein folding, transport, and degradation in living cells by radioactive pulse chase
Uploaded: 2019-02-12T13:30:00.000+00:00
Duration: 8 min 59 s
Description: Watch this Scientific Journal Video about Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase at JoVE.com

The authors thank all members of the Braakman lab, past and present, for their fruitful discussions and help in developing the methods presented in this article. This project has received funding from both the European Research Council under the European Union&#39;s Seventh Framework Programme (FP7/2007-2013) N&#176; 235649 and the Netherlands Organization of Scientific Research (NWO) under the ECHO-program N&#176; 711.012.008.

All radioactive reagents and procedures were handled in accordance with local Utrecht University radiation rules and regulations.
1. Pulse Chase
<ol>
	<li>Pulse Chase for Adherent Cells 
	NOTE: The volumes given here are based on 60 mm cell culture dishes. For 35 mm or 100 mm dishes, multiply the volumes by 1/2 or 2, respectively. This protocol uses a pulse time of 10 min and chase times of 0, 15, 30, 60, 120, and 240 min. These can be varied depending on the specific proteins being studied (discussed below).
	<ol>
		<li>Seed adherent cells (e.g., HEK 293, HeLa, CHO) in six 60&#160;mm cell culture dishes so that they will be subconfluent (80% - 90%) on the day of the pulse chase. Use at least one dish per time point and/or condition.</li>
		<li>If required, transfect the cells with commercially available transfection reagents according to the manufacturer&#39;s instructions; or, virally transduce8 the cells 1 day before the pulse-chase experiment with the appropriate construct for expression.</li>
		<li>Wash the dishes with 2 mL of wash buffer (Hank&#39;s balanced salt solution [HBSS]), add 2 mL of starvation medium (normal culture medium lacking methionine and cysteine and fetal bovine serum [FBS], such as&#160;minimum essential medium [MEM] without methionine and cysteine but with 10 mM HEPES, pH 7.4), and place the dishes in a 37 &#176;C humidified incubator with 5% CO2 for 15 min.</li>
		<li>Transfer the dishes to the racks in a prewarmed 37 &#176;C water bath so that they are in contact with water but do not float. Start a timer.</li>
		<li>At 40 s, aspirate the starvation medium, draw up 600 &#181;L of pulse solution (starvation media + 55 &#181;Ci/35 mm dish label, see the note preceding step 1.1.1) into the pipette, and add it gently to the center of the dish at exactly 1 min. Repeat this step at 1 min intervals for the remaining dishes.&#160;Start with the longest chase sample to save time during your experiment. 
		NOTE: When handling radioactive material, it is essential to follow appropriate precautions and local rules and regulations to prevent accidental exposure and/or contamination.</li>
		<li>At exactly 11 min and for all following dishes, except for the 0 min chase sample, add 2 mL of chase medium directly to the dish, aspirate it, and again, add 2 mL of chase medium. Repeat this step at 1 min intervals for remaining dishes. Transfer all dishes to a 37 &#176;C incubator.</li>
		<li>At exactly 16 min, add 2 mL of chase medium (normal culture medium + 10 mM HEPES [pH 7.4], 5 mM cysteine, and 5 mM methionine) directly to the 0 min sample dish on top of the pulse medium to stop labeling; then, aspirate immediately, transfer the dish to a cooled aluminum plate, and add 2 mL of ice-cold stop buffer (HBSS + 20 mM N-ethylmaleimide [NEM]). 
		NOTE: This is the 0 min chase sample. For this sample, proceed directly to step 1.1.9.</li>
		<li>Transfer each chase dish back to the water bath 2 min before each chase time (e.g., 24 min for a 15 min chase) and, at the exact chase time (e.g., 26 min for a 15 min chase), aspirate the chase media (or transfer it to a microcentrifuge tube if following protein secretion) and transfer the dish to a cooled aluminum plate. Add 2 mL of ice-cold stop buffer.</li>
		<li>Incubate all dishes on ice in the stop buffer for &#8805;5 min; then, aspirate the stop solution and wash it with 2 mL of ice-cold stop solution. Aspirate the wash and lyse dishes with 600 &#181;L of lysis buffer (phosphate-buffered saline [PBS] + nondenaturing detergent [see Table of Materials] + protease inhibitors + 20 mM NEM). Use a cell scraper to ensure that the dishes are lysed quantitatively.</li>
		<li>Transfer the lysate to a 1.5 mL microcentrifuge tube and centrifuge for 10 min at 15,000 - 20,000 x g and 4 &#176;C to pellet the nuclei.</li>
	</ol>
	</li>
	<li>Pulse Chase for Suspension Cells 
	NOTE: To ensure efficient labeling, cells should be pulsed at a concentration of 3 x 106 to 5 x 106 cells/mL, and chase volumes should be 4x the pulse volume. In the following example, we pulsed 5 x 106 cells in a volume of 1 mL for 10 min, to yield five chase time points (0, 15, 30, 60, and 120 min) of 1 mL, containing 1 x 106 cells per time point. All solutions are the same as in section 1.1.
	<ol>
		<li>Culture the suspension cells (e.g., 3T3, Jurkat) according to a previously published protocol9 so that there is a sufficient number of cells at the time of the experiment, at least 1 x 106 cells per time point and/or condition.</li>
		<li>If required, transfect cells with commercially available transfection reagents according to the manufacturer&#39;s instructions, or virally transduce8 the cells 1 day before the pulse-chase experiment with the appropriate construct for expression.</li>
		<li>Pellet 5 x 106 cells per condition in a 50 mL tube for 5 min at 250 x g at room temperature, wash them 1x with 5 mL of starvation media, pellet them again, and resuspend them in 1 mL of starvation media.</li>
		<li>Transfer the cells to a 37 &#176;C water bath and incubate them for 10 - 25 min. Agitate the tubes every 10 - 15 min to prevent the cells from settling at the bottom.</li>
		<li>Start the timer. At exactly 1 min, add 275 &#181;Ci (55 &#181;Ci/1 x 106 cells) of undiluted label directly to the tube containing cells and swirl to mix. 
		NOTE: When handling radioactive material, it is essential to follow appropriate precautions and local rules and regulations to prevent accidental exposure and/or contamination.</li>
		<li>At exactly 11 min, stop labeling by adding 4 mL of chase media. Mix the sample and immediately transfer 1 mL to a 15 mL tube on ice, containing 9 mL of ice-cold stop solution. 
		NOTE: This is the 0 min chase sample.</li>
		<li>Repeat these steps for each successive time point. Once all time points are collected, pellet the cells for 5 min at 250 x g at 4 &#176;C. Aspirate the medium (or transfer it to a new 15 mL tube if following protein secretion). Wash the cells with 5 mL of stop solution and pellet the cells for 5 min at 250 x g at 4 &#176;C.</li>
		<li>Aspirate the stop solution, completely lyse the cells with 300 &#181;L of ice-cold lysis buffer and incubate them for 20 min on ice to ensure a complete lysis. Transfer the lysate to a 1.5 mL microcentrifuge tube and centrifuge for 10 min at 15,000 - 20,000 x g at 4 &#176;C to pellet the nuclei.</li>
	</ol>
	</li>
</ol>
2. Immunoprecipitation
<ol>
	<li>Combine antibody (see the discussion) and 50 &#181;L of immunoprecipitation beads (e.g., protein A-Sepharose) (10% suspension [v/v] in lysis buffer + 0.25% bovine serum albumin [BSA]) in a microcentrifuge tube and incubate them at 4 &#176;C for ~30 min in a shaker.</li>
	<li>Pellet the beads for 1 min at 12,000 x g at room temperature and aspirate the supernatant. Add 200 &#181;L of lysate to the antibody-bead mixture and incubate at 4 &#176;C in a shaker for 1 h or head-over-head if the immunoprecipitation requires &#62;1 h.</li>
	<li>Pellet the beads for 1 min at 12,000 x g at room temperature. Aspirate the supernatant and add 1 mL of immunoprecipitation wash buffer. Place the sample in a shaker at room temperature for 5 min.</li>
	<li>Pellet the beads as described in step 2.3 and repeat the wash 1x. Then, aspirate the supernatant and resuspend the beads in 20 &#181;L of TE buffer, pH 6.8 (10 mM Tris-HCl, 1 mM EDTA). Vortex the sample.</li>
	<li>Add 20 &#181;L of 2x sample buffer without the reducing agent, vortex it, heat it for 5 min at 95 &#176;C, and vortex again. 
	NOTE: If only preparing reducing samples, 2 &#181;L of 500 mM DTT should be added at this point. Then, proceed to step 2.7.</li>
	<li>Pellet the beads as described in step 2.3. Transfer 19 &#181;L of the nonreduced supernatant to a fresh microcentrifuge tube containing 1 &#181;L of 500 mM DTT, centrifuge the sample, and vortex it before heating it for 5 min at 95 &#176;C again.</li>
	<li>Spin down the sample for 1 min at 12,000 x g; this is the reduced sample. Cool it down to room temperature and add 1.1 &#181;L of 1 M NEM to both the reduced and the nonreduced sample. Vortex and spin down the samples.</li>
</ol>
3. SDS-PAGE
<ol>
	<li>First, determine the appropriate SDS-PAGE resolving gel percentage for the protein of interest. For example, HIV-1 gp120, when deglycosylated, runs at ~60 kDa and is analyzed in a 7.5% gel.</li>
	<li>Prepare the resolving gel mixture without TEMED according to the manufacturer&#39;s instructions (x% acrylamide, 375 mM Tris-HCl [pH 8.8,] 0.1% SDS [w/v], and 0.05% ammonium persulfate [APS] [w/v]) and degas under vacuum for &#62;15 min. While the gel mixture is degassing, thoroughly clean the gel glass plates with 70% ethanol and lint-free tissues and place them into a casting apparatus.</li>
	<li>Add TEMED to the resolving gel mixture (at a final concentration of 0.005% [v/v]), mix thoroughly, and pipette between glass plates, leaving ~1.5 cm space for the stacking gel. Carefully overlay the gel with deionized H2O or isopropanol and leave it to polymerize.</li>
	<li>Once the resolving gel has polymerized, prepare the stacking gel mixture (4% acrylamide, 125 mM Tris-HCl [pH 6.8], 0.1% SDS [w/v], and 0.025% APS [w/v]).</li>
	<li>Flush the top of the resolving gel with deionized H2O and, then, remove all water. 
	NOTE: Use filter paper to remove the last drops.</li>
	<li>Add TEMED to the stacking gel mixture (0.005% [v/v]), mix thoroughly, and overlay the resolving gel with stacking gel and insert a 15-well comb. Once the stacking gel has polymerized, transfer it to a running chamber and fill the upper and lower chambers with running buffer (25 mM Tris-HCl, 192 mM glycine [pH 8.3], and 0.1% SDS [w/v]).</li>
	<li>Load 10 &#181;L of sample per lane in a 15-lane minigel. Avoid loading the samples in the first and the last lane on the gel and load the nonreducing sample buffer in all empty lanes to prevent the smiling of bands. Run the gels at constant a 25 mA/gel until the dye front is at the bottom of the gel.</li>
	<li>Remove the gels from the glass plates, stain the gels with protein-staining solution (10% acetic acid and 30% methanol in H2O + 0.25% brilliant blue R250 [w/v]) for 5 min with agitation, and then, destain for 30 min with destaining solution (staining solution without brilliant blue R250).</li>
	<li>Arrange the gels face-down on a plastic wrap and, then, place 0.4 mm chromatography paper on top of them. Place the gel sandwich chromatography paper-side down onto a gel dryer. Following the manufacturer&#39;s instructions, dry the gels for 2 h at 80 &#176;C.</li>
	<li>Transfer the dried gels to a cassette and overlay them with autoradiography film or phosphor screen. If using autoradiography film, this step must be performed in a dark room.</li>
</ol>

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N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=N-Glycan-based+ER+Molecular+Chaperone+and+Protein+Quality+Control+System:+The+Calnexin+Binding+Cycle.">N-Glycan-based ER Molecular Chaperone and Protein Quality Control System: The Calnexin Binding Cycle.</a> Traffic. 17 (4), 308-326 (2016).</li><li>Snapp, E. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+and+topology+around+the+cleavage+site+regulate+post-translational+cleavage+of+the+HIV-1+gp160+signal+peptide.">Structure and topology around the cleavage site regulate post-translational cleavage of the HIV-1 gp160 signal peptide.</a> Elife. 6, 26067 (2017).</li><li>Braakman, I., Hoover-Litty, H., Wagner, K. R., Helenius, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Folding+of+influenza+hemagglutinin+in+the+endoplasmic+reticulum.">Folding of influenza hemagglutinin in the endoplasmic reticulum.</a> Journal of Cell Biology. 114 (3), 401-411 (1991).</li><li>Jansens, A., van Duijn, E., Braakman, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coordinated+nonvectorial+folding+in+a+newly+synthesized+multidomain+protein.">Coordinated nonvectorial folding in a newly synthesized multidomain protein.</a> Science. 298 (5602), 2401-2403 (2002).</li><li>Land, A., Braakman, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Folding+of+the+human+immunodeficiency+virus+type+1+envelope+glycoprotein+in+the+endoplasmic+reticulum.">Folding of the human immunodeficiency virus type 1 envelope glycoprotein in the endoplasmic reticulum.</a> Biochimie. 83 (8), 783-790 (2001).</li><li>Singh, Y., Garden, O. 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N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=N-linked+glycans+direct+the+cotranslational+folding+pathway+of+influenza+hemagglutinin.">N-linked glycans direct the cotranslational folding pathway of influenza hemagglutinin.</a> Molecular Cell. 11 (1), 79-90 (2003).</li><li>Ferreira, L. R., Norris, K., Smith, T., Hebert, C., Sauk, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Association+of+Hsp47,+Grp78,+and+Grp94+with+procollagen+supports+the+successive+or+coupled+action+of+molecular+chaperones.">Association of Hsp47, Grp78, and Grp94 with procollagen supports the successive or coupled action of molecular chaperones.</a> Journal of Cellular Biochemistry. 56 (4), 518-526 (1994).</li><li>Hoelen, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+primary+folding+defect+and+rescue+of+DeltaF508+CFTR+emerge+during+translation+of+the+mutant+domain.">The primary folding defect and rescue of DeltaF508 CFTR emerge during translation of the mutant domain.</a> PLoS One. 5 (11), 15458 (2010).</li><li>Kleizen, B., van Vlijmen, T., de Jonge, H. R., Braakman, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Folding+of+CFTR+is+predominantly+cotranslational.">Folding of CFTR is predominantly cotranslational.</a> Molecular Cell. 20 (2), 277-287 (2005).</li><li>Hagiwara, M., Ling, J., Koenig, P. A., Ploegh, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Posttranscriptional+Regulation+of+Glycoprotein+Quality+Control+in+the+Endoplasmic+Reticulum+Is+Controlled+by+the+E2+Ub-Conjugating+Enzyme+UBC6e.">Posttranscriptional Regulation of Glycoprotein Quality Control in the Endoplasmic Reticulum Is Controlled by the E2 Ub-Conjugating Enzyme UBC6e.</a> Molecular Cell. 63 (5), 753-767 (2016).</li><li>Copeland, C. S., Doms, R. W., Bolzau, E. M., Webster, R. 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The authors have nothing to disclose.

Pulse-chase methods have been essential for developing scientists&#39; understanding of protein folding in intact cells. While we have attempted to provide a method that is as general as possible, this approach has the potential for almost limitless variations to study various processes that occur during the folding, the transport, and the life of proteins inside the cell.
When performing a pulse chase using adherent cells in dishes, it is essential to treat each dish the same as much as possible, as a separate dish is used for each time point and/or condition in an experiment. Especially for short pulse and chase times (&#60;5 min), it is essential to maintain a tight control over the pulse and chase times (with a digital timer). Pulse chases with suspension cells can help to reduce this variability as all samples are taken from a single tube. As some cell types adhere less well to culture dishes than others, care should be taken to ensure that the cells are not washed away during the various medium changes. This can be overcome by either using these cells in suspension or by coating the dishes with poly-L lysine or gelatin, for example, to adhere the cells to the culture dishes. To measure the reproducibility of labeling between dishes, a sample of lysate should be examined by SDS-PAGE to check all samples for identical total labeling and protein pattern. Alternatively, liquid scintillation counting of lysates will establish the total counts-per-minute (cpm) incorporated but will not provide information on the protein population labeled.
If a target protein labels poorly, the signal is increased by increasing the number of cells (or dishes) rather than by increasing the amount of label. Lengthening the pulse time is an option when the kinetics of the studied process will allow this. The ideal pulse time will vary with each target owing to factors such as the expression level, the transcription rate, the number of methionines/cysteines, and the folding rate of the protein. As such, experimentation must be undertaken to determine the best balance between the above-mentioned factors in each experiment while keeping the experimental aim in mind. If intermediates during folding are being studied, for example, it is desirable to pulse the label for as short a time as possible to have the starting material as close to an unfolded state as possible, while balancing expression levels. Alternatively, if the transport or degradation of a protein is the target of study, pulse times may be lengthened to provide the highest levels of signal possible without compromising kinetic information. In general, pulse times can range from 2 - 15 min when using this protocol without modification. Previous experimentation5 has demonstrated a lag time of ~10 s after the pulse before the incorporation of radioactivity into the total protein pool; this is important to keep in mind when deriving kinetic information from pulse-chase experiments.&#160;For short pulse times (&#60; 2 min), keep all the dishes on the water bath during the entire pulse time. If extended pulse times (&#62;1 h) are being used, it is advisable to increase the pulse volume by 1.5 times and place dishes on a rocker in a 37 &#176;C incubator during the pulse to prevent the dishes from drying out. While labeling solutions containing individual 35S-methionine/cysteine or a mix are available, the mix is preferred even when proteins that only contain methionine or cysteine are being labeled as both are needed to maintain general protein synthesis in the cell. If experimental conditions require specific methionine/cysteine labeling, the levels of nonradioactive methionine/cysteine during the pulse must be adjusted accordingly.
Chase times (and the number of time points) are determined in a similar fashion. A good starting place is to set the first chase time (after the 0 min chase) as equal to the pulse time and, then, double the length of time for each successive time point (e.g., 5, 10, 20, and 40 min). However, this must be optimized for each specific protein and question.
To prevent the activation of stress responses by starvation (which may affect protein synthesis and folding), ideally, some unlabeled methionine and cysteine should be added to the starvation and radiolabeling solutions. The quantity will depend on the cell line, the labeling time, the quantity of the radiolabel used, and media volumes, and will need testing. A good starting point is 1% of the amount of cysteine and methionine present in the cell culture medium, which can be increased with increasing pulse times. Starvation periods, whether with or without unlabeled amino acids, should be kept in the range of 15 - 30 min to ensure adequate label incorporation and prevent the activation of stress responses. When using an extended pulse (&#8805;1 h), starvation is not required.
The lysis buffer described here will be suitable for most purposes, but in principle, any buffer system, such as HEPES, Tris, or MES, will work. The detergent concentration needed to lyse cells will depend on the number of cells and detergent volume. The concentration should always be above the critical micelle concentration (CMC) and the quantity of detergent sufficient to lyse cells. An empirical rule of thumb is that 200 &#181;L of 0.5% nondenaturing detergent (see Table of Materials) is sufficient to lyse the cell equivalent of ~1 mg of protein (~1 x 106 cells). The salt concentration and detergents may also be varied according to the application, but in general, conditions that break open nuclei (high salt, &#62;0.1% SDS) should be avoided as the presence of free DNA will interfere with immunoprecipitation. If this cannot be avoided, for instance, when complete cell lysates including nuclei or pelleted proteins need to be analyzed, DNA can be sheared by passing the cells through a small gauge needle prior to immunoprecipitation. This is preferred over sonication so as to prevent excessive foaming and the production of radioactive aerosols.
For immunoprecipitation, both the optimal quantity of antibodies and the best wash buffer for each antibody-antigen combination must be tested thoroughly to achieve a balance between the desired antigen signal and background. Antibodies should always be present in excess over antigens to ensure quantitative immunoprecipitation; a good starting point is 1 &#181;g of antibodies per immunoprecipitation from a 35 mm dish/1 x 106 cells. The background can be decreased by increasing the detergent or salt concentrations. In particular, the addition of &#8804;0.1% SDS can help to greatly reduce the background but may also disrupt antibody-antigen interactions. To control for the specificity of antibodies during immunoprecipitation, identical cells lacking the target protein should be used when available. If this is not straightforward, such as when endogenous proteins are being analyzed, the overexpression or depletion of the antigen will work. To control for background (and specificity), either preimmune sera (where possible) or isotype controls should be used. To control for antigens binding directly to immunoprecipitation beads, beads without antibody should also be used. The conditions provided here should be considered as a starting point as each antibody-antigen pair will require an optimization of both wash buffer and wash conditions. In case of too much background, do not increase the number of washes but rather the time of washing and/or the composition of the wash buffer. If particularly sensitive interactions such as in coimmunoprecipitations have to be dealt with, it may be preferable to carry out all wash steps at 4 &#176;C, using ice-cold buffers.
One advantage of adherent cells over suspension cells is that drug treatments can be performed at virtually any point during the pulse chase. If used with chaperone inhibitors, for example, it is possible to differentiate its effects on the co- versus posttranslational phases of a folding process. This can be extended by adapting the lysis and immunoprecipitation wash buffers (discussed above) to allow the coimmunoprecipitation of protein-chaperone complexes during folding10,11,12.
The postlysis treatment of samples will widen the scope of questions that can be addressed. The limited protease digestion of target proteins before immunoprecipitation will provide additional conformational information and has been successfully used to monitor the folding of proteins, especially those lacking disulfide bonds13,14. Protein aggregation and complex formation can be monitored by sucrose density-gradient centrifugation prior to immunoprecipitation15,16.
To take full advantage of pulse chases, the availability of high-quality reagents is required for immunoprecipitations. When examining folding intermediates, it is essential to have antibodies that are capable of pulling down all forms of the protein under analysis. If specific antibodies to the target protein are not available, this can be substituted by using affinity tags. However, this severely restricts the usefulness of postlysis methods, such as the limited proteolysis, to directly probe the protein conformation, as only tagged fragments can be recovered.
The sensitivity of pulse chases is limited by the number of methionine and cysteine residues in the protein in question. Proteins with low numbers of methionine/cysteine residues coupled with a low expression level are undetectable in pulse-chase experiments. When performing postlysis modifications such as limited proteolysis, only methionine/cysteine-containing fragments will be detectable.
To conclude, given the kinetic detail that pulse chases provide, they are complementary to many other techniques and tools used to study molecular cell biology at the steady-state level. As such, they continue to be an invaluable component in the molecular biologist&#39;s toolbox.

The folding of even relatively simple proteins involves many different folding enzymes, molecular chaperones, and covalent modifications1. A complete reconstitution of these processes in vitro is practically impossible, given the vast number of different components involved. It is highly desirable, therefore, to study protein folding in vivo, in live cells. Radioactive pulse-chase techniques prove a powerful tool for studying the synthesis, folding, transport, and degradation of proteins in their natural environment.
The metabolic labeling of proteins during a short pulse with 35S-labeled methionine/cysteine, followed by a chase in the absence of a radioactive label, allows specific tracking of a population of newly synthesized proteins in the wider cellular milieu. Then, target proteins can be isolated via immunoprecipitation and analyzed via SDS-PAGE or other techniques. For many proteins, their journey through the cell is marked by modifications that are visible on SDS-PAGE gel. For example, the transport of glycosylated proteins from the endoplasmic reticulum (ER) to the Golgi complex is often accompanied by modifications of N-linked glycans or the addition of O-linked glycans2,3. These modifications cause large increases in the apparent molecular mass, which can be seen by mobility changes in SDS-PAGE. Maturation can also be marked by proteolytic cleavages, such as signal-peptide cleavage or the removal of pro-peptides, resulting in changes in the apparent molecular mass that can be followed easily on SDS-PAGE gel4. Radioactivity has considerable advantages over comparable techniques such as cycloheximide chases, where novel protein synthesis is prevented, as longer treatments are toxic to cells and do not exclude the majority of older, steady-state proteins from the analysis, as some proteins have half-lives of days. The comparison of proteins under both nonreducing and reducing conditions allows the analysis of disulfide bond formation, an important step in the folding of many secretory proteins4,5,6,7.
Here we describe a general method for the analysis of protein folding and transport in intact cells, using a radioactive pulse-chase approach. While we have aimed to provide the method as detailed as possible, the protocol has an almost limitless potential for adaptability and will allow optimization to study each reader&#39;s specific proteins.
Two alternative pulse-chase protocols, one for adherent cells (step 1.1 of the protocol presented here) and one for suspension cells (step 1.2 of the protocol presented here) are provided. The conditions provided here are sufficient to visualize a protein expressed with medium- to high-expression levels. If the reader is working with poorly expressed proteins or various posttreatment conditions, such as multiple immunoprecipitations, it is necessary to increase the dish size or cell number appropriately.
For suspension pulse chase, the chase samples taken at each time point are all taken from a single tube of cells. The wash steps after the pulse are omitted; instead, further incorporation of 35S is prevented by dilution with a high excess of unlabeled methionine and cysteine.
The presented protocols use radioactive 35S-labeled cysteine and methionine to follow cellular protein-folding processes. All operations with radioactive reagents should be performed using appropriate protective measures to minimize any exposure of the operator and the environment to radioactive radiation and be performed in a designated laboratory. As the pulse-chase labeling technique is relatively inefficient at short pulse times (&#60;15 min), less than 1% of the starting amount of radioactivity is incorporated in the newly synthesized proteins. After the enrichment of the target protein via immunoprecipitation, the sample for SDS-PAGE contains less than 0.05% of the starting amount of radioactivity.
Although the 35S methionine and cysteine labeling mix is stabilized, some decomposition, yielding volatile radioactive compounds, will occur. To protect the researcher and the apparatus, some precautions should be taken. The researcher should always obey the local radiation safety rules and may wear a charcoal nursing mask, besides a lab coat and (double) gloves. Stock vials with 35S methionine and cysteine should always be opened in a fume hood, or under a local aspiration point. Known laboratory contamination spots are centrifuges, pipettes, water baths, incubators, and shakers. The contamination of these areas is reduced by using pipette tips with a charcoal filter, positive-seal microcentrifuge tubes (see Table of Materials), aquarium charcoal sponges in water baths, charcoal filter papers glued in the pulse-chase dishes, charcoal guard in the aspiration system, and the placement of dishes containing charcoal grains in incubators and storage containers.

<table><tbody><tr><td>1.5 mL safeseal microcentrifuge tubes</td><td>Sarstedt</td><td>72.706.400</td><td></td></tr><tr><td>Acetic Acid</td><td>Sigma</td><td>A6283</td><td>glacial acetic acid</td></tr><tr><td>BAS Storage phosphor screen 20x25 cm</td><td>GE Life Sciences</td><td>28956475</td><td></td></tr><tr><td>Bromophenol Blue</td><td>Sigma</td><td>B8026</td><td>Molecular biology grade</td></tr><tr><td>Carestream Biomax MR films</td><td>Kodak</td><td>Z350370-50EA</td><td></td></tr><tr><td>Cell-culture media</td><td>Various</td><td>N/A</td><td>Normal cell culture media for specific cell-lines used</td></tr><tr><td>Cell-culture media, no methionine/cysteine</td><td>Various</td><td>N/A</td><td>Same media formulation as normal culture media e.g DMEM/MEM/RPMI, lacking methionine and cysteine</td></tr><tr><td>Charcoal filter paper</td><td>Whatman</td><td>1872047</td><td></td></tr><tr><td>Charcoal filtered pipette tips</td><td>Molecular bioproducts</td><td>5069B</td><td></td></tr><tr><td>Charcoal vacu-guard</td><td>Whatman</td><td>67221001</td><td></td></tr><tr><td>Coomassie Brilliant Blue R250</td><td>Sigma</td><td>112,553</td><td>for electrophoresis</td></tr><tr><td>Cysteine</td><td>Sigma</td><td>C7352</td><td>Molecular biology grade, Make 500 mM stock, store at -20&amp;nbsp;</td></tr><tr><td>Dithiothreitol (DTT)</td><td>Sigma</td><td>10197777001</td><td>Molecular biology grade</td></tr><tr><td>EasyTag Express35S Protein Labeling Mix</td><td>Perkin Elmer</td><td>NEG772014MC</td><td>Other size batches of label are available depending on useage</td></tr><tr><td>EDTA</td><td>Sigma</td><td>E1644</td><td>Molecular biology grade</td></tr><tr><td>Gel-drying equipment</td><td>Various</td><td>N/A</td><td></td></tr><tr><td>Glycerol</td><td>Sigma</td><td>G5516</td><td>Molecular biology grade</td></tr><tr><td>Grade 3 chromatography paper</td><td>GE Life Sciences</td><td>3003-917</td><td></td></tr><tr><td>Hank&#039;s Balanced Salt Solution (HBSS)</td><td>Gibco</td><td>24020117</td><td></td></tr><tr><td>Kimwipes delicate task wipes</td><td>VWR</td><td>21905-026</td><td></td></tr><tr><td>MES</td><td>Sigma&amp;nbsp;</td><td>M3671</td><td>Molecular biology grade</td></tr><tr><td>Methanol</td><td>Sigma</td><td>MX0490&amp;nbsp;</td><td></td></tr><tr><td>Methionine</td><td>Sigma</td><td>M5308</td><td>Molecular biology grade, Make 250 mM stock, store at -20</td></tr><tr><td>Minigel casting/running equipment</td><td>Various</td><td>N/A</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653</td><td>Molecular biology grade</td></tr><tr><td>N-ethylmaleimide</td><td>Sigma</td><td>E3876</td><td>Molecular biology grade, Make 1M stock in 100% ethanol, store at -20</td></tr><tr><td>PBS</td><td>Sigma</td><td>P5368</td><td>Molecular biology grade</td></tr><tr><td>Protein-A Sepharose fastflow beads</td><td>GE health-care</td><td>17-5280-04</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate (SDS)</td><td>Sigma</td><td>L3771</td><td>Molecular biology grade</td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>T8787</td><td>Molecular biology grade</td></tr><tr><td>Trizma base (Tris)</td><td>Sigma</td><td>T6066</td><td>Molecular biology grade</td></tr><tr><td>Typhoon IP Biomolecular imager</td><td>Amersham</td><td>29187194</td><td></td></tr><tr><td>Unwire Test Tube Rack 20 mm for waterbath</td><td>Nalgene</td><td>5970-0320PK</td><td></td></tr></tbody></table>

analysis of protein folding, transport, and degradation in living cells by radioactive pulse chase

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and the degradation of target proteins in live cells. By using short (pulse) radiolabeling times (&lt;30 min) and tightly controlled chase times, it is possible to label only a small fraction of the total protein pool and follow its folding. When combined with nonreducing/reducing SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoprecipitation with (conformation-specific) antibodies, folding processes can be examined in great detail. This system has been used to analyze the folding of proteins with a huge variation in properties such as soluble proteins, single and multi-pass transmembrane proteins, heavily N- and O-glycosylated proteins, and proteins with and without extensive disulfide bonding. Pulse-chase methods are the basis of kinetic studies into a range of additional features, including co- and posttranslational modifications, oligomerization, and polymerization, essentially allowing the analysis of a protein from birth to death. Pulse-chase studies on protein folding are complementary with other biochemical and biophysical methods for studying proteins in vitro by providing increased temporal resolution and physiological information. The methods as described within this paper are adapted easily to study the folding of almost any protein that can be expressed in mammalian or insect-cell systems.

The folding and secretion of HIV-1 gp120 from an adherent pulse chase is shown in Figure 2. The nonreducing gel (Cells NR in the figure) shows the oxidative folding of gp120. Immediately after the pulse labeling of 5 min (0 min chase) gp120 appears as a diffuse band higher in the gel, and as the chase progresses, the band migrates down the gel through even more diffused folding intermediates (IT) until it accumulates in the tight band (NT) that represents natively folded gp120. This occurs as the formation of disulfide bonds increases the compactness of the protein, causing it to migrate faster than the fully reduced protein. On the reducing gel (Cells R in the figure), the disulfide bonds on all forms have been reduced such that, at all chase times, they do not affect mobility. This allows the analysis of other modifications. The shift over time from Ru to Rc represents the posttranslational signal-peptide cleavage of gp120: the mobility increases due to the loss of the signal peptide, which increases during the chase as more proteins attain the native fold and lose their signal peptide. On both the nonreducing and reducing gel, the signal begins to decrease from ~1 h onward due to the secretion of gp120. This can be monitored by analyzing the media (Medium in the figure). A comparison of the nonreducing and reducing gels uncovers disulfide bond changes and signal-peptide removal. This was only possible because another modification, N-linked glycosylation, and glycan modifications, were removed from gp120 (and the analysis) by digestion with endoglycosidase H just before SDS-PAGE.
The trafficking of the &#181; heavy chain of immunoglobulin M (IgM) from a suspension pulse chase is shown in Figure 3. A shift over time from HCER to HCGolgi represents the trafficking of the &#181; chain from the ER to the Golgi, which precedes secretion from the cell. This change in molecular mass is caused by the modification of N-linked glycans in the Golgi.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58952/58952fig1.jpg" /> 
Figure 1: Schematic diagram of the protocol for pulse chase, immunoprecipitation, and SDS-PAGE. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58952/58952fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58952/58952fig2.jpg" /> 
Figure 2:&#160;Folding and secretion of HIV-1 gp120, determined by adherent pulse chase. Subconfluent 60&#160;mm dishes of HeLa cells expressing HIV-1 gp120 were pulse labeled for 10&#160;min and chased for the indicated times. After the immunoprecipitation of gp120, the samples were deglycosylated and analyzed using a 7.5% SDS-PAGE gel. IT = folding intermediates; NT = native gp120; Ru = reduced signal-peptide-uncleaved gp120; Rc = reduced signal-peptide-cleaved gp120; NR: = nonreducing; R = reducing. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58952/58952fig2large.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/58952/58952fig3.jpg" /> 
Figure 3: Trafficking of the heavy chain of IgM, determined by suspension pulse chase. 5 x 106 I.29 &#181;+ B cells were pulse labeled for 5 min and chased for the indicated times. After immunoprecipitation, the samples were analyzed using a 10% SDS-PAGE gel. HC = immunoglobulin &#181; heavy chain; LC = immunoglobulin light chain. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58952/58952fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase

Here we describe a protocol for a general pulse-chase method that allows the kinetic analysis of folding, transport, and degradation of proteins to be followed in live cells.

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and ...

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Unit 1

Transmembrane Transport in Endoplasmic Reticulum and Peroxisomes

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11.1K Views.  Utrecht University. De radioactieve pulsjacht met behulp van 35 S-gelabelde methionine en cysteïne is nog steeds de enige biochemische methoden om eiwitbiosynthese met de tijd in levende cellen te onderzoeken. Eiwitbiosynthese omvat zowel vertaling, vouwen en assemblage, als handel en degradatie. Het combineren van de radioactieve pulsjacht met analyse van co-en post-translation eiwitmodificaties, zoals disulfide bindingsvorming en acetylatie, biedt een gevoelige en kwantitatieve test om het geloof van het eiwi...

Video: Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Developmental Biology

Single-Cell Quantification of Protein Degradation Rates by Time-Lapse Fluorescence Microscopy in Adherent Cell Culture

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used approaches to determine protein half-life are either limited to population averages from lysed cells or require the use of protein synthesis inhibitors. This protocol describes a method to measure protein half-lives in single living adherent cells, using SNAP-tag fusion proteins in combination with fluorescence time-lapse microscopy. Any protein of interest fused to a SNAP-tag can be covalently bound by a fluorescent, cell permeable dye that is coupled to a benzylguanine derivative, and the decay of the labeled protein population can be monitored after washout of the residual dye. Subsequent cell tracking and quantification of the integrated fluorescence intensity over time results in an exponential decay curve for each tracked cell, allowing for determining protein degradation rates in single cells by curve fitting. This method provides an estimate for the heterogeneity of half-lives in a population of cultured cells, which cannot easily be assessed by other methods. The approach presented here is applicable to any type of cultured adherent cells expressing a protein of interest fused to a SNAP-tag. Here we use mouse embryonic stem (ES) cells grown on E-cadherin-coated cell culture plates to illustrate how single cell degradation rates of proteins with a broad range of half-lives can be determined.

This protocol describes a method to determine protein half-lives in single living adherent cells, using pulse labeling and fluorescence time-lapse imaging of SNAP-tag fusion proteins.

Proteins are in a dynamic state of synthesis and degradation and their half-lives can be adjusted under various circumstances. However, most commonly used ...

Monitoring the Assembly of a Secreted Bacterial Virulence Factor Using Site-specific Crosslinking

This article describes a method to detect and analyze dynamic interactions between a protein of interest and other factors in vivo. Our method is based on the amber suppression technology that was originally developed by Peter Schultz and colleagues1. An amber mutation is first introduced at a specific codon of the gene encoding the protein of interest. The amber mutant is then expressed in E. coli together with genes encoding an amber suppressor tRNA and an amino acyl-tRNA synthetase derived from Methanococcus jannaschii. Using this system, the photo activatable amino acid analog p-benzoylphenylalanine (Bpa) is incorporated at the amber codon. Cells are then irradiated with ultraviolet light to covalently link the Bpa residue to proteins that are located within 3-8 &#197;. Photocrosslinking is performed in combination with pulse-chase labeling and immunoprecipitation of the protein of interest in order to monitor changes in protein-protein interactions that occur over a time scale of seconds to minutes. We optimized the procedure to study the assembly of a bacterial virulence factor that consists of two independent domains, a domain that is integrated into the outer membrane and a domain that is translocated into the extracellular space, but the method can be used to study many different assembly processes and biological pathways in both prokaryotic and eukaryotic cells. In principle interacting factors and even specific residues of interacting factors that bind to a protein of interest can be identified by mass spectrometry.

This article illustrates the use of pulse-chase radio labeling in combination with site-specific photocrosslinking to monitor interactions between a protein of interest and other factors in E. coli. Unlike traditional chemical cross-linking methods, this approach generates high resolution &#8220;snapshots&#8221; of an ordered assembly pathway in a living cell.

This article describes a method to detect and analyze dynamic interactions between a protein of interest and other factors in vivo. Our method is based on ...

Immunology and Infection

Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using &#967;CRAC

The interaction between RNA-binding proteins (RBPs) and their RNA substrates exhibits fluidity and complexity. Within its lifespan, a single RNA can be bound by many different RBPs that will regulate its production, stability, activity, and degradation. As such, much has been done to understand the dynamics that exist between these two types of molecules. A particularly important breakthrough came with the emergence of &#8216;cross-linking and immunoprecipitation&#8217; (CLIP). This technique allowed stringent investigation into which RNAs are bound by a particular RBP. In short, the protein of interest is UV cross-linked to its RNA substrates in vivo, purified under highly stringent conditions, and then the RNAs covalently cross-linked to the protein are converted into cDNA libraries and sequenced. Since its conception, many derivative techniques have been developed in order to make CLIP amenable to particular fields of study. However, cross-linking using ultraviolet light is notoriously inefficient. This results in extended exposure times that make the temporal study of RBP-RNA interactions impossible. To overcome this issue, we recently designed and built much-improved UV irradiation and cell harvesting devices. Using these new tools, we developed a protocol for time-resolved analyses of RBP-RNA interactions in living cells at high temporal resolution: Kinetic CRoss-linking and Analysis of cDNAs (&#967;CRAC). We recently used this technique to study the role of yeast RBPs in nutrient stress adaptation. This manuscript provides a detailed overview of the &#967;CRAC method and presents recent results obtained with the Nrd1 RBP.

Kinetic cross-linking and analysis of cDNA is a method that allows investigation of the dynamics of protein-RNA interactions in living cells at high temporal resolution. Here the protocol is described in detail, including the growth of yeast cells, UV cross-linking, harvesting, protein purification, and next generation sequencing library preparation steps.

The interaction between RNA-binding proteins (RBPs) and their RNA substrates exhibits fluidity and complexity. Within its lifespan, a single RNA can be ...

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

Biology

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

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

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

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

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must be maintained to avoid deleterious consequences 1. External or internal factors that disrupt this balance can lead to protein aggregation, toxicity and cell death. In humans this is a major contributing factor to the symptoms associated with neurodegenerative disorders such as Huntington's, Parkinson's, and Alzheimer's diseases 10. It is therefore essential that the proteins involved in maintenance of proteostasis be identified in order to develop treatments for these debilitating diseases. This article describes techniques for monitoring in vivo protein folding at near-real time resolution using the model protein firefly luciferase fused to green fluorescent protein (FFL-GFP). FFL-GFP is a unique model chimeric protein as the FFL moiety is extremely sensitive to stress-induced misfolding and aggregation, which inactivates the enzyme 12. Luciferase activity is monitored using an enzymatic assay, and the GFP moiety provides a method of visualizing soluble or aggregated FFL using automated microscopy. These coupled methods incorporate two parallel and technically independent approaches to analyze both refolding and functional reactivation of an enzyme after stress. Activity recovery can be directly correlated with kinetics of disaggregation and re-solubilization to better understand how protein quality control factors such as protein chaperones collaborate to perform these functions. In addition, gene deletions or mutations can be used to test contributions of specific proteins or protein subunits to this process. In this article we examine the contributions of the protein disaggregase Hsp104 13, known to partner with the Hsp40/70/nucleotide exchange factor (NEF) refolding system 5, to protein refolding to validate this approach.

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

This article describes the use of a firefly luciferase-GFP fusion protein to investigate in vivo protein folding in Saccharomyces cerevisiae. Using this reagent, refolding of a model heat-denatured protein can be monitored simultaneously by fluorescence microscopy and an enzymatic assay to probe the roles of proteostasis network components in protein quality control.

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must ...

Assays for the Degradation of Misfolded Proteins in Cells

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded proteins may serve as potential therapeutic targets. To distinguish degradation of misfolding-prone proteins from other mechanisms that regulate their levels, one important method is to measure protein half-life in cells. However, this can be challenging because misfolding-prone proteins may exist in different forms, including the native form and misfolded forms of distinct characteristics. Here we describe assays to examine the half-life of misfolded proteins in mammalian cells using a highly aggregation-prone protein, Ataxin-1 with an extended polyglutamine (polyQ) stretch, and a conformationally unstable luciferase mutant as models. Cycloheximide chase is combined with cell fractionation to examine the turnover rate of misfolding-prone proteins in various cellular fractions. We further depict a fluorescence-based assay using an enhanced green fluorescence protein (EGFP)-fusion of the luciferase mutant, which can be adapted for high throughput screening on a microplate-reader.

This report describes protocols for measuring degradation rates of misfolded proteins by either western blot or fluorescence-based assays. The methods can be applied to analysis of other misfolded proteins and for high throughput screening.

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded ...

Fluorescence Microplate-Based Cycloheximide Chase Assay: A Technique to Monitor the Degradation Kinetics of Fluorescent Nuclear Misfolded Proteins

Source: Guo, L. et. al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/54266">Assays for the Degradation of Misfolded Proteins in Cells.</a> J. Vis. Exp. (2016)
This video describes an in vitro fluorescence-based microplate assay to study the degradation kinetics of a fluorescently-labeled misfolded luciferase mutant protein expressed in the nuclei of transfected mammalian cells. The assay involves the treatment of cells expressing the fluorescent mutant protein with a translation inhibitor and a proteasome inhibitor to assess the proteasome-mediated degradation of the misfolded protein.

Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase

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