This work was supported by Illinois State University School of Biological Sciences startup funds, Illinois State University New Faculty Initiative Grant, and the NIAID grant R15AI164585 (to J.-U. D.). G.M.A. was supported by the Illinois State University Undergraduate Research Support Program (to G.M.A.). K. P. H. was supported by a RISE fellowship provided by the German Academic Exchange Service (DAAD). The authors thank Dr. Uwe Landau and Dr. Carsten Meyer from Largentech Vertriebs GmbH for providing the AGXX powder. Figures 1, Figure 2, Figure 3, Figure 4, and Figure 5 were generated with Biorender.

1. Stress treatment of E. coli strains MG1655 and CFT073
<ol>
	<li>Inoculate 5 mL of lysogeny broth (LB) medium with a single colony of commensal E. coli strain MG1655 and uropathogenic E. coli (UPEC) strain CFT073, respectively, and incubate for 14-16 h (overnight) at 37 &#176;C and 300 rpm. 
	NOTE: Escherichia coli CFT073 is a human pathogen. Handling of CFT073 must be performed with appropriate biosafety measures in a Biosafety Level-2 certified lab.</li>
	<li>Dilute each strain into a 500 mL flask containing 70 mL of 3-(N-morpholino)propanesulfonic acid (MOPS)-glucose (MOPS-g) (Table 1) medium to an optical density at 600 nm (OD600) value of 0.1. Incubate at 37 &#176;C and 300 rpm until mid-log phase is reached (OD600 = 0.5-0.55).</li>
	<li>Transfer 20 mL of each culture into three prewarmed 125 mL flasks and incubate at 37 &#176;C and 300 rpm for 2 min. 
	NOTE: As timely processing of the samples is required, handle no more than 6 cultures at a time.</li>
	<li>Prepare an antimicrobial compound solution in MOPS-g medium at a concentration of 2 mg/mL. Add the antimicrobial to each culture to reach the indicated concentrations. For the untreated control, add the required volume of MOPS-g medium. 
	NOTE: Vortex the 2 mg/mL antimicrobial solution to allow an even distribution of the compound particles and avoid sedimentation.</li>
	<li>Incubate the cultures for 45 min at 37 &#176;C and 300 rpm.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62628/62628fig01.jpg" /> 
Figure 1: Escherichia coli stress treatment. Bacterial cultures are grown in MOPS-g and treated with the indicated concentrations of the silver-ruthenium-containing antimicrobial when the mid-log phase is reached. Abbreviations: LB = lysogeny broth; Ag-Ru = silver-ruthenium; MOPS-g = 3-(N-morpholino)propanesulfonic acid (MOPS)-glucose. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62628/62628fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Collecting bacterial cell samples
<ol>
	<li>After 45 min of stress treatment, determine the OD600 of each culture. For each sample, harvest cells equivalent to 4 mL of OD600 = 1 in 15 mL centrifuge tubes by centrifugation for 15 min at 3,000 &#215; g and 4 &#176;C.</li>
	<li>Completely remove the supernatant and resuspend the cell pellets in 50 &#181;L of ice-cold lysis buffer (Table 1). Incubate the samples for 30 min on ice. 
	NOTE: This lysis step degrades the peptidoglycan layer. Always use freshly prepared lysis buffer.</li>
	<li>Transfer the samples into 1.7 mL microcentrifuge tubes. Freeze at -80 &#176;C until further use.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62628/62628fig02.jpg" /> 
Figure 2: Bacterial sample collection. Cell samples are harvested by centrifugation and resuspended in lysis buffer followed by storage at -80 &#176;C. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62628/62628fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Extracting the insoluble protein aggregates
<ol>
	<li>Thaw samples on ice. 
	NOTE: The freeze-thaw cycle contributes to cell lysis.</li>
	<li>Add 360 &#181;L of ice-cold buffer A (Table 1) and mix gently by pipetting. 
	NOTE: The osmotic shock will also contribute to cell lysis.</li>
	<li>Transfer the sample to a 2 mL microcentrifuge tube containing ~200 &#181;L of 0.5 mm glass beads. Incubate for 30 min at 8 &#176;C in a thermomixer with shaking at 1,400 rpm. 
	NOTE: This step results in the physical disruption of the cell. A protease inhibitor can be used to minimize protein degradation. The disruption can be performed at 4 &#176;C. Note that the use of glass beads has been reported to induce aggregation of a small subset of proteins in yeast19.</li>
	<li>Incubate for 5 min on ice without shaking to settle the glass beads. Transfer 200 &#181;L of the cell lysate into 1.7 mL microcentrifuge tubes. 
	NOTE: Avoid the transfer of the glass beads.</li>
	<li>Centrifuge at 16,000 &#215; g and 4 &#176;C for 20 min. Collect the supernatant, which contains soluble proteins, and proceed to section 4.</li>
	<li>Resuspend the pellet in 200 &#181;L of ice-cold buffer A (Table 1) using the pipette. Centrifuge at 16,000 &#215; g and 4 &#176;C for 20 min. Carefully remove the supernatant altogether.</li>
	<li>Add 200 &#181;L of ice-cold buffer B (see Table 1 and the Table of Materials) and carefully resuspend the pellet by pipetting. 
	NOTE: The non-ionic detergent solubilizes membrane protein.</li>
	<li>Repeat the centrifugation at 16,000 &#215; g and 4 &#176;C for 20 min. Carefully remove the supernatant.</li>
	<li>Resuspend the pellet in 200 &#181;L of cold buffer A (Table 1) by pipetting. Centrifuge at 16,000 &#215; g and 4 &#176;C for 20 min. Completely remove the supernatant.</li>
	<li>Resuspend the pellet in 100 &#181;L of 1x reducing SDS sample buffer (Table 1) and boil for 5 min at 95 &#176;C in a thermomixer.</li>
	<li>Store the sample at -20 &#176;C to proceed later or immediately load on an SDS polyacrylamide gel for separation.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62628/62628fig03.jpg" /> 
Figure 3: Extraction of insoluble protein aggregates. The extraction of protein aggregates involves a series of steps including cell disruption, the separation of protein aggregates from soluble proteins, the solubilization of membrane proteins, and washing. Abbreviation: SDS = sodium dodecyl sulfate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62628/62628fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Soluble protein sample preparation
<ol>
	<li>Mix 1 volume of 100% trichloroacetic acid (TCA) with 4 volumes of soluble protein sample from step 3.6. 
	NOTE: Handling of TCA requires a fume hood and personal protective equipment and an approved waste disposal procedure.</li>
	<li>Incubate for 10 min at 4 &#176;C to allow for protein precipitation. 
	NOTE: White precipitate will appear very soon.</li>
	<li>Centrifuge to precipitate at 21,000 &#215; g and 4 &#176;C for 5 min and remove the supernatant. Wash the pellet with 200 &#181;L of ice-cold acetone to remove cellular debris. Centrifuge at 21,000 &#215; g and 4 &#176;C for 5 min and remove the supernatant. Repeat these actions in step 4.3 a total of three times.</li>
	<li>Place the microcentrifuge tubes with open lids in a thermomixer at 37 &#176;C to remove the remaining acetone from the pellet. 
	NOTE: Incubation of more than 5 min may reduce the solubility of the protein pellet.</li>
	<li>Add 100 &#181;L of 1x reducing SDS buffer (Table 1) and completely dissolve the pellet. Boil the sample for 5 min at 95 &#176;C.</li>
	<li>Store sample at -20 &#176;C to proceed later or immediately load on an SDS polyacrylamide gel for separation.</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62628/62628fig04.jpg" /> 
Figure 4: Preparation of soluble proteins. The preparation of soluble protein involves a precipitation step with trichloroacetic acid and repeated washing with ice-cold acetone. Abbreviations: TCA = trichloroacetic acid; SDS = sodium dodecyl sulfate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62628/62628fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Separation and visualization of extracted protein aggregates using SDS-PAGE
<ol>
	<li>Prepare a 12% SDS-polyacrylamide gel.
	<ol>
		<li>For two separating gels, pipette 5.1 mL of double-distilled water (ddH2O), 3.75 mL of Tris-HCl (pH 8.8), 7.5 mL of 20% (w/v) SDS, 6 mL of 30% acrylamide/bisacrylamide (29:1) solution, 75 mL of 10% w/v ammonium persulfate, and 10 mL of tetramethylethylenediamine (TEMED) into a 15 mL centrifuge tube and mix gently without introducing air bubbles. Pour the gel using a 1 mL pipet within the glass plates, leaving the upper 2 cm free of the mixture. Add 70% ethanol on the top of the separating gel and allow an even interface between the two layers.</li>
		<li>After polymerization of the separating gel, prepare the stacking gel by pipetting 1.535 mL of ddH2O, 625 mL of Tris-HCl (pH 6.8), 12.5 mL of 20% (w/v) SDS, 335 mL of 30% acrylamide/bisacrylamide (29:1) solution, 12.5 mL of 10% w/v ammonium persulfate, and 2.5 mL of TEMED. Remove the ethanol from the separating gels and add the stacking gel solution. Insert a comb with the desired number of pockets without introducing air bubbles. Allow polymerization for 20-30 min.</li>
	</ol>
	</li>
	<li>Load 4 &#181;L of each sample and protein ladder into separate wells and run the gel(s) in Tris-Glycine running buffer (Table 1) at 144 V for 45 min at room temperature. 
	NOTE: Stop the gel when the bromophenol band is about to migrate out of the gel.</li>
	<li>Stain the gel(s) in a prewarmed Fairbanks solution A (Table 1) for 30 min on a rocker.</li>
	<li>Decolor the gel(s) in a prewarmed Fairbanks solution D (Table 1) until the desired background (e.g., overnight) on a rocker.</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62628/62628fig05.jpg" /> 
Figure 5: Protein separation and visualization. The samples are separated by SDS-PAGE and visualized by Coomassie staining. Abbreviation: SDS-PAGE = sodium dodecyl sulfate-polyacrylamide gel electrophoresis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62628/62628fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have nothing to disclose.

This protocol describes an optimized methodology for the analysis of protein aggregate formation after treatment of different E. coli strains with a proteotoxic antimicrobial. The protocol allows the simultaneous extraction of insoluble and soluble protein fractions from treated and untreated E. coli cells. Compared to existing protocols for protein aggregate isolation from cells14,15,16,20, this method has several advantages: (i) only small culture volumes (4-8 mL) are needed; (ii) the cell disruption process does not rely on special equipment such as a French press, cell disruptor, or sonicator; and (iii) the protocol is easy to follow even for relatively early-career scientists in the field.
Bacteria encounter a myriad of stresses in their natural environment, and many of them represent a threat, particularly to proteins, the most abundant macromolecule in the cell21. The described methodology offers many potential applications. One of them is the possibility to investigate the efficacy by which a wide range of stresses (e.g., elevated temperatures, reactive oxygen species, reactive chlorine species) and chemical compounds affect protein homeostasis in bacteria, archaea, and even eukaryotic cells5,12,22,23. Our extraction was performed with the two distantly related E. coli strains, MG1655 and CFT073. However, it has also been successfully applied to study the role of specific gene products for protein homeostasis by comparing protein aggregate formation in wild-type and mutant strains11,12.
After considering appropriate modifications and troubleshooting of growth conditions and stressor concentrations, this protocol can also be used to determine protein aggregate formation in other Gram-negative5 and Gram-positive bacteria8. Notably, the gel bands separated after SDS-PAGE can be densitometrically analyzed (i.e., using ImageJ). This approach can also be used to analyze the impact of additional therapeutic compounds such as inhibitors of molecular chaperones24. Most intriguingly, it can be combined with mass spectrometric approaches to provide information on the type of protein species that are aggregation-sensitive under a specific stress condition, simply by determining their presence or absence in the insoluble protein fraction5,15.
The success of this protocol requires careful consideration of the stressor concentrations that cells are exposed to and the exposure. We, therefore, recommend performing a preliminary survival assay with increasing stressor concentrations to determine the proteotoxic concentration. Moreover, the stressor solutions should be prepared freshly before each experiment, and the sample collection times be kept consistent. One limitation of this method is the low number of samples that can concurrently be processed. This is mainly due to the time-sensitive handling of samples in section 1, which involves significant vortexing steps of the antimicrobial solution in between the addition of the compound to the cultures to avoid sedimentation. However, this may not be such an issue when different soluble stressors are applied. Moreover, the described procedure does not provide any time-resolved information on protein aggregate location and trajectory, which would require more advanced techniques such as fluorescence microscopy in combination with time-lapse microscopy25. In summary, the improved methodology is simple, easy to follow, inexpensive, and offers the potential for additional modification that allows a tailored approach for identifying proteotoxic compounds or bacterial stress response genes.

Bacteria are inevitably exposed to a myriad of environmental stresses, including low pH (e.g., in the mammalian stomach)1,2, reactive oxygen and chlorine species (ROS/RCS) (e.g., during oxidative burst in phagocytes)3,4,5, elevated temperatures (e.g., in hot springs or during heat-shock)6,7, and several potent antimicrobials (e.g., AGXX used in this protocol)8. Proteins are particularly vulnerable to any of these stressors, and exposure can provoke protein un-/misfolding that then seeds aggregation. All organisms employ protective systems that allow them to cope with protein misfolding9. However, severe stress can overwhelm the protein quality control machinery and disrupt the secondary and/or tertiary structure of proteins, which ultimately inactivates proteins. As a consequence, protein aggregates can severely impair critical cellular functions required for bacterial growth and survival, stress resistance, and virulence10. Therefore, research focusing on protein aggregation and its consequences in bacteria is a relevant topic due to its potential impact on infectious disease control.
Heat-induced protein unfolding and aggregation are often reversible7. In contrast, other proteotoxic stresses, such as oxidative stress, can cause irreversible protein modifications through the oxidation of specific amino acid side chains resulting in protein un-/misfolding and, eventually, protein aggregation4. Stress-induced formation of insoluble protein aggregates has been extensively studied in the context of molecular chaperones and their protective functions in yeast and bacteria11,12,13. Several protocols have been published that utilize a variety of biochemical techniques for the isolation and analysis of insoluble protein aggregates14,15,16,17. The existing protocols have mainly been used to study bacterial protein aggregation upon heat-shock and/or identification of molecular chaperones. While these protocols have certainly been an advancement to the field, there are some major inconveniences in the experimental procedures because they require (i) a large bacterial culture volume of up to 10 L14,17, (ii) complicated physical disruption processes, including the use of cell disruptors, French press, and/or sonication14,15,17, or (iii) time-consuming repeated washing and incubation steps15,16,17.
This paper describes a modified protocol that aims to address the limitations of the previous approaches and allows the analysis of the amount of protein aggregates formed in two different Escherichia coli strains after treatment with a proteotoxic antimicrobial surface coating. The coating is composed of metal-silver (Ag) and ruthenium (Ru)-conditioned with ascorbic acid, and its antimicrobial activity is achieved by the generation of reactive oxygen species8,18. Herein is a detailed description of the preparation of the bacterial culture after treatment with the antimicrobial compound and a comparison of protein aggregation status upon exposure of two E. coli strains with distinct susceptibility profiles to increasing concentration of the antimicrobial. The described method is inexpensive, fast, and reproducible and can be used to study protein aggregation in the presence of other proteotoxic compounds. In addition, the protocol can be modified to analyze the impact that specific gene deletions have on protein aggregation in a variety of different bacteria.

<table><tbody><tr><td>&lt;strong&gt;Chemicals/Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Acetone</td><td>Fisher Scientific</td><td>67-64-1</td><td></td></tr><tr><td>30% Acrylamide/Bisacrylamide solution 29:1</td><td>Bio-Rad</td><td>1610156</td><td></td></tr><tr><td>Ammonium persulfate</td><td>Millipore Sigma</td><td>A3678-100G</td><td></td></tr><tr><td>Benzonase nuclease</td><td>Sigma</td><td>E1014-5KU</td><td></td></tr><tr><td>Bluestain 2 Protein ladder, 5-245 kDa</td><td>GoldBio</td><td>P008-500</td><td></td></tr><tr><td>&amp;beta;-mercaptoethanol</td><td>Millipore Sigma</td><td>M6250-100ML</td><td></td></tr><tr><td>Bromophenol blue</td><td>GoldBio</td><td>B-092-25</td><td></td></tr><tr><td>Coomassie Brilliant Blue R-250</td><td>MP Biomedicals LLC</td><td>821616</td><td></td></tr><tr><td>D-Glucose</td><td>Millipore Sigma</td><td>G8270-1KG</td><td></td></tr><tr><td>D-Sucrose</td><td>Acros Organics</td><td>57-50-1</td><td></td></tr><tr><td>Ethylenediamine tetra acetic acid (EDTA)</td><td>Sigma-Aldrich</td><td>SLBT9686</td><td></td></tr><tr><td>Glacial Acetic acid</td><td>Millipore Sigma</td><td>ARK2183-1L</td><td></td></tr><tr><td>Glycerol, 99%</td><td>Sigma-Aldrich</td><td>G5516-1L</td><td></td></tr><tr><td>Glycine</td><td>GoldBio</td><td>G-630-1</td><td></td></tr><tr><td>Hydrochloric acid, ACS reagent</td><td>Sigma-Aldrich</td><td>320331-2.5L</td><td></td></tr><tr><td>Isopropanol (2-Propanol)</td><td>Sigma</td><td>402893-2.5L</td><td></td></tr><tr><td>LB broth (Miller)</td><td>Millipore Sigma</td><td>L3522-1KG</td><td></td></tr><tr><td>LB broth with agar (Miller)</td><td>Millipore Sigma</td><td>L2897-1KG</td><td></td></tr><tr><td>Lysozyme</td><td>GoldBio</td><td>L-040-25</td><td></td></tr><tr><td>10x MOPS Buffer</td><td>Teknova</td><td>M2101</td><td></td></tr><tr><td>Nonidet P-40</td><td>Thomas Scientific</td><td>9036-19-5</td><td></td></tr><tr><td>Potassium phosphate, dibasic</td><td>Sigma-Aldrich</td><td>P3786-1KG</td><td></td></tr><tr><td>Potassium phosphate, monobasic</td><td>Acros Organics</td><td>7778-77-0</td><td></td></tr><tr><td>Sodium dodecyl sulfate (SDS)</td><td>Sigma-Aldrich</td><td>L3771-500G</td><td></td></tr><tr><td>Tetramethylethylenediamine (TEMED)</td><td>Millipore Sigma</td><td>T9281-50ML</td><td></td></tr><tr><td>Thiamine</td><td>Sigma-Aldrich</td><td>T4625-100G</td><td></td></tr><tr><td>100% Trichloroacetic acid</td><td>Millipore Sigma</td><td>T6399-100G</td><td></td></tr><tr><td>Tris base</td><td>GoldBio</td><td>T-400-1</td><td></td></tr><tr><td>&lt;strong&gt;Material/Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Centrifuge tubes (15 mL)</td><td>Alkali Scientific</td><td>JABG-1019</td><td></td></tr><tr><td>Erlenmeyer flask (125 mL)</td><td>Carolina</td><td>726686</td><td></td></tr><tr><td>Erlenmeyer flask (500 mL)</td><td>Carolina</td><td>726694</td><td></td></tr><tr><td>Freezer: -80 &amp;deg;C</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Glass beads (0.5 mm)</td><td>BioSpec Products</td><td>1107-9105</td><td></td></tr><tr><td>Microcentrifuge</td><td>Hermle</td><td>Z216MK</td><td></td></tr><tr><td>Microcentriguge tubes (1.7 mL)</td><td>VWR International</td><td>87003-294</td><td></td></tr><tr><td>Microcentriguge tubes (2.0 mL)</td><td>Axygen Maxiclear Microtubes</td><td>MCT-200-C</td><td></td></tr><tr><td>Plastic cuvettes</td><td>Fischer Scientific</td><td>14-377-012</td><td></td></tr><tr><td>Power supply</td><td>ThermoFisher Scientific</td><td>EC105</td><td></td></tr><tr><td>Rocker</td><td>Alkali Scientific</td><td>RS7235</td><td></td></tr><tr><td>Shaking incubator (37 &amp;deg;C)</td><td>Benchmark Scientific</td><td></td><td></td></tr><tr><td>Small glass plate</td><td>Bio-Rad</td><td>1653311</td><td></td></tr><tr><td>Spacer plates (1 mm)</td><td>Bio-Rad</td><td>1653308</td><td></td></tr><tr><td>Spectrophotometer</td><td>Thermoscientific</td><td>3339053</td><td></td></tr><tr><td>Tabletop centrifuge for 15 mL centrifuge tubes</td><td>Beckman-Coulter</td><td></td><td></td></tr><tr><td>Vertical gel electrophoresis chamber</td><td>Bio-Rad</td><td>1658004</td><td></td></tr><tr><td>Vortexer</td><td>Fisher Vortex Genie 2</td><td>12-812</td><td></td></tr><tr><td>Thermomixer</td><td>Benchmark Scientific</td><td>H5000-HC</td><td></td></tr><tr><td>10 well comb</td><td>Bio-Rad</td><td>1653359</td><td></td></tr></tbody></table>

extraction and visualization of protein aggregates after treatment of escherichia coli with a proteotoxic stressor

The exposure of living organisms to environmental and cellular stresses often causes disruptions in protein homeostasis and can result in protein aggregation. The accumulation of protein aggregates in bacterial cells can lead to significant alterations in the cellular phenotypic behavior, including a reduction in growth rates, stress resistance, and virulence. Several experimental procedures exist for the examination of these stressor-mediated phenotypes. This paper describes an optimized assay for the extraction and visualization of aggregated and soluble proteins from different Escherichia coli strains after treatment with a silver-ruthenium-containing antimicrobial. This compound is known to generate reactive oxygen species and causes widespread protein aggregation. 
The method combines a centrifugation-based separation of protein aggregates and soluble proteins from treated and untreated cells with subsequent separation and visualization by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and Coomassie staining. This approach is simple, fast, and allows a qualitative comparison of protein aggregate formation in different E. coli strains. The methodology has a wide range of applications, including the possibility to investigate the impact of other proteotoxic antimicrobials on in vivo protein aggregation in a wide range of bacteria. Moreover, the protocol can be used to identify genes that contribute to increased resistance to proteotoxic substances. Gel bands can be used for the subsequent identification of proteins that are particularly prone to aggregation.

<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62628/62628fig06.jpg" /> 
Figure 6: Representative results of antimicrobial-induced protein aggregation in commensal Escherichia coli strain MG1655 and UPEC strain CFT073. E. coli strains MG1655 and CFT073 were grown at 37 &#176;C and 300 rpm to OD600= 0.5-0.55 in MOPS-g media before they were treated with the indicated concentrations (-, 0 mg/mL; +, 175 mg/mL; ++, 200 mg/mL) of the antimicrobial for 45 min. Soluble and insoluble protein samples were prepared as described in the protocol and Figure 1, Figure 2, Figure 3, and Figure 4 and visualized on a 12% SDS polyacrylamide gel (Figure 5). Protein aggregate formation (insoluble fraction) was increased in both strains in the presence of the antimicrobial while the amounts of soluble proteins were decreased. Overall, the antimicrobial had a much more potent effect on MG1655 than on CFT073. Abbreviations: M= Protein marker; UPEC = uropathogenic E. coli; MOPS-g = 3-(N-morpholino)propanesulfonic acid (MOPS)-glucose; SDS = sodium dodecyl sulfate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62628/62628fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Here, two E. coli strains were used that differ in their susceptibility to a proteotoxic silver-ruthenium-containing antimicrobial to demonstrate this protocol. Preliminary survival data revealed that the commensal E. coli strain MG1655 is significantly more sensitive to the ROS-generating antimicrobial than the UPEC strain CFT073 (data not shown). Both strains were grown in MOPS-g media at 37 &#176;C and 300 rpm. At the mid-log phase, the cells were either left untreated or treated with 175 &#181;g/mL and 200 &#181;g/mL of the antimicrobial, respectively, and incubated for 45 min. Subsequently, the cells were lysed, and cellular protein aggregates separated from the soluble proteins. Proteins in both fractions were then separated by SDS-PAGE and visualized by Coomassie staining. The insoluble fraction shown in Figure 6 represents the amount of protein aggregates formed, which was increased when cells were incubated in the presence of the antimicrobial compared to untreated cells. The increase in protein aggregate formation was independent of the strain background, although a much more pronounced increase in aggregate formation was detected in the more sensitive strain MG1655. Conversely, lower amounts of soluble proteins (soluble fraction) were observed after antimicrobial treatment of the cells compared to the untreated counterpart. This result was expected given the preliminary data that showed a substantially higher tolerance of the antimicrobial in CFT073 than MG1655.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Solutions</td>
			<td>Recipes</td>
		</tr>
		<tr>
			<td>Buffer A</td>
			<td>10 mM potassium phosphate (pH 6.5), 1 mM EDTA</td>
		</tr>
		<tr>
			<td>Buffer B</td>
			<td>Buffer A containing 2% Nonidet P-40. Can be stored in room temperature for later use.</td>
		</tr>
		<tr>
			<td>Fairbanks A (Staining solution)</td>
			<td>25% isopropanol, 10% Glacial Acetic acid, 1.4 g Coommassie R-250</td>
		</tr>
		<tr>
			<td>Fairbanks D (Destaining solution)</td>
			<td>10% Glacial acetic acid solution</td>
		</tr>
		<tr>
			<td>Lysis buffer</td>
			<td>10 mM potassium phosphate (pH 6.5), 1 mM EDTA, 20% sucrose can be prepared and stored at room temperature for long term use. Add 1 mg/mL lysozyme and 50 u/mL Benzonase fresh before use.&#160;</td>
		</tr>
		<tr>
			<td>MOPS-g media</td>
			<td>100 mL 10x MOPS, 10 mL 0.132 M K2HPO4, 10 mL 20% glucose, 0.5 mL 20 mM thiamine. Fill up to 1 L with ddH2O and sterile-filter</td>
		</tr>
		<tr>
			<td>1x SDS sample buffer</td>
			<td>6.5 mM Tris-HCl (pH 7), 10% glycerol, 2% SDS, 0.05% bromophenol blue and 2.5% &#946;-mercaptoethanol. Stored at -20 &#176;C.</td>
		</tr>
		<tr>
			<td rowspan="2">12% SDS polyacrylamide gel preparation (for 2 gels)</td>
			<td>Separating gel: 5.1 mL ddH2O, 3.75 mL Tris-HCl (pH 8.8), 75 mL 20% w/v SDS, 6 mL 30% Acrylamide/Bisacrylamide solution 29:1 solution, 75 mL 10% w/v ammonium persulfate, 10 mL TEMED</td>
		</tr>
		<tr>
			<td>Stacking gel: 1.535 mL ddH2O, 625 mL Tris-HCl (pH 6.8), 12.5 mL 20% w/v SDS, 335 mL 30% Acrylamide/Bisacrylamide solution 29:1 solution, 12.5 mL 10% w/v ammonium persulfate, 2.5 mL TEMED</td>
		</tr>
		<tr>
			<td>SDS running buffer</td>
			<td>25 mM Tris, 192 mM Glycine, 0.1% SDS in ddH2O. Store in room temperature.</td>
		</tr>
	</tbody>
</table>
Table 1: Buffer, Media, and Solutions. Recipes for buffer, media, and solutions used in this protocol.

Watch this Scientific Journal Video about Extraction and Visualization of Protein Aggregates after Treatment of Escherichia coli with a Proteotoxic Stressor at JoVE.com

Extraction and Visualization of Protein Aggregates after Treatment of Escherichia coli with a Proteotoxic Stressor

This protocol describes the extraction and visualization of aggregated and soluble proteins from Escherichia coli after treatment with a proteotoxic antimicrobial. Following this procedure allows a qualitative comparison of protein aggregate formation in vivo in different bacterial strains and/or between treatments.

Extraction and Visualization of Protein Aggregates after Treatment of Escherichia coli with a Proteotoxic Stressor

The exposure of living organisms to environmental and cellular stresses often causes disruptions in protein homeostasis and can result in protein ...

extraction-visualization-protein-aggregates-after-treatment

Nanyang Technological University

Research

JoVE Journal

Biochemistry

3.5K Views.  Illinois State University. Blootstelling aan stress kan eiwitaggregatie veroorzaken en resulteren in significante veranderingen in het fenotypische gedrag van bacteriën. De procedure die in deze video wordt beschreven, maakt de extractie en visualisatie van eiwitaggregaten na stressbehandeling mogelijk. In tegenstelling tot andere gepubliceerde procedures vereist dit protocol lagere celaantallen en onthoudt het zich van gecompliceerde fysieke verstoringsprocessen, evenals tijdrovende was- en incubatiestappen.Di...

Video: Extraction and Visualization of Protein Aggregates after Treatment of Escherichia coli with a Proteotoxic Stressor

Evaluation of the Impact of Protein Aggregation on Cellular Oxidative Stress in Yeast

Protein misfolding and aggregation into amyloid conformations have been related to the onset and progression of several neurodegenerative diseases. However, there is still little information about how insoluble protein aggregates exert their toxic effects in vivo. Simple prokaryotic and eukaryotic model organisms, such as bacteria and yeast, have contributed significantly to our present understanding of the mechanisms behind the intracellular amyloid formation, aggregates propagation, and toxicity. In this protocol, the use of yeast is described as a model to dissect the relationship between the formation of protein aggregates and their impact on cellular oxidative stress. The method combines the detection of the intracellular soluble/aggregated state of an amyloidogenic protein with the quantification of the cellular oxidative damage resulting from its expression using flow cytometry (FC). This approach is simple, fast, and quantitative. The study illustrates the technique by correlating the cellular oxidative stress caused by a large set of amyloid-&#946; peptide variants with their respective intrinsic aggregation propensities.

Protein aggregation elicits cellular oxidative stress. This protocol describes a method for monitoring the intracellular states of amyloidogenic proteins and the oxidative stress associated with them, using flow cytometry. The approach is used to study the behavior of soluble and aggregation-prone variants of the amyloid-&#946; peptide.

Protein misfolding and aggregation into amyloid conformations have been related to the onset and progression of several neurodegenerative diseases. ...

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

Assessment of Hsp70 Chaperone Activity Using Escherichia coli dnaK756 Cells

Escherichia coli -Based Complementation Assay to Study the Chaperone Function of Heat Shock Protein 70

Heat shock protein 70 (Hsp70) is a conserved protein that facilitates the folding of other proteins within the cell, making it a molecular chaperone. While Hsp70 is not essential for E. coli cells growing under normal conditions, this chaperone becomes indispensable for growth at elevated temperatures. Since Hsp70 is highly conserved, one way to study the chaperone function of Hsp70&#160;genes from various species is to heterologously express them in E. coli strains that are either deficient in Hsp70 or express a native Hsp70 that is functionally compromised. E. coli dnaK756&#160;cells are unable to support &#955; bacteriophage DNA. Furthermore, their native Hsp70 (DnaK) exhibits elevated ATPase activity while demonstrating reduced affinity for GrpE (Hsp70 nucleotide exchange factor). As a result, E. coli dnaK756&#160;cells grow adequately at temperatures ranging from 30 &#176;C to 37 &#176;C, but they die at elevated temperatures (&#62;40 &#176;C). For this reason, these cells serve as a model for studying the chaperone activity of Hsp70. Here, we describe a detailed protocol for the application of these cells to conduct a complementation assay, enabling the study of the in cellulo chaperone function of Hsp70.

Author Spotlight: Exploring Heat Shock Proteins in Malaria and Tuberculosis Infections

This protocol demonstrates the chaperone activity of heat shock protein 70 (Hsp70). E. coli dnaK756 cells serve as a model for the assay as they harbor a native, functionally impaired Hsp70, making them susceptible to heat stress. The heterologous introduction of functional Hsp70 rescues the growth deficiency of the cells.

Heat shock protein 70 (Hsp70) is a conserved protein that facilitates the folding of other proteins within the cell, making it a molecular chaperone. ...

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

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 ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we showed that D. discoideum is remarkably resilient to protein aggregation and can be used to gain insights into the cellular protein quality control system. However, the use of D. discoideum as a model system poses several challenges to microscopy-based experimental approaches, such as the high motility of the cells and their susceptibility to photo-toxicity. The latter proves to be especially challenging when studying protein homeostasis, as the phototoxic effects can induce a cellular stress response and thus alter to behavior of the protein quality control system. 
Temperature increase is a commonly used way to induce cellular stress. Here, we describe a temperature-controllable imaging protocol, which allows observing temperature-induced perturbations in D. discoideum. Moreover, when applied at normal growth temperature, this imaging protocol can also noticeably reduce photo-toxicity, thus allowing imaging with higher intensities. This can be particularly useful when imaging proteins with very low expression levels. Moreover, the high mobility of the cells often requires the acquisition of multiple fields of view to follow individual cells, and the number of fields needs to be balanced against the desired time interval and exposure time.

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The social amoebae Dictyostelium discoideum has recently been established as a system to study protein misfolding and proteostasis. Here, we describe a new imaging-based methodology to study temperature-induced protein aggregation and the cellular stress response in D. discoideum.

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we ...

Automating Aggregate Quantification in Caenorhabditis elegans

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This increased attention has called for the diversification and improvement of animal models capable of reproducing disease phenotypes observed in humans with PCDs. Though murine models have proven invaluable, they are expensive and are associated with laborious, low-throughput methods. Use of the Caenorhabditis elegans nematode model to study PCDs has been justified by its relative ease of maintenance, low cost, and rapid generation time, which allow for high-throughput applications. Additionally, high conservation between the C. elegans and human genomes makes this model an invaluable discovery tool. Nematodes that express fluorescently tagged tissue-specific polyglutamine (polyQ) tracts exhibit age- and polyQ length-dependent aggregation characterized by fluorescent foci. Such reporters are often employed as proxies to monitor changes in proteostasis across tissues. Manual aggregate quantification is time-consuming, limiting experimental throughput. Furthermore, manual foci quantification can introduce bias, as aggregate identification can be highly subjective. Herein, a protocol consisting of worm culturing, image acquisition, and data processing was standardized to support high-throughput aggregate quantification using C. elegans that express intestine-specific polyQ. By implementing a C. elegans-based image processing pipeline using CellProfiler, an image analysis software, this method has been optimized to separate and identify individual worms and enumerate their respective aggregates. Though the concept of automation is not entirely unique, the need to standardize such procedures for reproducibility, elimination of bias from manual counting, and increase throughput is high. It is anticipated that these methods can drastically simplify the screening process of large bacterial, genomic, or drug libraries using the C. elegans model.

Automating Aggregate Quantification in Caenorhabditis elegans

The following protocol describes the development and optimization of a high-throughput workflow for worm culturing, fluorescence imaging, and automated image processing to quantify polyglutamine aggregates as an assessment of changes in proteostasis.

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This ...

Misfolding-Prone Protein Degradation Assay: A Technique to Monitor Misfolded Protein Degradation Using Cycloheximide Treatment and Detergent Fractionation

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 a degradation assay for misfolded proteins using cycloheximide treatment along with detergent fractionation. This method aids in studying the dynamics of misfolded proteins and uncovering the in-depth mechanisms of protein turnover.

Source: Guo, L. et al., Assays for the Degradation of Misfolded Proteins in Cells. J. Vis. Exp. (2016).
This video describes a degradation assay for ...

Extraction and Visualization of Protein Aggregates after Treatment of Escherichia coli with a Proteotoxic Stressor

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