NBN is supported by a special Recruitment Grant from the Monash University Faculty of Medicine Nursing and Health Sciences with funding from the State Government of Victoria and the Australian Government. We thank Bernd Bukau (ZMBH, Heidelberg University, Germany) and Harm H. Kampinga (Department of Biomedical Sciences of Cells &#38; Systems, University of Groningen, The Netherlands) for their invaluable support and sharing of reagents, Holger Lorenz (ZMBH Imaging Facility, Heidelberg University, Germany) for his support with confocal microscopy and image processing, and Claire Hirst (ARMI, Monash University, Australia) for critical reading of the manuscript.

1. HeLa Cell Preparation
<ol>
	<li>Prepare the following materials: PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4), pH 7.4; DMEM, supplemented with 10% FCS and 1% Pen-Strep; 4% paraformaldehyde in PBS; 0.5% Triton-X100 in PBS; TBS-T (150 mM NaCl, 20 mM Tris, 0.05% Tween), pH 7.4; 0.0001% sterile poly-L-Lysine solution; 10-well diagnostic slides; a humid chamber; tissue paper; and Coplin slide-staining jars. 
	NOTE: To ensure optimal fixation efficiency, paraformaldehyde solutions should be prepared fresh before every experiment.</li>
	<li>Prepare a humid chamber by covering the bottom of a closed box with wet tissues. Place the humid chamber at 37 &#176;C before starting the experiment, to ensure that the chamber temperature is at 37 &#176;C while incubating the enzymatic reactions.</li>
	<li>Culture HeLa cells in T25 flasks, in 5 mL of DMEM (High Glucose, Glutamate and Sodium Pyruvate supplemented) supplemented with 10% FCS and 1% Pen-Strep, in a 37 &#176;C CO2 incubator containing 5% CO2. Dissociate adherent cells using 0.05% Trypsin-EDTA. After addition of fresh DMEM to dissociated cells, count cells using a cell counting chamber and grow on diagnostic slides inside a humid chamber.</li>
	<li>Sterilize diagnostic slides by UV-irradiation in a sterile cell culture hood for 30 min.</li>
	<li>Add 100 &#181;L of sterile filtered 0.0001% poly-L-lysine to each well required for the experiment. Incubate for 30 min. Wash away excess poly-L-lysine by washing each well 3x with 50 &#181;L ultrapure&#160;water.</li>
	<li>Trypsinize the HeLa cells and add approximately 15,000 cells to each well. If necessary, dilute cells to at least 30-50 &#181;L of DMEM per well.</li>
	<li>Grow cells in a humid chamber within a 37 &#176;C 5% CO2 incubator for approximately 24 h. Cells should be 60%&#8211;80% confluent before commencing the PLA. 
	NOTE: Too high a confluency decreases the absorption of reagents, decreasing the obtained signal at the end of the protocol.</li>
	<li>Remove medium by placing a tissue paper at the edge of the well. Wash wells 3x with 50 &#181;L of PBS. 
	NOTE: Cells are subject to detaching when liquids are added harshly. This can be prevented by not letting the wells dry completely before adding new liquid and by adding the new liquid gently to the edge of the well.</li>
	<li>Fix cells by adding 50 &#181;L of freshly prepared 4% paraformaldehyde in PBS to each well. Incubate for 10 min at room temperature.</li>
	<li>Wash slides 3x in PBS. Perform washes in a Coplin slide-staining jar containing 100 mL of PBS. For each wash, incubate for 5 min at room temperature without shaking.</li>
	<li>Permeabilize the cell membrane by submerging the slides in 100 mL of 0.5% Triton-X100 in PBS in a Coplin slide-staining jar. Incubate for 10 min at room temperature without shaking.</li>
	<li>Wash slides 3x in TBS-T. Perform washes in a Coplin slide-staining jar containing 100 mL of TBS-T. For each wash, incubate for 5 min at room temperature without shaking.</li>
	<li>After the last wash, remove excess buffer with a tissue paper. At this point the cells are ready for the Proximity Ligation Assay protocol that will be discussed in section 4.</li>
</ol>
2. S. cerevisiae Cell Preparation
<ol>
	<li>Prepare the following materials: 100 mM KPO4, pH 6.5, referred to as Wash Buffer; 37% formaldehyde; 4% paraformaldehyde in 100 mM KPO4, pH 6.5; 1.2 M sorbitol in 100 mM KPO4, pH 6.5; lyticase solution (500 &#181;g/mL lyticase, 20 mM &#946;-mercaptoethanol, 100 mM KPO4, pH 6.5); 0.0001% poly-L-Lysine solution; 1% Triton-X100 in 100 mM KPO4, pH 6.5; 10-well diagnostic slides; a humid chamber (prepared as in step 1.2); tissue paper; and Coplin slide-staining jars. 
	NOTE: To ensure optimal fixation efficiency, paraformaldehyde solutions should be prepared fresh before every experiment.</li>
	<li>Grow an overnight culture in non-selective Yeast Extract, Peptone and Dextrose (YPD) medium at 30 &#176;C while shaking. 
	NOTE: Depending on experimental requirements S. cerevisiae cells can be grown in Synthetic Complete (SC) medium or selective Synthetic Minimal (SM) medium instead.</li>
	<li>Dilute the stationary culture to an OD600 of 0.1 in 20 mL medium. Grow the cells at 30 &#176;C while shaking until the OD600 reaches 0.5.</li>
	<li>Transfer the 20 mL culture to a 50 mL centrifuge tube. Pellet the cells by centrifugation at 665 x g for 3 min. Remove the supernatant. Resuspend the cells in 5 mL of fresh medium.</li>
	<li>Fix the cells by adding 550 &#181;L of 37% formaldehyde to the culture. Incubate at room temperature for 15 min.</li>
	<li>Pellet the cells by centrifugation at 665 x g for 3 min. Remove the supernatant. Resuspend the pellet in 1 mL of freshly prepared 4% paraformaldehyde in Wash Buffer. Incubate for 45 min at room temperature.</li>
	<li>During the incubation, prepare the diagnostic slides by adding 100 &#181;L of 0.01% poly-L-lysine solution to each well. Incubate the slides for 30 min at room temperature.</li>
	<li>After 30 min, wash away excess poly-L-lysine with ultrapure water and let the slides air dry. The dry slides are ready for use.</li>
	<li>Wash cells twice with 1 mL of Wash Buffer. Perform washes by centrifugation of the cells at 665 x g for 3 min. Remove the supernatant and resuspend the cells in Wash Buffer.</li>
	<li>Pellet the cells by centrifugation at 665 x g for 3 min. Remove the supernatant. Resuspend the cells in 1 mL of 1.2 M sorbitol in Wash Buffer.</li>
	<li>Pellet the cells by centrifugation at 665 x g for 3 min. Remove the supernatant. Resuspend the pellet in 250 &#181;L of freshly prepared Lyticase solution to digest the cell wall. Incubate the cells in Lyticase solution for 15 min at 30 &#176;C while shaking.</li>
	<li>After digestion, wash cells 3x by centrifugation at 665 x g for 3 min and remove the supernatant. Resuspend the cells in 250 &#181;L of 1.2 M sorbitol in Wash Buffer. 
	NOTE: Because cells are fragile after cell wall digestion, resuspend them very carefully to not damage the cells.</li>
	<li>Add 20 &#181;L of resuspended cells to the poly-L-lysine coated slides. Allow them to attach to the slides for 30 min. Wash away non-adherent cells by washing the wells 3x with 50 &#181;L of Wash Buffer.</li>
	<li>Permeabilize the cell membrane by washing 3x with 50 &#181;L of 1% Triton-X in Wash Buffer. 
	NOTE: At this point the cells are ready for the proximity ligation assay protocol that will be discussed in section 4.</li>
</ol>
3. E. coli Cell Preparation
<ol>
	<li>Prepare the following materials: PBS-T (140 mM NaCl, 2 mM KCl, 8 mM K2HPO4, 1.5 mM KH2PO4, 0.05% Tween-20), pH 7.4; lysozyme solution (2 mg/mL lysozyme, 25 mM Tris-HCl pH 8.0, 50 mM Glucose, 10 mM EDTA); 0.0001% poly-L-Lysine solution; 99% ice-cold methanol; 99% room temperature methanol; 99% acetone; 10-well diagnostic slides; a humid chamber (prepared as in step 1.1.1); tissue paper; and Coplin slide-staining jars.</li>
	<li>Grow an overnight culture in Luria-Bertani (LB) medium at 30 &#176;C while shaking.</li>
	<li>Dilute the stationary culture to an OD600 of 0.02 in fresh LB medium. Grow the cells at 30 &#176;C while shaking until the OD600 reaches 0.4 for log phase cells.</li>
	<li>Approximately 15 min before cells reach an OD600 of 0.4, prepare poly-L-lysine coated slides by adding 100 &#181;L of 0.0001% poly-L-lysine to each well. Incubate the slides for 30 min at room temperature.</li>
	<li>After 30 min wash away excess poly-L-lysine with ultrapure water and let slides air dry. The dry slides are ready for use.</li>
	<li>When the cells reach an OD600 of 0.4, transfer 1 mL of the culture to a sterile microcentrifuge tube and pellet cells at 2,650 x g for 2 min.</li>
	<li>Resuspend cells in 50 &#181;L of LB medium.</li>
	<li>Fix cells by adding 1 mL of ice-cold 99% methanol. Mix very gently by hand. Incubate cells for 30 min at -20 &#176;C.</li>
	<li>After fixation add 20 &#181;L of the cells to the poly-L-lysine coated slides. Let the slides air dry for 30 min.</li>
	<li>Add 50 &#181;L of freshly prepared lysozyme solution to each well to digest the cell wall. Incubate in a humid chamber for 30 min at 25 &#176;C.</li>
	<li>Remove the lysozyme solution from the wells by adding a tissue paper to the edge of the well. Wash slides 3x in 100 mL of PBS-T. Perform each wash in a Coplin slide-staining jar for 30 s, without shaking.</li>
	<li>Remove wash buffer from the slides by tapping the slides on a tissue paper.</li>
	<li>Permeabilize the cell membranes by adding 50 &#181;L of 99% methanol to each well. Incubate for 1 min at room temperature.</li>
	<li>Remove methanol by placing a tissue paper to the edge of the well.</li>
	<li>Add 50 &#181;L of 99% acetone to each well. Incubate for 1 min.</li>
	<li>Remove excess acetone by placing a tissue paper to the edge of the well. Allow slides to air dry. At this point the cells are ready for the proximity ligation assay protocol that will be discussed in section 4.</li>
</ol>
4. Proximity Ligation Assay
<ol>
	<li>Prepare the following materials: Blocking Buffer; Antibody Dilution Buffer; Wash Buffer &#8216;A&#8217; &#8211; (10 mM Tris, 150 mM NaCl, 0.05% Tween-20) pH 7.4; Wash Buffer &#8216;B&#8217; &#8211; (200 mM Tris, 100 mM NaCl) pH 7.4; Anti-Rabbit Secondary Antibody PLUS; Anti-Mouse Secondary Antibody MINUS; 5x Ligation Buffer; ligase; 5x Amplification Buffer (Orange: &#955;ex&#160;554&#160;nm; &#955;em&#160;576&#160;nm); polymerase; ultrapure water; and mounting medium containing DAPI. 
	NOTE: The PLA detection reagents are also available in the variants Green (&#955;ex 495 nm; &#955;em 527 nm), Red (&#955;ex 594 nm; &#955;em 624 nm), FarRed (&#955;ex 644 nm; &#955;em 669 nm) or Brightfield (horseradish peroxidase (HRP) conjugated).</li>
	<li>Block the cells by adding a drop of Blocking Buffer to each well. Incubate for 30 min at 37 &#176;C in a humid chamber.</li>
	<li>Prepare antibody solutions by diluting stock antibodies in Antibody Dilution Buffer. For each well 40 &#181;L of antibody solution is required.</li>
	<li>Remove blocking buffer by placing a tissue paper at the edge of the well. Add 40 &#181;L of antibody diluted in Antibody Dilution Buffer to each well. Incubate for 60 min in a humid chamber at 37 &#176;C or overnight at 4 &#176;C.</li>
	<li>Remove antibody solution from the wells by placing a tissue paper at the edge of the well. Wash slides 2x in 100 mL of Wash Buffer &#8216;A&#8217; in a Coplin slide-staining jar for 5 min, without shaking.</li>
	<li>During the wash steps, dilute 5x secondary antibody probes, anti-rabbit PLUS &#38; anti-mouse MINUS (the species specificity of the probes depends on the primary antibodies used), in Antibody Dilution Buffer. Prepare 40 &#181;L of antibody solution per well.</li>
	<li>Add 40 &#181;L of secondary antibody solution to each well. Incubate for 60 min at 37 &#176;C in a humid chamber.</li>
	<li>Remove secondary antibody solution from the wells by placing a tissue paper at the edge of the well. Wash slides 2x in 100 mL of Wash Buffer &#8216;A&#8217; in a Coplin slide-staining jar for 5 min, without shaking.</li>
	<li>During the washes prepare 40 &#181;L of ligation mixture per well, by mixing 8 &#181;L of 5x Ligation Buffer, 31 &#181;L of ultrapure water and 1 &#181;L of ligase.</li>
	<li>Add 40 &#181;L of ligation mixture to each well. Incubate for 30 min at 37 &#176;C in a humid chamber.</li>
	<li>Remove ligation mixture from the wells by placing a tissue paper at the edge of the well. Wash slides 2x in 100 mL of Wash Buffer &#8216;A&#8217; in a Coplin slide-staining jar for 2 min, without shaking.</li>
	<li>During the washes prepare 40 &#181;L of amplification mixture per well, by mixing 8 &#181;L of 5x amplification solution, 31.5 &#181;L of ultrapure water, and 0.5 &#181;L of polymerase. 
	NOTE: The 5x amplification contains fluorescent probes. Protect this mixture from light. Also, protect the slides from light during each of the following steps. If using translucent Coplin slide-staining jars and humid chambers, wrap them in aluminum foil.</li>
	<li>Add 40 &#181;L of amplification mixture per well. Incubate for 100 min at 37 &#176;C in a humid chamber.</li>
	<li>Remove amplification mixture from the wells. Wash slides 2x in 100 mL of Wash Buffer &#8216;B&#8217; in a Coplin slide-staining jar for 10 min without shaking.</li>
	<li>Wash slides in 100 mL of Wash Buffer &#8216;B&#8217; diluted 1:100 in ultrapure water in a Coplin slide-staining jar for 30 s.</li>
	<li>Add 20 &#181;L of DAPI containing mounting medium per well to the slides. Close slides with a coverslip and seal slides with nail polish.</li>
	<li>If imaging, immediately incubate DAPI containing mounting medium for 10&#8211;15 min, while protected from light. If not, store the slides at -20 &#176;C for up to 1 week, protected from light.</li>
</ol>
5. Detection
<ol>
	<li>Use confocal microscopy to acquire images of HeLa, S. cerevisiae and E. coli cells with 20x/0.8 NA, 63x/1.4 NA and 100x/1.4 NA Plan Apochromat objectives, respectively. Excite DNA-stained DAPI with a 405 nm pulsed diode laser. For the PLA signal (for this study) excite with a 561 nm solid-state laser.</li>
</ol>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mycobacterial+stationary+phase+induced+by+low+oxygen+tension:+cell+wall+thickening+and+localization+of+the+16-kilodalton+alpha-crystallin+homolog.">Mycobacterial stationary phase induced by low oxygen tension: cell wall thickening and localization of the 16-kilodalton alpha-crystallin homolog.</a> Journal of Bacteriology. 180 (4), 801-808 (1998).</li><li>Werner-Washburne, M., Braun, E., Johnston, G. C., Singer, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stationary+phase+in+the+yeast+Saccharomyces+cerevisiae.">Stationary phase in the yeast Saccharomyces cerevisiae.</a> Microbiological Reviews. 57 (2), 383-401 (1993).</li></ol>

The authors have nothing to disclose.

Co-immunoprecipitation and co-localization based approaches have been used as long-standing methods to characterize protein assembles. The detection of transiently formed specific chaperone complexes is a major challenge with such conventional methods, and as a result, previous findings are largely restricted to qualitative&#160; interpretations. The cell lysis-based co-immunoprecipitation techniques often require cross-linking to stabilize protein-protein interactions.&#160;Cell lysis increases the risk of disrupting transiently formed chaperone complexes, while cross-linking could introduce non-native interactions, particularly driven by the inherent &#8220;stickiness&#8221; of most chaperones. When analyzing the Hsp70 chaperone system using co-immunoprecipitation, potential artifacts could&#160;arise from (a) high expression levels of the chaperone and (b)&#160;solubility&#160;issues pertaining to membrane or protein aggregate&#160;bound chaperone machineries. Besides, co-immunoprecipitation-based techniques provide no information on cellular localization of the captured protein assemblies, which could be important for delineating the associated functions. The drawbacks of using co-immunoprecipitation techniques are somewhat reduced in co-localization-based approaches that preserve&#160;protein localization information. However, the co-localization of two or more proteins could indicate either direct protein-protein association or partitioning of proteins into the same microdomain in cells. Therefore, co-localization analysis is, at best, speculative in predicting protein interactions and lacks any proximity value. Due to the bulk and indiscriminate detection of the targeted proteins, the technique is highly limited in studying specific protein assemblies in cells. This is particularly problematic when the targeted protein(s) could, in parallel, form a wide range of distinct&#160;protein assemblies as in the case with&#160;the Hsp70 chaperone system. Compared to these conventional methodologies, PLA is a more refined in situ technique developed to robustly detect and quantify native protein associations in cells with preserved protein localization information. A protein interaction is revealed by this assay based on the proximity (approximately 10-30 nm) between the targeted proteins. PLA is &#34;tunable&#34; in that the range of the proximity distance&#160;could be reduced to obtain a more stringent readout by (a) decreasing the length of the oligonucleotide tags attached to the PLA probes and/or (b) conjugating the tags directly to primary antibodies.&#160;Care should be taken when interpreting a positive PLA signal. Ideally, a protein interaction should be confirmed with two or more independent methodologies. The intensity of the fluorescent signal in PLA is not as deterministically related to protein cluster size or separation distance between the interacting proteins as is&#160;the case with FRET. However, PLA has a high degree of specificity and sensitivity, and could&#160;even be used for analyzing very low abundant proteins such as growth factors or&#160;cytokines and their interactions in rare cell types or clinical specimens13.&#160;
The Hsp70 chaperone partners with multiple co-chaperones (e.g., JDPs and NEFs) during its cellular lifetime14&#160;to drive distinct biological functions. The sheer number of probable Hsp70-JDP-NEF machine configurations and their dynamic behavior in cells have largely hampered a detailed understanding of the specific roles of these chaperone machines. Biological tools that allow selective tracing of distinct Hsp70 assemblies are, therefore, required to dissect the overall wiring of this chaperone network in cells. The transiently formed JDP-JDP and JDP-Hsp70 chaperone complexes7 were effectively captured in cells with PLA, indicating that this technique is suitable for studying molecular interactions with high dissociation constants (e.g., &#62;3 &#956;M&#160;Kd). Though, in the current work, the assay is primarily used as a &#8220;yes&#8221; or &#8220;no&#8221; type binary indicator for chaperone interaction, the users can employ&#160;this technique to obtain semi-quantitative readouts of the degree of interaction by digitally counting the fluorescence signal intensities15. However, due to the non-linear amplification of&#160;the PLA signal, caution should be exercised when interpreting PLA data in a quantitative manner16. Despite the aforementioned advantages, this method has certain limitations. One of the main disadvantages of PLA is that the assay requires cell fixation thus largely limiting its ability to resolve temporal dynamics of protein complexation in live cells. In comparison, in vivo FRET17 or Bioluminescence Resonance Energy Transfer (BRET)18 allows the monitoring of similar protein interactions in a spatio-temporal manner in living cells with relatively smaller proximity values (&#60;10 nm). In contrast to PLA, a&#160;FRET signal exhibits a strict linear correlation with protein expression levels16 making FRET the gold standard in quantitative analysis of protein interactions. However, unlike FRET&#8217;s dependence on the enigmatic orientation factor &#954;2, PLA is not influenced by the orientation of the PLA probes, which increases the probability of capturing a protein complex19. In terms of user-friendliness, FRET and BRET require unique expertise thus generally limiting the accessibility of these methodologies to the broader research community. Further, both of these techniques require modification of the interacting proteins by attaching bulky luminescent/fluorescent protein tags that could potentially interfere with protein function and complex formation.
The following steps (1-4) require careful consideration for the successful implementation of PLA in cells. (1) Antibody selection: The commercially available PLA kit is compatible with primary antibodies raised against mouse, rabbit and goat only. PLA requires highly specific primary antibodies that do not bind to off targets. Additionally, it is important to select primary antibodies that are compatible with applications such as immunohistochemistry (IHC),&#160;enzyme-linked immunosorbent assay (ELISA), and/or immunoprecipitation (IP) to ensure that the antibodies&#160;could recognize antigenic amino acid sequences exposed on the surface of folded proteins. Prior to use, testing of all primary antibodies for their specificity is highly recommended. This can be performed via western blot analysis of whole cell lysates. In cases where the targeted proteins have&#160;homologs with small variations in size and sequence similarity (e.g., Hsp70 and JDP paralogs), the specificity of the antibodies could&#160;be tested with gene knockdown/knockout approaches or by probing against&#160;purified homologs to rule out any potential cross recognition. If suitable antibodies are not available, the targeted proteins could be tagged with epitopes that have antibodies of acceptable quality (Figure 2F). (2) Fixation and permeabilization of cells: A treatment of 4% paraformaldehyde and 0.1-0.5% Triton-X100 could be used to fix and permeabilize cells, respectively. The use of 4% paraformaldehyde is a better treatment for preserving protein-protein interactions and cellular ultrastructure compared to fixative conditions employing&#160;99% methanol20,21,22. Fixing cells with 99% methanol, however, yields less cytoplasmic background staining, and perhaps is more suitable for specific cases such as detection of cytoskeleton associated protein assemblies21,22. Efficient permeabilization of both the plasma membrane as well as organelle membranes to increase antibody accessibility could be achieved with non-ionic surfactant Triton-X100. Triton-X100 can, however, non-specifically remove proteins from the plasma membrane23,24. Therefore, for the analysis of protein assemblies associated with cellular membranes, alternative detergents such as saponin or&#160;digitonin that targets sterols to permeabilize membranes could be applied21,22,25. (3) Disruption of the cell wall: Prior to membrane permeabilization, an additional step involving specific lytic enzymes is needed to increase the penetrability of antibodies in cell types with cell walls (e.g. fungi, plant and bacterial cells). For example, we employed lyticase, which degrades&#160;&#946;-glucan&#160;to disrupt the cell wall of S. cerevisiae26. Similarly, the E. coli cell wall was disrupted using lysozyme, an enzyme that targets peptidoglycans27,28. The cell wall composition and lytic enzyme sensitivity varies with different growth conditions26 and culture confluence29,30. Therefore, the concentration of the lytic enzymes and digestion times may vary and care has to be taken to prevent over- or underdigestion of cells. After the cell wall digestion, the cells are relatively fragile and require crowding agents such as sorbitol for them to remain intact during the washing steps. (4) DNA amplification: The DNA ligation and the rolling circle PCR reaction steps&#160;are sensitive to temperature and humidity fluctuations. To ensure robust DNA amplification and reproducibility of the assay, these reactions are required to be performed at 37 &#176;C in a humidity chamber. Importantly, the proccessed cells should be prevented from drying out while performing the assay&#160;to avoid any non-specific antibody binding and DNA amplification events that could lead to an increased in background signal.
All things considered, the implementation of this assay does not require unique expertise and sophisticated instrumentation. The successful monitoring of JDP-JDP and JDP-Hsp70 chaperone complexes illustrate&#160;the potential application of this technique to trace transiently formed protein assemblies&#160;in all cell types.&#160;Our implementation of the technique in yeast and bacteria significantly increases the applicability of PLA to study various biological processes mediated by distinct protein assemblies in a broad range of organisms. Further, our work highlights PLA as a promising new protein interaction tool to study evolutionary changes occurring at a molecular level across species8.

A vast amount of genomic information remains uninterpretable due to our incomplete understanding of cellular interactomes. Conventional protein-protein interaction detection methodologies such as protein co-immunoprecipitation with/without chemical cross-linking and protein co-localization, though widely used, pose a range of disadvantages. Some&#160; of the main disadvantages include poor quantification of the interactions and the potential introduction of non-native binding events.&#160;In comparison, emerging proximity-based techniques provide an alternative and a powerful approach for capturing protein interactions in cells. The proximity ligation assay (PLA)1, now available as a proprietary kit, employs&#160;antibodies to specifically target protein complexes based on the proximity of the interacting subunits.
PLA is initiated by the&#160; formation of a scaffold consisting of primary and secondary antibodies with small DNA tags (PLA probes) on the surface of the targeted protein complex (Figure 1, steps 1-3). Next, determined by the proximity of the DNA tags, a circular DNA molecule is generated via hybridization with connector oligonucleotides (Figure 1, step 4). The formation of the circular DNA is completed by a DNA ligation step. The newly formed circular piece of DNA serves as a template for the subsequent&#160;rolling circle amplification (RCA)-based polymerase chain reaction (PCR) primed by one of the conjugated oligonucleotide tags. This generates a single-stranded concatemeric DNA-molecule attached to the protein complex via the antibody scaffold (Figure 1, step 6). The concatemeric DNA molecule is visualized using fluorescently labeled oligonucleotides that hybridize to multiple unique sequences scattered across the amplified DNA&#160;(Figure 1, step 7)2. The generated PLA signal, which appears as a fluorescent dot (Figure 1, step 7), corresponds to the location of the targeted protein complex in the cell. As a result, the assay could detect protein complexes with high spatial accuracy. The technique is not limited to simply capturing protein interactions, but could also be utilized to detect single molecules or protein modifications on proteins with high sensitivity1,2.
Hsp70 forms a highly versatile chaperone system fundamentally important for maintaining cellular protein homeostasis by participating in an array of&#160;housekeeping and stress-associated functions. Housekeeping activities of the Hsp70 chaperone system include&#160;de novo protein folding, protein translocation across cellular membranes, assembly and disassembly of protein complexes, regulation of protein activity and linking different protein folding/quality control machineries3. The same chaperone system also refolds misfolded/unfolded proteins, prevents protein aggregation, promotes protein disaggregation and cooperates with cellular proteases to degrade terminally misfolded/damaged proteins to facilitate cellular repair after proteotoxic stresses4,5. To achieve this functional diversity, the Hsp70 chaperone relies on partnering co-chaperones of the JDP family and nucleotide exchange factors (NEFs) that fine-tune the Hsp70&#8217;s ATP-dependent allosteric control of substrate binding and release3,6. Further, the JDP co-chaperones play a vital role in&#160;selecting&#160;substrates for&#160;this versatile chaperone system.&#160;Members of this family are subdivided into three classes (A, B and C) based on their structural homology to the prototype JDP, the E. coli DnaJ. Class A JDPs contain an N-terminal J-domain, which interacts with Hsp70, a glycine-phenylalanine rich region, a substrate binding region consisting of a Zinc finger-like region (ZFLR) and two &#946;-barrel domains, and a C-terminal dimerization domain. JDPs with an N-terminal J-domain and a glycine-phenylalanine rich region, but lacking the ZFLR, fall into class B. In general, members of these two classes are&#160;involved in chaperoning functions. Members falling under the catchall class C, which contains JDPs that only share the J-domain4, recruit Hsp70s to perform a variety of non-chaperoning functions. The important role of JDPs as interchangeable substrate recognition &#8220;adaptors&#8221; of the Hsp70 system is reflected by an expansion of the family members during evolution. For example, humans have over 42 distinct JDP members4. These JDPs function as monomers, homodimers and/or homo/hetero oligomers4,5. Recently, a functional cooperation via transient complex formation between class A (e.g., H. sapiens&#160;DNAJA2; S. cerevisiae Ydj1) and class B (e.g., H. sapiens DNAJB1; S. cerevisiae Sis1) eukaryotic JDPs was reported to promote efficient recognition of amorphous protein aggregates in vitro7,8. These mixed class JDP complexes presumably assemble on the surface of aggregated proteins to facilitate the formation of Hsp70- and Hsp70+Hsp100-based protein disaggregases7,8,9,10. The critical evidence to support the existence of these transiently formed mixed class JDP complexes in eukaryotic cells was provided with PLA8.
PLA is increasingly employed for assessing protein interactions in metazoa, primarily in mammalian cells. Here, we report the successful expansion of this&#160;technique to monitor transiently formed chaperone complexes in eukaryotic and prokaryotic unicellular organisms such as the budding yeast S. cerevisiae and the bacterium E. coli.&#160;Importantly, this expansion highlights the potential use of PLA in detecting and analyzing microbes that infect human and animal cells.

<table><tbody><tr><td>37% Formaldehyde</td><td>Merck</td><td>103999</td><td></td></tr><tr><td>Acetone</td><td>Sigma-Aldrich</td><td>32201</td><td></td></tr><tr><td>anti-DNAJA2 antibody</td><td>Abcam</td><td>ab157216</td><td></td></tr><tr><td>anti-DNAJB1 Antibody</td><td>Enzo Life Sciences</td><td>ADI-SPA-450</td><td></td></tr><tr><td>anti-DnaK antibody</td><td>In house</td><td></td><td></td></tr><tr><td>anti-mCherry antibody</td><td>Abcam</td><td>ab125096</td><td></td></tr><tr><td>anti-Sis1 Antibody</td><td>Cosmo Bio Corp</td><td>COP-080051</td><td></td></tr><tr><td>anti-Ydj1 antibody</td><td>StressMarq Biosciences</td><td>&amp;nbsp;SMC-150,</td><td></td></tr><tr><td>anti-YFP antibody</td><td>In house</td><td></td><td></td></tr><tr><td>Coplin slide-staining jar</td><td>Sigma-Aldrich</td><td>S5516</td><td></td></tr><tr><td>Diagnostic slides</td><td>Marienfeld</td><td>1216530</td><td></td></tr><tr><td>DMEM</td><td>Thermo-Fischer</td><td>31966021</td><td></td></tr><tr><td>DuoLink In Situ Detection Reagents Orange</td><td>Sigma-Aldrich</td><td>DUO92007</td><td></td></tr><tr><td>DuoLink In Situ Mounting Medium + DAPI</td><td>Sigma-Aldrich</td><td>DUO82040</td><td></td></tr><tr><td>DuoLink In Situ PLA Probe Anti-Mouse MINUS</td><td>Sigma-Aldrich</td><td>DUO92004</td><td></td></tr><tr><td>DuoLink In Situ PLA Probe Anti-Rabbit PLUS</td><td>Sigma-Aldrich</td><td>DUO92002</td><td></td></tr><tr><td>DuoLink In Situ Wash Buffers, Fluorescence</td><td>Sigma-Aldrich</td><td>DUO82049</td><td></td></tr><tr><td>Fetal Calf Serum</td><td>Thermo-Fischer</td><td>10082147</td><td></td></tr><tr><td>Lysozyme</td><td>Sigma-Aldrich</td><td>62971</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>32213</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>P6148</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Thermo-Fischer</td><td>15070063</td><td></td></tr><tr><td>Poly-L-Lysine</td><td>Sigma-Aldrich</td><td>P47-07</td><td></td></tr><tr><td>Sorbitol</td><td>Sigma-Aldrich</td><td>S7547</td><td></td></tr><tr><td>Triton-X100</td><td>Merck</td><td>108643</td><td></td></tr><tr><td>Trypsin</td><td>Thermo-Fischer</td><td>25300096</td><td></td></tr><tr><td>Tween-20</td><td>Sigma-Aldrich</td><td>P1379</td><td></td></tr><tr><td>Zymolase 100T / / Lyticase</td><td>United States Biological</td><td>Z1004</td><td></td></tr></tbody></table>

in situ monitoring of transiently formed molecular chaperone assemblies in bacteria, yeast, and human cells

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

Our previous in vitro studies using purified proteins revealed that a subset of human class A and class B JDPs form transient mixed class JDP complexes to efficiently target a broad range of aggregated proteins and possibly facilitate the assembly of Hsp70-based protein disaggregases7. We employed PLA to determine whether mixed class (A+B) JDP complexes occur in&#160;human cervical cancer cells (HeLa). Human JDPs DNAJA2 (class A) and DNAJB1 (class B) were targeted with highly specific primary antibodies and secondary PLA probes (Figure 2A-C). The appearance of red fluorescent punctae&#160;indicated the presence of mixed class DNAJA2 and DNAJB1 complexes in HeLa cells (Figure 2C) confirming our previous biochemical findings7. Each punctum represents an individual protein interaction event in the HeLa cell cytosol/nucleus.
Previous biochemical results obtained from F&#246;rster resonance energy transfer (FRET), which detects protein interactions as a readout of the amount of energy transferred from an excited donor fluorophore attached to one protein to a suitable acceptor fluorophore attached to the second protein, indicated that similar mixed class complexes could also occur between yeast JDPs, in vitro8. Confirming our biochemical findings, we observed the formation of mixed class complexes between Ydj1 (class A) and Sis1 (class B) in unicellular eukaryote S. cerevisiae (baker&#8217;s yeast) cells with PLA (Figure 2F). In yeast, due to their small cell size and the coalescence of multiple punctae, the individual fluorescent dots generated by PLA could often be less distinguishable.&#160;A considerable&#160;decrease in fluorescent punctae formation was observed after knockout or knockdown of interacting JDPs in both human and yeast cells8, which&#160;demonstrates the high specificity of PLA in capturing these co-chaperone complexes in different cell types. In contrast to their eukaryotic counterparts, the prokaryotic (E. coli) JDPs lacked the ability to form mixed class JDP complexes and functionally cooperate to boost protein disaggregation, in vitro8. In agreement with the biochemical analysis, we did not observe mixed class JDP complex formation between bacterial DnaJ-YFP&#160;(class A) and CbpA-mCherry&#160;(class B), the only&#160;JDPs in E. coli (Figure 2I). However, our PLA setup was able to capture other chaperone assemblies involving the two JDPs. For example, we detected chaperone complexes between DnaJ-YFP&#160;and DnaK (bacterial Hsp70) (Figure 2L) and CbpA-mCherry and DnaK8 (data not shown). These two&#160;bacterial chaperone complexes are extensively characterized both in vitro and in vivo8,11,12 thus confirming our PLA results. Together, these observations demonstrate the ability of PLA to robustly capture transiently formed chaperone machineries in unicellular/multicellular eukaryotic and prokaryotic cells. The technical controls lacking a primary antibody against one of the interacting JDPs/chaperones, but containing both the mouse and rabbit derived secondary PLA probes, showed little to no background fluorescence signal (Figure 2A,B,D,E,G,H,J,K) indicating a lack of false positive signal amplification in our experimental setup.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60172/60172fig01.jpg" /> 
Figure 1: Schematic representation of the chronological steps of the Proximity Ligation Assay.&#160;(1) Complex between protein A and protein B. (2) Binding of primary (1&#176;) antibodies (green and light brown) to proteins A and B. The two primary antibodies must be raised in different host species (e.g. mouse, rabbit or goat). (3) Primary antibodies are recognized by species-specific secondary (2&#176;) antibodies covalently attached with DNA oligo tags (PLA probes). (4) When proteins A and B are in complex, the bound PLA probes are in close proximity to facilitate the DNA oligos to hybridize with connector DNA strands, which results in the formation of a circular DNA molecule. Subsequently, a DNA ligase facilitates the joining of DNA strands together by catalyzing the formation of a phosphodiester bond. (5) Initiation of the Rolling Circle Amplification (RCA) of the circular DNA molecule by a bacterial DNA polymerase at 37&#176; C. RCA reaction is primed by one of the antibody-conjugated DNA oligo tags. (6) Generation of a single-stranded concatemeric DNA molecule attached to one of the secondary antibodies. (7) Hybridization of fluorescently-labeled complementary oligonucleotide probes to a unique repetitive sequence in the concatemeric DNA molecule. After the hybridization step, the concatemeric DNA molecule could be visualized as a bright fluorescent dot at the location of the targeted protein complex in fixed cells. Bottom&#160;denote the prokaryotic and eukaryotic cell types in which the PLA technique is applicable. <a href="https://www.jove.com/files/ftp_upload/60172/60172fig01large.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/60172/60172fig02.jpg" /> 
Figure 2. Molecular chaperone assemblies captured by PLA in prokaryotic and eukaryotic cells.&#160;(A-C) Detection of mixed class JDP complexes formed between DNAJA2 (class A) and DNAJB1 (class B) in human cervical cancer cell line HeLa. PLA performed with (A) anti-DNAJA2 antibody alone (negative technical control); (B) anti-DNAJB1 antibody alone (negative technical control); (C) anti-DNAJB1 and anti-DNAJA2 antibodies together. The appearance of multiple fluorescent dots in panel C (positive signal) indicates the presence of protein complexes formed between DNAJA2 and DNAJB1. Each red fluorescent dot/punctum represents a single interaction. Nuclei (DNA) stained with DAPI (cyan). (D-F) Detection of mixed class JDP complexes formed between Ydj1 (class A) and Sis1 (class B) in S. cerevisiae cells. (D) PLA performed with anti-Ydj1 antibody alone (negative technical control); (E) PLA performed with anti-Sis1 antibody alone (negative technical control); (F) PLA performed with anti-Ydj1 and anti-Sis1 antibodies together. The positive fluorescent signal denotes the presence of Ydj1 and Sis1 complexes in S. cerevisiae. Yeast nuclei stained with DAPI (cyan). (G-L) Detection of chaperone complexes formed between prokaryotic Hsp70 (DnaK) and JDPs (DnaJ and CbpA) in E. coli cells. Due to the lack of specific primary antibodies against prokaryotic JDPs, the E. coli DnaJ (class A) and CbpA (class B) were tagged with YFP and mCherry, respectively. (G, J) PLA performed with anti-YFP alone (negative technical control); (H) PLA performed with anti-mCherry alone (negative technical control). (I) PLA performed with anti-YFP and anti-mCherry antibody. The lack of fluorescent dot formation indicates no complex formation between DnaJ and CbpA. (K) PLA performed with anti-DnaK antibody alone (negative technical control). (L) PLA performed with anti-YFP and anti-DnaK antibodies together. The positive fluorescent signal indicates complex formation between DnaK and DnaJ. Bacterial DNA stained with DAPI (cyan). In addition to containing a single primary antibody, all the negative technical controls were performed in the presence of the respective secondary PLA probes. Scale bars = 10 &#181;m.

Watch this Scientific Journal Video about In Situ Monitoring of Transiently Formed Molecular Chaperone Assemblies in Bacteria, Yeast, and Human Cells at JoVE.com

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

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

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

in-situ-monitoring-transiently-formed-molecular-chaperone-assemblies

Nanyang Technological University

Research

JoVE Journal

Biology

7.0K Views.  University of Groningen. Cognate J-domain proteins cooperate with the Hsp70 chaperone to assist in a myriad of biological processes ranging from protein folding to degradation. Here, we describe an in situ proximity ligation assay, which allows the monitoring of these transiently formed chaperone machineries in bacterial, yeast and human cells.

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

Detection of the pH-dependent Activity of Escherichia coli Chaperone HdeB In Vitro and In Vivo

Bacteria are frequently exposed to environmental changes, such as alterations in pH, temperature, redox status, light exposure or mechanical force. Many of these conditions cause protein unfolding in the cell and have detrimental impact on the survival of the organism. A group of unrelated, stress-specific molecular chaperones have been shown to play essential roles in the survival of these stress conditions. While fully folded and chaperone-inactive before stress, these proteins rapidly unfold and become chaperone-active under specific stress conditions. Once activated, these conditionally disordered chaperones bind to a large number of different aggregation-prone proteins, prevent their aggregation and either directly or indirectly facilitate protein refolding upon return to non-stress conditions. The primary approach for gaining a more detailed understanding about the mechanism of their activation and client recognition involves the purification and subsequent characterization of these proteins using in vitro chaperone assays. Follow-up in vivo stress assays are absolutely essential to independently confirm the obtained in vitro results.
This protocol describes in vitro and in vivo methods to characterize the chaperone activity of E. coli HdeB, an acid-activated chaperone. Light scattering measurements were used as a convenient read-out for HdeB&#39;s capacity to prevent acid-induced aggregation of an established model client protein, MDH, in vitro. Analytical ultracentrifugation experiments were applied to reveal complex formation between HdeB and its client protein LDH, to shed light into the fate of client proteins upon their return to non-stress conditions. Enzymatic activity assays of the client proteins were conducted to monitor the effects of HdeB on pH-induced client inactivation and reactivation. Finally, survival studies were used to monitor the influence of HdeB&#39;s chaperone function in vivo.

Detection of the pH-dependent Activity of Escherichia coli Chaperone HdeB In Vitro and In Vivo

This study describes biophysical, biochemical and molecular techniques to characterize the chaperone activity of Escherichia coli HdeB under acidic pH conditions. These methods have been successfully applied for other acid-protective chaperones such as HdeA and can be modified to work for other chaperones and stress conditions.

Bacteria are frequently exposed to environmental changes, such as alterations in pH, temperature, redox status, light exposure or mechanical force. Many ...

Biochemistry

Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of reactive oxygen species, and more. These fluctuations can lead to a widespread protein unfolding, aggregation, and cell death. Therefore, cells have evolved a dynamic and stress-specific network of molecular chaperones, which maintain a "healthy" proteome during stress conditions. ATP-independent chaperones constitute one major class of molecular chaperones, which serve as first-line defense molecules, protecting against protein aggregation in a stress-dependent manner. One feature these chaperones have in common is their ability to utilize structural plasticity for their stress-specific activation, recognition, and release of the misfolded client. 
In this paper, we focus on the functional and structural analysis of one such intrinsically disordered chaperone, the bacterial redox-regulated Hsp33, which protects proteins against aggregation during oxidative stress. Here, we present a toolbox of diverse techniques for studying redox-regulated chaperone activity, as well as for mapping conformational changes of the chaperone, underlying its activity. Specifically, we describe a workflow which includes the preparation of fully reduced and fully oxidized proteins, followed by an analysis of the chaperone anti-aggregation activity in vitro using light-scattering, focusing on the degree of the anti-aggregation activity and its kinetics. To overcome frequent outliers accumulated during aggregation assays, we describe the usage of Kfits, a novel graphical tool which allows easy processing of kinetic measurements. This tool can be easily applied to other types of kinetic measurements for removing outliers and fitting kinetic parameters. To correlate the function with the protein structure, we describe the setup and workflow of a structural mass spectrometry technique, hydrogen-deuterium exchange mass spectrometry, that allows the mapping of conformational changes on the chaperone and substrate during different stages of Hsp33 activity. The same methodology can be applied to other protein-protein and protein-ligand interactions.

One of the most challenging stress conditions that organisms encounter during their lifetime involves the accumulation of oxidants. During oxidative stress, cells heavily rely on molecular chaperones. Here, we present methods used to investigate the redox-regulated anti-aggregation activity, as well as to monitor structural changes governing the chaperone function using HDX-MS.

Living organisms regularly need to cope with fluctuating environments during their life cycle, including changes in temperature, pH, the accumulation of ...

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 ...

Studies of Chaperone-Cochaperone Interactions using Homogenous Bead-Based Assay

Targeting the heat shock protein 90 (Hsp90)-cochaperone interactions provides the possibility to specifically regulate Hsp90-dependent intracellular processes. The conserved MEEVD pentapeptide at the C-terminus of Hsp90 is responsible for the interaction with the tetratricopeptide repeat (TPR) motif of co-chaperones. FK506-binding protein (FKBP) 51 and FKBP52 are two similar TPR-motif co-chaperones involved in steroid hormone-dependent diseases with different functions. Therefore, identifying molecules specifically blocking interactions between Hsp90 and FKBP51 or FKBP52 provides a promising therapeutic potential for several human diseases. Here, we describe the protocol for an amplified luminescent proximity homogenous assay to probe interactions between Hsp90 and its partner co-chaperones FKBP51 and FKBP52. First, we have purified the TPR motif-containing proteins FKBP51 and FKBP52 in glutathione S-transferase (GST)-tagged form. Using the glutathione-linked donor beads with GST-fused TPR-motif proteins and the acceptor beads coupled with a 10-mer C-terminal peptide of Hsp90, we have probed protein-protein interactions in a homogeneous environment. We have used this assay to screen small molecules to disrupt Hsp90-FKBP51 or Hsp90-FKBP52 interactions and identified potent and selective Hsp90-FKBP51 interaction inhibitors.

This protocol presents a technique for probing protein-protein interactions using glutathione-linked donor beads with GST-fused TPR-motif co-chaperones and acceptor beads coupled with an Hsp90-derived peptide. We have used this technique to screen small molecules to disrupt Hsp90-FKBP51 or Hsp90-FKBP52 interactions and identified potent and selective Hsp90-FKBP51 interaction inhibitors.

Targeting the heat shock protein 90 (Hsp90)-cochaperone interactions provides the possibility to specifically regulate Hsp90-dependent intracellular ...

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. ...

Intracellular Refolding Assay

This protocol describes a method to measure the enzymatic activity of molecular chaperones in a cell-based system and the possible effects of compounds with inhibitory/stimulating activity. Molecular chaperones are proteins involved in regulation of protein folding1 and have a crucial role in promoting cell survival upon stress insults like heat shock2, nutrient starvation and exposure to chemicals/poisons3. For this reason chaperones are found to be involved in events like tumor development, chemioresistance of cancer cells4 as well as neurodegeneration5. Design of small molecules able to inhibit or stimulate the activity of these enzymes is therefore one of the most studied strategies for cancer therapy7 and neurodegenerative disorders9. The assay here described offers the possibility to measure the refolding activity of a particular molecular chaperone and to study the effect of compounds on its activity. In this method the gene of the molecular chaperone investigated is transfected together with an expression vector encoding for the firefly luciferase gene. It has been already described that denaturated firefly luciferase can be refolded by molecular chaperones10,11. As normalizing transfection control, a vector encoding for the renilla luciferase gene is transfected. All transfections described in this protocol are performed with X-treme Gene 11 (Roche) in HEK-293 cells. In the first step, protein synthesis is inhibited by treating the cells with cycloheximide. Thereafter protein unfolding is induced by heat shock at 45&deg;C for 30 minutes. Upon recovery at 37&deg;C, proteins are re-folded into their active conformation and the activity of the firefly luciferase is used as read-out: the more light will be produced, the more protein will have re-gained the original conformation. Non-heat shocked cells are set as reference (100% of refolded luciferase).

In this protocol a method to measure intracellular protein refolding after heat shock is described. This method can be used to study foldases like molecular chaperones and their co-factors or compounds able to influence their activity. Firefly luciferase activity is used as reporter to measure chaperone refolding activity.

This protocol describes a method to measure the enzymatic activity of molecular chaperones in a cell-based system and the possible effects of compounds ...

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 ...

Adenofection: A Method for Studying the Role of Molecular Chaperones in Cellular Morphodynamics by Depletion-Rescue Experiments

Cellular processes such as mitosis and cell differentiation are governed by changes in cell shape that largely rely on proper remodeling of the cell cytoskeletal structures. This involves the assembly-disassembly of higher-order macromolecular structures at a given time and location, a process that is particularly sensitive to perturbations caused by overexpression of proteins. Methods that can preserve protein homeostasis and maintain near-to-normal cellular morphology are highly desirable to determine the functional contribution of a protein of interest in a wide range of cellular processes. Transient depletion-rescue experiments based on RNA interference are powerful approaches to analyze protein functions and structural requirements. However, reintroduction of the target protein with minimum deviation from its physiological level is a real challenge. Here we describe a method termed adenofection that was developed to study the role of molecular chaperones and partners in the normal operation of dividing cells and the relationship with actin remodeling. HeLa cells were depleted of BAG3 with siRNA duplexes targeting the 3&#39;UTR region. GFP-tagged BAG3 proteins were reintroduced simultaneously into &#62;75% of the cells using recombinant adenoviruses coupled to transfection reagents. Adenofection enabled to express BAG3-GFP proteins at near physiological levels in HeLa cells depleted of BAG3, in the absence of a stress response. No effect was observed on the levels of endogenous Heat Shock Protein chaperones, the main stress-inducible regulators of protein homeostasis. Furthermore, by adding baculoviruses driving the expression of fluorescent markers at the time of cell transduction-transfection, we could dissect mitotic cell dynamics by time-lapse microscopic analyses with minimum perturbation of normal mitotic progression. Adenofection is applicable also to hard-to-infect mouse cells, and suitable for functional analyses of myoblast differentiation into myotubes. Thus adenofection provides a versatile method to perform structure-function analyses of proteins involved in sensitive biological processes that rely on higher-order cytoskeletal dynamics.

We describe a method for depletion-rescue experiments that preserves cellular integrity and protein homeostasis. Adenofection enables functional analyses of proteins within biological processes that rely on finely tuned actin-based dynamics, such as mitotic cell division and myogenesis, at the single-cell level.

Cellular processes such as mitosis and cell differentiation are governed by changes in cell shape that largely rely on proper remodeling of the cell ...

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

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Chapters in this video

Detection of the pH-dependent Activity of Escherichia coli Chaperone HdeB In Vitro and In Vivo

Defining Hsp33's Redox-regulated Chaperone Activity and Mapping Conformational Changes on Hsp33 Using Hydrogen-deuterium Exchange Mass Spectrometry

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

Monitoring Protein-RNA Interaction Dynamics In Vivo at High Temporal Resolution Using χCRAC

Studies of Chaperone-Cochaperone Interactions using Homogenous Bead-Based Assay

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

Intracellular Refolding Assay

4D Imaging of Protein Aggregation in Live Cells

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Adenofection: A Method for Studying the Role of Molecular Chaperones in Cellular Morphodynamics by Depletion-Rescue Experiments