This work was supported by the Russian Science Foundation projects 19-74-30026 (method development and validation) and 19-74-10099 (protein-protein interaction assays); and by the Ministry of Science and Higher Education of the Russian Federation-grant 075-15-2019-1661 (analysis of representative protein-protein interactions).

The schematic representation of the method workflow is shown in Figure 1.
1. Co-transformation of E. coli
<ol>
	<li>Prepare expression vectors for target proteins using standard cloning methods. 
	NOTE: Typically, a good starting point is to use conventional pGEX/pMAL vectors bearing an ampicillin resistance gene and ColE1 origin for the expression of GST/MBP-tagged proteins and a compatible vector with p15A or RSF origin and kanamycin resistance to express 6xHis-tagged proteins, in some cases combined with either Thioredoxin or SUMO-tag to increase solubility. Usually, several combinations of tags need to be tested prior to the experiment. The described method itself is convenient for batch testing the expression conditions of target proteins. It is important to note that most Rosetta strains already contain the plasmid with p15A origin for expressing the tRNAs for rare codon;, thus if using such strains is a possible option, the p15A plasmids should be avoided. See the Table of Materials for details.
	<ol>
		<li>Grow bacteria of an appropriate strain in Luria-Bertani (LB) media at 37 &#176;C to an optical density (OD) of 0.1-0.2. BL21(DE3) strain was used for examples in this study. 
		NOTE: It is recommended to use freshly prepared competent cells to achieve efficient co-transformation with two vectors. If more than two vectors need to be co-transformed, it is better to transform them sequentially to achieve good transformation efficiency. Electroporation is a good alternative.</li>
		<li>Centrifuge 1.0 mL of the bacterial suspension for 1 min at 9,000 x g at 4 &#176;C, and discard the supernatant.</li>
		<li>Add 0.5 mL of ice-cold buffer transformation buffer (TB) (10 mM MOPS [pH 6.7], 250 mM KCl, 55 mM MnCl2, and 15 mM CaCl2) and incubate for 10 min on ice.</li>
		<li>Centrifuge for 30 s at 8,000 x g at 4 &#176;C, and discard the supernatant.</li>
		<li>Add 100 &#181;L of TB buffer, add 100 ng of each vector, and incubate for 30 min on ice. Separately transform single vectors to study protein behavior without co-expression. Additionally, co-transform expression vectors in pairs with empty co-expression vectors for non-specific binding controls. 
		&#8203;NOTE: In the examples provided in this study, pGEX/pMAL vectors with corresponding cDNAs fused to GST/MBP cDNAs were used in combination with compatible pACYC-derived vectors bearing cDNA encoding partner protein domains fused to Thioredoxin cDNA.</li>
		<li>Heat at 42 &#176;C for 150 s, then chill for 1 min on ice.</li>
		<li>Add 1 mL of liquid LB media without antibiotics and incubate at 37 &#176;C for 90 min. Plate on LB-agar plates containing 0.5% glucose and corresponding antibiotics (common concentrations are: 50 mg/L ampicillin; 20 mg/L kanamycin; 50 mg/L streptomycin; 35 mg/L chloramphenicol). Incubate the plates overnight at 37 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Expression
<ol>
	<li>Flush the cells from the plate with 2 mL of liquid LB media into 50 mL of LB media with corresponding antibiotics (common concentrations are: 50mg/L ampicillin; 20 mg/L kanamycin; 50 mg/L streptomycin; 35 mg/L chloramphenicol). Add metal ions or other known co-factors (in the examples provided in this study, 0.2 mM ZnCl2 was added to the media). Store an aliquot with 20% glycerol at -70 &#176;C for a subsequent repeat of the experiment. 
	NOTE: It is recommended to flush several colonies directly from the plate to exclude the possibility of bad expression in a single isolated clone due to occasional recombination events between two plasmids.</li>
	<li>Grow the cells with a constant rotation of 220 rpm at 37 &#176;C to an OD of 0.5-0.7, cool to room temperature (RT), and add isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) to 1 mM. Store a 20 &#181;L aliquot of cell suspension as a control of the un-induced sample.</li>
	<li>Incubate the cells with a constant rotation of 220 rpm at 18 &#176;C overnight. 
	NOTE: The optimal time and temperature of incubation may vary; 18 &#176;C overnight works best for most proteins and is advised to be tried by default. Reduce the incubation time to 2-3 h if a strong non-specific binding is observed.</li>
	<li>Divide the bacterial suspension into two parts (or more if more than two different tags were used) and store a 20 &#181;L aliquot of the cell suspension to confirm protein expression. Centrifuge at 4,000 x g for 15 min. 
	NOTE: Pause point: bacterial pellets can be stored at -70 &#176;C for at least 6 months.</li>
</ol>
3. Pulldown assay
NOTE: The detailed procedures are described for proteins tagged with either 6xHis or MBP/GST. All procedures are performed at 4&#176;C.
<ol>
	<li>Resuspend the bacterial pellets in 1 mL of ice-cold lysis buffer with protease inhibitors and reducing agents (see below) added immediately prior to the experiment. Avoid dithiotreitol (DTT) when using metal-chelating resins since it strips out metal ions. Adjust the buffer composition for the tested proteins. The common recipes of lysis buffers found suitable for most proteins are:
	<ol>
		<li>6xHis-pulldown: Mix 30 mM HEPES (pH 7.5), 400 mM NaCl, 10 mM Imidazole, 0.1% NP40, 10% [w/w] glycerol, 5 mM beta-mercaptoethanol, 1 mM phenylmethylsfulfonylfluoride (PMSF), and a 1:1,000 dilution of the protease inhibitor cocktail (see Table of Materials).</li>
		<li>GST- or MBP-pulldown: Mix 20 mM Tris (pH 7.5 at 25 &#176;C), 150 mM NaCl, 10 mM KCl, 10 mM MgCl2, 0.1 mM ZnCl2, 0.1% NP40, 10% [w/w] glycerol, 5 mM DTT, 1 mM PMSF, and a 1:1,000 dilution of the protease inhibitor cocktail (see Table of Materials).</li>
	</ol>
	</li>
	<li>Disrupt the cells by sonication on ice. Store a 20 &#181;l aliquot for electrophoresis. 
	NOTE: Typically, 20-25 pulses of 5 s with 15 s intervals with 20 W output power are required per sample. The appropriate sonication power should be adjusted for each instrument to avoid overheating and ensure total cell disruption. To achieve better performance, it is strongly recommended to use high-throughput multi-tip sonicator probes.</li>
	<li>Centrifuge at 20,000 x g for 30 min. Collect 20 &#181;L of the clarified lysate for subsequent SDS-PAGE analysis.</li>
	<li>Equilibrate the resin (50 &#181;L for each sample) with 1 mL of ice-cold lysis buffer for 10 min, centrifuge at 2,000 x g for 30 s, and discard the supernatant.</li>
	<li>Add cell lysates (total protein concentration: 20-50 mg/mL) to the resin, incubate for 10 min at a constant rotation of 15 rpm, centrifuge at 2,000 x g for 30 s, and discard the supernatant. Collect 20 &#181;L of the unbound fraction for subsequent SDS-PAGE analysis.</li>
	<li>Add 1 mL of ice-cold wash buffer and incubate for 1 min. Centrifuge at 2,000 x g for 30 s, and discard the supernatant. The common recipes for wash buffers are:
	<ol>
		<li>6xHis-pulldown: Mix 30 mM HEPES (pH 7.5), 400 mM NaCl, 30 mM Imidazole, 0.1% NP40, 10% [w/w] glycerol, and 5 mM beta-mercaptoethanol.</li>
		<li>GST- or MBP-pulldown: Mix 20 mM Tris (pH 7.5 at 25 &#176;C), 500 mM NaCl, 10 mM KCl, 10 mM MgCl2, 0.1 mM ZnCl2, 0.1% NP40, 10% [w/w] glycerol, and 5 mM DTT.</li>
	</ol>
	</li>
	<li>Perform two long washes: add 1 mL of ice-cold wash buffer, incubate for 10-30 min with a constant rotation of 15 rpm, centrifuge at 2,000 x g for 30 s, and discard the supernatant.</li>
	<li>Add 1 mL of ice-cold wash buffer, incubate for 1 min, centrifuge at 2,000 x g for 30 s, and discard the supernatant.</li>
	<li>Elute the bound proteins with 50 &#956;L of the elution buffer in a shaker at 1,200 rpm for 10 min. The common recipes of elution buffers are:
	<ol>
		<li>6xHis-pulldown: Mix 30 mM HEPES (pH 7.5), 400 mM NaCl, 300 mM Imidazole, and 5 mM beta-mercaptoethanol.</li>
		<li>GST-pulldown: 20 mM Tris (pH 7.5 at 25 &#176;C), 150 mM NaCl, 50 mM Glutathione (adjusted to pH 7.5 with basic Tris), and 5 mM DTT.</li>
		<li>MBP-pulldown: Mix 20 mM Tris (pH 7.5 at 25 &#176;C), 150 mM NaCl, 40 mM maltose, and 5 mM DTT.</li>
	</ol>
	</li>
	<li>Analyze the eluted proteins with SDS-PAGE. 
	NOTE: The percentage of acrylamide should be adjusted to the size of the proteins. In the examples provided in this study, 12% acrylamide gels were used, running in tris-glycine-SDS buffer (2 mM Tris, 250 mM Glycine, 0.1% SDS) at constant voltage of 180 V. Gels were stained by boiling in 0.2% Coomassie blue R250, 10% acetic acid, and 30% isopropanol, and de-stained by boiling in 10% acetic acid. The amount of loaded protein should not be equal, since various amounts of interacting proteins can be pulled down in different experiments.</li>
</ol>

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The authors declare no competing interests.

The described method allows the testing of protein-protein interactions that cannot be efficiently assembled in vitro and require co-expression. The method is one of the few suitable approaches for studying heterodimerizing proteins, which are also capable of homodimerization since, when purified separately, such proteins form stable homodimers which most often cannot dissociate and re-associate during the experiment3,11.
The workflow of the described method is more time-efficient compared to traditional pulldown because it lacks the time-consuming steps of separate purification of interacting proteins and their following incubation. Another advantage is a higher reproducibility, due to a significantly smaller number of steps and a shorter period of time in which proteins exist within an artificial in vitro environment while being exposed to proteolysis and oxidation. The method was successfully applied for studying several protein-protein interactions when other in vitro techniques were found to be unsuitable3,11,27. Parallel co-precipitation with different affinity tags provides protein expression level control. Since the method is easier and faster than conventional pulldown, it can be used instead of it, even if co-expression is not absolutely required. The speed of cell disruption is a non-obvious bottleneck which can significantly increase the total time and can lead to a deviation in results, due to the various amount of time between the first and last samples being exposed to in vitro conditions and proteolysis. Thus, using high-throughput multi-tip sonicator probes is strongly recommended; see the Table of Materials for examples. 
Using magnetic beads could decrease the non-specific binding and further speed up the method, reducing the time required for washing steps. Using vectors with compatible origins has an advantage over polycistronic expression as they provide a convenient combinatorial approach to test multiple protein interactions with the same target and do not require producing a new vector for each combination.
A disadvantage of the method is the relatively high probability of false-positive results, as proteins are co-expressed in bacteria at high concentrations. Thus, unexpected interactions discovered by this method should be treated with caution and tested with an independent assay, such as the Y2A or cellular techniques24, which also uses co-expression3,11,28. High-throughput bioinformatic approaches can also be used to analyze and validate complex protein-protein interaction networks29. Another peculiar obstacle is that protein complexes may have completely different biochemical properties compared to separate proteins; the complex might be insoluble while its components are present in the solution. This problem can typically be solved by fusing the proteins to the appropriate solubility tag. MBP is the most effective, while NusA is another good all-around alternative22. GST-tag was found to be efficient with many zinc-coordinating domains, although since it is dimeric, it should be avoided when working with oligomeric domains. On the contrary, small monomeric domains such as Thioredoxin and SUMO work well with multimeric protein domains.
Critical steps of the described method are the proper time of protein expression (a shorter time is required if non-specific binding is observed, while longer incubation times may be required to improve poor protein expression), fast cell disruption, proper choice of buffers, and maintaining the constant temperature during the protocol. Excessive incubation times after induction may result in protein aggregation in bacteria, leading to false positive results. On the other hand, some proteins require more time to be expressed in sufficient quantities. Temperature fluctuations during the sonication and wash steps may lead to changes in buffer pH and protein precipitation. Improper buffer choice also may result in non-specific protein aggregation.
In the case of poor protein expression, different solubility tags should be attempted, as well as an increased incubation time after induction. If a strong non-specific association is observed, all necessary negative controls should be run to determine the possible non-specific association of proteins with resin or the affinity tag. In case the controls do not work, different combinations of affinity tags should be attempted and different buffers used. Many proteins tend to non-specifically bind to the metal-chelating resin-like ZADs mentioned in this article; in such cases, MBP/GST-pulldown, in general, would be sufficient without a reciprocal experiment that can be used only for protein expression control. If negative controls work well, the protein expression time should be decreased, and the buffer system and reducing agent changed or the reducing agent concentration increased, especially when working with cysteine-rich proteins. In the case of poor reproducibility of results, attention should be paid to the cell disruption step, which should be performed fast but without overheating. The temperature should be monitored throughout the whole experiment.
This method can be easily modified to study multiprotein complexes or ribonucleoproteins. Another possible application is the batch testing of protein complex expression and purification conditions for subsequent studies.

Conventional pulldown is widely used to study protein-protein interactions1. However, purified proteins often do not interact effectively in vitro2,3, and some of them are insoluble without their binding partner4,5. Such proteins might require co-translational assembly or the presence of molecular chaperones5,6,7,8,9. Another limitation of conventional pulldown is the testing of possible heteromultimerization activity between domains that can exist as stable homo-oligomers assembled co-translationally8,10, as many of them cannot dissociate and re-associate in vitro during the incubation time. Co-expression was found to be useful in overcoming such problems3,11. Co-expression using compatible vectors in bacteria was successfully used to purify large multi-subunit macromolecular complexes, inclduing polycomb repressive complex PRC212, RNA polymerase II mediator head module13, bacteriophage T4 baseplate14, SAGA complex deubiquitinylase module15,16, and ferritin17. Replication origins commonly used for co-expression are ColE1, p15A18, CloDF1319, and RSF20. In the commercially available Duet expression system, these origins are combined with different antibiotic resistance genes and convenient multiple cloning sites to produce polycistronic vectors, allowing the expression of up to eight proteins. These origins have different copy numbers and can be used in varying combinations to achieve balanced expression levels of target proteins21. To test protein-protein interactions, various affinity tags are used; the most common are 6xHistidine, glutathione-S-transferase (GST), and maltose-binding protein (MBP), each of which has a specific affinity to the corresponding resin. GST and MBP also enhance the solubility and stability of tagged proteins22.
A number of methods involving protein co-expression in eukaryotic cells have also been developed, the most prominent of which is yeast two-hybrid assay (Y2H)23. Y2H assay is cheap, easy, and allows the testing of multiple interactions; however, its workflow takes more than 1 week to complete. There are also a few less frequently used mammalian cell-based assays, for example, fluorescent two-hybrid assay (F2H)24 and cell array protein-protein interaction assay (CAPPIA)25. F2H assay is relatively fast, allowing to observe protein interactions in their native cellular environment, but involves using expensive imaging equipment. All these methods have an advantage over prokaryotic expression providing the native eukaryotic translation and folding environment; however, they detect interaction indirectly, either by transcriptional activation or by fluorescent energy transfer, which often produces artifacts. Also, eukaryotic cells may contain other interaction partners of proteins of interest, which can interfere with the testing of binary interactions between proteins of higher eukaryotes.
The present study describes a method for the bacterial co-expression of differentially tagged proteins followed by conventional pulldown techniques. The method allows studying interactions between target proteins that require co-expression. It is more time-efficient compared to traditional pulldown, allowing batch testing of multiple targets, which makes it advantageous in most cases. Co-expression using compatible vectors is more convenient than polycistronic co-expression since it does not require a laborious cloning step.

<table><tbody><tr><td>8-ELEMENT probe</td><td>Sonics</td><td>630-0586</td><td>The high throughput 8-element sonicator probes</td></tr><tr><td>Agar</td><td>AppliChem</td><td>A0949</td><td></td></tr><tr><td>Amylose resin</td><td>New England Biolabs</td><td>E8021</td><td>Resin for purification of MBP-tagged proteins</td></tr><tr><td>Antibiotics</td><td>AppliChem</td><td>A4789 (kanamycin); A0839 (ampicillin)</td><td></td></tr><tr><td>Beta-mercaptoethanol</td><td>AppliChem</td><td>A1108</td><td></td></tr><tr><td>BL21(DE3)&amp;nbsp;</td><td>Novagen</td><td>69450-M</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>AppliChem</td><td>A4689</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5415R (Z605212)</td><td></td></tr><tr><td>Glutathione</td><td>AppliChem</td><td>A9782</td><td></td></tr><tr><td>Glutathione agarose</td><td>Pierce</td><td>16100</td><td>Resin for purification of GST-tagged proteins</td></tr><tr><td>Glycerol</td><td>AppliChem</td><td>A2926</td><td></td></tr><tr><td>HEPES&amp;nbsp;</td><td>AppliChem</td><td>A3724</td><td></td></tr><tr><td>HisPur Ni-NTA Superflow Agarose</td><td>Thermo Scientific</td><td>25214</td><td>Resin for purification of 6xHis-tagged proteins</td></tr><tr><td>Imidazole</td><td>AppliChem</td><td>A1378</td><td></td></tr><tr><td>IPTG</td><td>AppliChem</td><td>A4773</td><td></td></tr><tr><td>KCl</td><td>AppliChem</td><td>A2939</td><td></td></tr><tr><td>LB</td><td>AppliChem</td><td>414753</td><td></td></tr><tr><td>Maltose</td><td>AppliChem</td><td>A3891</td><td></td></tr><tr><td>MOPS</td><td>AppliChem</td><td>A2947</td><td></td></tr><tr><td>NaCl</td><td>AppliChem</td><td>A2942</td><td></td></tr><tr><td>NP40</td><td>Roche</td><td>11754599001</td><td></td></tr><tr><td>pACYCDuet-1</td><td>Sigma-Aldrich</td><td>71147</td><td>Vector for co-expression of proteins with p15A replication origin</td></tr><tr><td>pCDFDuet-1</td><td>Sigma-Aldrich</td><td>71340</td><td>Vector for co-expression of proteins with CloDF13 replication origin</td></tr><tr><td>PMSF</td><td>AppliChem</td><td>A0999</td><td></td></tr><tr><td>Protease Inhibitor Cocktail VII</td><td>Calbiochem</td><td>539138</td><td>Protease Inhibitor Cocktail</td></tr><tr><td>pRSFDuet-1</td><td>Sigma-Aldrich</td><td>71341</td><td>Vector for co-expression of proteins with RSF replication origin</td></tr><tr><td>SDS&amp;nbsp;</td><td>AppliChem</td><td>A2263</td><td></td></tr><tr><td>Tris&amp;nbsp;</td><td>AppliChem</td><td>A2264</td><td></td></tr><tr><td>VC505 sonicator</td><td>Sonics</td><td>CV334</td><td>Ultrasonic liquid processor</td></tr><tr><td>ZnCl&lt;sub&gt;2&lt;/sub&gt;</td><td>AppliChem</td><td>A6285</td><td></td></tr></tbody></table>

pulldown assay coupled with co-expression in bacteria cells as a time-efficient tool for testing challenging protein-protein interactions

Pulldown is an easy and widely used protein-protein interaction assay. However, it has limitations in studying protein complexes that do not assemble effectively in vitro. Such complexes may require co-translational assembly and the presence of molecular chaperones; either they form stable oligomers which cannot dissociate and re-associate in vitro or are unstable without a binding partner. To overcome these problems, it is possible to use a method based on the bacterial co-expression of differentially tagged proteins using a set of compatible vectors followed by the conventional pulldown techniques. The workflow is more time-efficient compared to traditional pulldown because it lacks the time-consuming steps of separate purification of interacting proteins and their following incubation. Another advantage is a higher reproducibility due to a significantly smaller number of steps and a shorter period of time in which proteins that exist within the in vitro environment are exposed to proteolysis and oxidation. The method was successfully applied for studying a number of protein-protein interactions when other in vitro techniques were found to be unsuitable. The method can be used for batch testing protein-protein interactions. Representative results are shown for studies of interactions between BTB domain and intrinsically disordered proteins, and of heterodimers of zinc-finger-associated domains.

The described method was used routinely with many different targets. Presented here are some representative results which likely cannot be obtained using conventional pulldown techniques. The first is the study of specific ZAD (Zinc-finger-associated domain) dimerization11. ZADs form stable and specific dimers, with heterodimers possible only between closely related domains within paralogous groups. The dimers formed by these domains are stable and do not dissociate for at least a few days; thus, mixing purified ZADs does not produce any detectable binding. At the same time, the co-expression of MBP and Thioredoxin-6xHis-fused ZADs shows good and reproducible homo-dimerization activity (Figure 2A), appearing as an additional band in the SDS-PAGE results of the MBP-pulldown assay. A small portion of heterodimers can be seen with M1BP co-expressed with another domain; this interaction was not confirmed with Y2A and most likely is a result of non-specific association due to high protein concentration, since these domains are cysteine-rich and extremely aggregation-prone. Notably, in this case, 6xHis-pulldown is inappropriate as ZADs are metal-coordinating domains that non-specifically bind to the metal-chelating resin. Such activity should be carefully examined in a parallel experiment.
Another example is the competition assay between the ENY2 protein and its binding partners Sgf11 (1-83aa) and the zinc-finger domain (460-631 aa) of the CTCF protein26. When expressed alone, the ENY2 protein forms dimers that prevent it from interacting with its native binding partners. Presumably, both the Sgf11 and CTCF proteins bind to the same molecular surface of ENY2, making their interaction mutually exclusive. In the co-expression assay, 6xHis-tagged ENY2 interacted both with GST-tagged Sgf11 and MBP-CTCF, but no GST-Sgf11 was present in MBP-pulldowns and vice versa, (Figure 2B). These results suggest that no triple complex can be formed, and interactions are mutually exclusive. These data were independently confirmed with other assays and support the different functional roles of ENY2 in these complexes. Attention should be brought to the fact that large affinity tags can impose steric hindrances by themselves, preventing complex formation; therefore, the conclusion should not be based solely on co-expression data.
A step-by-step comparison of the workflows of conventional and coupled co-expression pulldowns is shown in Figure 3A. Co-expression coupled pulldown is at least two times more time-efficient, even with a small number of samples, and allows good scalability. The results of utilizing both techniques to study the same interaction between the BTB domain of the CP190 protein (1-126aa) and the GST-tagged C-terminal domain (610-818aa) of Drosophila CTCF protein (dCTCF) are shown in Figure 3B. Both methods show good efficiency and reproducibility (assays were performed in three replicates); for this case, co-expression coupled pulldown showed lower non-specific binding, as can be seen in the control samples with GST protein alone.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64541/64541fig01.jpg" /> 
Figure 1: The schematic representation of the protocol. The schematic shows the method workflow employed in this study. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64541/64541fig01large.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/64541/64541fig02.jpg" /> 
Figure 2: Representative results. (A) Study of the Zinc-finger associated domain (ZAD) homodimerization in MBP- and 6xHis-pulldown assays. ZADs fused either with MBP (40 kDa) or with 6xHis-Thioredoxin (20 kDa) were co-expressed in bacteria cells and affinity purified with amylose resin (binds MBP-tagged proteins) or with Ni-NTA resin (binds 6xHis-tagged proteins). Co-purified proteins were visualized with SDS-PAGE followed by Coomassie staining. MBP-pulldown results are shown in upper panels, 6xHis-pulldown results are in lower panels (used only as protein expression control since many ZADs bind non-specifically to Ni-NTA). (B) Study of mutually exclusive interactions between ENY2 and Sgf11/CTCF proteins. GST-tagged Sgf11 (1-81aa), MBP-tagged CTCF zinc-finger domain (460-631aa), and 6xHis-tagged ENY2 proteins were co-expressed in various combinations and affinity purified with amylose resin, glutathione resin (binds GST-tagged proteins), or with Ni-NTA resin.Co-purified proteins were visualized with SDS-PAGE followed by Coomassie staining. The figure in panels A and B have been modified with permission from Bonchuk et al.11 and Bonchuk et al.26. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64541/64541fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64541/64541fig03.jpg" /> 
Figure 3: Comparison of workflows of conventional and coupled co-expression pulldown assays. (A) Step-by-step comparison of required time intervals in the workflow of the conventional pulldown assay compared to pulldown coupled to co-expression. (B) Comparison of two different pulldown techniques in studying the interaction between the dCTCF C-terminal domain (610-818aa) and the BTB domain of the CP190 protein (1-126aa) in GST-pulldown assays. dCTCF (610-818aa) fused with GST (25kDa) or GST alone were either co-expressed in bacteria cells or incubated in vitro with Thioredoxin-6xHis-tagged CP190 BTB and affinity purified with glutathione resin. Three independent replicates of each assay are shown. Co-purified proteins were visualized with SDS-PAGE followed by Coomassie staining. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64541/64541fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Pulldown Assay Coupled with Co-Expression in Bacteria Cells as a Time-Efficient Tool for Testing Challenging Protein-Protein Interactions at JoVE.com

Pulldown Assay Coupled with Co-Expression in Bacteria Cells as a Time-Efficient Tool for Testing Challenging Protein-Protein Interactions

Here, we describe a method for the bacterial co-expression of differentially tagged proteins using a set of compatible vectors, followed by the conventional pulldown techniques to study protein complexes that cannot assemble in vitro.

Pulldown is an easy and widely used protein-protein interaction assay. However, it has limitations in studying protein complexes that do not assemble ...

pulldown-assay-coupled-with-co-expression-bacteria-cells-as-time

Nanyang Technological University

Research

JoVE Journal

Biochemistry

2.8K Views.  Russian Academy of Sciences. Deze methode maakt het mogelijk om formaties van eiwitcomplexen te bestuderen die co-expressie vereisen voor efficiënte testmonsters, zoals de vorming van heterodimeren. Het belangrijkste eindpotentieel van deze methode is het benutten van de co-expressie van differentieel gelabelde bestudeerde eiwitten met behulp van combinaties van compatibele vectoren om efficiënte complexe assemblage te bevorderen, samen met een tijdefficiënte workflow en schaalbaarheid die snelle patch-testen van meer...

Video: Pulldown Assay Coupled with Co-Expression in Bacteria Cells as a Time-Efficient Tool for Testing Challenging Protein-Protein Interactions

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

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level. There are several methods, both in vitro and in vivo, to evaluate protein binding, and at least two methods that complement the shortcomings of each other should be conducted to obtain reliable insights. 
For an in vivo assay, the bimolecular fluorescence complementation (BiFC) assay represents the most popular and least invasive approach that enables to detect protein-protein interaction within living cells, as well as identify the intracellular localization of the interacting proteins 1,2. In this assay, non-fluorescent N- and C-terminal halves of GFP or its variants are fused to tested proteins, and when the two fusion proteins are brought together due to the tested proteins&#x2019; interactions, the fluorescent signal is reconstituted3-6. Because its signal is readily detectable by epifluorescence or confocal microscopy, BiFC has emerged as a powerful tool of choice among cell biologists for studying about protein-protein interactions in living cells 3. This assay, however, can sometimes produce false positive results. For example, the fluorescent signal can be reconstituted by two GFP fragments arranged as far as 7 nm from each other due to close packing in a small subcellular compartment, rather that due to specific interactions7.
Due to these limitations, the results obtained from live cell imaging technologies should be confirmed by an independent approach based on a different principle for detecting protein interactions. Co-immunoprecipitation (Co-IP) or glutathione transferase (GST) pull-down assays represent such alternative methods that are commonly used to analyze protein-protein interactions in vitro. However, iIn these assays, however, the tested proteins must be readily soluble in the buffer that supportsused for the binding reaction. Therefore, specific interactions involving an insoluble protein cannot be assessed by these techniques.
 Here, we illustrate the protocol for the protein membrane overlay binding assay, which circumvents this difficulty. In this technique, interaction between soluble and insoluble proteins can be reliably tested because one of the proteins is immobilized on a membrane matrix. This method, in combination with in vivo experiments, such as BiFC, provides a reliable approach to investigate and characterize interactions faithfully between soluble and insoluble proteins. In this article, binding between Tobacco mosaic virus (TMV) movement protein (MP), which exerts multiple functions during viral cell-to-cell transport8-14, and a recently identified plant cellular interactor, tobacco ankyrin repeat-containing protein (ANK) 15, is demonstrated using this technique.

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Testing protein-protein interaction is indispensable for dissection of protein functionality. Here, we introduce an in vitro protein-protein binding assay to probe a membrane-immobilized protein with a soluble protein. This assay provides a reliable method to test interaction between an insoluble protein and a protein in solution.

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level.  There are several ...

Biology

Measuring Transcellular Interactions through Protein Aggregation in a Heterologous Cell System

Protein interactions at cellular interfaces dictate a multitude of biological outcomes ranging from tissue development and cancer progression to synapse formation and maintenance. Many of these fundamental interactions occur in trans and are typically induced by heterophilic or homophilic interactions between cells expressing membrane anchored binding pairs. Elucidating how disease relevant mutations disrupt these fundamental protein interactions can provide insight into a myriad of cell biology fields. Many protein-protein interaction assays do not typically disambiguate between cis and trans interactions, which potentially leads to an overestimation of the extent of binding that is occurring in vivo and involve labor intensive purification of protein and/or specialized monitoring equipment. Here, we present an optimized simple protocol that allows for the observation and quantification of only trans interactions without the need for lengthy protein purifications or specialized equipment. The HEK cell aggregation assay involves the mixing of two independent populations of HEK cells, each expressing membrane-bound cognate ligands. After a short incubation period, samples are imaged and the resulting aggregates are quantified.

Here, we present an optimized protocol to rapidly and semiquantitatively measure ligand-receptor interactions in trans in a heterologous cell system using fluorescence microscopy.

Protein interactions at cellular interfaces dictate a multitude of biological outcomes ranging from tissue development and cancer progression to synapse ...

Neuroscience

Identification of Protein Interaction Partners in Mammalian Cells Using SILAC-immunoprecipitation Quantitative Proteomics

Quantitative proteomics combined with immuno-affinity purification, SILAC immunoprecipitation, represent a powerful means for the discovery of novel protein:protein interactions. By allowing the accurate relative quantification of protein abundance in both control and test samples, true interactions may be easily distinguished from experimental contaminants. Low affinity interactions can be preserved through the use of less-stringent buffer conditions and remain readily identifiable. This protocol discusses the labeling of tissue culture cells with stable isotope labeled amino acids, transfection and immunoprecipitation of an affinity tagged protein of interest, followed by the preparation for submission to a mass spectrometry facility. This protocol then discusses how to analyze and interpret the data returned from the mass spectrometer in order to identify cellular partners interacting with a protein of interest. As an example this technique is applied to identify proteins binding to the eukaryotic translation initiation factors: eIF4AI and eIF4AII.

SILAC immunoprecipitation experiments represent a powerful means for discovering novel protein:protein interactions. By allowing the accurate relative quantification of protein abundance in both control and test samples, true interactions may be easily distinguished from experimental contaminants, and low affinity interactions preserved through use of less-stringent buffer conditions.

Quantitative proteomics combined with immuno-affinity purification, SILAC immunoprecipitation, represent a powerful means for the discovery of novel ...

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

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

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

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

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

The Multifaceted Benefits of Protein Co-expression in Escherichia coli

We report here that the expression of protein complexes in vivo in Escherichia coli can be more convenient than traditional reconstitution experiments in vitro. In particular, we show that the poor solubility of Escherichia coli DNA polymerase III &#949; subunit (featuring 3&#8217;-5&#8217; exonuclease activity) is highly improved when the same protein is co-expressed with the &#945; and &#952; subunits (featuring DNA polymerase activity and stabilizing &#949;, respectively). We also show that protein co-expression in E. coli can be used to efficiently test the competence of subunits from different bacterial species to associate in a functional protein complex. We indeed show that the &#945; subunit of Deinococcus radiodurans DNA polymerase III can be co-expressed in vivo with the &#949; subunit of E. coli. In addition, we report on the use of protein co-expression to modulate mutation frequency in E. coli. By expressing the wild-type &#949; subunit under the control of the araBAD promoter (arabinose-inducible), and co-expressing the mutagenic D12A variant of the same protein, under the control of the lac promoter (inducible by isopropyl-thio-&#946;-D-galactopyranoside, IPTG), we were able to alter the E. coli mutation frequency using appropriate concentrations of the inducers arabinose and IPTG. Finally, we discuss recent advances and future challenges of protein co-expression in E. coli.

The Multifaceted Benefits of Protein Co-expression in Escherichia coli

Protein co-expression is a powerful alternative to the reconstitution in vitro of protein complexes, and is of help in performing biochemical and genetic tests in vivo. Here we report on the use of protein co-expression in Escherichia coli to obtain protein complexes, and to tune the mutation frequency of cells.

We report here that the expression of protein complexes in vivo in Escherichia coli can be more convenient than traditional reconstitution experiments in ...

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel protein-protein interaction pairs and monitoring protein interaction dynamics are of particular interest for revealing how plants respond to environmental factors and/or developmental signals. A plethora of approaches have been developed to examine protein-protein interactions, either in vitro or in vivo. Among them, the recently established luciferase complementation imaging (LCI) assay is the simplest and fastest method for demonstrating in vivo protein-protein interactions. In this assay, protein A or protein B is fused with the amino-terminal or carboxyl-terminal half of luciferase, respectively. When protein A interacts with protein B, the two halves of luciferase will be reconstituted to form a functional and active luciferase enzyme. Luciferase activity can be recorded with a luminometer or CCD-camera. Compared with other approaches, the LCI assay shows protein-protein interactions both qualitatively and quantitatively. Agrobacterium infiltration in Nicotiana benthamiana leaves is a widely used system for transient protein expression. With the combination of LCI and transient expression, these approaches show that the physical interaction between COP1 and SPA1 was gradually reduced after jasmonate treatment.

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

This manuscript describes an easy and rapid experimental procedure for determining protein-protein interactions based on the measurement of luciferase activity.

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel ...

Co-Immunoprecipitation and Pull-Down Assays

Co-immunoprecipitation (CoIP) and pull-down assays are closely related methods to identify stable protein-protein interactions. These methods are related to immunoprecipitation, a method for separating a target protein bound to an antibody from unbound proteins. In CoIP, an antibody-bound protein is itself bound to another protein that does not bind with the antibody, this is followed by a separation process that preserves the protein-protein complex. The difference in pull-down assays is that affinity-tagged bait proteins replace antibodies, and affinity chromatography is used to isolate protein-protein complexes.
This video explains CoIP, pull-down assays, and their implementation in the laboratory. A step-by-step protocol for each technique is covered, including the reagents, apparatus, and instruments used to purify and analyze bound proteins. Additionally, the applications section of this video describes a procedure to study how myxovirus proteins inhibit influenza nucleoprotein, an investigation into the role of calcium ions in calmodulin via a pull-down assay, and a modified pull-down assay for characterizing transient protein interactions.
<div class="chapter_text" id="script_0">
Protein-protein interactions play a significant role in a wide variety of biological functions. The majority of protein-protein interactions and their biological effects have yet to be identified. Co-immunoprecipitation, or CoIP, and pull-down assays are two closely related methods for the identification of stable protein-protein interactions. This video will cover the principles of the two assays, their general laboratory procedures, and applications of these techniques.
</div>
<div class="chapter_text" id="script_1">
CoIP and pull-down assays are variants of immunoprecipitation, a method for selectively isolating a protein species from a complex solution. In an immunoprecipitation experiment, an antibody specific to a target protein is allowed to form an immune complex with that target in the sample. The complex is then captured on a solid support, typically protein A bound to a sepharose bead. Any proteins not captured are removed by centrifugation steps. The protein is then released from the antibody and solid support by boiling in reducing SDS-PAGE sample-loading buffer.
Co-immunoprecipitation is conducted in the same manner, except that intact protein complexes are captured onto the solid support. The antibody binds to the target protein, which, in turn, is bound to another protein that is not targeted by the antibody. As in immunoprecipitation, the protein complex is released from the antibody and solid support by boiling in reducing SDS-PAGE sample loading buffer.
Pull-down assays are similar to co-immunoprecipitation, differing only in the use of a &#34;bait&#34; protein, as opposed to an antibody. Through molecular biology techniques, this bait protein is engineered with an affinity tag, such as a series of histidine residues. These affinity tags are described in &#34;Chromatography-based Biomolecule Separations.&#34; The protein is then captured on an immobilized affinity ligand specific for the tag. The captured protein is then incubated with a sample that contains proteins that form complexes with the &#34;bait&#34;. The protein complex is released from the affinity support by washing with a solution containing a competitive analyte specific for the tag on the &#34;bait&#34; protein. Pull-down assays are useful for confirming protein interactions predicted by co-immunoprecipitation, and for discovering unknown protein interactions.
Now that the principles of co-immunoprecipitation and pull-down assays have been discussed, let&#39;s look at their laboratory procedures.
</div>
<div class="chapter_text" id="script_2">
First let&#39;s discuss co-immunoprecipitation. To a series of microfuge tubes, the following is added: PBS buffer, and a 50% solution of protein A-sepharose, a resin-protein complex that binds to the antibody. The microfuge tubes are rotated to ensure proper distribution, and then the resin is washed with additional PBS buffer. Cell lysate, containing the desired protein, and 2 &#956;g of antibody are added to the microfuge tubes, and the mixture is rotated for 1 h at 4 &#176;C. The beads are pelleted by centrifugation, the supernatant is discarded, and the beads re-washed three times with buffer to remove non-bound protein. The beads containing the antibody-protein complex are in reducing SDS-PAGE sample-loading buffer, for removal of the complex from the antibody and for analysis by SDS-PAGE and immunoblotting.
</div>
<div class="chapter_text" id="script_3">
Now we will discuss the procedure for pull-down assays: The &#34;bait&#34; protein is expressed in a plasmid with the appropriate affinity tag. After reaching log-phase growth, the cells are lysed, and then centrifuged. Suspended streptavidin-sepharose beads, which capture biotin-tagged &#34;bait&#34; protein, are pipetted into a microfuge tube. The beads are then centrifuged and the supernatant carefully removed by aspiration. The beads are then washed with buffer, centrifuged, and the supernatant removed.
Cells containing the putative &#34;prey&#34; protein, which has an affinity for the &#34;bait&#34; protein, are harvested by centrifugation. The supernatant is then added to the microfuge tube containing the resin, and incubated at 4 &#176;C for 3 h on a shaker. The resin is then centrifuged, the supernatant removed, and the resin washed to remove non-bound protein. Elution buffer is added to the resin, and the mixture is incubated at room temperature for 30 min on a shaker. The resin is then centrifuged, and the supernatant containing the desired complex is analyzed by immunoblotting.
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Now that we&#39;ve reviewed the procedures, let&#39;s look at some of the useful applications of co-immunoprecipitation and pull-down assays.
Co-immunoprecipitation can be useful in better understanding of enzymes&#39; mechanism of action. Myxovirus-, or Mx-, resistance proteins inhibit a wide range of viruses, including influenza A, for which the mechanism is poorly understood. Co-immunoprecipitation was used to study the interaction between mouse Mx1 protein and influenza nucleoprotein.
Pull-down assays have proven useful in studying the effects of second messengers, which are proteins that communicate a signal from the cellular environment. They are a component of a signaling pathway where multiple proteins interact in response to environmental cues. Calcium ions act as secondary messengers by binding to calmodulin, which, in turn, binds to a wide variety of proteins, mediating many types of biological responses. Without the calcium, the protein can&#39;t bind to the calmodulin. A pull-down assay was conducted to test the ability of proteins to bind to calmodulin in the presence or absence of calcium ions.
Co-immunoprecipitation and pull-down assays are generally used for analyzing stable or strong protein interactions, but not transient ones. A recent development in pull-down assays, the HaloTag, has simplified the study of transient protein interactions; HaloTag is a genetically-encoded protein fusion tag, fused to the protein of interest, capable of chemically reacting with a haloalkane solid support. If functional analysis is desired, the protein complex, minus the tag, in its entirety could then be isolated by incubating with tobacco etch virus protease.
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You&#39;ve just watched JoVE&#39;s video on co-immunoprecipitation and pull-down assays. This video described the principles of the two methods, the general laboratory procedures, and some of their applications.
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Co-immunoprecipitation (CoIP) and pull-down assays are closely related methods to identify stable protein-protein interactions. These methods are related ...

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Tandem Affinity Purification Assay to Study Protein-Protein Interactions

Source: Ma, Z., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/55236">Tandem Affinity Purification of Protein Complexes from Eukaryotic Cells.</a> J. Vis. Exp. (2017).
This video demonstrates tandem affinity purification &#8212; a technique to purify protein complexes from eukaryotic cells to study protein-protein interaction. The complex contains two proteins labeled with different epitope tags. Upon the purification of the complex using two resins with an affinity for the two different tags in a sequential manner, the presence of both proteins in the elute confirms the interaction between the two.

Source: Ma, Z., et al. Tandem Affinity Purification of Protein Complexes from Eukaryotic Cells. J. Vis. Exp. (2017).
This video demonstrates tandem ...

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Biomolecular Interaction Detection Techniques

Protein-protein interactions

Bimolecular Complementation Affinity Purification to Isolate Two Interacting Proteins

Source: Hastings, J. F. et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/57109">Dissecting Multi-protein Signaling Complexes by Bimolecular Complementation Affinity Purification (BiCAP).</a> J. Vis. Exp. (2018)
In this video, we have demonstrated bimolecular complementation affinity purification, BiCAP, to detect and isolate specific protein dimers using conformation-specific single-domain antibodies.

Source: Hastings, J. F. et al. Dissecting Multi-protein Signaling Complexes by Bimolecular Complementation Affinity Purification (BiCAP). J. Vis. Exp. ...

Pulldown Assay Coupled with Co-Expression in Bacteria Cells as a Time-Efficient Tool for Testing Challenging Protein-Protein Interactions

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