The authors thank diverse Twitter users who raised questions about the protocol presented in the paper describing the VNp technology. Figure 1A was generated using icons from flaticon.com. This work was supported by the University of Kent and funding from the Biotechnology and Biological Sciences Research Council (BB/T008/768/1 and BB/S005544/1).

The bacterial work undertaken follows the local, national, and international biosafety containment regulations befitting the particular biosafety hazard level of each strain.
1. Selection of different VNps
<ol>
	<li>Identify the VNp sequences. 
	NOTE: For the present study, three VNp sequences have been identified1 that result in maximal yield and vesicular export of the proteins examined to date: VNp2, VNp6, and VNp15. It is not currently clear why certain VNp variants perform more efficiently with some proteins than others; therefore, it is recommended that fusions are generated between a new protein of interest with each VNp variant (i.e., VNp2, 6, or 15). 
	VNp2:&#160; &#160;MDVFMKGLSKAKEGVVAAAEKTKQGVAEA 
	AGKTKEGVL 
	VNp6:&#160; &#160;MDVFKKGFSIADEGVVGAVEKTDQGVTEA 
	AEKTKEGVM 
	VNp15:&#160; &#160;MDVFKKGFSIADEGVVGAVE 
	Plasmids that allow expression of the protein of interest with different VNp amino terminal fusions have been made available commercially (see Table of Materials).</li>
	<li>Design a cloning strategy to insert the gene of interest at the 3&#39; end of the cDNA encoding for the VNp in one of these constructs, or adapt an existing plasmid by integrating synthesized VNp cDNA upstream of the first ATG codon of the gene encoding for the protein of interest. Use the methods as described in1.</li>
	<li>For toxic proteins, use a vector with a repressible expression promoter or a promotor with minimal uninduced expression noise.</li>
	<li>Clone the VNp sequence tag at the amino-terminal of the fusion protein. Ensure that the affinity tags, protease cleavage sequences, etc., and the protein of interest are located on the carboxyl side of the VNp tag. It is recommended to separate the VNp from the downstream peptide with a flexible linker region, such as two or three repeats of a -G-G-S-G- polypeptide sequence (Figure 1). 
	&#8203;NOTE: Use plasmids with an antibiotic selection that does not target peptidoglycan, which weakens the cell surface and reduces vesicle yield. Kanamycin and chloramphenicol (see Table of Materials) were the preferred antibiotics used for this study.</li>
</ol>
2. Bacterial cell culture and protein induction
NOTE: Bacterial strains typically used in this protocol are either Escherichia coli BL21 (DE3) or W3110. E. coli cells are cultured in lysogeny broth (LB) (10 g/L tryptone; 10 g/L NaCl; 5 g/L yeast extract) or terrific broth (TB) (12 g/L tryptone; 24 g/L yeast extract; 4 mL/L 10% glycerol; 17 mM KH2PO4; 72 mM K2HPO4, salts autoclaved separately) media (see Table of Materials). Example images showing each step of the protein induction and subsequent isolation and purification process are shown in Figure 2.
<ol>
	<li>Culture 5 mL LB starters from fresh bacterial transformations at 37 &#176;C to saturation and use them to inoculate 25 mL of TB in a 500 mL conical flask, all with appropriate antibiotic selection.</li>
	<li>The surface area:volume ratio is an important factor in optimizing this system. Use as large a volume flask as possible (e.g., 5 L flask containing 1 L of culture; for optimization runs, use 25 mL of media in a 500 mL flask).</li>
	<li>Incubate the larger shaking flask cultures in an incubator at 37 &#176;C with shaking at 200 rpm (&#8805;25 mm orbital throw) until a 600 nm optical density value (OD600) of 0.8-1.0 is reached. 
	NOTE: Vesiculation is optimal when cells are grown at 37 &#176;C. However, some recombinant proteins require expression in lower temperatures. If this is the case for the protein of interest, the VNp6 tag must be used, as this allows high-yield vesicle export at temperatures down to 25 &#176;C.</li>
	<li>To induce recombinant protein expression from the T7 promoter, add isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) to a final concentration of up to 20 &#181;g/mL (84 &#181;M) (see Table of Materials). The induction of recombinant protein expression must occur at the late-log phase (i.e., typical OD600 of 0.8-1.0) for the production of vesicles. 
	&#8203;NOTE: The length of the induction period may differ between proteins, with some reaching maximum production at 4 h and others overnight (18 h). To date, maximum vesicle export has been obtained in overnight cultures.</li>
</ol>
3. Recombinant vesicle isolation
<ol>
	<li>Pellet the cells by centrifugation at 3,000 x g (4 &#176;C) for 20 min.</li>
	<li>To sterilize vesicle-containing media for long-term storage, pass the cleared culture media through a sterile and detergent-free 0.45 &#181;m polyethersulfone (PES) filter (see Table of Materials). 
	NOTE: To test the exclusion of viable cells from the vesicle-containing filtrate, plate onto LB agar and incubate overnight at 37 &#176;C.</li>
	<li>To concentrate vesicles into a smaller volume, pass the sterile vesicle-containing media through a sterile and detergent-free 0.1 &#181;m mixed cellulose esters (MCE) filter (see Table of Materials).</li>
	<li>Gently wash the membrane with 0.5-1 mL of sterile PBS using a cell scraper or plastic spreader to carefully remove vesicles from the membrane. Transfer to a fresh microfuge tube. 
	&#8203;NOTE: Purified vesicles can be stored in sterile media or phosphate-buffered saline (PBS) at 4 &#176;C. There are examples of recombinant proteins stored in these vesicles for 6 months, in this way, with no loss in enzymatic activity.</li>
</ol>
4. Soluble protein release from isolated vesicles
<ol>
	<li>Once protein-containing vesicles have been isolated into the sterile media/buffer, subject vesicular lipid membranes to sonication using an appropriate schedule for the apparatus (e.g., 6x 20 s on and off cycles) and centrifuge at 39,000 x g (4 &#176;C) for 20 min to remove vesicle debris. 
	NOTE: Osmotic shock or detergent treatment can be used as an alternative to break open the vesicles, but consideration must be given to the impact upon protein functionality and/or downstream application.</li>
	<li>If VNp fusion remains cytosolic and does not release into the media, isolate the protein using standard protocols (e.g., resuspend the cell pellets in 5 mL of an appropriate extraction buffer (20 mM tris, 500 mM NaCl), sonicate, and remove the cell debris by centrifugation).</li>
</ol>
5. Protein concentration determination
<ol>
	<li>Determine the concentration of proteins by gel densitometry analysis of triplicate samples1. Run alongside bovine serum albumin (BSA) loading standards on coomassie-stained sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gels. Scan and analyze the gels using appropriate software (e.g., Image J; see Table of Materials).</li>
</ol>
6. Visualization of vesicle formation and isolated vesicles by fluorescence microscopy
NOTE: If the cells contain fluorescently labeled VNp fusion or membrane markers, live cell imaging can be used to follow vesicle formation. Alternatively, fluorescent lipid dyes can be used to visualize vesicles to confirm production and purification.
<ol>
	<li>Cell mounting
	<ol>
		<li>Induce VNp fusion expression for several hours before mounting onto the coverslip.</li>
		<li>Agarose pad method (Figure 3A-C): pipette the cells onto a thin (&#60;1 mm), circular LB-agarose (2%) pad that has been allowed to form and set on a clean glass slide. Allow the cells to settle and equilibrate and place a 50 mm x 25 mm coverslip onto the pad and cells. Hold the coverslip in place with spacers and adhesive tape.</li>
		<li>Polyethyleneimine (PEI) method (Figure 3D-F): spread 20 &#181;L of 0.05% PEI (in dH2O) onto a coverslip with a pipette tip and leave for 3-5 min to bind to glass without allowing to dry. Add 50 &#181;L of cell culture and leave for 5-10 min to ensure bacteria have associated with the PEI-coated surface4. Wash the coverslip with 100 &#181;L of media before placing it onto the slide and hold it in place with spacers and adhesive tape.</li>
	</ol>
	</li>
	<li>Mounting vesicles
	<ol>
		<li>Pipette purified vesicles onto a thin (&#60;1 mm), circular LB-agarose (2%) pad that has been allowed to form and set on a clean glass slide. Once the liquid has dried, place a 50 mm x 25 mm coverslip onto the pad and vesicles. Hold the coverslip in place with spacers and adhesive tape.</li>
		<li>The fluorescent lipid dye FM4-64 is able to stain membranes5, and therefore can be used to visualize vesicles. Add FM4-64 (see Table of Materials) to purified vesicles at a final concentration of 2 &#181;M (from a 2 mM stock dissolved in dimethyl sulfoxide [DMSO]) and image after a 10 min incubation. This is especially useful for identifying vesicles containing non-fluorescently labeled cargoes5.</li>
		<li>Rinse the coverslips with the same media used to culture the observed cells. 
		NOTE: Some complex media (e.g., TB) can exhibit autofluorescence, which may result in excess background signal.</li>
	</ol>
	</li>
	<li>Imaging vesicles 
	&#8203;NOTE: Example microscopy images of VNp recombinant vesicles can be seen in Figure 4.
	<ol>
		<li>Mount the slide onto an inverted microscope (see Table of Materials) using an oil immersion objective and leave for 2-3 min to allow the sample to settle and the temperature to equilibrate. 
		NOTE: All live cell imaging for each sample must be completed within 30 min of mounting cells onto coverslips to minimize the impact of phototoxicity and anaerobic stress. For this reason, single-plane images are preferred to z-stacks.</li>
		<li>Use appropriate light sources (e.g., light emitting diode [LED] or halogen bulb; see Table of Materials) and filter combinations for the fluorescent protein(s)/dye(s) being used6.</li>
		<li>Use a high magnification (i.e., 100x or 150x) and high numerical aperture (i.e., NA &#8805;1.4) lens for imaging the microbial cells and vesicles.</li>
		<li>Determine the minimal light intensity required to visualize fluorescence signals from cells and / or vesicles. This may require some adjustment of the exposure and gain settings for the camera in use. 
		NOTE: Typical exposure times from current complementary metal-oxide semiconductor (CMOS) cameras vary between 50-200 ms depending upon the imaging system.</li>
		<li>For single frame images, use three-image averaging to reduce hardware-dependent random background noise.</li>
		<li>For time-lapse imaging, allow 3-5 min between individual frames. 
		NOTE: Depending upon the microscope setup, the focus may need to be intermittently adjusted throughout longer time-lapse experiments.</li>
	</ol>
	</li>
</ol>

Production and Imaging of Recombinant Protein-Filled Vesicles from E. coli

<ol>
	<li>Eastwood, T. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-yield+vesicle-packaged+recombinant+protein+production+from+E.+coli.">High-yield vesicle-packaged recombinant protein production from E. coli.</a> Cell Reports Methods. 3 (2), 100396 (2023).</li><li>Makino, T., Skretas, G., Georgiou, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strain+engineering+for+improved+expression+of+recombinant+proteins+in+bacteria.">Strain engineering for improved expression of recombinant proteins in bacteria.</a> Microbial Cell Factories. 10, 32 (2011).</li><li>Peng, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Factors+influencing+recombinant+protein+secretion+efficiency+in+gram-positive+bacteria:+Signal+peptide+and+beyond.">Factors influencing recombinant protein secretion efficiency in gram-positive bacteria: Signal peptide and beyond.</a> Frontiers in Bioengineering and Biotechnology. 7, 139 (2019).</li><li>Lewis, K., Klibanov, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surpassing+nature:+rational+design+of+sterile-surface+materials.">Surpassing nature: rational design of sterile-surface materials.</a> Trends in Biotechnology. 23 (7), 343-348 (2005).</li><li>Betz, W. J., Mao, F., Smith, C. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+exocytosis+and+endocytosis.">Imaging exocytosis and endocytosis.</a> Current Opinion in Neurobiology. 6 (3), 365-371 (1996).</li><li>Mulvihill, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+cell+imaging+in+fission+yeast.">Live cell imaging in fission yeast.</a> Cold Spring Harbor Protocols. 2017 (10), (2017).</li><li>Shaner, N. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+bright+monomeric+green+fluorescent+protein+derived+from+Branchiostoma+lanceolatum.">A bright monomeric green fluorescent protein derived from Branchiostoma lanceolatum.</a> Nature Methods. 10 (5), 407-409 (2013).</li><li>Shen, Y., Chen, Y., Wu, J., Shaner, N. C., Campbell, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+of+mCherry+variants+with+long+Stokes+shift,+red-shifted+fluorescence,+and+low+cytotoxicity.">Engineering of mCherry variants with long Stokes shift, red-shifted fluorescence, and low cytotoxicity.</a> PloS One. 12 (2), e0171257 (2017).</li><li>Periz, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+highly+dynamic+F-actin+network+regulates+transport+and+recycling+of+micronemes+in+Toxoplasma+gondii+vacuoles.">A highly dynamic F-actin network regulates transport and recycling of micronemes in Toxoplasma gondii vacuoles.</a> Nature Communications. 10 (1), 4183 (2019).</li><li>Qiu, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Uniform+patchy+and+hollow+rectangular+platelet+micelles+from+crystallizable+polymer+blends.">Uniform patchy and hollow rectangular platelet micelles from crystallizable polymer blends.</a> Science. 352 (6286), 697-701 (2016).</li></ol>

The authors declare no competing financial interests or other conflicts of interest.

The amino-terminal peptide-tagged method for the production of recombinant proteins described above is a simple process, which consistently yields large amounts of protein that can be efficiently isolated and/or stored for months.
It is important to highlight the key steps in the protocol that are required for optimal use of this system. Firstly, the VNp tag1 must be located at the N-terminus, followed by the protein of interest and any appropriate tags. It is also important to avoid using antibiotics that target the peptidoglycan layer, such as ampicillin.
In terms of growth conditions, rich media (e.g., LB or TB media) and a high surface area:volume ratio is necessary to maximize vesicle production. The optimal temperature for the production of extracellular vesicles is 37 &#176;C, but the conditions typically required for expression of the protein of interest must be considered too. For lower induction temperatures, VNp6 must be used. Crucially, induction of the T7 promoter must be achieved using no greater than 20 &#181;g/mL (84 &#181;M) IPTG once the cells reach an OD600 of 0.8-1.0. Proteins expressed using the system reach maximum vesicle production at either 4 h or after overnight induction.
Despite the simplicity of this protocol, it requires optimization. VNp variant fusion, expression temperatures, and induction time periods may differ depending on the protein of interest. Furthermore, there is a need to optimize the purification and subsequent concentration of extracellular vesicles from the media. The current procedure is not scalable and can be time-consuming. These are the limitations of this methodology.
The VNp technology has many advantages over traditional methods2. It allows the vesicular export of diverse proteins, with the maximum size successfully expressed to date being 175 kDa for vesicles that remain internal and 85 kDa for those that are exported. Furthermore, this technology can significantly increase the yield of recombinant proteins with a range of physical properties and activities. Exported vesicles containing the protein of interest can be isolated by simple filtration from the precleared media and can subsequently be stored, in sterile culture media or buffer, at 4 &#176;C for several months.
The applications for this system are diverse, from discovery science to applied biotechnology and medicine (e.g., through the production of functional therapeutics)3. The ease of production, downstream processing, and high yield are all&#160;attractive qualities in these areas and especially in industry.

The Gram-negative bacteria E. coli is an attractive system for recombinant protein production on both industrial and academic scale. It is not only cost-effective and straightforward to culture in batches to high densities, but a broad-spectrum of reagents, strains, tools, and promoters have been established to promote the generation of functional proteins in E. coli1. Additionally, synthetic biology techniques are now overcoming obstacles typically related to the application of post-translational modifications and folding of complex proteins2. The ability to target the secretion of recombinant proteins into culture media is attractive for improving yield and reducing manufacturing costs. Controlled packaging of user-defined proteins into membrane vesicles assists the development of products and technologies within the applied biotechnology and medical industries. Until now, there has been a lack of widely applicable methods for secreting recombinant proteins from E. coli 3.
Eastwood et al.&#160;have recently developed a peptide tagging-based method for producing and isolating recombinant protein-containing vesicles from E. coli1. This Vesicle Nucleating peptide (VNp) allows the production of extracellular bacterial membrane vesicles, into which the recombinant protein of choice can be targeted to simplify purification and storage of the target protein, and allows significantly higher yields than normally allowed from shaking flask cultures. Yields of close to 3 g of recombinant protein per liter of flask culture have been reported, with &#62;100x higher yields than those obtained with equivalent proteins lacking the VNp tag. These recombinant protein-enriched vesicles can be rapidly purified and concentrated from the culture media and provide a stable environment for storage. This technology represents a major breakthrough in E. coli recombinant protein production. The vesicles compartmentalize toxic and disulfide-bond containing proteins in a soluble and functional form, and support the simple, efficient, and rapid purification of vesicle-packaged, functional proteins for long-term storage or direct processing1.
The major advantages this technology presents over current techniques are: (1) the applicability to a range of sizes (1 kDa to &#62;100 kDa) and types of protein; (2) facilitating inter- and intra- protein disulfide-bond formation; (3) applicable&#160;to multiprotein complexes; (4) can be used with a range of promoters and standard lab E. coli strains; (5) the generation of yields of proteins from shaking flasks normally only seen with fermentation cultures; proteins are exported and packaged into membrane bound vesicles that (6) provide a stable environment for storage of the active soluble protein; and (7) simplifies downstream processing and protein purification. This simple and cost-effective recombinant protein tool is likely to have a positive impact on the biotechnology and medical industries, as well as discovery science.
Here, a detailed protocol, developed over several years, describes the optimal conditions to produce recombinant protein-filled vesicles from bacteria with the VNp technology. Example images of this system in practice are shown, with a fluorescent protein being expressed, allowing the presence of vesicles during different stages of the production, purification, and concentration to be visualized. Finally, guidance is provided on how to use live cell imaging to validate the production of VNp fusion-containing vesicles from the bacteria.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Ampicillin</td>
			<td>Melford</td>
			<td>69-52-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Chloramphenicol</td>
			<td>Acros Organics (Thermofisher Scientific)</td>
			<td>56-75-7</td>
			<td></td>
		</tr>
		<tr>
			<td>E. coli BL21 (DE3)</td>
			<td>Lab Stock</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>E. coli DH10&#946;</td>
			<td>Lab Stock</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Filters for microscope</td>
			<td>Chroma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>FM4-64</td>
			<td>Molecular Probes (Invitrogen)</td>
			<td>T-3166</td>
			<td>Dissolved in DMSO, stock concentration 2 mM</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>Open Source</td>
			<td></td>
			<td>Downloaded from: https://imagej.net/ij/index.html</td>
		</tr>
		<tr>
			<td>Inverted microscope</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropyl &#946;-D-1-thiogalactopyranoside (IPTG)</td>
			<td>Melford</td>
			<td>367-93-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Kanamycin sulphate</td>
			<td>Gibco (Thermofisher Scientific)</td>
			<td>11815-024</td>
			<td></td>
		</tr>
		<tr>
			<td>LED light source for micrscope</td>
			<td>Cairn Research Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Lysogeny Broth (LB) / LB agar</td>
			<td>Lab Stock</td>
			<td>N/A</td>
			<td>10 g/L Tryptone; 10 g/L NaCl; 5 g/L Yeast Extract (1.5 g/L agar)</td>
		</tr>
		<tr>
			<td>Metamorph imaging software</td>
			<td>Molecular Devices</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MF-Millipore Membrane filter (0.1 &#181;m, MCE)</td>
			<td>Merck</td>
			<td>VCWP04700</td>
			<td></td>
		</tr>
		<tr>
			<td>Millipore Express PLUS membrane filter (0.45 &#181;m, PES)</td>
			<td>Merck</td>
			<td>HPWP04700</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS)</td>
			<td>Lab Stock</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasmids allowing expression of protein of interest with different VNp amino terminal fusions&#160;</td>
			<td>Addgene</td>
			<td></td>
			<td>https://www.addgene.org/Dan_Mulvihill/</td>
		</tr>
		<tr>
			<td>Terrific Broth (TB)</td>
			<td>Lab Stock</td>
			<td>N/A</td>
			<td>12 g/L Tryptone; 24 g/L Yeast Extract; 4 ml/L 10% glycerol; 17 mM KH2PO4 72 mM K2HPO4</td>
		</tr>
	</tbody>
</table>

optimized production and analysis of recombinant protein-filled vesicles from e. coli

This innovative system, using a short peptide tag, that exports multiple recombinant proteins in membrane bound vesicles from E. coli, provides an effective solution to a range of problems associated with bacterial recombinant protein expression. These recombinant vesicles compartmentalise proteins within a micro-environment that facilitates the production of otherwise challenging, toxic, insoluble, or disulfide-bond containing proteins from bacteria. Protein yield is increased considerably when compared to typical bacterial expression in the absence of the vesicle-nucleating peptide tag. The release of vesicle-packaged proteins supports isolation from the culture medium&#160;and permits long-term active protein storage. This technology gives rise to increased yields of vesicle-packaged, functional proteins for simplified downstream processing for a diverse range of applications from applied biotechnology to discovery science and medicine. In the present article and the associated video, a detailed protocol of the method is provided, which highlights key steps in the methodology to maximize recombinant protein-filled vesicle production.

BL21 DE3 E. coli containing the VNp6-mNeongreen expression construct were grown to&#160;late-log phase (Figure 2A). VNp6-mNeongreen expression was induced by the addition of IPTG to the culture (20 &#181;g/mL or 84 &#181;M final concentration), which was subsequently left to grow overnight at 37 &#176;C with vigorous shaking (200 rpm, &#8805;25 mm orbital throw). The following morning, the culture displayed mNeongreen fluorescence7 (Figure 2B), which remained visible in the media after the removal of bacterial cells by centrifugation (Figure 2C). The presence of VNp-mNeongreen within the culture and cleared culture media was confirmed by SDS-PAGE (Figure 2D). The mNeongreen-containing vesicles were isolated onto a 0.1 &#181;m MCE filter (Figure 2E) and resuspended in PBS (Figure 2F). The purified vesicles were subsequently mounted on an agarose pad (Figure 3A-C) and imaged using widefield fluorescence microscopy (Figure 4A). The presence of vesicle membranes was confirmed using the lipophilic fluorescent dye FM4-64 (Figure 4B). The E. coli cells expressing the inner membrane protein CydB fused to mNeongreen (green) and VNp6-mCherry2 (magenta)8 show vesicle production and cargo insertion in live bacterial cells (Figure 4C). Figure 4A,B were captured using a widefield fluorescence microscope, while Figure 4C was acquired using structured illumination microscopy (SIM), using methods described previously9,10.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65442/65442fig01.jpg" /> 
Figure 1: Summary of VNp technology from designing a cloning strategy to the purification and storage of extracellular vesicles. (A) Schematic of a typical VNp fusion protein. VNp at the NH2 terminus, followed by a flexible linker and an appropriate combination of affinity and fluorescence tags (Tag1, Tag 2, protease cleavage site [e.g., TEV]) and protein of interest. (B) Schematic diagram summarising the protocol for the expression and purification of recombinant protein-filled membrane vesicles from E. coli. <a href="https://www.jove.com/files/ftp_upload/65442/65442fig01large.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/65442/65442fig02.jpg" /> 
Figure 2: Stages of production and purification of VNp6-mNg vesicles. Cultures of E. coli cells containing the VNp-mNeongreen expression construct in blue light either before (A) or after (B) IPTG-induced expression of the fusion protein. The cells from (B) were removed by centrifugation, leaving VNp-mNeongreen-filled vesicles in the media (C). (D) Equivalent samples from A, B, and C were analyzed by SDS-PAGE and coomassie staining. The vesicles were isolated onto a 0.1 &#181;m filter (E) and subsequently washed off into an appropriate volume of buffer (F). <a href="https://www.jove.com/files/ftp_upload/65442/65442fig02large.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/65442/65442fig03.jpg" /> 
Figure 3: Cell mounting procedure for imaging vesicles and vesicle production. (A-C) The agarose pad method and (D-F) the PEI method for mounting E. coli cells onto the coverslip. <a href="https://www.jove.com/files/ftp_upload/65442/65442fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65442/65442fig04.jpg" /> 
Figure 4: Microscopy images of VNp recombinant vesicles. Green (A) and red (B) emission images from different fields of FM4-64 labeled VNp6-mNeongreen-containing vesicles mounted on an agarose pad. (C) Imaging E. coli cells expressing the inner membrane protein CydB fused to mNeongreen (green) and VNp6-mCherry2 (magenta) shows vesicle production and cargo insertion in live bacterial cells. (A,B) were imaged using a widefield fluorescence microscope, while (C) was acquired using structured illumination microscopy (SIM). Scale bars: (A,B) = 10 &#181;m; (C) = 1 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65442/65442fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about The Production of Recombinant Proteins at JoVE.com

Author Spotlight: The Production of Recombinant Proteins 

The present protocol describes a detailed method for the bacterial production of recombinant proteins, including typically insoluble or disulfide-bond containing proteins, packaged inside extracellular membrane-bound vesicles. This has the potential to be applied to versatile areas of scientific research, including applied biotechnology and medicine.

Optimized Production and Analysis of Recombinant Protein-Filled Vesicles from E. coli

This innovative system, using a short peptide tag, that exports multiple recombinant proteins in membrane bound vesicles from E. coli, provides an ...

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Research

JoVE Journal

Biology

6.6K Views.  University of Kent. The present protocol describes a detailed method for the bacterial production of recombinant proteins, including typically insoluble or disulfide-bond containing proteins, packaged inside extracellular membrane-bound vesicles. This has the potential to be applied to versatile areas of scientific research, including applied biotechnology and medicine.

Video: Author Spotlight: The Production of Recombinant Proteins 

Transformation of Plasmid DNA into E. coli Using the Heat Shock Method

Transformation of plasmid DNA into E. coli using the heat shock method is a basic technique of molecular biology. It consists of inserting a foreign plasmid or ligation product into bacteria. This video protocol describes the traditional method of transformation using commercially available chemically competent bacteria from Genlantis. After a short incubation in ice, a mixture of chemically competent bacteria and DNA is placed at 42&deg;C for 45 seconds (heat shock) and then placed back in ice. SOC media is added and the transformed cells are incubated at 37&deg;C for 30 min with agitation. To be assured of isolating colonies irrespective of transformation efficiency, two quantities of transformed bacteria are plated. This traditional protocol can be used successfully to transform most commercially available competent bacteria. The turbocells from Genlantis can also be used in a novel 3-minute transformation protocol, described in the instruction manual.

Transformation of plasmid DNA into E. coli using the heat shock method is a basic technique of molecular biology. It consists of inserting a foreign ...

Visualization of UV-induced Replication Intermediates in E. coli using Two-dimensional Agarose-gel Analysis

Inaccurate replication in the presence of DNA damage is responsible for the majority of cellular rearrangements and mutagenesis observed in all cell types and is widely believed to be directly associated with the development of cancer in humans. DNA damage, such as that induced by UV irradiation, severely impairs the ability of replication to duplicate the genomic template accurately. A number of gene products have been identified that are required when replication encounters DNA lesions in the template. However, a remaining challenge has been to determine how these proteins process lesions during replication in vivo. Using Escherichia coli as a model system, we describe a procedure in which two-dimensional agarose-gel analysis can be used to identify the structural intermediates that arise on replicating plasmids in vivo following UV-induced DNA damage. This procedure has been used to demonstrate that replication forks blocked by UV-induced damage undergo a transient reversal that is stabilized by RecA and several gene products associated with the RecF pathway. The technique demonstrates that these replication intermediates are maintained until a time that correlates with the removal of the lesions by nucleotide excision repair and replication resumes.

Visualization of UV-induced Replication Intermediates in E. coli using Two-dimensional Agarose-gel Analysis

We present a procedure by which two-dimensional agarose-gel analysis can be used to identify the structure of replication intermediates that occur following UV irradiation.

Inaccurate replication in the presence of DNA damage is responsible for the majority of cellular rearrangements and mutagenesis observed in all cell types ...

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular signatures, or to evolve proteins in the laboratory. Here, we present a protocol that allows the generation of large (>1011) mutant libraries for a given target sequence. This method is based on replication of a ColE1 plasmid encoding the desired sequence by a low-fidelity variant of DNA polymerase I (LF-Pol I). The target plasmid is transformed into a mutator strain of E. coli and plated on solid media, yielding between 0.2 and 1 mutations/kb, depending on the location of the target gene. Higher mutation frequencies are achieved by iterating this process of mutagenesis. Compared to alternative methods of mutagenesis, our protocol stands out for its simplicity, as no cloning or PCR are involved. Thus, our method is ideal for mutational labeling of plasmids or other Pol I templates or to explore large sections of sequence space for the evolution of activities not present in the original target. The tight spatial control that PCR or randomized oligonucleotide-based methods offer can also be achieved through subsequent cloning of specific sections of the library. Here we provide protocols showing how to create a random mutant library and how to establish drug-based selections in E. coli to identify mutants exhibiting new biochemical activities.

Mutagenesis and Functional Selection Protocols for Directed Evolution of Proteins in E. coli

Here we demonstrate a simple protocol to create a random mutant library for a given target sequence. We show how this method, which is performed in vivo in Escherichia coli, can be coupled with functional selections to evolve new enzymatic activities.

The efficient generation of genetic diversity represents an invaluable molecular tool that can be used to label DNA synthesis, to create unique molecular ...

Assembly of Nucleosomal Arrays from Recombinant Core Histones and Nucleosome Positioning DNA

Core histone octamers that are repetitively spaced along a DNA molecule are called nucleosomal arrays. Nucleosomal arrays are obtained in one of two ways: purification from in vivo sources, or reconstitution in vitro from recombinant core histones and tandemly repeated nucleosome positioning DNA. The latter method has the benefit of allowing for the assembly of a more compositionally uniform and precisely positioned nucleosomal array. Sedimentation velocity experiments in the analytical ultracentrifuge yield information about the size and shape of macromolecules by analyzing the rate at which they migrate through solution under centrifugal force. This technique, along with atomic force microscopy, can be used for quality control, ensuring that the majority of DNA templates are saturated with nucleosomes after reconstitution. Here we describe the protocols necessary to reconstitute milligram quantities of length and compositionally defined nucleosomal arrays suitable for biochemical and biophysical studies of chromatin structure and function.

A method is presented for the reconstitution of model nucleosomal arrays from recombinant core histones and tandemly repeated nucleosome positioning DNA. We also describe how sedimentation velocity experiments in the analytical ultracentrifuge, and atomic force microscopy (AFM) are used to monitor the extent of nucleosomal array saturation after reconstitution.

Core histone octamers that are repetitively spaced along a DNA molecule  are called nucleosomal arrays. Nucleosomal arrays are obtained in one of two  ...

Efficient Production and Purification of Recombinant Murine Kindlin-3 from Insect Cells for Biophysical Studies

Kindlins are essential coactivators, with talin, of the cell surface receptors&#160;integrins and also participate in integrin outside-in signalling, and the control of gene transcription in the cell nucleus. The kindlins are ~75 kDa multidomain proteins and bind to an NPxY motif and upstream T/S cluster of the integrin &#946;-subunit cytoplasmic tail. The hematopoietically-important kindlin isoform, kindlin-3, is critical for platelet aggregation during thrombus formation, leukocyte rolling in response to infection and inflammation and osteoclast podocyte formation in bone resorption. Kindlin-3&#39;s role in these processes has resulted in extensive cellular and physiological studies. However, there is a need for an efficient method of acquiring high quality milligram quantities of the protein for further studies. We have developed a protocol, here described, for the efficient expression and purification of recombinant murine kindlin-3 by use of a baculovirus-driven expression system in Sf9 cells yielding sufficient amounts of high purity full-length protein to allow its biophysical characterization. The same approach could be taken in the study of the other mammalian kindlin isoforms.

Kindlins are fundamental to cell adhesion through integrins but studies of them have been hampered by the difficulty encountered in expressing them recombinantly in bacterial hosts. We describe here methods for their efficient production in baculovirus-infected insect cells.

Kindlins are essential coactivators, with talin, of the cell surface receptors&#160;integrins and also participate in integrin outside-in signalling, and ...

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. However, purifying proteins is sometimes challenging since many proteins are expressed in an insoluble form. When working with difficult or multiple targets it is therefore recommended to use high throughput (HTP) protein expression screening on a small scale (1-4&#160;ml cultures) to quickly identify conditions for soluble expression. To cope with the various structural genomics programs of the lab, a quantitative (within a range of 0.1-100 mg/L culture of recombinant protein) and HTP protein expression screening protocol was implemented and validated on thousands of proteins. The protocols were automated with the use of a liquid handling robot but can also be performed manually without specialized equipment.
Disulfide-rich venom proteins are gaining increasing recognition for their potential as therapeutic drug leads. They can be highly potent and selective, but their complex disulfide bond networks make them challenging to produce. As a member of the FP7 European Venomics project (<a href="http://www.venomics.eu">www.venomics.eu</a>), our challenge is to develop successful production strategies with the aim of producing thousands of novel venom proteins for functional characterization. Aided by the redox properties of disulfide bond isomerase DsbC, we adapted our HTP production pipeline for the expression of oxidized, functional venom peptides in the E. coli cytoplasm. The protocols are also applicable to the production of diverse disulfide-rich proteins. Here we demonstrate our pipeline applied to the production of animal venom proteins. With the protocols described herein it is likely that soluble disulfide-rich proteins will be obtained in as little as a week. Even from a small scale, there is the potential to use the purified proteins for validating the oxidation state by mass spectrometry, for characterization in pilot studies, or for sensitive micro-assays.

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

A protocol for the quantitative, high throughput expression screening and analytical purification of fusion proteins from small-scale Escherichia coli cultures is described and applied to the expression of disulfide-rich animal venom protein targets.

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. ...

Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli

Elastin-like polypeptides are repetitive biopolymers that exhibit a lower critical solution temperature phase transition behavior, existing as soluble unimers below a characteristic transition temperature and aggregating into micron-scale coacervates above their transition temperature. The design of elastin-like polypeptides at the genetic level permits precise control of their sequence and length, which dictates their thermal properties. Elastin-like polypeptides are used in a variety of applications including biosensing, tissue engineering, and drug delivery, where the transition temperature and biopolymer architecture of the ELP can be tuned for the specific application of interest. Furthermore, the lower critical solution temperature phase transition behavior of elastin-like polypeptides allows their purification by their thermal response, such that their selective coacervation and resolubilization allows the removal of both soluble and insoluble contaminants following expression in Escherichia coli. This approach can be used for the purification of elastin-like polypeptides alone or as a purification tool for peptide or protein fusions where recombinant peptides or proteins genetically appended to elastin-like polypeptide tags can be purified without chromatography. This protocol describes the purification of elastin-like polypeptides and their peptide or protein fusions and discusses basic characterization techniques to assess the thermal behavior of pure elastin-like polypeptide products.

Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli

Elastin-like polypeptides are stimulus-responsive biopolymers with applications ranging from recombinant protein purification to drug delivery. This protocol describes the purification and characterization of elastin-like polypeptides and their peptide or protein fusions from Escherichia coli using their lower critical solution temperature phase transition behavior as a simple alternative to chromatography.

Elastin-like polypeptides are repetitive biopolymers that exhibit a lower critical solution temperature phase transition behavior, existing as soluble ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Purification and Refolding to Amyloid Fibrils of (His)6-tagged Recombinant Shadoo Protein Expressed as Inclusion Bodies in E. coli

The Escherichia coli expression system is a powerful tool for the production of recombinant eukaryotic proteins. We use it to produce Shadoo, a protein belonging to the prion family. A chromatographic method for the purification of (His)6-tagged recombinant Shadoo expressed as inclusion bodies is described. The inclusion bodies are solubilized in 8 M urea and bound to a Ni2+-charged column to perform ion affinity chromatography. Bound proteins are eluted by a gradient of imidazole. Fractions containing Shadoo protein are subjected to size exclusion chromatography to obtain a highly purified protein. In the final step purified Shadoo is desalted to remove salts, urea and imidazole. Recombinant Shadoo protein is an important reagent for biophysical and biochemical studies of protein conformation disorders occurring in prion diseases. Many reports demonstrated that prion neurodegenerative diseases originate from the deposition of stable, ordered amyloid fibrils. Sample protocols describing how to fibrillate Shadoo into amyloid fibrils at acidic and neutral/basic pHs are presented. The methods on how to produce and fibrillate Shadoo can facilitate research in laboratories working on prion diseases, since it allows for production of large amounts of protein in a rapid and low cost manner.

Purification and Refolding to Amyloid Fibrils of His6-tagged Recombinant Shadoo Protein Expressed as Inclusion Bodies in E. coli

A two-step chromatographic method is described for the purification of recombinant Shadoo protein expressed as inclusion bodies in Escherichia coli, as well as a protocol to fibrillate purified Shadoo into amyloid structures.

The Escherichia coli expression system is a powerful tool for the production of recombinant eukaryotic proteins. We use it to produce Shadoo, a protein ...

Production of Human Norovirus Protruding Domains in E. coli for X-ray Crystallography

The norovirus capsid is composed of a single major structural protein, termed VP1. VP1 is subdivided into a shell (S) domain and a protruding (P) domain. The S domain forms a contiguous scaffold around the viral RNA, whereas the P domain forms viral spikes on the S domain and contains determinants for antigenicity and host-cell interactions. The P domain binds carbohydrate structures, i.e., histo-blood group antigens, which are thought to be important for norovirus infections. In this protocol, we describe a method for producing high quality norovirus P domains in high yields. These proteins can then be used for X-ray crystallography and ELISA in order to study antigenicity and host-cell interactions.
The P domain is firstly cloned into an expression vector and then expressed in bacteria. The protein is purified using three steps that involve immobilized metal-ion affinity chromatography and size exclusion chromatography. In principle, it is possible to clone, express, purify, and crystallize proteins in less than four weeks, which makes this protocol a rapid system for analyzing newly emerging norovirus strains.

Production of Human Norovirus Protruding Domains in E. coli for X-ray Crystallography

Here, we describe a method to express and purify high quality norovirus protruding (P) domains in E. coli for use in X-ray crystallography studies. This method can be applied to other calicivirus P domains, as well as non-structural proteins, i.e., viral protein genome-linked (VPg), protease, and RNA dependent RNA polymerase (RdRp).

The norovirus capsid is composed of a single major structural protein, termed VP1. VP1 is subdivided into a shell (S) domain and a protruding (P) domain. ...