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><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>&lt;em&gt;E. coli &lt;/em&gt;BL21 (DE3)</td><td>Lab Stock</td><td>N/A</td><td></td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; DH10&amp;beta;</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 &amp;beta;-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 &amp;micro;m, MCE)</td><td>Merck</td><td>VCWP04700</td><td></td></tr><tr><td>Millipore Express PLUS membrane filter (0.45 &amp;micro;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&amp;nbsp;</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 KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; 72 mM K&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</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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6.8K 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 

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique applications for OMV where traditional nanoparticles are proving too difficult to synthesize. To date, all Gram-negative bacteria have been shown to produce OMV demonstrating packaging of a variety of cargo that includes small molecules, peptides, proteins and genetic material. Based on their diverse cargo, OMV are implicated in many biological processes ranging from cell-cell communication to gene transfer and delivery of virulence factors depending upon which bacteria are producing the OMV. Only recently have bacterial OMV become accessible for use across a wide range of applications through the development of techniques to control and direct packaging of recombinant proteins into OMV. This protocol describes a method for the production, purification, and use of enzyme packaged OMV providing for improved overall production of recombinant enzyme, increased vesiculation, and enhanced enzyme stability. Successful utilization of this protocol will result in the creation of a bacterial strain that simultaneously produces a recombinant protein and directs it for OMV encapsulation through creating a synthetic linkage between the recombinant protein and an outer membrane anchor protein. This protocol also details methods for isolating OMV from bacterial cultures as well as proper handling techniques and things to consider when adapting this protocol for use for other unique applications such as: pharmaceutical drug delivery, medical diagnostics, and environmental remediation.

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

A protocol for the production, purification, and use of enzyme packaged outer membrane vesicles (OMV) providing for enhanced enzyme stability for implementation across diverse applications is presented.

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique ...

Biochemistry

Large-scale Production of Recombinant RNAs on a Circular Scaffold Using a Viroid-derived System in Escherichia coli

With increasing interest in RNA biology and the use of RNA molecules in sophisticated biotechnological applications, the methods to produce large amounts of recombinant RNAs are limited. Here, we describe a protocol to produce large amounts of recombinant RNA in Escherichia coli based on co-expression of a chimeric molecule that contains the RNA of interest in a viroid scaffold and a plant tRNA ligase. Viroids are relatively small, non-coding, highly base-paired circular RNAs that are infectious to higher plants. The host plant tRNA ligase is an enzyme recruited by viroids that belong to the family Avsunviroidae, such as Eggplant latent viroid (ELVd), to mediate RNA circularization during viroid replication. Although ELVd does not replicate in E. coli, an ELVd precursor is efficiently transcribed by the E. coli RNA polymerase and processed by the embedded hammerhead ribozymes in bacterial cells, and the resulting monomers are circularized by the co-expressed tRNA ligase reaching a remarkable concentration. The insertion of an RNA of interest into the ELVd scaffold enables the production of tens of milligrams of the recombinant RNA per liter of bacterial culture in regular laboratory conditions. A main fraction of the RNA product is circular, a feature that facilitates the purification of the recombinant RNA to virtual homogeneity. In this protocol, a complementary DNA (cDNA) corresponding to the RNA of interest is inserted in a particular position of the ELVd cDNA in an expression plasmid that is used, along the plasmid to co-express eggplant tRNA ligase, to transform E. coli. Co-expression of both molecules under the control of strong constitutive promoters leads to production of large amounts of the recombinant RNA. The recombinant RNA can be extracted from the bacterial cells and separated from the bulk of bacterial RNAs taking advantage of its circularity.

Large-scale Production of Recombinant RNAs on a Circular Scaffold Using a Viroid-derived System in Escherichia coli

Here, we present a protocol to produce large amounts of recombinant RNA in Escherichia coli by co-expressing a chimeric RNA that contains the RNA of interest in a viroid scaffold and a plant tRNA ligase. The main product is a circular molecule that facilitates purification to homogeneity.

With increasing interest in RNA biology and the use of RNA molecules in sophisticated biotechnological applications, the methods to produce large amounts ...

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine exchangers in plants have not been identified. Here, we outline a method to characterize polyamine antiporters using membrane vesicles generated from the lysis of Escherichia coli cells heterologously expressing a plant antiporter. First, we heterologously expressed AtBAT1 in an E. coli strain deficient in polyamine and arginine exchange transporters. Vesicles were produced using a French press, purified by ultracentrifugation and utilized in a membrane filtration assay of labeled substrates to demonstrate the substrate specificity of the transporter. These assays demonstrated that AtBAT1 is a proton-mediated transporter of arginine, &#947;-aminobutyric acid (GABA), putrescine and spermidine. The mutant strain that was developed for the assay of AtBAT1 may be useful for the functional analysis of other families of plant and animal polyamine exchangers. We also hypothesize that this approach can be used to characterize many other types of antiporters, as long as these proteins can be expressed in the bacterial cell membrane. E. coli is a good system for the characterization of novel transporters, since there are multiple methods that can be employed to mutagenize native transporters.

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

We describe a method for the characterization of proton-driven membrane transporters in membrane vesicle preparations produced by heterologous expression in E. coli and lysis of cells using a French press.

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine ...

Generic Protocol for Optimization of Heterologous Protein Production Using Automated Microbioreactor Technology

A core business in industrial biotechnology using microbial production cell factories is the iterative process of strain engineering and optimization of bioprocess conditions. One important aspect is the improvement of cultivation medium to provide an optimal environment for microbial formation of the product of interest. It is well accepted that the media composition can dramatically influence overall bioprocess performance. Nutrition medium optimization is known to improve recombinant protein production with microbial systems and thus, this is a rewarding step in bioprocess development. However, very often standard media recipes are taken from literature, since tailor-made design of the cultivation medium is a tedious task that demands microbioreactor technology for sufficient cultivation throughput, fast product analytics, as well as support by lab robotics to enable reliability in liquid handling steps. Furthermore, advanced mathematical methods are required for rationally analyzing measurement data and efficiently designing parallel experiments such as to achieve optimal information content.
The generic nature of the presented protocol allows for easy adaption to different lab equipment, other expression hosts, and target proteins of interest, as well as further bioprocess parameters. Moreover, other optimization objectives like protein production rate, specific yield, or product quality can be chosen to fit the scope of other optimization studies. The applied Kriging Toolbox (KriKit) is a general tool for Design of Experiments (DOE) that contributes to improved holistic bioprocess optimization. It also supports multi-objective optimization which can be important in optimizing both upstream and downstream processes.

This manuscript describes a generic approach for tailor-made design of microbial cultivation media. This is enabled by an iterative workflow combining Kriging-based experimental design and microbioreactor technology for sufficient cultivation throughput, which is supported by lab robotics to increase reliability and speed in liquid handling media preparation.

A core business in industrial biotechnology using microbial production cell factories is the iterative process of strain engineering and optimization of ...

Bioengineering

CFPS Protocols for Micro-Compartmentalized Synthetic Cells

Cell-Free Protein Synthesis System for Building Synthetic Cells

The Cell-Free Protein Synthesis (CFPS) system has been widely employed to facilitate the bottom-up assembly of synthetic cells. It serves as the host for the core machinery of the Central Dogma, standing as an optimal chassis for the integration and assembly of diverse artificial cellular mimicry systems. Despite its frequent use in the fabrication of synthetic cells, establishing a tailored and robust CFPS system for a specific application remains a nontrivial challenge. In this methods paper, we present a comprehensive protocol for the CFPS system, routinely employed in constructing synthetic cells. This protocol encompasses key stages in the preparation of the CFPS system, including the cell extract, template preparation, and routine expression optimization utilizing a fluorescent reporter protein. Additionally, we show representative results by encapsulating the CFPS system within various micro-compartments, such as monolayer droplets, double-emulsion vesicles, and chambers situated atop supported lipid bilayers. Finally, we elucidate the critical steps and conditions necessary for the successful assembly of these CFPS systems in distinct environments. We expect that our approach will facilitate the establishment of good working practices among various laboratories within the continuously expanding synthetic cell community, thereby accelerating progress in the field of synthetic cell development.

Author Spotlight: Optimizing CFPS Systems for Synthetic Cell Construction

This protocol describes the Cell-Free Protein Synthesis (CFPS) system used in constructing synthetic cells. It outlines key stages with representative results in different micro-compartments. The protocol aims to establish best practices for diverse laboratories in the synthetic cell community, advancing progress in synthetic cell development.

The Cell-Free Protein Synthesis (CFPS) system has been widely employed to facilitate the bottom-up assembly of synthetic cells. It serves as the host for ...

Preparing Protein Producing Synthetic Cells using Cell Free Bacterial Extracts, Liposomes and Emulsion Transfer

The bottom-up assembly approach for construction of synthetic cells is an effective tool for isolating and investigating cellular processes in a cell mimicking environment. Furthermore, the development of cell-free expression systems has demonstrated the ability to reconstitute the protein production, transcription and translation processes (DNA&#8594;RNA&#8594;protein) in a controlled manner, harnessing synthetic biology. Here we describe a protocol for preparing a cell-free expression system, including the production of a potent bacterial lysate and encapsulating this lysate inside cholesterol-rich lipid-based giant unilamellar vesicles (GUVs) (i.e., stable liposomes), to form synthetic cells. The protocol describes the methods for preparing the components of the synthetic cells including the production of active bacterial lysates, followed by a detailed step-by-step preparation of the synthetic cells based on a water-in-oil emulsion transfer method. These facilitate the production of millions of synthetic cells in a simple and affordable manner with a high versatility for producing different types of proteins. The obtained synthetic cells can be used to investigate protein/RNA production and activity in an isolated environment, in directed evolution, and also as a controlled drug delivery platform for on-demand production of therapeutic proteins inside the body.

This protocol describes the method, materials, equipment and steps for bottom-up preparation of RNA and protein producing synthetic cells. The inner aqueous compartment of the synthetic cells contained the S30 bacterial lysate encapsulated within a lipid bilayer (i.e., stable liposomes), using a water-in-oil emulsion transfer method.

The bottom-up assembly approach for construction of synthetic cells is an effective tool for isolating and investigating cellular processes in a cell ...

A Convenient and General Expression Platform for the Production of Secreted Proteins from Human Cells

Recombinant protein expression in bacteria, typically E. coli, has been the most successful strategy for milligram quantity expression of proteins. However, prokaryotic hosts are often not as appropriate for expression of human, viral or eukaryotic proteins due to toxicity of the foreign macromolecule, differences in the protein folding machinery, or due to the lack of particular co- or post-translational modifications in bacteria. Expression systems based on yeast (P. pastoris or S. cerevisiae) 1,2, baculovirus-infected insect (S. frugiperda or T. ni) cells 3, and cell-free in vitro translation systems 2,4 have been successfully used to produce mammalian proteins. Intuitively, the best match is to use a mammalian host to ensure the production of recombinant proteins that contain the proper post-translational modifications. A number of mammalian cell lines (Human Embryonic Kidney (HEK) 293, CV-1 cells in Origin carrying the SV40 larget T-antigen (COS), Chinese Hamster Ovary (CHO), and others) have been successfully utilized to overexpress milligram quantities of a number of human proteins 5-9. However, the advantages of using mammalian cells are often countered by higher costs, requirement of specialized laboratory equipment, lower protein yields, and lengthy times to develop stable expression cell lines. Increasing yield and producing proteins faster, while keeping costs low, are major factors for many academic and commercial laboratories.
Here, we describe a time- and cost-efficient, two-part procedure for the expression of secreted human proteins from adherent HEK 293T cells. This system is capable of producing microgram to milligram quantities of functional protein for structural, biophysical and biochemical studies. The first part, multiple constructs of the gene of interest are produced in parallel and transiently transfected into adherent HEK 293T cells in small scale. The detection and analysis of recombinant protein secreted into the cell culture medium is performed by western blot analysis using commercially available antibodies directed against a vector-encoded protein purification tag. Subsequently, suitable constructs for large-scale protein production are transiently transfected using polyethyleneimine (PEI) in 10-layer cell factories. Proteins secreted into litre-volumes of conditioned medium are concentrated into manageable amounts using tangential flow filtration, followed by purification by anti-HA affinity chromatography. The utility of this platform is proven by its ability to express milligram quantities of cytokines, cytokine receptors, cell surface receptors, intrinsic restriction factors, and viral glycoproteins. This method was also successfully used in the structural determination of the trimeric ebolavirus glycoprotein 5,10.
In conclusion, this platform offers ease of use, speed and scalability while maximizing protein quality and functionality. Moreover, no additional equipment, other than a standard humidified CO2 incubator, is required. This procedure may be rapidly expanded to systems of greater complexity, such as co-expression of protein complexes, antigens and antibodies, production of virus-like particles for vaccines, or production of adenoviruses or lentiviruses for transduction of difficult cell lines.

In the post-human genomics era, the availability of recombinant proteins in native conformations is crucial to structural, functional and therapeutic research and development. Here, we describe a test- and large-scale protein expression system in human embryonic kidney 293T cells that can be used to produce a variety of recombinant proteins.

Recombinant protein expression in bacteria, typically E. coli, has been the most successful strategy for milligram quantity expression of proteins. ...

The MultiBac Protein Complex Production Platform at the EMBL

Proteomics research revealed the impressive complexity of eukaryotic proteomes in unprecedented detail. It is now a commonly accepted notion that proteins in cells mostly exist not as isolated entities but exert their biological activity in association with many other proteins, in humans ten or more, forming assembly lines in the cell for most if not all vital functions.1,2 Knowledge of the function and architecture of these multiprotein assemblies requires their provision in superior quality and sufficient quantity for detailed analysis. The paucity of many protein complexes in cells, in particular in eukaryotes, prohibits their extraction from native sources, and necessitates recombinant production. The baculovirus expression vector system (BEVS) has proven to be particularly useful for producing eukaryotic proteins, the activity of which often relies on post-translational processing that other commonly used expression systems often cannot support.3 BEVS use a recombinant baculovirus into which the gene of interest was inserted to infect insect cell cultures which in turn produce the protein of choice. MultiBac is a BEVS that has been particularly tailored for the production of eukaryotic protein complexes that contain many subunits.4 A vital prerequisite for efficient production of proteins and their complexes are robust protocols for all steps involved in an expression experiment that ideally can be implemented as standard operating procedures (SOPs) and followed also by non-specialist users with comparative ease. The MultiBac platform at the European Molecular Biology Laboratory (EMBL) uses SOPs for all steps involved in a multiprotein complex expression experiment, starting from insertion of the genes into an engineered baculoviral genome optimized for heterologous protein production properties to small-scale analysis of the protein specimens produced.5-8 The platform is installed in an open-access mode at EMBL Grenoble and has supported many scientists from academia and industry to accelerate protein complex research projects.

Protein complexes catalyze key cellular functions. Detailed functional and structural characterization of many essential complexes requires recombinant production. MultiBac is a baculovirus/insect cell system particularly tailored for expressing eukaryotic proteins and their complexes. MultiBac was implemented as an open-access platform, and standard operating procedures developed to maximize its utility.

Proteomics  research revealed the impressive complexity of eukaryotic proteomes in  unprecedented detail. It is now a commonly accepted notion that ...

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. ...

Recombinant Protein Expression for Structural Biology in HEK 293F Suspension Cells: A Novel and Accessible Approach

The expression and purification of large amounts of recombinant protein complexes is an essential requirement for structural biology studies. For over two decades, prokaryotic expression systems such as E. coli have dominated the scientific literature over costly and less efficient eukaryotic cell lines. Despite the clear advantage in terms of yields and costs of expressing recombinant proteins in bacteria, the absence of specific co-factors, chaperones and post-translational modifications may cause loss of function, mis-folding and can disrupt protein-protein interactions of certain eukaryotic multi-subunit complexes, surface receptors and secreted proteins. The use of mammalian cell expression systems can address these drawbacks since they provide a eukaryotic expression environment. However, low protein yields and high costs of such methods have until recently limited their use for structural biology. Here we describe a simple and accessible method for expressing and purifying milligram quantities of protein by performing transient transfections of suspension grown HEK (Human Embryonic Kidney) 293F cells.

The expression of recombinant proteins by mammalian systems is becoming an attractive method for producing protein complexes for structural biology. Here we present a simple yet highly efficient expression system using suspension grown mammalian cells to purify protein complexes for structural studies.

The expression and purification of large amounts of recombinant protein complexes is an essential requirement for structural biology studies. For over two ...

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

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Chapters in this video

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

Large-scale Production of Recombinant RNAs on a Circular Scaffold Using a Viroid-derived System in Escherichia coli

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Generic Protocol for Optimization of Heterologous Protein Production Using Automated Microbioreactor Technology

Cell-Free Protein Synthesis System for Building Synthetic Cells

Preparing Protein Producing Synthetic Cells using Cell Free Bacterial Extracts, Liposomes and Emulsion Transfer

A Convenient and General Expression Platform for the Production of Secreted Proteins from Human Cells

The MultiBac Protein Complex Production Platform at the EMBL

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

Recombinant Protein Expression for Structural Biology in HEK 293F Suspension Cells: A Novel and Accessible Approach