This study was supported by a research grant from National Institutes of Health grants (NIH) R01HL147095, R01HL141917, and R01HL136431, Kowa Company, Ltd., and the European Union&#39;s Horizon 2020 Research and Innovation Program under the Marie Sk&#322;odowska-Curie Grant Agreement No. 101023041 (R. Cahalane). Figure 1 was created with Biorender.com. The current cleaning protocol was developed by modifying a recommended tube cleaning protocol presented at the International Society of Extracellular Vesicles 2023 Education Day (https://www.youtube.com/watch?v=DOebcOes6iI). Many thanks to Dr. Kathryn Howe and Dr. Sneha Raju from the University Health Network (University of Toronto, Canada) for the original carotid tissue EV samples.

1. Tube cleaning
NOTE: The EV isolation procedure uses both capped and uncapped polycarbonate UC tubes (detailed below). The same procedure was followed for both capped and uncapped tubes. In the case of capped tubes, the lid parts were cleaned individually and reassembled post drying and pre-storage.
<ol>
	<li>Following the initial use of the polycarbonate tubes and removal of the sample, immediately submerge UC tubes in tap water to prevent drying of the sample to the side of the tube.</li>
	<li>Transfer and submerge tubes in 0.1% sodium dodecyl sulfate (SDS) in tap water for 10 min.</li>
	<li>One tube at a time, decant most, but not all, of the SDS solution from the tube and use a cotton swab to thoroughly wipe the area of the tube where the EV pellet was located. 
	NOTE: Do not use anything with bristles. This protocol used normal cotton swabs; however, specialized, low-lint cotton swabs can be used for cleaning the tubes.</li>
	<li>Rinse at least 3x with hot tap water (~50 &#176;C). Make sure to fill and decant the tube completely.</li>
	<li>Rinse 1x with demineralized water using a spray bottle or a narrow-mouthed wash bottle.</li>
	<li>Rinse 1x with 70% ethanol using a spray bottle.</li>
	<li>Leave the tube upside down to dry. 
	NOTE: Tube racks for fluorescence activated cell sorting work well for drying the UC tubes.</li>
	<li>Place the completely dried tubes in a clean jar; they are ready for reuse. 
	NOTE: Currently, the protocol is validated for re-use up to 2x.</li>
</ol>
2. EV enrichment
NOTE: The following EV isolation and proteomics protocol was used for both the original tissue EV isolation, as well as the &#34;mock samples&#34; used to obtain the blank (never-used) and cleaned tube samples. The original samples were resuspended tissue-entrapped EVs from carotid plaques of patients who underwent carotid endarterectomies. Surgical specimens were collected from the University Health Network (Toronto, Canada) and was approved by the institutional ethics committee. All patients provided informed consent for sample collection and data analysis, in accordance with the Declaration of Helsinki. The mock samples were pseudo-pellets obtained from treating blank and cleaned tubes with the same solutions and processing steps as the EV isolation samples.
<ol>
	<li>Defrost EV samples or mock samples in PBS on ice.</li>
	<li>Make NTE buffer (10 mM Tris-HCl, 1 mM EDTA, 0.137 M NaCl pH 7.4) and sterile filter.</li>
	<li>Spin the samples at 10,000 &#215; g for 10 min at 4 &#176;C.</li>
	<li>Move the supernatant into a 1 mL open-top polycarbonate tube.</li>
	<li>Centrifuge the supernatant at 100,000&#215; g for 1 h at 4 &#176;C.</li>
	<li>Resuspend the pellet in 150 &#181;L of sterile-filtered NTE buffer with protease inhibitor.</li>
	<li>Transfer to a 1.5 mL microcentrifuge tube and add NTE buffer to reach a total sample volume of 1.5 mL.</li>
	<li>Keep the samples on ice.</li>
</ol>
3. Preparation of density gradients and density gradient separation
NOTE: This EV isolation protocol has been previously described by Blaser et al.9 In brief:
<ol>
	<li>Prepare five solutions of Iodixanol Density Gradient Medium (10%, 15%, 20%, 25%, and 30%) with NTE buffer as a diluent.</li>
	<li>Layer the five density solutions sequentially in capped ultracentrifuge tube(s) with 30% at the bottom and 10% on the top.</li>
	<li>Secure the caps on the tube(s) and move gently to a stable horizontal position for 1 h at room temperature.</li>
	<li>After 1 h, move the tube(s) gently to a vertical position on ice for 30 min to cool the gradient.</li>
	<li>Add 1.5 mL of the sample or mock sample (NTE buffer) to the top of the prepared density gradient tube(s).</li>
	<li>Ultracentrifuge tube(s) at 250,000 &#215; g for 40 min at 4 &#176;C.</li>
	<li>Remove the top fractions of the density gradient (2.4 mL) to uncapped ultracentrifuge tube(s). Fill to 9 mL with sterile-filtered NTE buffer.</li>
	<li>Pellet the EVs by ultracentrifuging at 100,000 &#215; g for 1 h at 4 &#176;C. 
	NOTE: Circle the location of the visible EV pellet or expected location with a marker. This will allow you to clean the area of the pellet with the swab directly.</li>
	<li>Aspirate the supernatant while moving the tube into an upside-down position for 3 min. Remove the remaining droplets with a cotton swab before turning the tube right side up.</li>
	<li>Resuspend the EV pellet in 12 &#181;L of lyse buffer in the referenced kit.</li>
</ol>
4. Peptide preparation for proteomics
NOTE: Proteomics sample preparation was performed according to the kit protocol with in-house modifications for low-abundance protein samples. The modified protocol is as follows:
<ol>
	<li>Lyse, denature, reduce, and alkylate the samples by heating in lyse buffer at 95 &#176;C for 10 min.</li>
	<li>Digest the samples using the Lys-C/trypsin digest solution for 2 h at 37 &#176;C using 10 &#181;L of lyse solution from the sample and 10 &#181;L of the digest solution.</li>
	<li>Purify peptides through a cartridge by washing hydrophilic and hydrophobic contaminants out and eluting off purified peptides using kit solutions.</li>
	<li>Dry the purified peptides in a speed vacuum at 45 &#176;C for 45 min or until dry. 
	NOTE: Avoid overdrying of the peptides.</li>
	<li>Store samples at -80 &#176;C until the time of mass spectrometry sequencing.</li>
	<li>Add 17 &#181;L of LC-load solution to the dried peptides.</li>
	<li>Leave the samples on ice for 1 h.</li>
	<li>Vortex the samples briefly and then sonicate in an ultrasonic waterbath for 5 min.</li>
	<li>Centrifuge the LC-load samples at 16,000 &#215; g for 1 min, aspirate 15 &#181;L from the top, and separate into a new tube.</li>
</ol>
5. Liquid chromatography and tandem mass spectrometry sequencing
<ol>
	<li>Inject 2 &#181;L of LC-load peptide solution onto a mass spectrometer.</li>
	<li>Fractionate the peptides using a dual-column setup.</li>
	<li>Heat the column at a constant temperature of 45 &#730;C.</li>
	<li>Run the analytical gradient at 300 nL/min from 5% to 21% Solvent B (acetonitrile/0.1% formic acid) for 60 min, followed by 10 min of 21% to 30% Solvent B, and another 10 min of a 95%-5% jigsaw wash.</li>
	<li>Set the MS1 to 120 K resolution (maximum injection time, 25 ms), subject the top precursor ions (within a 3.0 s cycle time) to HCD isolation width 1.2 m/z, dynamic exclusion enabled (60 s), and set the MS2 resolution to 60 K (maximum injection time, auto; normalized collision energy, 24%, 26%, and 28%).</li>
	<li>Set the MS1 and MS2 acquisition range to 400 m/z-1,200 m/z and 120 m/z-1,200 m/z, respectively.</li>
</ol>
6. Peptide/protein identification
<ol>
	<li>Search the acquired peptide spectra with a proteomic search software&#160;using a commercial search algorithm against a human uniProt database.</li>
	<li>Set the digestion enzyme to trypsin and allow up to two missed cleavages.</li>
	<li>Set the precursor tolerance to 10 ppm and the fragment tolerance window to 0.02 Da.</li>
	<li>Set methionine oxidation and N-terminal acetylation as variable modifications and cysteine carbamidomethylation as fixed modification.</li>
	<li>Filter the peptides based on a false discovery rate (FDR) threshold of 1.0% as calculated by the Percolator algorithm.</li>
	<li>Only consider peptides assigned to one given protein group and not detected in any other protein group as unique and use them for further analyses.</li>
	<li>Only include a minimum of two unique peptides for each protein in the analyses.</li>
	<li>Maximize IDs with precursor ion detection.</li>
	<li>Perform chromatographic alignment with a maximum retention time shift of 10 min, mass tolerance of 10 ppm, and signal-to-noise minimum of 5.</li>
	<li>Normalize peptide abundance by total peptide amount.</li>
</ol>

Efficient Recycling of Polycarbonate Ultracentrifuge Tubes for Proteomics

<ol><li>van Niel, G., D'Angelo, G., Raposo, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Shedding+light+on+the+cell+biology+of+extracellular+vesicles%5BTitle%5D)+AND+%22Nature+Reviews+Molecular+Cell+Biology%22%5BJournal%5D)">Shedding light on the cell biology of extracellular vesicles.</a> Nature Reviews Molecular Cell Biology. 19 (4), 213-228 (2018).</li>
<li>Théry, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Minimal+information+for+studies+of+extracellular+vesicles+2018+%28MISEV2018%29%3A+a+position+statement+of+the+International+Society+for+Extracellular+Vesicles+and+update+of+the+MISEV2014+guidelines%5BTitle%5D)+AND+%22Journal+of+Extracellular+Vesicles%22%5BJournal%5D)">Minimal information for studies of extracellular vesicles 2018 (MISEV2018): a position statement of the International Society for Extracellular Vesicles and update of the MISEV2014 guidelines.</a> Journal of Extracellular Vesicles. 7 (1), 1535750(2018).</li>
<li>Witwer, K. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Standardization+of+sample+collection%2C+isolation+and+analysis+methods+in+extracellular+vesicle+research%5BTitle%5D)+AND+%22Journal+of+Extracellular+Vesicles%22%5BJournal%5D)">Standardization of sample collection, isolation and analysis methods in extracellular vesicle research.</a> Journal of Extracellular Vesicles. 2 (1), 20360(2013).</li>
<li>Konoshenko, M. Y., Lekchnov, E. A., Vlassov, A. V., Laktionov, P. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Isolation+of+extracellular+vesicles%3A+general+methodologies+and+latest+trends%5BTitle%5D)+AND+%22BioMed+Research+International%22%5BJournal%5D)">Isolation of extracellular vesicles: general methodologies and latest trends.</a> BioMed Research International. 2018, e854347(2018).</li>
<li>Kowal, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Proteomic+comparison+defines+novel+markers+to+characterize+heterogeneous+populations+of+extracellular+vesicle+subtypes%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences%22%5BJournal%5D)">Proteomic comparison defines novel markers to characterize heterogeneous populations of extracellular vesicle subtypes.</a> Proceedings of the National Academy of Sciences. 113 (8), E968-E977 (2016).</li>
<li>Karimi, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Detailed+analysis+of+the+plasma+extracellular+vesicle+proteome+after+separation+from+lipoproteins%5BTitle%5D)+AND+%22Cellular+and+Molecular+Life+Sciences%22%5BJournal%5D)">Detailed analysis of the plasma extracellular vesicle proteome after separation from lipoproteins.</a> Cellular and Molecular Life Sciences. 75 (15), 2873-2886 (2018).</li>
<li>Lischnig, A., Bergqvist, M., Ochiya, T., Lässer, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Quantitative+proteomics+identifies+proteins+enriched+in+large+and+small+extracellular+vesicles%5BTitle%5D)+AND+%22Molecular+%26+Cellular+Proteomics%22%5BJournal%5D)">Quantitative proteomics identifies proteins enriched in large and small extracellular vesicles.</a> Molecular & Cellular Proteomics. 21 (9), 100273(2022).</li>
<li>van Herwijnen, M. J. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Comprehensive+proteomic+analysis+of+human+milk-derived+extracellular+vesicles+unveils+a+novel+functional+proteome+distinct+from+other+milk+components%5BTitle%5D)+AND+%22Molecular+%26+Cellular+Proteomics%22%5BJournal%5D)">Comprehensive proteomic analysis of human milk-derived extracellular vesicles unveils a novel functional proteome distinct from other milk components.</a> Molecular & Cellular Proteomics. 15 (11), 3412-3423 (2016).</li>
<li>Blaser, M. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Multiomics+of+tissue+extracellular+vesicles+identifies+unique+modulators+of+atherosclerosis+and+calcific+aortic+valve+stenosis%5BTitle%5D)+AND+%22Circulation%22%5BJournal%5D)">Multiomics of tissue extracellular vesicles identifies unique modulators of atherosclerosis and calcific aortic valve stenosis.</a> Circulation. 148 (8), 661-678 (2023).</li>
<li>Kristensen, K., Henriksen, J. R., Andresen, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Adsorption+of+cationic+peptides+to+solid+surfaces+of+glass+and+plastic%5BTitle%5D)+AND+%22PLOS+ONE%22%5BJournal%5D)">Adsorption of cationic peptides to solid surfaces of glass and plastic.</a> PLOS ONE. 10 (5), e0122419(2015).</li>
<li>Use and care of centrifuge tubes and bottles. Beckman Coulter. , Available from: 
 <a target="_blank" href="https://www.beckman.com/supplies/tubes-and-bottles">https://www.beckman.com/supplies/tubes-and-bottles</a> (2014).</li>
<li>R&D & Technology Department chemical resistance of polycarbonate products. PALRAM. , Available from: 
 <a target="_blank" href="http://ilgbcatalog.org/wp-content/uploads/2015/02/PC-chemical-resistance.pdf">http://ilgbcatalog.org/wp-content/uploads/2015/02/PC-chemical-resistance.pdf</a> (2015).</li>
<li>Wang, L., Zhang, J., Hou, S., Sun, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+simple+method+for+quantifying+polycarbonate+and+polyethylene+terephthalate+microplastics+in+environmental+samples+by+liquid+chromatography-tandem+mass+spectrometry%5BTitle%5D)+AND+%22Environmental+Science+%26+Technology+Letters%22%5BJournal%5D)">A simple method for quantifying polycarbonate and polyethylene terephthalate microplastics in environmental samples by liquid chromatography-tandem mass spectrometry.</a> Environmental Science & Technology Letters. 4 (12), 530-534 (2017).</li>
<li>Schittek, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dermcidin%3A+a+novel+human+antibiotic+peptide+secreted+by+sweat+glands%5BTitle%5D)+AND+%22Nature+Immunology%22%5BJournal%5D)">Dermcidin: a novel human antibiotic peptide secreted by sweat glands.</a> Nature Immunology. 2 (12), 1133-1137 (2001).</li>
<li>Blaydon, D. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mutations+in+CSTA%2C+encoding+Cystatin+A%2C+underlie+exfoliative+ichthyosis+and+reveal+a+role+for+this+protease+inhibitor+in+cell-cell+adhesion%5BTitle%5D)+AND+%22American+Journal+of+Human+Genetics%22%5BJournal%5D)">Mutations in CSTA, encoding Cystatin A, underlie exfoliative ichthyosis and reveal a role for this protease inhibitor in cell-cell adhesion.</a> American Journal of Human Genetics. 89 (4), 564-571 (2011).</li>
<li>Henry, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Hornerin+is+a+component+of+the+epidermal+cornified+cell+envelopes%5BTitle%5D)+AND+%22FASEB+Journal%22%5BJournal%5D)">Hornerin is a component of the epidermal cornified cell envelopes.</a> FASEB Journal. 25 (5), 1567-1576 (2011).</li>
<li>Bolling, M. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Generalized+ichthyotic+peeling+skin+syndrome+due+to+FLG2+mutations%5BTitle%5D)+AND+%22The+Journal+of+Investigative+Dermatology%22%5BJournal%5D)">Generalized ichthyotic peeling skin syndrome due to FLG2 mutations.</a> The Journal of Investigative Dermatology. 138 (8), 1881-1884 (2018).</li>
<li>Jin, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((DAMP+molecules+S100A9+and+S100A8+activated+by+IL-17A+and+house-dust+mites+are+increased+in+atopic+dermatitis%5BTitle%5D)+AND+%22Experimental+Dermatology%22%5BJournal%5D)">DAMP molecules S100A9 and S100A8 activated by IL-17A and house-dust mites are increased in atopic dermatitis.</a> Experimental Dermatology. 23 (12), 938-941 (2014).</li>
<li>Gläser, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+antimicrobial+protein+psoriasin+%28S100A7%29+is+upregulated+in+atopic+dermatitis+and+after+experimental+skin+barrier+disruption%5BTitle%5D)+AND+%22Journal+of+Investigative+Dermatology%22%5BJournal%5D)">The antimicrobial protein psoriasin (S100A7) is upregulated in atopic dermatitis and after experimental skin barrier disruption.</a> Journal of Investigative Dermatology. 129 (3), 641-649 (2009).</li>
<li>Molhave, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((House+dust+in+seven+Danish+offices%5BTitle%5D)+AND+%22Atmospheric+Environment%22%5BJournal%5D)">House dust in seven Danish offices.</a> Atmospheric Environment. 34, 4767-4779 (1999).</li>
<li>Buschow, S. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((MHC+class+II-associated+proteins+in+B-cell+exosomes+and+potential+functional+implications+for+exosome+biogenesis%5BTitle%5D)+AND+%22Immunology+and+Cell+Biology%22%5BJournal%5D)">MHC class II-associated proteins in B-cell exosomes and potential functional implications for exosome biogenesis.</a> Immunology and Cell Biology. 88 (8), 851-856 (2010).</li>
</ol>

The authors have no conflicts of interest to disclose.

Here we describe and validate a protocol for cleaning polycarbonate UC tubes for EV enrichment and proteomic applications. We demonstrated the successful removal of residual protein from the previous UC tube sample compared with a cleaned pseudo-pellet analysis below the limit of detection of this mass spectrometry acquisition protocol and showed the proteomic similarity of blank never-used UC tube compared to cleaned UC tube pseudo pellets.
First, to prevent the inadvertent adsorption of protein to the inner polycarbonate wall, UC tubes were immediately submerged in H2O10. Hand washing the polycarbonate UC tubes is recommended by the manufacturers11. Other detergents, such as Lipsol, should be used with caution. It is a proprietary laboratory detergent composed of a mixture of detergent, surfactant, and chelating agents with a non-neutral pH. Therefore, for this step, a mass spectrometry-compatible detergent, 0.1% SDS, was used and washing was not performed for more than 10 min11. In our experience, the use of bristle cleaning tools can lead to tube scratching if caution is not taken. To mitigate potential scratches, which could provide a reservoir for residual protein, a cotton swab was used to wipe the inside of the polycarbonate tube, with a focus on the area of EV pellet formation. Repeated autoclaving can reduce the polycarbonate tube life11; instead, three brief hot water rinses were performed. It should also be noted that although the manufacturers suggest non-thermal sterilization methods, a specific temperature to be avoided is not described. Tubes should always be inspected for signs of deformation or cracking before re-use, as per manufacturers&#39; recommendations. Flammable substances are not recommended to be used near operating centrifuges; thus, we dry the tubes overnight after the final 70% ethanol rinse step.
No significant polymer contamination was observed by manual assessment of chromatograms. SDS is not expected to degrade polycarbonate, and brief exposures to hot water (&#60;60 &#176;C) have no effect, while longer exposures do12. Further, previous studies have indicated that polycarbonate species elute at a retention time prior to collection based on our acetonitrile gradient13. While we cannot be certain that polymers are not present, with our acquisition methods, we are confident that the proteome is not substantively impacted.
When re-using labware for label-free proteomics experiments, residual protein contaminants from the previous use must be specifically considered. Further, EV proteomics are often conducted with very low-yielding samples, sometimes yielding protein amounts in the &#60;100 ng range. These amounts of starting protein are suitable for mass-spectrometry but substantially increase the impact even small amounts of residual protein will have on the overall proteome. Given the sensitivity of advancing mass spectrometry sequencing, contaminants from human handling and manufacturing will be identified. In this protocol, we show a substantial reduction in the number of identified proteins by 98% following cleaning. The remaining proteins were almost exclusively shared between the blank never-used tubes and the clean tubes. These results suggest that the identified proteins were contaminants from the proteomics preparation itself or residual elements from tube manufacturing, not carryover from the previous use. By far, the most abundant three identified proteins by orders of magnitude in the blank and cleaned tubes were human keratin type I (KRT1, KRT10, KRT9), found in hair and skin, generally recognized in these types of preparations as contaminants from human handling (https://www.thegpm.org/crap/). Nine of the shared 18 proteins were other keratins. Also included as a recognized human contaminant is dermcidin (DCD), a protein found in human sweat14. Other identified proteins are also found in the epidermis (CSTA15, HRNR16, FLG217) or previously identified in dust mites (S100A7, S100A8, S100A9)18,19. It should be noted that a substantial component of &#34;dust&#34; is human skin20. The two unique proteins identified in the cleaned tubes have both been implicated in EVs, however, are very minor compared to other identified EV proteins1,21. The proteins identified in the samples are consistent with the preparation and indicate a very clear overlap between blank and cleaned tubes, supporting re-use.
EVs have an important role in biological processes, including but not limited to cell-cell communication and the formation of biologic mineralization. EVs are ubiquitous in the human body and are a rapidly growing area of basic science due to their potential as biomarkers, therapeutics, and sources of information regarding (patho)physiology. Enriching these particles for experimentation using differential UC involves specialized polycarbonate tubes. Single use and disposal of these thermoplastics contribute to higher experimental consumable costs and the global plastic pollution burden. Herein, we presented and validated a protocol for cleaning and re-using polycarbonate UC tubes to perform EV isolation suitable for proteomics experiments. Ultimately, this will reduce the waste produced by EV laboratories and lower the cost associated with such experiments.

Extracellular vesicles (EVs) are lipid-bilayer-delimited particles released from cells that carry biologically active cargo, such as protein, and participate in various biological processes, including cell-cell communication and the formation of biologic mineralization1. These particles are found in all body fluids and tissues, and their biological activities and uses are a rapidly evolving field of scientific research. Isolation and validation of these nanoparticles present various challenges due to their small size and bio-similarity to other particles, such as liposomes and protein aggregates. The most recent International Society of Extracellular Vesicles guidelines, Minimal information for studies of extracellular vesicles 2018 (MISEV2018), are the recognized standard for EV scientific research2.
Various methods, often in tandem, must be used for EV isolation depending on the upstream source and the downstream applications. As of 2015, the most common primary isolation method for EVs was differential ultracentrifugation (UC)2. In principle, initial lower speed centrifugation separates&#160;larger or denser unwanted components, such as cells and cell debris, leaving EVs in the supernatant. Subsequently, UC uses very high centrifugal force to pellet out and thus separate and purify EVs from other particles that are smaller or less dense but which may also contain dense non-EV particles. Most protocols will often use, at one step or another, UC to isolate EVs from a fluid3. Further, UC is used in other methods of EV isolation, such as density gradient ultracentrifugation (DGUC), which uses mediums such as OptiprepTM iodixanol and centrifugal force to separate EVs according to their buoyant density4. Other methods of EV isolation exist3.
Considering the rapidly evolving understanding of the biological processes dictated by EVs and their potential as biomarkers and information regarding pathophysiology, discovery-based analyses such as proteomics have gained traction5,6,7,8. EVs are small, and, depending on the source, have a low yield of protein (&#60;1 &#956;g) when compared to whole tissue or cell lysate. Isolation of EVs for proteomics analysis requires special considerations, such as the removal of non-EV protein contaminants from upstream liquid or tissue, consideration of EV protein degradation during the isolation process, and the utilization of methods that create protein solutions that are chemically compatible with peptide preparation and mass spectrometry analyses.
Research laboratory consumables are often plastic and disposable in nature. These single-use materials contribute to the global plastic pollution crisis and consumable costs. Specialized polycarbonate and polystyrene UC tubes are designed to withstand the high centrifugal forces required to pellet EVs in laboratory applications. While it is possible to sterilize and disinfect UC tubes for re-use, proteomic analysis, especially those of low protein yields such as EVs, requires special attention. Not only is sufficient removal of residual protein from the previous use paramount, chemical compatibility with mass-spectroscopy and plastic-derived polymer contamination must also be considered.
Here we present a cleaning protocol of polycarbonate tubes using detergents suitable for mass-spectrometry and perform experiments to validate its successful removal of residual protein below limits of detection and lack of detectable polymer contaminants. To validate the cleaning protocol for EV proteomic applications using both UC and DGUC purposes, we obtained tubes from isolations of human vasculature tissue EVs with a combined UC and DGUC protocol. Tubes were cleaned using the protocol described, and the experimental process was repeated without samples comparing a blank never-used UC tube and a cleaned UC tube. Ultimately, the validation of a UC tube cleaning protocol suitable for the enrichment of EVs will reduce the waste produced by EV laboratories and lower the cost associated with such experiments.

<table><tbody><tr><td>10 mL Open-Top Thickwall Polycarbonate Tube</td><td>Beckman Coulter Life Sciences</td><td>355630</td><td>uncapped ultracentrifuge tube(s)&amp;nbsp;</td></tr><tr><td>10.4 mL Polycarbonate Bottle with Cap Assembly</td><td>Beckman Coulter Life Sciences</td><td>355603</td><td>capped ultracentrifuge tube(s)&amp;nbsp;</td></tr><tr><td>an Acclaim PepMap 100 C18 HPLC Columns, 75 &amp;micro;m x 70 mm; and an EASY-Spray HPLC Column, 75 &amp;micro;m x 250 mm</td><td>ThermoFisher Scientific</td><td>164946 and ES902</td><td>Dual column setup</td></tr><tr><td>Critical Swab Swab, Cotton Head</td><td>VWR</td><td>89031-270</td><td>cotton swab</td></tr><tr><td>Exploris 480 fronted with EASY-Spray Source, coupled to an Easy-nLC1200 HPLC pump. &amp;nbsp;</td><td>ThermoFisher Scientific</td><td>BRE725533</td><td>Mass spectrometer</td></tr><tr><td>Human UniProt database (101043 entries, updated January 2022)</td><td>NA</td><td>NA</td><td>Human database</td></tr><tr><td>MilliQ water</td><td></td><td></td><td>water</td></tr><tr><td>PreOmics iST kit &amp;nbsp;</td><td>PreOmics</td><td>P.O.00027</td><td>commercial protein sample preparation kit&amp;nbsp;</td></tr><tr><td>Proteome Discoverer &amp;nbsp;package (PD, Version 2.5)</td><td>ThermoFisher Scientific</td><td>NA</td><td>Proteomic search software</td></tr><tr><td>SEQUEST-HT search algorithm&amp;nbsp;</td><td>NA</td><td>NA</td><td>Search algorithm</td></tr><tr><td>Sodium Dodecyl Sulfate (20%)</td><td>Boston BioProducts</td><td>BM-230</td><td>detergent</td></tr></tbody></table>

polycarbonate ultracentrifuge tube re-use in proteomic analyses of extracellular vesicles

Single-use laboratory plastics exacerbate the pollution crisis and contribute to consumable costs. In extracellular vesicle (EV) isolation, polycarbonate ultracentrifuge (UC) tubes are used to endure the associated high centrifugal forces. EV proteomics is an advancing field and validated re-use protocols for these tubes are lacking. Re-using consumables for low-yield protein isolation protocols and downstream proteomics requires reagent compatibility with mass spectroscopy acquisitions, such as the absence of centrifuge tube-derived synthetic polymer contamination, and sufficient removal of residual proteins.
This protocol describes and validates a method for cleaning polycarbonate UC tubes for re-use in EV proteomics experiments. The cleaning process involves immediate submersion of UC tubes in H2O to prevent protein drying, washing in 0.1% sodium dodecyl sulfate (SDS) detergent, rinsing in hot tap water, demineralized water, and 70% ethanol. To validate the UC tube re-use protocol for downstream EV proteomics, used tubes were obtained following an experiment isolating EVs from cardiovascular tissue using differential UC and density gradient separation. Tubes were cleaned and the experimental process was repeated without EV samples comparing blank never-used UC tubes to cleaned UC tubes. The pseudo-EV pellets obtained from the isolation procedures were lysed and prepared for liquid chromatography-tandem mass spectrometry using a commercial protein sample preparation kit with modifications for low-abundance protein samples.
Following cleaning, the number of identified proteins was reduced by 98% in the pseudo-pellet versus the previous EV isolation sample from the same tube. Comparing a cleaned tube against a blank tube, both samples contained a very small number of proteins (&#8804;20) with 86% similarity. The absence of polymer peaks in the chromatograms of the cleaned tubes was confirmed. Ultimately, the validation of a UC tube cleaning protocol suitable for the enrichment of EVs will reduce the waste produced by EV laboratories and lower the experimental costs.

To validate the cleaning protocol (Figure 1), two experiments were performed. First, the proteome of the &#34;mock sample&#34; from the cleaned tube was compared against the proteome of the tissue EV sample from the tube&#39;s initial use to determine the carryover of identified proteins. Representative chromatograms show a reduction in peak heterogeneity following cleaning of the tubes (Figure 2). In the original EV isolation, 806 proteins were identified with two or more unique peptides. Following cleaning, the number of identified proteins was reduced by 98% to 19 proteins. Twelve proteins were shared in both, and seven proteins were unique to the cleaned tube (Figure 3A). Using a literature review, a list of characteristic EV proteins was compiled as previously published9. Of the listed 24 characteristic EV proteins identified in the original sample, only one was identified after cleaning (Table 1). Of the shared proteins, MFGE8, a characteristic EV protein was the most differentially enriched protein in the EV sample. In comparison two contaminant proteins, KRT14 and KRT5, were the most differentially enriched in the post-cleaned sample (Figure 3B).
To confirm the removal of protein from the tubes after washing, an SDS-PAGE gel was performed with a BSA standard curve from 1 &#181;g (typical low end of EV isolation yield) to 16.5 ng, and then blank and cleaned tube &#34;mock&#34; EV pellets in triplicate. All standard curve bands were visible however the samples were not, indicating cleaning removal of at least 98% of protein and no observable difference between the blank and cleaned tube (Figure 4A). To further assess the specific proteins identified in the cleaned tubes compared to a blank never-used tube, a direct comparison was conducted (N = 4, group). The chromatograms of the blank and cleaned tubes were very similar (Figure 4B). In the blank never-used tubes, 19 proteins were identified. Similarly, in the cleaned tube, 20 proteins were identified, with 18 shared within the two groups (Figure 4C). Ranking all the proteins by their abundance, contaminant keratin proteins KRT1, KRT9, and KRT10 are the most abundant in both groups (Figure 4D). In the cleaned tube, one characteristic EV protein was identified: lactadherin (MFGE8). There was high similarity between the clean versus blank tubes in the number of triggered MS2 sequencing events at each 5 min retention time bin (Figure 4E), the number of peptides sequenced, and the confidence in which they were sequenced (Figure 4F), as well as large overlap between the peptides (Figure 4G). Taken together, these results indicate no discernable impact of tube re-use with this cleaning protocol on the proteome.
The monomer of polycarbonate is 254.3 MH1+, which is below the MS1 acquisition range. Chromatograms were assessed for the presence of singly charged dimers and trimers, which were not detected (data not shown, N = 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66126/66126fig01.jpg" /> 
Figure 1: Polycarbonate ultracentrifuge tube cleaning protocol schematic. Abbreviations: UC = ultracentrifuge; SDS = sodium dodecyl sulfate. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66126/66126fig01large.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/66126/66126fig02.jpg" /> 
Figure 2: Representative comparison of the previous EV sample proteome to the &#34;mock sample&#34; from the cleaned tube (N = 2). Abbreviations: EV = extracellular vesicle; TIC-MS = total ion chromatography-mass spectrometry. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66126/66126fig02large.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/66126/66126fig03v2.jpg" /> 
Figure 3: Comparison of proteins identified in the EV sample to the &#34;mock&#34; sample obtained from the cleaned tubes. (A) Venn diagram protein identifications in the previous EV sample proteome to the &#34;mock sample&#34; from the cleaned tube (N = 2). (B)&#160;Differential enrichment of shared proteins ranked by log 2-fold change. Abbreviation: EV = extracellular vesicle, FC = fold change <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66126/66126fig03largev2.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/66126/66126fig04.jpg" /> 
Figure 4: Comparison of blank never-used tubes to cleaned tubes. (A) An SDS-PAGE gel was performed with a BSA standard curve from 1 &#181;g to 16.5 ng, and then blank and cleaned tube &#34;mock&#34; EV pellets in triplicate.&#160;(B) Representative chromatograms for&#160;blank and cleaned tubes. (C) Venn diagram of the shared or unique proteins from blank and cleaned tubes (N = 4). (D) All proteins identified in each group ranked according to abundance for (i) blank and (ii) cleaned tubes. Median values are plotted. (E) Histogram of all triggered MS2 events by retention time summed by group (N = 4). (F) Number of peptide IDs in each group by confidence, where high confidence peptides are distinct from intragroup matched peptides, where the identification confidence is lower but bolstered by matching to other spectra, mean/SD. (G) Number of unique and shared peptides between groups. Abbreviations: EV = extracellular vesicle; TIC-MS = total ion chromatography-mass spectrometry. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66126/66126fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Characteristic EV proteins found in pre versus post UC tube cleaning analysis. Abbreviations: UC = ultracentrifuge; EV = extracellular vesicle. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66126/Table 1.zip" target="_blank">Please click here to download this Table.</a>

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Author Spotlight: Effective Reuse of Polycarbonate Tubes for Extracellular Vesicle Isolation

A detailed protocol is provided for cleaning and re-using polycarbonate ultracentrifuge tubes to perform extracellular vesicle isolation suitable for proteomics experiments.

Polycarbonate Ultracentrifuge Tube Re-Use in Proteomic Analyses of Extracellular Vesicles

Single-use laboratory plastics exacerbate the pollution crisis and contribute to consumable costs. In extracellular vesicle (EV) isolation, polycarbonate ...

polycarbonate-ultracentrifuge-tube-re-use-proteomic-analyses

Nanyang Technological University

Research

JoVE Journal

Biology

901 Views.  Brigham and Women’s Hospital, Harvard Medical School. We bestuderen hart- en vaatziekten, met name de rol van extracellulaire blaasjes bij de pathologische verkalking van cardiovasculaire weefsels, zoals kleppen en bloedvaten. Onderzoekers in onze groep hebben onlangs methoden gedefinieerd om weefsel en gevangen extracellulaire blaasjes uit fibro-verkalkt cardiovasculair weefsel te extraheren om hun proteomische profiel te bepalen. Onze groep heeft aangetoond dat deze extracellulaire blaasjes een belangrijke eiwitlading bevatten voor het initië...

Video: Auteur in de schijnwerpers: effectief hergebruik van polycarbonaatbuizen voor extracellulaire blaasjesisolatie

Protein Digestion, Ultrafiltration, and Size Exclusion Chromatography to Optimize the Isolation of Exosomes from Human Blood Plasma and Serum

Exosomes, a type of nanovesicle released from all cell types, can be isolated from any bodily fluid. The contents of exosomes, including proteins and RNAs, are unique to the cells from which they are derived and can be used as indicators of disease. Several common enrichment protocols, including ultracentrifugation, yield exosomes laden with soluble protein contaminants. Specifically, we have found that the most abundant proteins within blood often co-purify with exosomes and can confound downstream proteomic studies, thwarting the identification of low abundance biomarker candidates. Of additional concern is irreproducibility of exosome protein quantification due to inconsistent representation of non-exosomal protein levels. The protocol detailed here was developed to remove non-exosomal proteins that co-purify along with exosomes, adding rigor to the exosome purification process. Five methods were compared using paired blood plasma and serum from five donors. Analysis using nanoparticle tracking analysis and micro bicinchoninic acid protein assay revealed that a combined protocol utilizing ultrafiltration and size exclusion chromatography yielded the optimal vesicle enrichment and soluble protein removal. Western blotting was used to verify that the expected abundant blood proteins, including albumin and apolipoproteins, were depleted.

Here, we present a protocol to purify exosomes from both plasma and serum with reduced co-purification of non-exosomal blood proteins. The optimized protocol includes ultrafiltration, protease treatment, and size exclusion chromatography. Enhanced purification of exosomes benefits downstream analyses, including more accurate quantification of vesicles and proteomic characterization.

Exosomes, a type of nanovesicle released from all cell types, can be isolated from any bodily fluid. The contents of exosomes, including proteins and ...

Biochemistry

Extraction of Extracellular Vesicles from Whole Tissue

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or prognostic biomarker assays, and likely serve as important players in the progression of a vast spectrum of diseases. Current research is focused on the characterization of vesicles secreted from multiple cell and tissue types in order to better understand the role of EVs in the pathogenesis of conditions including neurodegeneration, inflammation, and cancer. However, globally consistent and reproducible techniques to isolate and purify vesicles remain in progress. Moreover, methods for extraction of EVs from solid tissue ex vivo are scarcely described. Here, we provide a detailed protocol for extracting small EVs of interest from whole fresh or frozen tissues, including brain and tumor specimens, for further characterization. We demonstrate the adaptability of this method for multiple downstream analyses, including electron microscopy and immunophenotypic characterization of vesicles, as well as quantitative mass spectrometry of EV proteins.

Here, we provide a detailed protocol to isolate small extracellular vesicles (EVs) from whole tissues, including brain and tumor specimens. This method offers a reproducible technique to extract EVs from solid tissue for further downstream analyses.

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or ...

Cancer Research

Evaluation of the Storage Stability of Extracellular Vesicles

Extracellular vesicles (EVs) are promising targets in current research, to be used as drugs, drug-carriers, and biomarkers. For their clinical development, not only their pharmaceutical activity is important but also their production needs to be evaluated. In this context, research focuses on the isolation of EVs, their characterization, and their storage. The present manuscript aims at providing a facile procedure for the assessment of the effect of different storage conditions on EVs, without genetic manipulation or specific functional assays. This makes it possible to quickly get a first impression of the stability of EVs under a given storage condition, and EVs derived from different cell sources can be compared easily. The stability measurement is based on the physicochemical parameters of the EVs (size, particle concentration, and morphology) and the preservation of the activity of their cargo. The latter is assessed by the saponin-mediated encapsulation of the enzyme beta-glucuronidase into the EVs. Glucuronidase acts as a surrogate and allows for an easy quantification via the cleavage of a fluorescent reporter molecule. The present protocol could be a tool for researchers in the search for storage conditions that optimally retain EV properties to advance EV research toward clinical application.

Here we present a readily applicable protocol to assess the storage stability of extracellular vesicles, a group of naturally occurring nanoparticles produced by cells. The vesicles are loaded with glucuronidase as a model enzyme and stored under different conditions. After storage, their physicochemical parameters and the activity of the encapsulated enzyme are evaluated.

Extracellular vesicles (EVs) are promising targets in current research, to be used as drugs, drug-carriers, and biomarkers. For their clinical ...

Bioengineering

Purification of High Yield Extracellular Vesicle Preparations Away from Virus

One of the major hurdles in the field of extracellular vesicle (EV) research today is the ability to achieve purified EV preparations in a viral infection setting. The presented method is meant to isolate EVs away from virions (i.e., HIV-1), allowing for a higher efficiency and yield compared to conventional ultracentrifugation methods. Our protocol contains three steps: EV precipitation, density gradient separation, and particle capture. Downstream assays (i.e., Western blot, and PCR) can be run directly following particle capture. This method is advantageous over other isolation methods (i.e., ultracentrifugation) as it allows for the use of minimal starting volumes. Furthermore, it is more user friendly than alternative EV isolation methods requiring multiple ultracentrifugation steps. However, the presented method is limited in its scope of functional EV assays as it is difficult to elute intact EVs from our particles. Furthermore, this method is tailored towards a strictly research-based setting and would not be commercially viable.

This protocol isolates extracellular vesicles (EVs) away from virions with high efficiency and yield by incorporating EV precipitation, density gradient ultracentrifugation, and particle capture to allow for a streamlined workflow and a reduction of starting volume requirements, resulting in reproducible preparations for use in all EV research.

One of the major hurdles in the field of extracellular vesicle (EV) research today is the ability to achieve purified EV preparations in a viral infection ...

Immunology and Infection

Flow Cytometric Analysis of Extracellular Vesicles from Cell-conditioned Media

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for phenotypic analyses of extracellular vesicles (EV) including exosomes (Exo) in the peripheral blood and other body fluids. The small size of EV mandates the use of dedicated instruments having a detection threshold around 50-100 nm. Alternatively, EV can be bound to latex microbeads that can be detected by FC. Microbeads, conjugated with antibodies that recognize EV-associated markers/Cluster of Differentiation CD63, CD9, and CD81 can be used for EV capture. Exo isolated from CM can be analyzed with or without pre-enrichment by ultracentrifugation. This approach is suitable for EV analyses using conventional FC instruments. Our results demonstrate a linear correlation between Mean Fluorescence Intensity (MFI) values and EV concentration. Disrupting EV through sonication dramatically decreased MFI, indicating that the method does not detect membrane debris. We report an accurate and reliable method for the analysis of EV surface antigens, which can be easily implemented in any laboratory.

The protocol describes a reproducible method designed for use with cell culture supernatants to detect surface epitopes on small extracellular vesicles (EV). It utilizes specific EV immunoprecipitation using beads coupled with antibodies that recognize surface antigen CD9, CD63, and CD81. The method is optimized for downstream flow cytometry analysis.

Flow cytometry (FC) is the method of choice for semi-quantitative measurement of cell-surface antigen markers. Recently, this technique has been used for ...

Isolation and Characterization of Human Adipocyte-Derived Extracellular Vesicles using Filtration and Ultracentrifugation

Extracellular vesicles (EVs) are lipid enclosed envelopes that carry biologically active material such as proteins, RNA, metabolites and lipids. EVs can modulate the cellular status of other cells locally in tissue microenvironments or through liberation into peripheral blood. Adipocyte-derived EVs are elevated in the peripheral blood and show alterations in their cargo (RNA and protein) during metabolic disturbances, including obesity and diabetes. Adipocyte-derived EVs can regulate the cellular status of neighboring vascular cells, such as endothelial cells and adipose tissue resident macrophages to promote adipose tissue inflammation. Investigating alterations in adipocyte-derived EVs in vivo is complex because EVs derived from peripheral blood are highly heterogenous and contain EVs from other sources, namely platelets, endothelial cells, erythrocytes and muscle. Therefore, the culture of human adipocytes provides a model system for the study of adipocyte derived EVs. Here, we provide a detailed protocol for the extraction of total small EVs from cell culture media of human gluteal and abdominal adipocytes using filtration and ultracentrifugation. We further demonstrate the use of Nanoparticle Tracking Analysis (NTA) for quantification of EV size and concentration and show the presence of EV-protein tumor susceptibility gene 101 (TSG101) in the gluteal and abdominal adipocyte derived-EVs. Isolated EVs from this protocol can be used for downstream analysis, including transmission electron microscopy, proteomics, metabolomics, small RNA-sequencing,&#160;microarrays and can be&#160;utilized in functional in vitro/in vivo studies.

We describe the isolation of human adipocyte-derived extracellular vesicles (EVs) from gluteal and abdominal adipose tissue using filtration and ultracentrifugation. We characterize the isolated adipocyte-derived EVs by determining their size and concentration by Nanoparticle Tracking Analysis and by western blotting for the presence of EV-protein tumor susceptibility gene 101 (TSG101).

Extracellular vesicles (EVs) are lipid enclosed envelopes that carry biologically active material such as proteins, RNA, metabolites and lipids. EVs can ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...

Isolation and Characterization of Cyanobacterial Extracellular Vesicles

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential biotechnology models. Recent work has demonstrated that both marine and freshwater cyanobacteria produce extracellular vesicles -&#160;small membrane-bound structures released from the outer surface of the microbes. While vesicles likely contribute to diverse biological processes, their specific functional roles in cyanobacterial biology remain largely unknown. To encourage and advance research in this area, a detailed protocol is presented for isolating, concentrating, and purifying cyanobacterial extracellular vesicles. The current work discusses methodologies that have successfully isolated vesicles from large cultures of Prochlorococcus, Synechococcus, and Synechocystis. Methods for quantifying and characterizing vesicle samples from these strains are presented. Approaches for isolating vesicles from aquatic field samples are also described. Finally, typical challenges encountered with cyanobacterial vesicle purification, methodological considerations for different downstream applications, and the trade-offs between approaches are also discussed.

The present protocol providesdetailed descriptions for isolation, concentration,and characterization of extracellular vesicles from cyanobacterial cultures. Approaches for purifying vesicles from cultures at different scales, trade-offs among methodologies, and considerations for working with field samples are also discussed.

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential ...

Extracellular Vesicle Extraction: A Technique to Isolate Extracellular Vesicles from Whole Tissue Specimens Using Differential Centrifugation

Source: Hurwitz, S. N. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/59143/extraction-of-extracellular-vesicles-from-whole-tissue">Extraction of Extracellular Vesicles from Whole Tissue.</a> J. Vis. Exp. (2019)
This video describes the technique of extracting extracellular vesicles from fresh or frozen tissues. The isolated extracellular vesicles can be used for further analysis to uncover their significant roles in disease pathogenesis.

Source: Hurwitz, S. N. et al. Extraction of Extracellular Vesicles from Whole Tissue. J. Vis. Exp. (2019)
This video describes the technique of extracting ...

JoVE Encyclopedia of Experiments

Cancers of the Nervous System

In vitro studies

A Size Exclusion Chromatography Technique to Purify Mycobacterial Extracellular Vesicles

Source: Ryan, J. M., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/63895">Mycobacterium tuberculosis Extracellular Vesicle Enrichment through Size Exclusion Chromatography.</a> J. Vis. Exp. (2022).
This video demonstrates the size exclusion chromatography (SEC) technique for purifying bacterial extracellular vesicles (EVs) from the culture filtrate of Mycobacterium tuberculosis (Mtb). The technique utilizes a porous resin-packed column to obtain purified fractions of EVs. The porous particles selectively allow smaller proteins to enter the pores, resulting in slower elution. At the same time, EVs are excluded from entering the pores due to their larger size and are eluted first.

1. Preparation of crude Mtb EV concentrate
NOTE: For detailed procedures on the cultivation of Mtb and preparation of culture filtrate protein (CFP), see ...

Polycarbonate Ultracentrifuge Tube Re-Use in Proteomic Analyses of Extracellular Vesicles

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