The NanoMatFutur Junior Research program from the Federal Ministry of Education and Research, Germany (grant number 13XP5029A) supported this work. Maximilian Richter was supported by Studienstiftung des Deutschen Volkes (German Academic Scholarship Foundation) through a Ph.D. fellowship.

1. Cell culture and the production of cell-conditioned medium
<ol>
	<li>Generally, cultivate cells under the individual conditions required for the respective cell line.</li>
	<li>Cultivate the cells for 24-72 h in serum-free conditions or in medium containing EV-depleted fetal bovine serum (FBS). 
	NOTE: If EV-depleted FBS is used, employ a method proven to efficiently deplete the serum, to prevent contamination with bovine serum-derived EVs26.</li>
	<li>Collect the medium from the flasks. Centrifuge at 300 x g for 10 min to pellet the cells. Carefully collect the cell-conditioned medium (CCM), without disturbing the pelleted cells. Preferably, use the CCM directly, or store it overnight at 4 &#176;C. 
	NOTE: It is always preferable to use freshly produced CCM. If storage for longer time periods cannot be circumvented, all relevant parameters should be recorded in accordance with MISEV2018 guidelines26, and the potential biases of the results acquired need to be taken into consideration.</li>
	<li>Example protocol for HUVECs
	<ol>
		<li>Cultivate HUVEC cells for 120 h in EGM-2 medium containing FBS and other supplements.</li>
		<li>Cultivate HUVEC cells for 48 h in EBM-2 basal medium free of any additional supplements.</li>
		<li>Collect the medium from the flasks and perform the centrifugation step as indicated above (step 1.3). Typically, use 100 mL of medium for one EV-isolation.</li>
	</ol>
	</li>
</ol>
2. Ultracentrifugation of CCM
<ol>
	<li>Immediately before ultracentrifugation (UC), centrifuge the CCM for 15 min at 3,000 x g and 4 &#176;C to remove cell debris and large agglomerates.</li>
	<li>Carefully transfer the supernatant to the UC tubes. If using a fixed angle rotor, mark the orientation of the tubes in the centrifuge to facilitate the retrieval of the EV pellet after the UC. Centrifuge for 2 h at 120,000 x g, with a k-factor of 259.4.</li>
	<li>After UC, carefully discard the supernatant using a serological pipet, to avoid the disturbance of the pelleted EVs. 
	NOTE: The pellet might be invisible.</li>
	<li>Add 200 &#181;L of 0.2 &#181;m-filtered phosphate-buffered saline (PBS) to the first tube and use PBS and the residual supernatant to resuspend the pellet by pipetting up and down. Transfer the resulting EV suspension to the next tube of the respective sample and use it for the resuspension. Proceed this way to resuspend all EVs of the sample in a final volume of approximately 300-350 &#181;L.</li>
	<li>After resuspension, confirm the presence of particles by nanoparticle tracking analysis (NTA). Use the settings optimized for the given EV type, such as the settings below (step 2.5.1). 
	NOTE: The papers of Gardiner et al.27 and Vestad et al.28 contain valuable information on how to optimize the parameters for measuring EVs.
	<ol>
		<li>To reproduce the results described below, use instruments (e.g., NanoSight LM14) equipped with a green laser. Record three videos of 30 s with a screen gain of 1.0 and a camera level of 13. For analysis, use a screen gain of 1.0 and a detection threshold of 5.</li>
	</ol>
	</li>
	<li>Use the pellet immediately, if possible; otherwise, store it at 4 &#176;C overnight.</li>
</ol>
3. Glucuronidase encapsulation into EVs
<ol>
	<li>To the resuspended pellet, add beta-glucuronidase (10 mg/mL in PBS) to a final concentration of 1.5 mg/mL and saponin (10 mg/mL in H2O) to a final concentration of 0.1 mg/mL. Mix well by vortexing for 3 s.</li>
	<li>Incubate for 10 min at room temperature with intermitted mixing by gently flicking the tube. After incubation, directly purify by size-exclusion chromatography (SEC) (see section 5). 
	NOTE: Do not refreeze glucuronidase samples once thawed, to prevent enzyme degradation due to freezing.</li>
</ol>
4. Liposome production
<ol>
	<li>To prepare liposomes for comparison with EVs, dissolve 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) in a 2:3 molar ration in chloroform to a final concentration of 5 mM. Prepare 1 mL aliquots in high-performance liquid chromatography (HPLC) vials and let them dry overnight to form a lipid film. 
	CAUTION: Chloroform is toxic and suspected to be cancerogenic. Take proper precautions when handling it.</li>
	<li>Rehydrate the lipid-film with 1 mL of PBS containing 1.5 mg/mL glucuronidase. Heat it to 42 &#176;C and vortex for 1 min. Heat the extruder assembly to 42 &#176;C and extrude the lipid suspension 11x through a 200 nm polycarbonate membrane. Directly purify by SEC (see section 5).</li>
</ol>
5. Purification by SEC
<ol>
	<li>Prepare the SEC column using the following protocol.
	<ol>
		<li>Use only fresh purified water and freshly prepared buffers. Filter all buffers through a 0.2 &#181;m membrane filter and degas them to prevent the formation of air bubbles in the column.</li>
		<li>For the preparation of an SEC column, use agarose gel filtration-based matrix (e.g., Sepharose Cl-2b) or another SEC medium suitable to separate EVs and liposomes from protein impurities and excess enzyme. First, remove the 20% EtOH solution the medium is stored in, to prevent air bubble formation in the column. To this end, centrifuge the SEC medium at 3,000 x g for 10 min, remove the EtOH, and replace it with degassed water.</li>
		<li>Fill a glass column (with an inner diameter of 10 mm) with the SEC medium to the 15 mL mark. 
		NOTE: Volumes will differ for columns with different dimensions. Make sure to let the gel settle completely.</li>
		<li>Before a run, equilibrate the column with at least two column volumes of PBS. To store the column, first wash it with one column volume of water, followed by at least two column volumes of 20% EtOH. After storage, wash the column first with one column volume of water before equilibrating with PBS.</li>
		<li>Use up to 400 &#181;L of EV or liposome suspension in one separation. Collect fractions of 1 mL. After SEC, either store the purified EVs (see section 7) or subject them to a glucuronidase assay (see section 6).</li>
	</ol>
	</li>
	<li>Confirm the separation of EVs and liposomes from contaminating proteins and free glucuronidase. To this end, correlate the particle concentrations of the collected fractions with the protein concentration and the glucuronidase activity.
	<ol>
		<li>Assess the particle concentration by NTA (see 2.5)</li>
		<li>Assess the protein content by bicinchoninic acid (BCA) assay or another suitable protein quantification assay. Perform the assay according to the manufacturer&#8217;s protocol.</li>
		<li>Assess the glucuronidase activity by glucuronidase assay (see section 6).</li>
	</ol>
	</li>
	<li>Optionally, assess the EV morphology by transmission electron microscopy (TEM) and scanning electron microscopy (SEM).
	<ol>
		<li>For the preparation of TEM samples, add 10 &#181;L of EV suspension to a TEM grid, incubate for 10 min, and then blot away any excess liquid using a filter paper. Perform the fixation for 10 min with 10 &#181;L of 4% paraformaldehyde and blot away any excess. Wash 3x with water. Stain the vesicles by 20 s incubation with 30 &#181;L of 1% phosphor-tungstic acid hydrate. After blotting away the excess, dry the vesicles overnight. Visualize by TEM. 
		CAUTION: Phospho-tungstic acid is highly caustic; thus, protect skin and eyes.</li>
		<li>For SEM, fix the previously prepared TEM samples onto carbon disks and sputter them with a 50 nm thick gold layer. Visualize by SEM.</li>
	</ol>
	</li>
</ol>
6. Glucuronidase assay
<ol>
	<li>To allow a comparison between different samples and storage conditions by correlating particle number and enzyme activity, first measure the particle concentration of the sample by NTA (see step 2.5).</li>
	<li>Prepare a working solution of fluorescein di-&#946;-D-glucuronide by adding 1 &#181;L of the compound (10 mg/mL in H2O) to 199 &#181;L of PBS. Add 25 &#181;L of this solution to 125 &#181;L of purified EVs to get a final concentration of 8.3 &#181;g/mL. Pipet the sample into a black 96-well plate. Measure time point 0 h with a plate reader, using 480 nm as excitation and 516 nm as emission wavelength.</li>
	<li>Cover the plate tightly (e.g., with transparent plastic foil used for PCR plates) to minimize evaporation and incubate in the dark for 18 h at 37 &#176;C. Measure the fluorescein production using the plate reader parameters listed in step 6.2.</li>
</ol>
7. Storage of EVs and liposomes
NOTE: For all storage purposes, it is advisable to use low-binding tubes to reduce EV loss due to adsorption.
<ol>
	<li>Follow the parameters in this section to reproduce the representative results given below. Use samples consisting of 400 &#181;L of EV suspension.
	<ol>
		<li>Store at 4 &#176;C or -80 &#176;C or proceed to steps listed below.</li>
		<li>Lyophilize the EVs.
		<ol>
			<li>Add trehalose (40 mg/mL in H2O) up to a final concentration of 4 mg/mL to the purified EVs. Freeze the samples at -80 &#176;C for at least 1 h.</li>
			<li>Lyophilize the samples using the following parameters. For main drying, set the shelf temperature to 15 &#176;C and pressure to 0.180 mbar and leave the samples to dry for 46 h. For final drying, set the shelf temperature to 25 &#176;C and pressure to 0.0035 mbar and leave the samples to dry for 2 h. Store the lyophilized samples at 4 &#176;C.</li>
			<li>To rehydrate the samples, add H2O, equal to the amount of EV suspension present in the beginning (typically, 400 &#181;L). Do not use any buffer for rehydration.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
8. Analysis after storage
<ol>
	<li>To assess the enzyme activity, first remove the free glucuronidase, which may have leaked from the EVs during storage. Achieve this by an additional step of purification that is carried out either by SEC (see step 5) or asymmetric flow field-flow fractionation (AF4) (see step 8.1.2). 
	NOTE: Please be advised that both methods lead to a dilution of the EV sample; thus, use sufficient EV concentrations before storage to avoid moving below the NTA quantification limit. Expect a 1:10 dilution of the particles due to SEC or AF4.
	<ol>
		<li>For SEC purification, follow the protocol described above (see section 5).</li>
		<li>Perform AF4 purification.
		<ol>
			<li>Set up the instrument using a small channel with a 350 &#181;m spacer and a 30 kD molecular weight cut-off cellulose membrane. Place a 0.1 &#181;m pore size filter between the HPLC pump and the AF4 channel. Use freshly prepared 0.1 &#181;m-filtered PBS as the mobile phase to reduce the particle load and noise in the light-scattering detectors.</li>
			<li>Detect proteins using a UV detector set to 280 nm. To detect particles, use multiangle light scattering with the laser set to 658 nm.</li>
			<li>Use the following run method. Pre-focus for 1 min with a focus flow of 1 mL/min; then, inject 300 &#181;L of the sample at a rate of 0.2 mL/min and keep up the focus flow for 10 min. After the injection, elute the sample at 1 mL/min while applying a cross flow that decreases from 2 mL/min to 0.1 mL/min over the course of 8 min. Elute for another 10 min without cross flow. Collect fractions of 1 mL, starting after 12.5 min and continuing until 27 min.</li>
		</ol>
		</li>
		<li>Perform the glucuronidase assay as described above (see section 6).</li>
	</ol>
	</li>
	<li>To assess the size and concentration, use the NTA as described above (see step 2.5).</li>
	<li>Optionally, perform TEM and SEM to assess the morphology of the EVs after storage (see 5.3).</li>
</ol>

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B., 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=Impact+of+cell+culture+parameters+on+production+and+vascularization+bioactivity+of+mesenchymal+stem+cell-derived+extracellular+vesicles.">Impact of cell culture parameters on production and vascularization bioactivity of mesenchymal stem cell-derived extracellular vesicles.</a> Bioengineering &amp; Translational Medicine. 2 (2), 170-179 (2017).</li><li>Gardiner, 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=Techniques+used+for+the+isolation+and+characterization+of+extracellular+vesicles:+results+of+a+worldwide+survey.">Techniques used for the isolation and characterization of extracellular vesicles: results of a worldwide survey.</a> Journal of Extracellular Vesicles. 5 (1), 32945 (2016).</li><li>Zhang, 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=Identification+of+distinct+nanoparticles+and+subsets+of+extracellular+vesicles+by+asymmetric+flow+field-flow+fractionation.">Identification of distinct nanoparticles and subsets of extracellular vesicles by asymmetric flow field-flow fractionation.</a> Nature Cell Biology. 20 (3), 332-343 (2018).</li><li>Linares, R., Tan, S., Gounou, C., Arraud, N., Brisson, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-speed+centrifugation+induces+aggregation+of+extracellular+vesicles.">High-speed centrifugation induces aggregation of extracellular vesicles.</a> Journal of Extracellular Vesicles. 4 (1), 29509 (2015).</li><li>Lobb, R. 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=Optimized+exosome+isolation+protocol+for+cell+culture+supernatant+and+human+plasma.">Optimized exosome isolation protocol for cell culture supernatant and human plasma.</a> Journal of Extracellular Vesicles. 4, 27031 (2015).</li></ol>

The authors have nothing to disclose.

In this manuscript, we present a comprehensive protocol to study the stability of EVs under different storage conditions. With the combination of encapsulated glucuronidase as a functional readout and the evaluation of the physicochemical parameters of the EVs, the protocol allows for a straightforward storage stability evaluation of EVs and the comparison of EVs from different cell lines. SEM and TEM as complementary methods allow an insight into changes of the EVs on the single-particle level. The results presented here showed a tendency of the EVs and liposomes to aggregate due to lyophilization without a cryoprotectant (Figure 6). However, this was only observed in the SEM experiments. While EM imaging was not conducted with samples that were preserved with trehalose, the literature suggests that this cryoprotectant might indeed reduce the aggregation of EVs29. It is also possible to assess, if a given storage condition differentially affects EV size, the recovery rate and encapsulated molecules. Such an effect is exemplified by the results obtained for HUVEC EVs (Figure 1). While the particle recovery for 4 &#176;C and -80 &#176;C was better than for the lyophilized samples, the recovery of active encapsulated glucuronidase was best for the lyophilized samples. Lyophilization of EVs could be the more favorable storage condition, for instance, when analyzing EV biomarkers, where the focus is more on obtaining and preserving intact cargo rather than on intact vesicles.
In their recent paper on the lyophilization of EVs22, Charoenviriyakul et al. followed a comparable approach. Regarding the physicochemical parameters, they focused on the polydispersity index (PDI) and the zeta potential, as changes in zeta potential can correlate with reduced colloidal stability. However, the zeta potential cannot solely explain observed changes in stability30, which was also reflected in their results. They also compared the protein and RNA content of stored EVs by polyacrylamide gel electrophoresis. This technique can very well indicate substantial changes in the protein or RNA content of the vesicles. However, it requires much larger amounts of material than the method presented here. To assess the effect of storage on the EV cargo, Charoenviriyakul et al. heterologously expressed an enzyme and DNA species each in their EV producer cell line and monitored their activity and integrity in the EVs. However, such heterologous expression is not suitable if EVs from different sources are to be compared, as it requires a substantial amount of time for each new cell line, while the simple saponin-mediated encapsulation can be readily applied.
It is crucial to remove any nonencapsulated enzyme, as shown in Figure 3. EV-encapsulated glucuronidase reacts much slower with its substrate than free enzyme in solution, leading to an overestimation of the encapsulation efficiency. Clean separation is especially important for the analysis after storage, as the storage conditions might affect the activity of free glucuronidase in the sample differently and might lead to leakage from damaged EVs. This was apparent in our experiments (Figure 5), as the application of AF4 before the glucuronidase assay managed to remove dye not encapsulated in the vesicles. Thus, it made it possible to get clear results on the effect of the storage methods discussed here, while without AF4, the activity loss due to storage would have been underestimated.
Another important consideration is the isolation and characterization of the EVs to be tested for their stability. Although the described technique enables the comparison of EVs from different sources, this is only possible if all of them are prepared by the same isolation method so the results are not distorted18,31,32. In this context, cell culture conditions need to be taken into consideration also, as they can impact the quantity and bioactivity of the isolated EVs33. To obtain results that can be compared to other published results, it is advisable to consult the recently updated &#34;Minimal information for studies of extracellular vesicles&#34;&#160;that contains guidelines for the harmonization of EV research26.
In the protocol presented here, we used SEC or AF4 for removing the free enzyme after storage. Other methods could be applied, such as gradient ultracentrifugation, UC, or ultrafiltration34. Compared to centrifugal methods, SEC and AF4 are less time-consuming (e.g., roughly 1.5 h for SEC including column equilibration versus up to 16 h or more for gradient UC) and they can completely separate free proteins from the EVs in comparison to normal UC, where there always remains residual supernatant with the pellet. Moreover, SEC15 and AF435 are mild methods, which induce less shear stress on the EVs. In comparison, the forces imposed upon EVs during UC might lead to alterations of the particles, such as aggregation and alterations in size34,36.
A disadvantage of SEC and AF4 is the dilution of the samples. Thus, it is required to isolate sufficient amounts of EVs to maintain the concentrations required for NTA measurements after multiple purification steps. EV fractions may be concentrated using centrifugal filters but, depending on the filter material and the protocol, there could be a loss of vesicles37.
The limitation of the protocol discussed here is that it only monitors the enzyme activity of exogenously encapsulated glucuronidase, neither taking into account encapsulated RNA and DNA nor the EV surface proteins that might be important for functionality18. For the research of new EV therapeutics, this technique cannot replace assays on functionality but compliment them and give an indication about vesicle stability. It could be of great use if, for example, EVs derived from biofluids are to be stored for later analysis of their vesicle content.
In the future, the scope of this protocol could be expanded to also include EVs derived from other organisms, for instance, bacterial outer membrane vesicles. Another interesting next step would be to test the encapsulation of nucleic acids by saponin incubation and to also look into the stability of DNA or RNA cargos.
In conclusion, this procedure offers a simple method for the assessment of the storage stability of EVs from various mammalian cell sources by integrating both a physicochemical and a functional readout. This detailed protocol will allow EV researchers a straightforward determination of suitable storage conditions for their vesicles.

EVs are membrane-bound nanoparticles produced by nearly all cell types. For mammalian cells, EVs can be subdivided into two main groups with distinct production pathways1,2. Membrane vesicles, with a size range from roughly 100-1,000 nm, are produced by direct budding from the cell membrane. Exosomes, sized 30-200 nm, are derived from multivesicular bodies formed by inward budding into endosomes that subsequently fuse with the cell membrane to release multiple exosomes at once. The main function of these vesicles is the transport of information between cells3. For this purpose, cargos such as RNA, DNA, and proteins are actively sorted into them. EVs can convey a variety of effects on their targets, with implications for both health and disease state. On one side, they mediate positive effects such as tissue regeneration, antigen presentation, or antibiotic effects, which makes them auspicious targets for their development as therapeutics4,5. On the other side, EVs can promote tumor vascularization6, induce bystander effects in stress responses7, and might play a role in autoimmune diseases8 and inflammatory diseases9. Thus, they might be a key component to a better understanding of many pathological effects. However, the presence of altered EVs in manifold diseases, such as cancer10,11,12 and cardiovascular disorders13, and their easy accessibility in blood and urine makes them ideal biomarkers. Finally, their good biocompatibility14 and their inherent targeting ability make EVs also interesting for drug delivery15. In this manuscript, we describe a protocol for the evaluation of the storage stability of EVs derived from mammalian cells, an important property that is still little investigated.
For the clinical development of EVs, there are still many obstacles to surmount16, including the evaluation of their therapeutic effects, production, purification, and storage17. While -80 &#176;C is widely seen as the gold standard for EV storage18, the required freezers are expensive, and maintaining the required cold chain from the production to the patient can be challenging. Moreover, some reports indicate that storage at -80 &#176;C still not optimally preserves EVs and induces a loss in EV functionality19,20. Other methods, such as freeze-drying21,22 or spray-drying23, have been proposed as potential alternatives to the frozen storage of EVs.
The optimal way of assessing storage stability would be to test the EVs in functional assays or by the evaluation of a specific marker, for instance, their antibacterial activity19. This is possible when the desired effect of the vesicles is known and when one distinct group of EVs is to be studied. If EVs from different cell sources are to be compared (e.g., for drug encapsulation) or if there is no known functional readout, it is no longer possible to assess changes due to storage in a direct manner.
On the other hand, simply evaluating changes in their physicochemical parameters, such as size, particle recovery, and protein concentration, does not always predict changes in EV activity, as has been shown in a recent patent20.
Here, we provide a readily applicable protocol for measuring the storage stability of EVs by assessing their physicochemical parameters combined with the activity of an encapsulated beta-glucuronidase enzyme as a surrogate for the cargo of the EVs. The loading of the enzyme is done by saponin incubation, a mild method established with EVs from different sources21,24,25. Saponin forms transient pores in the EV membrane, which allows enzyme uptake into the vesicle. As enzymes are prone to lose their activity if subjected to unfavorable storage conditions, they are an ideal surrogate for the evaluation of the preservation of functional cargoes of the EVs.
We have demonstrated that the application of this protocol to EVs derived from human mesenchymal stem cells (MSCs), human umbilical vein endothelial cells (HUVECs), and human adenocarcinoma alveolar epithelial cells (A549) indeed result in great differences in storage stability between different cell lines, which should be taken into consideration when choosing the EV source21.

<table><tbody><tr><td>1,2 dimyristoyl-sn glycero-3-phospho-choline (DMPC)</td><td>Sigma-Aldrich</td><td>P2663-25MG</td><td></td></tr><tr><td>1,2-dipalmitoyl-sn-glycero-3-phospho-choline (DPPC)</td><td>Sigma-Aldrich</td><td>P4329-25MG</td><td></td></tr><tr><td>225 cm&amp;sup2; cell culture flasks</td><td>Corning</td><td>431082</td><td>Used with 25 ml of medium</td></tr><tr><td>30 kDa regenerated cellulose membrane</td><td>Wyatt Technology Europe</td><td>1854</td><td></td></tr><tr><td>350 &amp;micro;m spacer</td><td>Wyatt Technology Europe</td><td></td><td></td></tr><tr><td>Automated fraction collector</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Beta-glucuronidase</td><td>Sigma-Aldrich</td><td>G7646-100KU</td><td></td></tr><tr><td>Chloroform</td><td>Fisher scientific</td><td>C/4966/17</td><td></td></tr><tr><td>Column oven</td><td>Hitachi High-Technologies Europe</td><td></td><td></td></tr><tr><td>D-(+)-Trehalose dihydrate</td><td>Sigma-Aldrich</td><td>T9531-10G</td><td></td></tr><tr><td>DAWN HELEOS II, Multi-angle light scattering detector</td><td>&amp;nbsp;Wyatt Technology Europe</td><td></td><td></td></tr><tr><td>Durapore Membrane filter, PVDF,&amp;nbsp; 0,1&amp;nbsp;&amp;micro;m, 47&amp;nbsp;mm</td><td>Merck</td><td>VVLP04700</td><td>Used for the preparation of buffers for AF4</td></tr><tr><td>EBM-2</td><td>Lonza Verviers, S.p.r.</td><td>CC-3156</td><td>Endothelial Cell Growth basal medium, used for the serum free culture of HUVEC cells</td></tr><tr><td>Eclipse dualtec</td><td>Wyatt Technology Europe</td><td></td><td></td></tr><tr><td>EGM-2</td><td>Lonza Verviers, S.p.r.</td><td>CC-3162</td><td>Endothelial Cell Growth medium, used for the normal culture of HUVEC cells</td></tr><tr><td>ELISA Plate Sealers</td><td>R&amp;amp;D Systems</td><td>DY992</td><td>used for sealing of 96-well plates for the glucuronidase assay</td></tr><tr><td>Ethanol</td><td>Fisher scientific</td><td>E/0665DF/17</td><td></td></tr><tr><td>Extruder Set With Holder/Heating Block</td><td>Avanti Polar Lipids</td><td>610000-1EA</td><td></td></tr><tr><td>Filter support</td><td>Avanti Polar Lipids</td><td>610014-1EA</td><td>used for liposome preparation</td></tr><tr><td>Fluorescein di-&amp;beta;-D-glucoronide</td><td>Thermo Fisher Scientific</td><td>F2915</td><td></td></tr><tr><td>Gibco PBS-tablets+CA10:F36</td><td>Thermo Fisher Scientific</td><td>18912014</td><td></td></tr><tr><td>Hettich Universal 320 R</td><td>Andreas Hettich GmbH &amp;amp; Co.KG</td><td></td><td>Used for pelleting cells at 300 g</td></tr><tr><td>Hettich Rotina 420 R</td><td>Andreas Hettich GmbH &amp;amp; Co.KG</td><td></td><td>Used for pelleting larger debris at 3000 g</td></tr><tr><td>HUVEC cells</td><td>Lonza Verviers, S.p.r.</td><td>C2517A</td><td></td></tr><tr><td>Kimble&amp;nbsp; FlexColumn 1X30CM</td><td>Kimble</td><td>420401-1030</td><td></td></tr><tr><td>Lyophilizer ALPHA 2-4 LSC</td><td>Christ</td><td></td><td></td></tr><tr><td>Microcentrifuge Tubes, Polypropylene</td><td>VWR international</td><td>525-0255</td><td>the tubes used for all EV-handling, found to be more favorable than comparable products from other suppliers regarding particle recovery</td></tr><tr><td>Nanosight LM14 equipped with a green laser</td><td>Malvern Pananalytical</td><td></td><td></td></tr><tr><td>Nanosight-software version 3.1</td><td>Malvern Pananalytical</td><td></td><td></td></tr><tr><td>Nucleopore 200 nm track-etch polycarbonate membranes</td><td>Whatman/GE Healthcare</td><td>110406</td><td>used for liposome preparation</td></tr><tr><td>PEEK Inline filter holder</td><td>Wyatt Technology Europe</td><td></td><td></td></tr><tr><td>Phosphotungstic acid hydrate</td><td>Sigma-Aldrich</td><td>79690-25G</td><td></td></tr><tr><td>Polycarbonate bottles for ultracentrifugation</td><td>Beckman Coulter</td><td>355622</td><td></td></tr><tr><td>QuantiPro BCA Assay Kit</td><td>Sigma-Aldrich</td><td>QPBCA-1KT</td><td></td></tr><tr><td>Saponin</td><td>Sigma-Aldrich</td><td>47036</td><td></td></tr><tr><td>Scanning electron microscopy Zeiss EVO HD 15</td><td>Carl Zeiss AG</td><td></td><td></td></tr><tr><td>Sepharose Cl-2b</td><td>GE Healthcare</td><td>17014001</td><td></td></tr><tr><td>SEM copper grids with carbon film</td><td>Plano</td><td>S160-4</td><td></td></tr><tr><td>Small AF4 channel</td><td>Wyatt Technology Europe</td><td></td><td></td></tr><tr><td>Sputter-coater Q150R ES</td><td>Quorum Technologies</td><td></td><td></td></tr><tr><td>Transmission electron microscopy JEOL JEM 2011</td><td>Oxford Instruments</td><td></td><td></td></tr><tr><td>Type 45 Ti ultracentrifugation rotor</td><td>Beckman Coulter</td><td>339160</td><td></td></tr><tr><td>Ultimate 3000 Dionex autosampler</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Ultimate 3000 Dionex isocratic pump</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Ultimate 3000 Dionex online vacuum degasser</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Ultracentrifuge OptimaTM L-90 K</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>UV detector</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Whatman 0.2 &amp;micro;m pore size mixed cellulose filter</td><td>Whatman/GE Healthcare</td><td>10401712</td><td>Used for the filtration of all buffers used with the EVs and in SEC</td></tr></tbody></table>

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.

Figure 1 displays the storage characteristics of EVs isolated from HUVECs. EVs were isolated by UC, glucuronidase was encapsulated, and after SEC, the purified EVs were evaluated for their physicochemical properties by NTA. A sample of the vesicles was subsequently subjected to AF4 purification and the glucuronidase activity was measured.
The vesicles were then stored for 7 d at 4 &#176;C or -80 &#176;C and at 4 &#176;C in lyophilized form, in the latter case with the addition of 4% trehalose. After storage, the vesicles were again measured by NTA, and after AF4, the remaining glucuronidase activity per particle was assessed.
The working principle of AF4 is based on the combination of laminar flow and an orthogonal crossflow through the porous membrane at the bottom of the AF4 channel, which differentially affects particles according to their size (Figure 2). Larger particles tend to be located closer to the membrane while smaller particles are located further up in the channel. Due to the parabolic flow profile of the laminar flow, particles further away from the membrane travel faster toward the detector, leading to a particle-separation by size. In a typical AF4 experiment, the injected particles are first focused on the membrane, by applying a cross flow without a longitudinal flow through the channel (Figure 2A). After injection and focusing, the elution starts by simultaneously applying a cross flow and a longitudinal flow to fractionate and elute the different subsets of particles (Figure 2B), which, in our case, were EVs and free glucuronidase.
Figure 3 illustrates the separation of nanoparticles (EVs or liposomes) from contaminating proteins and nonencapsulated glucuronidase. The SEC elution profile of liposomes (Figure 3A) purified after their preparation (see section 4 of the protocol) showed the separation of the particles with encapsulated glucuronidase from the free enzyme, detected both through BCA assay and enzyme activity. In the present example, fraction 6 and 7 would be chosen for further experiments as they contained the highest particle concentrations and to prevent possible contaminations with free glucuronidase. AF4 was also successful in separating EVs from free glucuronidase as demonstrated in Figure 3B, with a higher degree of separation than SEC, making contamination of the fractions containing vesicles less probable.
In Figure 4, a control experiment was performed to ensure that the enzyme activity measured for the vesicles was indeed linked to the encapsulation of glucuronidase into EVs and not caused by enzyme aggregates. These aggregates might have been formed due to the incubation of glucuronidase with saponin and lead to false positive results. To verify the fractions where vesicles would elute, purified EVs without encapsulated glucuronidase were subjected to SEC on the same column as the sample just containing saponin and the enzyme.
When glucuronidase was incubated with saponin and subsequently purified by SEC, no enzyme activity was found in the fractions typically containing EVs. While small amounts of particles were found to elute at the same time as EVs (making up &#60;0.1% of the particles recovered from a typical EV pellet), there was no correlating enzyme activity. These results indicate that active enzyme recovered in the fractions containing vesicles was encapsulated in them.
Figure 5 compares the recovery of active enzyme after storage with or without additional AF4 purification to remove nonencapsulated dye. With AF4 purification, there is generally a lower recovery of enzyme per particle, with the highest effect for storage at 4 &#176;C, where recovery drops by two-thirds. Thus, omitting this additional purification step can lead to wrong assumptions about the enzyme stability.
Figure 6 shows the effect of lyophilization without a cryoprotectant on vesicles derived from MSCs, A549 cells, and liposomes, compared with freezing at -80 &#176;C. The particles were imaged by TEM and SEM as described above (see step 5.3 of the protocol). The MSCs and A549 EVs did not exhibit big differences in shape in TEM images, comparing the two storage conditions. In the SEM pictures, however, the lyophilized samples displayed aggregates not found in the -80 &#176;C samples. Liposomes also displayed aggregates in the SEM picture, while in TEM, lyophilization appeared to induce a size increase of liposomes. Lyophilization without cryoprotectants also reduced the recovery of intact glucuronidase after storage (Figure 7). The addition of trehalose as a cryoprotectant increased the recovery of the active enzyme in a dose-dependent manner.
Figure 8 demonstrates the conversion of fluorescein di-&#946;-D-glucuronide to free fluorescein, taking place in the glucuronidase assay. While the educt was nonfluorescent at 516 nm, fluorescein is highly fluorescent at this wavelength. This allowed for a straightforward enzyme activity assay with high sensitivity.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59584/59584fig1.jpg" /> 
Figure 1: Storage stability of HUVEC EVs. (A) Particle recovery compared to before storage, (B) mean size, and (C) the normalized glucuronidase activity per particle of EVs isolated from HUVECs. Vesicles were stored for 7 d at 4 &#176;C and -80 &#176;C and at 4 &#176;C after lyophilization with 4% trehalose. The mean size and particle recovery were measured by NTA; the glucuronidase activity per particle was calculated combining NTA data and the results of the glucuronidase assay and normalized to before storage. Mean &#177; SD, n = 3, *p &#60; 0.05 (one-way ANOVA followed by Tukey&#8217;s post hoc test). Modified from Frank et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig1large.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/59584/59584fig2.jpg" /> 
Figure 2: Working principle of the AF4. (A) EVs and free glucuronidase are focused after injection by applying a cross flow. (B) Afterward, EVs and free enzyme are eluted separately by combining the flow through the channel with a cross flow. Modified from Frank et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig2large.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/59584/59584fig3.jpg" /> 
Figure 3: Separation of EVs and liposomes from free glucuronidase. (A) Representative SEC separation of glucuronidase-loaded liposomes, free glucuronidase, and protein contaminants. The graph displays the particle concentration (red), protein concentration (blue), and glucuronidase activity (green). (B) Demonstration of the separation of EVs from free glucuronidase. Untreated EVs, EVs spiked with 0.05 mg/mL glucuronidase, free glucuronidase (0.5 mg/mL), and EVs loaded with glucuronidase and purified by SEC (EV glucuronidase) were injected and analyzed by UV at 280 nm and 90&#176; light scattering. Free enzyme eluted separately from EVs. Modified from Frank et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig3large.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/59584/59584fig4.jpg" /> 
Figure 4: Control experiments for the purification of EVs from free glucuronidase. 400 &#181;L of native EVs or 400 &#181;L of 1.5 mg/mL glucuronidase in PBS incubated for 10 min with 0.1 mg/mL saponin were purified by SEC. (A) Particle concentrations of the collected fractions of vesicles and glucuronidase, respectively, and the enzyme activity of the purified glucuronidase. (B) UV absorption at 280 nm measured in the same experiment. The first small peak for glucuronidase corresponds with the grey line in panel A. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59584/59584fig5.jpg" /> 
Figure 5: The effect of additional purification steps after EV storage. Remaining glucuronidase activity per particle compared to d0 with or without an additional AF4 purification step after storage. HUVEC EVs were stored for 7 d at 4 &#176;C or -80 &#176;C and lyophilized with 4% trehalose. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59584/59584fig6.jpg" /> 
Figure 6: TEM and SEM pictures of EVs. TEM and SEM pictures of EVs from (A) MSCs and (B) A549 cells and (C) liposomes. Samples were stored for 14 d at -80 &#176;C or lyophilized without the addition of trehalose. Arrows indicate the presence of morphologically altered particles in the TEM pictures and aggregates in the SEM pictures. Modified from Frank et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/59584/59584fig7.jpg" /> 
Figure 7: The effect of trehalose on enzyme recovery after lyophilization. Comparison of the enzyme activity after lyophilization for 14 days, with 0%, 1%, and 4% trehalose with the sample before storage (0 days). Modified from Frank et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/59584/59584fig8.jpg" /> 
Figure 8: The enzymatic cleavage of fluorescein di-&#946;-D-glucuronide by glucuronidase. In the scheme, the reaction underlying the detection of glucuronidase is explained. Nonfluorescent fluorescein di-&#946;-D-glucuronide is cleaved by glucuronidase. Through the removal of the sugar residues, fluorescein regains its fluorescent properties. The fluorescence measured after the incubation period correlates with the amount of active enzyme present and is the readout of the glucuronidase assay. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59584/59584fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Evaluation of the Storage Stability of Extracellular Vesicles at JoVE.com

Evaluation of the Storage Stability of Extracellular Vesicles

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 ...

evaluation-of-the-storage-stability-of-extracellular-vesicles

Nanyang Technological University

Research

JoVE Journal

Bioengineering

14.0K Views.  Biogenic Nanotherapeutics group (BION). De opslagstabiliteit van extra cellulaire akeltjes is een belangrijke parameter voor hun prospectief therapeutisch gebruik. Ons protocol biedt een gemakkelijk toepasbare tool om te evalueren hoe opslag de vesicles beïnvloedt. Het belangrijkste voordeel van onze techniek is dat door de invoering van glucuronidase als een exogene marker, extracellulaire vesikels afgeleid van verschillende oudercellen een gemakkelijk te vergelijken met betrekking tot hun storvermogen.Asymmetrische stroom...

Video: Evaluation of the Storage Stability of Extracellular Vesicles

Isolation of Tissue Extracellular Vesicles from the Liver

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in cell-to-cell communication by shuttling bioactive molecules such as RNA or protein from one cell to another. Most studies of EVs have been performed in cell culture models or in EVs isolated from body fluids. There is emerging interest in the isolation of EVs from tissues to study their contribution to physiological processes and how they are altered in disease. The isolation of EVs with sufficient yield from tissues is technically challenging because of the need for tissue dissociation without cellular damage. This method describes a procedure for the isolation of EVs from mouse liver tissue. The method involves a two-step process starting with in situ collagenase digestion followed by differential ultra-centrifugation. Tissue perfusion using collagenase provides an advantage over mechanical cutting or homogenization of liver tissue due to its increased yield of obtained EVs. The use of this two-step process to isolate EVs from the liver will be useful for the study of tissue EVs.

This is a protocol to isolate tissue extracellular vesicles (EVs) from the liver. The protocol describes a two-step process involving collagenase perfusion followed by differential ultracentrifugation to isolate liver tissue EVs.

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in ...

Biochemistry

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular communication. EVs consist of three populations of vesicles, namely microvesicles (MVs), exosomes, and apoptotic bodies. The limiting membrane of EVs is crucially involved in the interactions with the recipient cells, which could lead to the transfer of biologically active molecules to the recipient cells and, consequently, affect their behavior. The freeze-fracture electron microscopy technique is used to study the internal organization of biological membranes. Here, we present a protocol for MV isolation from cultured cancerous urothelial cells and the freeze-fracture of MVs in the steps of rapid freezing, fracturing, making and cleaning the replicas, and analyzing them with transmission electron microscopy. The results show that the protocol for isolation yields a homogenous population of EVs, which correspond to the shape and size of MVs. Intramembrane particles are found mainly in the protoplasmic face of the limiting membrane. Hence, freeze-fracture is the technique of choice to characterize the MVs' diameter, shape, and distribution of membrane proteins. The presented protocol is applicable to other populations of EVs.

We present a protocol for the isolation and freeze-fracturing of extracellular vesicles (EVs) originating from cancerous urothelial cells. The freeze-fracture technique revealed the EVs' diameter and shape and-as a unique feature-the internal organization of the EV membranes. These are of immense importance in understanding how EVs interact with the recipient membranes.

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular ...

Cancer Research

Labeling of Extracellular Vesicles for Monitoring Migration and Uptake in Cartilage Explants

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their cellular source, such as platelet lysate (PL). When used as treatment, EVs are expected to enter the target cells and effect a response from these. In this research, PL-derived EVs have been studied as a cell-free treatment for osteoarthritis (OA). Thus, a method was set up to label EVs and test their uptake on cartilage explants. PL-derived EVs are labeled with the lipophilic dye PKH26, washed twice through a column, and then tested in an in vitro inflammation-driven&#160;OA model for 5 h after particle quantification by nanoparticle tracking analysis (NTA). Hourly, cartilage explants are fixed, paraffined, cut into 6 &#181;m sections to mount on slides, and observed under a confocal microscope. This allows verification of whether EVs enter the target cells (chondrocytes) during this period and analyze their direct effect.

Here, we present a protocol to label platelet lysate-derived extracellular vesicles to monitor their migration and uptake in cartilage explants used as a model for osteoarthritis.

Extracellular vesicles (EVs) are used in different studies to prove their potential as a cell-free treatment due to their cargo derived from their ...

Isolation And Dendritic Cell-Uptake of Small Extracellular Vesicles from Echinococcus granulosus

The secretion of extracellular vesicles by cestodes is crucial for enabling cellular communication not only among parasites but also with host tissues. In particular, small extracellular vesicles (sEVs) act as nano-carriers transferring natural antigens, which are critical in host immunomodulation and parasite survival. This article presents a step-by-step protocol to isolate sEVs from larval stage cultures of Echinococcus granulosus and analyzes their uptake by dendritic cells obtained from murine bone marrow, which acquire adhesion and antigen presentation capacity during their maturation after one week of in vitro culture. This article provides comprehensive information for generating, purifying, and quantifying sEVs using ultracentrifugation alongside parallel analyses of dynamic light scattering and transmission electron microscopy. Additionally, a detailed experimental protocol is outlined for isolating and cultivating mouse bone marrow cells and driving their differentiation into dendritic cells using Flt3L. These dendritic cells can present antigens to na&#239;ve T cells, thereby modulating the type of immune response in vivo. Thus, alternative protocols, including confocal microscopy and flow cytometry analysis, are proposed to check the acquired maturational phenotype of dendritic cells previously exposed to parasitic sEVs. Finally, it is worth noting that the described protocol can be applied as a whole or in individual parts to carry out parasite in vitro culture, isolate extracellular vesicles, generate bone marrow-derived dendritic cell cultures, and perform uptake assays with these cells.

Here, we describe in vitro culture conditions, isolation, and increased generation of extracellular vesicles (EVs) from Echinococcus granulosus. The small EVs were characterized by dynamic light scattering and transmission electron microscopy. The uptake by bone marrow-derived dendritic cells and their phenotypic modulation were studied using confocal microscopy and flow cytometry.

The secretion of extracellular vesicles by cestodes is crucial for enabling cellular communication not only among parasites but also with host tissues. In ...

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 ...

Biology

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 ...

Isolation and Analysis of Traceable and Functionalized Extracellular Vesicles from the Plasma and Solid Tissues

Circulating and tissue-resident extracellular vesicles (EVs) represent promising targets as novel theranostic biomarkers, and they emerge as important players in the maintenance of organismal homeostasis and the progression of a wide spectrum of diseases. While the current research focuses on the characterization of endogenous exosomes with the endosomal origin, microvesicles blebbing from the plasma membrane have gained increasing attention in health and sickness, which are featured by an abundance of surface molecules recapitulating the membrane signature of parent cells. Here, a reproducible procedure is presented based on differential centrifugation for extracting and characterizing EVs from the plasma and solid tissues, such as the bone. The protocol further describes subsequent profiling of surface antigens and protein cargos of EVs, which are thus traceable for their derivations and identified with components related to potential function. This method will be useful for correlative, functional, and mechanistic analysis of EVs in biological, physiological, and pathological studies.

The present protocol describes a method to extract extracellular vesicles from the peripheral blood and solid tissues with subsequent profiling of surface antigens and protein cargos.

Circulating and tissue-resident extracellular vesicles (EVs) represent promising targets as novel theranostic biomarkers, and they emerge as important ...

Isolation, Characterization, and Therapeutic Application of Extracellular Vesicles from Cultured Human Mesenchymal Stem Cells

Extracellular vesicles (EVs) are heterogeneous membrane nanoparticles released by most cell types, and they are increasingly recognized as physiological regulators of organismal homeostasis and important indicators of pathologies; in the meantime, their immense potential to establish accessible and controllable disease therapeutics is emerging. Mesenchymal stem cells (MSCs) can release large amounts of EVs in culture, which have shown promise to jumpstart effective tissue regeneration and facilitate extensive therapeutic applications with good scalability and reproducibility. There is a growing demand for simple and effective protocols for collecting and applying MSC-EVs. Here, a detailed protocol is provided based on differential centrifugation to isolate and characterize representative EVs from cultured human MSCs, exosomes, and microvesicles for further applications. The adaptability of this method is shown for a series of downstream approaches, such as labeling, local transplantation, and systemic injection. The implementation of this procedure will address the need for simple and reliable MSC-EVs collection and application in translational research.

The present protocol describes the differential centrifugation for isolating and characterizing representative EVs (exosomes and microvesicles) from cultured human MSCs. Further applications of these EVs are also explained in this article.

Extracellular vesicles (EVs) are heterogeneous membrane nanoparticles released by most cell types, and they are increasingly recognized as physiological ...

Cardiovascular Biomarker Studies Using Plasma Extracellular Vesicle Proteome

Isolation, Characterization, and Proteomic Analysis of Plasma-Derived Extracellular Vesicles for Cardiovascular Biomarker Discovery

Extracellular vesicles (EV) are cell-derived, lipid bilayer-enclosed, non-replicable nanoparticles. EV currently gain attention in cardiovascular research due to their role in regulating intercellular communication, potentially serving as valuable biomarkers for cardiovascular disease. However, the EV proteome and its potential as a biomarker in cardiovascular diagnostics remain poorly understood. This protocol presents a standardized method for the isolation and quantification of plasma-derived EV and the analysis of their protein cargo using plasma samples from patients presenting to the Chest Pain Unit of a large university hospital. Following routine phlebotomy, EV are isolated from plasma by differential ultracentrifugation. The enrichment of specific EV marker proteins in EV isolates is visualized by immunoblotting, and average size distribution and plasma EV concentrations are quantified by nanoparticle tracking analysis. Finally, ultra-performance liquid chromatography-tandem mass spectrometry is employed for label-free analysis of the EV proteome. This protocol thus provides a comprehensive approach to study and use plasma-derived EV as potential carriers of critical biological information as well as to explore their potential as novel biomarkers.

Author Spotlight: Advancing the Analysis of Plasma Extracellular Vesicle Proteome for Cardiovascular Biomarker Studies

This article describes a methodology for the isolation, characterization, and quantification of human plasma-derived extracellular vesicles (EV) and presents a workflow for label-free analysis of the EV proteome using liquid chromatography-tandem mass spectrometry (LC-MS/MS).

Extracellular vesicles (EV) are cell-derived, lipid bilayer-enclosed, non-replicable nanoparticles. EV currently gain attention in cardiovascular research ...

Medicine

Fluorescence-Based Sorting of Adipose-Derived Extracellular Vesicles

Setting a Successful Sorting for Extracellular Vesicle Isolation

Extracellular vesicles (EVs) released by mesenchymal stromal cells (MSCs) contain a set of microRNAs with regenerative and anti-inflammatory roles. Therefore, purified MSC-EVs are envisioned as a next-generation therapeutic option for a wide array of diseases. In this protocol, we report the strategy for successfully sorting EVs from the supernatant of adipose-derived MSCs (ASCs), often used in orthopedics regenerative medicine applications.
First, we described the sample preparation, focusing on EV isolation and labeling steps with carboxyfluorescein succinimidyl ester (CFSE) for fluorescence detection; subsequently, we detailed the sorting process, which constitutes the main part of the protocol. 
In addition to the rules defined by MISEV 2023 and MIFlowCyt EV guidelines, we applied specific experimental conditions concerning nozzle size, frequency, and sheath pressure. Morphological parameters are established using beads of diameters selected to cover the theoretical range of EV size. After ASC-EVs sorting, we performed a purity check of the sorted fraction by re-analyzing it with the sorter and verifying the EV size distribution with the nanoparticle tracking analysis technique.
Due to the increasing importance of EVs, having a pure population to study and characterize is becoming crucial. Here, we demonstrate a winner strategy to set up sorting to achieve this goal.

This protocol provides a detailed description of sorting extracellular vesicles (EVs) released by mesenchymal stromal cells. In particular, it focuses on the instrument setting and the optimization of the sorting conditions. The goal is to sort extracellular vesicles while preserving their characteristics.

Extracellular vesicles (EVs) released by mesenchymal stromal cells (MSCs) contain a set of microRNAs with regenerative and anti-inflammatory roles. ...

Evaluation of the Storage Stability of Extracellular Vesicles

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