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 her...

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 border="0" cellpadding="0" cellspacing="0" style="width:933px;" width="933">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:333px;">1,2 dimyristoyl-sn glycero-3-phospho-choline (DMPC)</td>
			<td style="width:315px;">Sigma-Aldrich</td>
			<td style="width:119px;">P2663-25MG</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:333px;">1,2-dipalmitoyl-sn-glycero-3-phospho-choline (DPPC)</td>
			<td style="width:315px;">Sigma-Aldrich</td>
			<td>P4329-25MG</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">225 cm&#178; cell culture flasks</td>
			<td style="width:315px;">Corning</td>
			<td>431082</td>
			<td>Used with 25 ml of medium</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">30 kDa regenerated cellulose membrane</td>
			<td style="width:315px;">Wyatt Technology Europe</td>
			<td>1854</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">350 &#181;m spacer</td>
			<td style="width:315px;">Wyatt Technology Europe</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Automated fraction collector</td>
			<td style="width:315px;">Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Beta-glucuronidase</td>
			<td style="width:315px;">Sigma-Aldrich</td>
			<td style="width:119px;">G7646-100KU</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Chloroform</td>
			<td style="width:315px;">Fisher scientific</td>
			<td>C/4966/17</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Column oven</td>
			<td style="width:315px;">Hitachi High-Technologies Europe</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">D-(+)-Trehalose dihydrate</td>
			<td style="width:315px;">Sigma-Aldrich</td>
			<td>T9531-10G</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:333px;">DAWN HELEOS II, Multi-angle light scattering detector</td>
			<td style="width:315px;">&#160;Wyatt Technology Europe</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Durapore Membrane filter, PVDF,&#160; 0,1&#160;&#181;m, 47&#160;mm</td>
			<td style="width:315px;">Merck</td>
			<td>VVLP04700</td>
			<td>Used for the preparation of buffers for AF4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">EBM-2</td>
			<td style="width:315px;">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 height="21">
			<td height="21" style="height:21px;width:333px;">Eclipse dualtec</td>
			<td style="width:315px;">Wyatt Technology Europe</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">EGM-2</td>
			<td style="width:315px;">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 height="21">
			<td height="21" style="height:21px;">ELISA Plate Sealers</td>
			<td>R&#38;D Systems</td>
			<td>DY992</td>
			<td>used for sealing of 96-well plates for the glucuronidase assay</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Ethanol</td>
			<td style="width:315px;">Fisher scientific</td>
			<td>E/0665DF/17</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Extruder Set With Holder/Heating Block</td>
			<td style="width:315px;">Avanti Polar Lipids</td>
			<td>610000-1EA</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Filter support</td>
			<td style="width:315px;">Avanti Polar Lipids</td>
			<td>610014-1EA</td>
			<td>used for liposome preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Fluorescein di-&#946;-D-glucoronide</td>
			<td style="width:315px;">Thermo Fisher Scientific</td>
			<td>F2915</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Gibco PBS-tablets+CA10:F36</td>
			<td style="width:315px;">Thermo Fisher Scientific</td>
			<td>18912014</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Hettich Universal 320 R</td>
			<td>Andreas Hettich GmbH &#38; Co.KG</td>
			<td style="width:119px;"></td>
			<td>Used for pelleting cells at 300 g</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Hettich Rotina 420 R</td>
			<td style="width:315px;">Andreas Hettich GmbH &#38; Co.KG</td>
			<td style="width:119px;"></td>
			<td>Used for pelleting larger debris at 3000 g</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">HUVEC cells</td>
			<td style="width:315px;">Lonza Verviers, S.p.r.</td>
			<td style="width:119px;">C2517A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Kimble&#160; FlexColumn 1X30CM</td>
			<td style="width:315px;">Kimble</td>
			<td>420401-1030</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Lyophilizer ALPHA 2-4 LSC</td>
			<td style="width:315px;">Christ</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Microcentrifuge Tubes, Polypropylene</td>
			<td>VWR international</td>
			<td style="width:119px;">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 height="21">
			<td height="21" style="height:21px;width:333px;">Nanosight LM14 equipped with a green laser</td>
			<td style="width:315px;">Malvern Pananalytical</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Nanosight-software version 3.1</td>
			<td style="width:315px;">Malvern Pananalytical</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:333px;">Nucleopore 200 nm track-etch polycarbonate membranes</td>
			<td>Whatman/GE Healthcare</td>
			<td align="right">110406</td>
			<td>used for liposome preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">PEEK Inline filter holder</td>
			<td style="width:315px;">Wyatt Technology Europe</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Phosphotungstic acid hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>79690-25G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Polycarbonate bottles for ultracentrifugation</td>
			<td style="width:315px;">Beckman Coulter</td>
			<td style="width:119px;">355622</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">QuantiPro BCA Assay Kit</td>
			<td style="width:315px;">Sigma-Aldrich</td>
			<td>QPBCA-1KT</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Saponin</td>
			<td style="width:315px;">Sigma-Aldrich</td>
			<td>47036</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Scanning electron microscopy Zeiss EVO HD 15</td>
			<td style="width:315px;">Carl Zeiss AG</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Sepharose Cl-2b</td>
			<td style="width:315px;">GE Healthcare</td>
			<td>17014001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">SEM copper grids with carbon film</td>
			<td style="width:315px;">Plano</td>
			<td>S160-4</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Small AF4 channel</td>
			<td style="width:315px;">Wyatt Technology Europe</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Sputter-coater Q150R ES</td>
			<td style="width:315px;">Quorum Technologies</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:333px;">Transmission electron microscopy JEOL JEM 2011</td>
			<td style="width:315px;">Oxford Instruments</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Type 45 Ti ultracentrifugation rotor</td>
			<td style="width:315px;">Beckman Coulter</td>
			<td>339160</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Ultimate 3000 Dionex autosampler</td>
			<td style="width:315px;">Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Ultimate 3000 Dionex isocratic pump</td>
			<td style="width:315px;">Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Ultimate 3000 Dionex online vacuum degasser</td>
			<td style="width:315px;">Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Ultracentrifuge OptimaTM L-90 K</td>
			<td style="width:315px;">Beckman Coulter</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">UV detector</td>
			<td style="width:315px;">Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:333px;">Whatman 0.2 &#181;m pore size mixed cellulose filter</td>
			<td>Whatman/GE Healthcare</td>
			<td style="width:119px;">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...

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

Research

JoVE Journal

Bioengineering

13.9K Views.  Biogenic Nanotherapeutics group (BION). 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.

Video: Evaluation of the Storage Stability of Extracellular Vesicles

Magnetic Resonance Elastography Methodology for the Evaluation of Tissue Engineered Construct Growth

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use of destructive assessment is not acceptable. A proposed alternative is the use of an imaging process called magnetic resonance elastography. Elastography is a nondestructive method for determining the engineered outcome by measuring local mechanical property values (i.e., complex shear modulus), which are essential markers for identifying the structure and functionality of a tissue. As a noninvasive means for evaluation, the monitoring of engineered constructs with imaging modalities such as magnetic resonance imaging (MRI) has seen increasing interest in the past decade1. For example, the magnetic resonance (MR) techniques of diffusion and relaxometry have been able to characterize the changes in chemical and physical properties during engineered tissue development2. The method proposed in the following protocol uses microscopic magnetic resonance elastography (&mu;MRE) as a noninvasive MR based technique for measuring the mechanical properties of small soft tissues3. MRE is achieved by coupling a sonic mechanical actuator with the tissue of interest and recording the shear wave propagation with an MR scanner4. Recently, &mu;MRE has been applied in tissue engineering to acquire essential growth information that is traditionally measured using destructive mechanical macroscopic techniques5. In the following procedure, elastography is achieved through the imaging of engineered constructs with a modified Hahn spin-echo sequence coupled with a mechanical actuator. As shown in Figure 1, the modified sequence synchronizes image acquisition with the transmission of external shear waves; subsequently, the motion is sensitized through the use of oscillating bipolar pairs. Following collection of images with positive and negative motion sensitization, complex division of the data produce a shear wave image. Then, the image is assessed using an inversion algorithm to generate a shear stiffness map6. The resulting measurements at each voxel have been shown to strongly correlate (R2&gt;0.9914) with data collected using dynamic mechanical analysis7. In this study, elastography is integrated into the tissue development process for monitoring human mesenchymal stem cell (hMSC) differentiation into adipogenic and osteogenic constructs as shown in Figure 2.

The procedure demonstrates the methodology of magnetic resonance elastography for monitoring the engineered outcome of adipose and osteogenic tissue engineered constructs through noninvasive local assessment of the mechanical properties using microscopic magnetic resonance elastography (&mu;MRE).

Traditional mechanical testing often results in the destruction of the sample, and in the case of long term tissue engineered construct studies, the use ...

Constant Pressure-controlled Extrusion Method for the Preparation of Nano-sized Lipid Vesicles

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform to study protein-protein and protein-lipid interactions3, monitor drug delivery4,5, and encapsulation4. Phospholipids naturally create curved lipid bilayers, distinguishing itself from a micelle.6 Liposomes are traditionally classified by size and number of bilayers, i.e. large unilamellar vesicles (LUVs), small unilamellar vesicles (SUVs) and multilamellar vesicles (MLVs)7. In particular, the preparation of homogeneous liposomes of various sizes is important for studying membrane curvature that plays a vital role in cell signaling, endo- and exocytosis, membrane fusion, and protein trafficking8. Several groups analyze how proteins are used to modulate processes that involve membrane curvature and thus prepare liposomes of diameters &lt;100 - 400 nm to study their behavior on cell functions3. Others focus on liposome-drug encapsulation, studying liposomes as vehicles to carry and deliver a drug of interest9. Drug encapsulation can be achieved as reported during liposome formation9. Our extrusion step should not affect the encapsulated drug for two reasons, i.e. (1) drug encapsulation should be achieved prior to this step and (2) liposomes should retain their natural biophysical stability, securely carrying the drug in the aqueous core. These research goals further suggest the need for an optimized method to design stable sub-micron lipid vesicles.
Nonetheless, the current liposome preparation technologies (sonication10, freeze-and-thaw10, sedimentation) do not allow preparation of liposomes with highly curved surface (i.e. diameter &lt;100 nm) with high consistency and efficiency10,5, which limits the biophysical studies of an emerging field of membrane curvature sensing. Herein, we present a robust preparation method for a variety of biologically relevant liposomes.
Manual extrusion using gas-tight syringes and polycarbonate membranes10,5 is a common practice but heterogeneity is often observed when using pore sizes &lt;100 nm due to due to variability of manual pressure applied. We employed a constant pressure-controlled extrusion apparatus to prepare synthetic liposomes whose diameters range between 30 and 400 nm. Dynamic light scattering (DLS)10, electron microscopy11 and nanoparticle tracking analysis (NTA)12 were used to quantify the liposome sizes as described in our protocol, with commercial polystyrene (PS) beads used as a calibration standard. A near linear correlation was observed between the employed pore sizes and the experimentally determined liposomes, indicating high fidelity of our pressure-controlled liposome preparation method. Further, we have shown that this lipid vesicle preparation method is generally applicable, independent of various liposome sizes. Lastly, we have also demonstrated in a time course study that these prepared liposomes were stable for up to 16 hours. A representative nano-sized liposome preparation protocol is demonstrated below.

This protocol describes an extrusion method for preparing lipid vesicles of sub-micron sizes with a high degree of homogeneity. This method uses a pressure-controlled system with controlled nitrogen flow rates for liposome preparation. The lipid preparation1,2, liposome extrusion, and size characterization will be presented herein.

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform ...

The Encapsulation of Cell-free Transcription and Translation Machinery in Vesicles for the Construction of Cellular Mimics

As interest shifts from individual molecules to systems of molecules, an increasing number of laboratories have sought to build from the bottom up cellular mimics that better represent the complexity of cellular life. To date there are a number of paths that could be taken to build compartmentalized cellular mimics, including the exploitation of water-in-oil emulsions, microfluidic devices, and vesicles. Each of the available options has specific advantages and disadvantages. For example, water-in-oil emulsions give high encapsulation efficiency but do not mimic well the permeability barrier of living cells. The primary advantage of the methods described herein is that they are all easy and cheap to implement. Transcription-translation machinery is encapsulated inside of phospholipid vesicles through a process that exploits common instrumentation, such as a centrifugal evaporator and an extruder. Reactions are monitored by fluorescence spectroscopy. The protocols can be adapted for recombinant protein expression, the construction of cellular mimics, the exploration of the minimum requirements for cellular life, or the assembly of genetic circuitry.

Herein we describe simple methods for the preparation of vesicles, the encapsulation of transcription and translation machinery, and the monitoring of protein production. The resulting cell-free systems can be used as a starting point from which to build increasingly complex cellular mimics.

As interest shifts from individual molecules to systems of molecules, an increasing number of laboratories have sought to build from the bottom up ...

Quantification and Size-profiling of Extracellular Vesicles Using Tunable Resistive Pulse Sensing

Extracellular vesicles (EVs), including &#8216;microvesicles&#8217; and &#8216;exosomes&#8217;, are highly abundant in bodily fluids. Recent years have witnessed a tremendous increase in interest in EVs. EVs have been shown to play important roles in various physiological and pathological processes, including coagulation, immune responses, and cancer. In addition, EVs have potential as therapeutic agents, for instance as drug delivery vehicles or as regenerative medicine. Because of their small size (50 to 1,000 nm) accurate quantification and size profiling of EVs is technically challenging.
This protocol describes how tunable resistive pulse sensing (tRPS) technology, using the qNano system, can be used to determine the concentration and size of EVs. The method, which relies on the detection of EVs upon their transfer through a nano sized pore, is relatively fast, suffices the use of small sample volumes and does not require the purification and concentration of EVs. Next to the regular operation protocol an alternative approach is described using samples spiked with polystyrene beads of known size and concentration. This real-time calibration technique can be used to overcome technical hurdles encountered when measuring EVs directly in biological fluids.

Extracellular vesicles play important roles in physiological and pathological processes, including coagulation, immune responses, and cancer or as potential therapeutic agents in drug delivery or regenerative medicine. This protocol presents methods for the quantification and size characterization of isolated and non-isolated extracellular vesicles in various fluids using tunable resistive pulse sensing.

Extracellular vesicles (EVs), including &#8216;microvesicles&#8217; and &#8216;exosomes&#8217;, are highly abundant in bodily fluids. Recent years have ...

Paper-based Devices for Isolation and Characterization of Extracellular Vesicles

Extracellular vesicles (EVs), membranous particles released from various types of cells, hold a great potential for clinical applications. They contain nucleic acid and protein cargo and are increasingly recognized as a means of intercellular communication utilized by both eukaryote and prokaryote cells. However, due to their small size, current protocols for isolation of EVs are often time consuming, cumbersome, and require large sample volumes and expensive equipment, such as an ultracentrifuge. To address these limitations, we developed a paper-based immunoaffinity platform for separating subgroups of EVs that is easy, efficient, and requires sample volumes as low as 10 &#956;l. Biological samples can be pipetted directly onto paper test zones that have been chemically modified with capture molecules that have high affinity to specific EV surface markers. We validate the assay by using scanning electron microscopy (SEM), paper-based enzyme-linked immunosorbent assays (P-ELISA), and transcriptome analysis. These paper-based devices will enable the study of EVs in the clinic and the research setting to help advance our understanding of EV functions in health and disease.

This protocol details a method to isolate extracellular vesicles (EVs), small membranous particles released from cells, from as little as 10 &#956;l serum samples. This approach circumvents the need for ultracentrifugation, requires only a few minutes of assay time, and enables the isolation of EVs from samples of limited volumes.

Extracellular vesicles (EVs), membranous particles released from various types of cells, hold a great potential for clinical applications. They contain ...

Fabrication of Extracellular Matrix-derived Foams and Microcarriers as Tissue-specific Cell Culture and Delivery Platforms

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus of this article is on the methods for fabricating ECM-derived porous foams and microcarriers for use as biologically-relevant substrates in advanced 3D in vitro cell culture models or as pro-regenerative scaffolds and cell delivery systems for tissue engineering and regenerative medicine. Using decellularized tissues or purified insoluble collagen as a starting material, the techniques can be applied to synthesize a broad array of tissue-specific bioscaffolds with customizable geometries. The approach involves mechanical processing and mild enzymatic digestion to yield an ECM suspension that is used to fabricate the three-dimensional foams or microcarriers through controlled freezing and lyophilization procedures. These pure ECM-derived scaffolds are highly porous, yet stable without the need for chemical crosslinking agents or other additives that may negatively impact cell function. The scaffold properties can be tuned to some extent by varying factors such as the ECM suspension concentration, mechanical processing methods, or synthesis conditions. In general, the scaffolds are robust and easy to handle, and can be processed as tissues for most standard biological assays, providing a versatile and user-friendly 3D cell culture platform that mimics the native ECM composition. Overall, these straightforward methods for fabricating customized ECM-derived foams and microcarriers may be of interest to both biologists and biomedical engineers as tissue-specific cell-instructive platforms for in vitro and in vivo applications.

The tissue-specific extracellular matrix (ECM) is a key mediator of cell function. This article describes methods for synthesizing pure ECM-derived foams and microcarriers that are stable in culture without the need for chemical crosslinking for applications in advanced 3D in vitro cell culture models or as pro-regenerative bioscaffolds.

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus ...

Primary Clarification of CHO Harvested Cell Culture Fluid using an Acoustic Separator

Primary clarification is an essential step in a biomanufacturing process for the initial removal of cells from therapeutic products within the harvested cell culture fluid. While traditional methods like centrifugation or filtration are widely implemented for cell removal, the equipment for these processes have large footprints and operation can involve contamination risks and filter fouling. Additionally, traditional methods may not be ideal for continuous bioprocessing schemes for primary clarification. Thus, an alternate application using acoustic (sound) waves was investigated to continuously separate cells from the cell culture fluid. Presented in this study is a detailed protocol for using a bench-scale acoustic wave separator (AWS) for the primary separation of culture fluid containing a monoclonal IgG1 antibody from a CHO cell bioreactor harvest. Representative data are presented from the AWS and demonstrate how to achieve effective cell clarification and product recovery. Finally, potential applications for AWS in continuous bioprocessing are discussed. Overall, this study provides a practical and general protocol for the implementation of AWS in primary clarification for CHO cell cultures and further describes its application potential in continuous bioprocessing.

Presented here is a protocol for the primary clarification of CHO cell culture using an acoustic separator. This protocol can be used for the primary clarification of shake flask cultures or bioreactor harvests and has the potential application for continuous clarification of the cell bleed material during perfusion bioreactor operations.

Primary clarification is an essential step in a biomanufacturing process for the initial removal of cells from therapeutic products within the harvested ...

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 ...

Platelet-Derived Extracellular Vesicle Functionalization of Ti Implants

Extracellular Vesicles (EVs) are biological nanovesicles that play a key role in cell communication. Their content includes active biomolecules such as proteins and nucleic acids, which present great potential in regenerative medicine. More recently, EVs derived from Platelet Lysate (PL) have shown an osteogenic capability comparable to PL. Besides, biomaterials are frequently used in orthopedics or dental restoration. Here, we provide a method to functionalize Ti surfaces with PL-derived EVs in order to improve their osteogenic properties.
EVs are isolated from PL by size exclusion chromatography, and afterward Ti surfaces are functionalized with PL-EVs by drop casting. Functionalization is proven by EVs release and its biocompatibility by the lactate dehydrogenase (LDH) release assay.

Here, we present a method for the isolation of Extracellular Vesicles (EVs) derived from the platelet lysates (PL) and their use for coating titanium (Ti) implant surfaces. We describe the drop casting coating method, the EVs release profile from the surfaces, and in vitro biocompatibility of EVs coated Ti surfaces.

Extracellular Vesicles (EVs) are biological nanovesicles that play a key role in cell communication. Their content includes active biomolecules such as ...

Improving Reproducibility to Meet Minimal Information for Studies of Extracellular Vesicles 2018 Guidelines in Nanoparticle Tracking Analysis

Nanoparticle tracking analysis (NTA) has been one of several characterization methods used for extracellular vesicle (EV) research since 2006. Many consider that NTA instruments and their software packages can be easily utilized following minimal training and that size calibration is feasible in-house. As both NTA acquisition and software analysis constitute EV characterization, they are addressed in Minimal Information for Studies of Extracellular Vesicles 2018 (MISEV2018). In addition, they have been monitored by Transparent Reporting and Centralizing Knowledge in Extracellular Vesicle Research (EV-TRACK) to improve the robustness of EV experiments (e.g., minimize experimental variation due to uncontrolled factors).
Despite efforts to encourage the reporting of methods and controls, many published research papers fail to report critical settings needed to reproduce the original NTA observations. Few papers report the NTA characterization of negative controls or diluents, evidently assuming that commercially available products, such as phosphate-buffered saline or ultrapure distilled water, are particulate-free. Similarly, positive controls or size standards are seldom reported by researchers to verify particle sizing. The Stokes-Einstein equation incorporates sample viscosity and temperature variables to determine particle displacement. Reporting the stable laser chamber temperature during the entire sample video collection is, therefore, an essential control measure for accurate replication. The filtration of samples or diluents is also not routinely reported, and if so, the specifics of the filter (manufacturer, membrane material, pore size) and storage conditions are seldom included. The International Society for Extracellular Vesicle (ISEV)&#39;s minimal standards of acceptable experimental detail should include a well-documented NTA protocol for the characterization of EVs. The following experiment provides evidence that an NTA analysis protocol needs to be established by the individual researcher and included in the methods of publications that use NTA characterization as one of the options to fulfill MISEV2018 requirements for single vesicle characterization.

Nanoparticle tracking analysis (NTA) is a widely used method to characterize extracellular vesicles. This paper highlights NTA experimental parameters and controls plus a uniform method of analysis and characterization of samples and diluents necessary to supplement the guidelines proposed by MISEV2018 and EV-TRACK for reproducibility between laboratories.

Nanoparticle tracking analysis (NTA) has been one of several characterization methods used for extracellular vesicle (EV) research since 2006. Many ...