This work was supported by the National Science Fund for Distinguished Young Scholars (82025024);&#160;the Key project of the National Natural Science Foundation of China (82230080);&#160;the National Key R&#38;D Program of China (2021YFA1300604); the National Natural Science Foundation of China (81871735, 82272438, and 82002245);&#160;Guangdong&#160;Natural Science Fund&#160;for&#160;Distinguished Young Scholars (2023B1515020058);&#160;the Natural Science Foundation of Guangdong Province (2021A1515011639);&#160;the Major State Basic Research Development Program of Natural Science Foundation of Shandong Province in China (ZR2020ZD11);&#160;the Post-doctoral Science Foundation (2022M720059); the Outstanding Youths Development Scheme of Nanfang Hospital, Southern Medical University (2022J001).

The Ethics Committee of Nanfang Hospital, Southern Medical University, sanctioned this study, which was conducted with the informed consent of the participants. All methods employed herein adhered to the standard operating guidelines furnished by the International Human Microbiome Standards (IHMS:&#160;http://www.microbiome-standards.org/). All subsequent liquid handling procedures were mandated to be carried out within a biosafety cabinet or an ultra-clean bench.
1. Collection and aliquoting of fecal samples
<ol>
	<li>Distribute a stool sampler, a sealed bag, and an iced box, and provide comprehensive instructions to the participants on how to procure and preserve the samples.</li>
	<li>Instruct each participant to collect their fecal sample using the provided sampler, and to transport it to the lab at a temperature of 4 &#176;C within a 24-h window.</li>
	<li>Upon receipt at the lab, use a sterile spoon to aliquot less than 3.5 g of feces into a pre-weighed 50 mL centrifuge tube. Notate the fecal sample&#39;s weight on the tube for subsequent pre-treatment procedures. 
	PAUSE POINT: If immediate processing of the samples is not feasible, allocate the sample for long-term storage at -80 &#176;C after appropriate notation.</li>
</ol>
2. Feces sample preparation
<ol>
	<li>Chill 500 mL of phosphate-buffered saline (PBS) at 4 &#176;C. Filter the pre-chilled PBS through a 0.22 &#181;m Polyethersulfone (PES) filter using a 50 mL syringe into ten 50 mL centrifugation tubes.</li>
	<li>Position two 50 mL tubes on ice, each containing 3.5 g of fecal samples. Add 35 mL of PBS to each tube. 
	NOTE: Calculate the required amount of PBS for feces dissolution, ensuring a maximum sample concentration of 10% (w/v). If processing additional samples, employ more tubes as needed.</li>
	<li>Shake the samples at 300 rpm for 2 h at 4 &#176;C, or place them for 8 h at least at 4 &#176;C until the feces are completely suspended visibly.</li>
</ol>
3. Differential speed centrifugation
<ol>
	<li>Chill the high-speed refrigerated centrifuge to 4 &#176;C.</li>
	<li>Adjust the weight of the two tubes containing the samples prepared in Step 2.3 with PBS to reach a total weight of 0.1 g.</li>
	<li>Centrifuge the samples at 3, 000 &#215; g for 20 min at 4 &#176;C.</li>
	<li>Carefully pipette the supernatant into two clean 50 mL centrifuge tubes, leaving approximately 1 mL above the pellet. Use a disposable plastic Pasteur pipette for this process.</li>
	<li>Adjust the weight of the two tubes with PBS to reach a total weight within &#177; 0.1 g.</li>
	<li>Centrifuge the transferred supernatant at 12, 000 &#215; g for 30 min at 4 &#176;C.</li>
	<li>Aspirate the supernatant using a 20 mL syringe, remove the syringe needle, and filter it through 0.22 &#181;m filters into 50 mL tubes. At this point, the supernatant volume should be approximately 30 mL. 
	&#8203;NOTE: If a significant amount of impurity is still visible after the 12,000 &#215; g centrifugation, it is necessary to repeat the centrifugation step at 12,000 &#215; g for 30 min at 4 &#176;C. Failure to do so may result in a significant loss of BEVs due to pore blockage during filtration.</li>
</ol>
4. Ultra-high-speed centrifugation
<ol>
	<li>Clean the rotor and bucket by wiping them with 75% (v/v) alcohol to eliminate any residual contamination.</li>
	<li>Position a 38.5 mL ultracentrifuge tube in the tube holder. 
	NOTE: Select the appropriate ultracentrifuge tube based on the sample volume. Avoid ultraviolet radiation and unsuitable chemical reagents for sterilizing centrifugal tubes as indicated in the product manual.</li>
	<li>Transfer approximately 30 mL of the filtered fecal supernatant from Step 3.7 into the ultracentrifuge tube.</li>
	<li>Fill the ultracentrifuge tube with approximately 8 mL of PBS, leaving a 3 mm gap from the tube&#39;s opening.</li>
	<li>Place the two ultracentrifuge tubes with the samples in opposing ultracentrifugation buckets, for example, bucket 1 corresponds to 4 (1-4), 2-5, 3-6.</li>
	<li>Adjust the weight of the two buckets with PBS to achieve a total weight within &#177; 0.005 g.</li>
	<li>Install all buckets onto the rotor, regardless of whether the tubes are loaded.</li>
	<li>Initiate the vacuum and centrifuge the samples at 160,000 &#215; g for 70 min at 4 &#176;C.</li>
	<li>Release the chamber vacuum, and open the door once&#160;Ready&#160;is displayed on the&#160;Home&#160;page of the instrument.&#160;</li>
	<li>Remove the rotor from the ultracentrifuge.</li>
	<li>Move the buckets from the rotor to the rack, and retrieve the tubes using a nipper.</li>
	<li>Discard the supernatant, which will contain visible brown pellets at the bottom.</li>
	<li>Resuspend the pellets by repeatedly pipetting them up and down using a 1,000 &#181;L pipette with 1 mL of prechilled (4 &#176;C) PBS until completely suspended. Fill the tube with approximately 37 mL of PBS, leaving a 3 mm gap from the opening.</li>
	<li>Perform another ultracentrifugation following Steps 4.5-4.11 at 160,000 &#215; g for 70 min at 4 &#176;C to remove partially attached contaminating components from the tube walls.</li>
	<li>Discard the supernatant, invert the two ultracentrifuge tubes for 5 min, and clear any remaining solution on the inner wall with wipers. 
	NOTE: Cleaning the inner wall minimizes interference from residual contamination adhering to the ultracentrifuge tube wall. Choose wipers that will not influence the BEV analysis, particularly regarding particle size.</li>
	<li>Resuspend the pellets in each tube by pipetting them up and down with 1.2 mL of pre-chilled (4 &#176;C) PBS using a 1,000 &#181;L pipette.</li>
	<li>Transfer 1.2 mL of the PBS/BEV solution from one ultracentrifuge tube to a clean 1.5 mL microtube. 
	PAUSE POINT: The PBS/BEV solution collected from one ultracentrifuge tube can proceed to Step 6. Store the solution from the other tube at -80 &#176;C.</li>
</ol>
5. Solution preparation for density gradient centrifugation
<ol>
	<li>Preparation of 0.02 M HEPES buffer
	<ol>
		<li>Combine 0.477 g of HEPES powder and 0.8 g of NaCl with 90 mL of autoclaved deionized water.</li>
		<li>Adjust the pH to 7.2 by adding 1 M sodium hydroxide (NaOH). Bring the volume to 100 mL with deionized water and filter the solution through 0.22 &#181;m PES membranes. 
		CAUTION: Sodium hydroxide is a strong caustic alkali. Operators must handle and prepare it carefully in a fume cupboard.</li>
	</ol>
	</li>
	<li>Preparation of density gradient buffer
	<ol>
		<li>Calculate the required volume of each density gradient buffer based on the number of ultracentrifuge tubes for density gradient centrifugation (Step 6) (In this protocol, the volumes for 60%, 50%, 40%, 20%, and 10% gradient solutions were 2.5 mL, 3 mL, 6 mL, 6 mL, and 6 mL, respectively).</li>
		<li>For the preparation of a 50% (w/v) iodixanol working solution, mix the 0.02 M HEPES buffer and 60% (w/v) iodixanol stock solution in a volume ratio of 1:5 (0.5 mL:2.5 mL). 
		NOTE: Use a disposable 20 mL syringe to withdraw the iodixanol stock solution and avoid introducing air.</li>
		<li>To prepare iodixanol buffers with different concentrations, combine the 50% iodixanol working solution from Step 5.2.2 with the HEPES buffer obtained in Step 5.1 using a 1,000 &#181;L pipette according to the proportions shown in Table 1. 
		&#8203;NOTE: Perform Step 5 on an ultra-clean bench with the lights off. Store the opened iodixanol stock solution in the refrigerator at 4 &#176;C to prevent bacterial growth. Keep the iodixanol solution and HEPES buffer protected from light.</li>
	</ol>
	</li>
</ol>
6. Establishment of a density gradient centrifugation system
<ol>
	<li>Top-down DGC mode
	<ol>
		<li>Combine 500 &#181;L of PBS/BEV solution isolated from Step 4 with 3 mL of PBS. Gently mix the solutions using a 1,000 &#181;L pipette to obtain 3.5 mL of PBS/BEV solution.</li>
		<li>Place a 31 mL ultracentrifuge tube on a foam plate with holes and label it as &#34;&#8595;&#34;.</li>
		<li>Vertically add 3 mL of the 50% iodixanol solution to the bottom of the tube using a 1,000 &#181;L pipette.</li>
		<li>Tilt the tube to a 70&#176; angle and position a tube holder or other support slightly above the level of the foam plate, below the opening of the tube.</li>
		<li>Add 3 mL of 40% iodixanol solution on top of the 50% iodixanol solution using a 1,000 &#181;L pipette.</li>
		<li>Add 3 mL of 20% iodixanol solution on top of the 40% iodixanol solution using a 1,000 &#181;L pipette.</li>
		<li>Add 3 mL of 10% iodixanol solution on top of the 20% iodixanol solution using a 1,000 &#181;L pipette.</li>
		<li>Add 3.5 mL of PBS/BEV solution from Step 6.1.1 on top of the 10% iodixanol solution using a 1,000 &#181;L pipette.</li>
		<li>Gently return the tubes to an upright position. 
		NOTE: After pipetting, the stratification should be visible.</li>
	</ol>
	</li>
	<li>Bottom-up DGC mode
	<ol>
		<li>Place a 31 mL ultracentrifuge tube on a foam plate with holes and label it as &#34;&#8593;&#34;.</li>
		<li>Vertically add 2.5 mL of the 60% iodixanol stock solution to the bottom of the tube. Mix it gently with 500 &#181;L of PBS/BEV solution isolated from Step 4 using a 1,000 &#181;L pipette to obtain 3 mL of 50% iodixanol/BEV solution.</li>
		<li>Tilt the tube to a 70&#176; angle and position a tube holder or other support slightly above the level of the foam plate, below the opening of the tube.</li>
		<li>Add 3 mL of the 40% iodixanol solution on top of the 50% iodixanol/BEV solution using a 1,000 &#181;L pipette.</li>
		<li>Add 3 mL of the 20% iodixanol solution on top of the 40% iodixanol solution using a 1000 &#181;L pipette.</li>
		<li>Add 3 mL of the 10% iodixanol solution on top of the 20% iodixanol solution using a 1000 &#181;L pipette.</li>
		<li>Add 3.5 mL of PBS on top of the 10% iodixanol solution using a 1,000 &#181;L pipette.</li>
		<li>Gently return the tubes to an upright position.</li>
		<li>At this point, 200 &#181;L of the suspension obtained in Step 4.17 remained, which can be analyzed subsequently as a group named &#34;UC&#34;. 
		&#8203;NOTE: When transferring solutions in Step 6, always keep the pipette tip against the ultracentrifuge tube wall perpendicular to the axis of the tube.</li>
	</ol>
	</li>
</ol>
7. Density gradient centrifugation and fraction collection
<ol>
	<li>Adjust the weight of the two tubes with PBS to achieve a total weight within &#177; 0.005 g.</li>
	<li>Place the tubes in the buckets according to Step 4.5 and subject them to centrifugation at 160,000 &#215; g for 7 h at 4 &#176;C.</li>
	<li>Collect the fractions using a pipettor (1000 &#181;L) from top to bottom against the side wall in the following order: 3 mL, 2 mL, 1 mL, 1 mL, 1 mL, 1 mL, 1 mL, 1 mL, 1.5 mL, and 3 mL into a 38.5 mL ultracentrifuge tube. 
	&#8203;NOTE: Maintain the line of sight at the same level as the liquid surface.</li>
	<li>Conduct ultracentrifugation for each fraction obtained in Step 7.3 according to Step 4 at 160,000 &#215; g for 70 min at 4 &#176;C.</li>
	<li>Remove the supernatant and invert the ultracentrifuge tubes for 5 min.</li>
	<li>Resuspend the pellets in 200 &#181;L of pre-chilled (4 &#176;C) PBS using a 200 &#181;L pipette.</li>
	<li>Transfer the PBS/BEV solution into a clean 1.5 mL microtube.</li>
	<li>Label the fractions from F1 to F10 and analyze them based on Step 8. 
	PAUSE POINT: If the samples obtained in Step 7.8 cannot be processed immediately, store them at -80 &#176;C.</li>
</ol>
8. Characterization and quantitative analysis of the collected fractions
<ol>
	<li>Determine the absorbance values (OD 340 nm) of each fraction using a microplate detector with a blank control to calculate their corresponding density. 
	NOTE: Replace PBS/BEV solution (Step 6.1.1 and Step 6.2.2) with PBS to establish a blank control.
	<ol>
		<li>Prepare 100 &#181;L each of 0%, 5%, 10%, 12.5%, and 20% iodixanol solution diluted by 0.02 M HEPES based on 50% stock solution (Step 5.2.2) as the standards, corresponding to densities of 1.0058 g/mL, 1.0318 g/mL, 1.058 g/mL, 1.0708 g/mL, and 1.111 g/mL, respectively.</li>
		<li>Add 50 &#181;L of each fraction from the blank control (prepared in Step 7.3) and 50 &#181;L of each standard solution (prepared in Step 8.1.1) to duplicate wells of a 96-well plate.</li>
		<li>Set the wavelength to 340 nm, measure the endpoint optical density, and calculate the density of each fraction.</li>
		<li>Identify F4-F9 as the range of BEV fractions based on their density.</li>
	</ol>
	</li>
	<li>Determine the protein concentration of the BEV fractions using the bicinchoninic acid assay (BCA).
	<ol>
		<li>Dilute the substance tenfold in PBS provided by the manufacturer to prepare a 0.5 &#181;g/&#181;L standard protein solution.</li>
		<li>Add the standard protein solution (prepared in Step 8.2.1) with varying volumes according to the reagent instructions, and supplement each well of a 96-well plate with PBS to a total volume of 20 &#181;L.</li>
		<li>Add 20 &#181;L of the samples prepared in Step 7.8 and the samples left after UC defined in Step 6.2.9 to each well.</li>
		<li>Mix reagents A and B in a ratio of 50:1, and transfer 200 &#181;L to each well.</li>
		<li>Incubate the plate in a 37 &#176;C water bath for 30 min.</li>
		<li>Measure the absorbance value using a microplate reader, calculate the protein concentration (&#181;g/&#181;L), and determine the protein content (&#181;g) of the samples based on the standard curve (x: optical density; y: protein concentration) generated from the values of the standard solutions diluted in Step 8.2.2.</li>
	</ol>
	</li>
	<li>Characterize the BEV fractions defined in Step 8.1.4 using transmission electron microscopy (TEM) to verify the presence of BEVs. All procedures below should be performed on ice.
	<ol>
		<li>Apply 10 &#181;L of each isolated F4-F9 fraction from Step 7.8 and the samples left after UC defined in Step 6.2.9 onto Formvar/Carbon supported copper grids for 20 min, and blot with filter paper.</li>
		<li>Rinse the samples with 100 &#181;L of PBS three times for 1 min each, and blot with filter paper.</li>
		<li>Fix the samples with 100 &#181;L of 1% (w/v) glutaraldehyde for 5 min and blot with filter paper.</li>
		<li>Wash the grids with 100 &#181;L of PBS ten times for 2 min each, and blot with filter paper.</li>
		<li>Stain the grids with 50 &#181;L of 1.5% (w/v) uranyl acetate for 10 min, and blot with filter paper. 
		CAUTION: Uranyl acetate is radioactive and extremely toxic when in contact with the skin or inhaled. Handle solutions with uranyl acetate in a fume cupboard and follow appropriate safety measures.</li>
		<li>Transfer the grids onto a 1% (w/v) methylcellulose drop for 5 min and blot with filter paper.</li>
		<li>Store the air-dried grids in a dark, dust-free environment until observation.</li>
		<li>Capture images and analyze them using a TEM.</li>
	</ol>
	</li>
	<li>Characterize the BEV fractions defined in Step 8.1.4 using nanoparticle tracking analysis (NTA) to count the number of particles.
	<ol>
		<li>Calibrate the system using 110 nm polystyrene particles.</li>
		<li>Wash the sample pool using PBS.</li>
		<li>Dilute the samples from different fractions acquired in Step 7.8 and the samples left after UC defined in Step 6.2.9 to a concentration in the range of 105-109 particles/mL, with an optimized concentration of 107 particles/mL.</li>
		<li>Record and analyze at 11 positions with three replications, maintaining the temperature around 23-30 &#176;C.</li>
	</ol>
	</li>
	<li>Analyze the protein contents of the samples by coomassie brilliant blue staining (CBBS) and western blotting (WB).
	<ol>
		<li>Dilute the BEVs samples from different fractions obtained in Step 7.8 and the samples left after UC defined in Step 6.2.9 using PBS and 5 &#215; loading buffer to a final concentration of 0.5 or 1 &#181;g/&#181;L, if quantitation is needed, ensuring sufficient sample loading of 20 &#181;L (10 &#181;g or 20 &#181;g) of each well in Step 8.5.2. Boil the samples at 95 &#176;C for 10 min.</li>
		<li>Assemble the prefabricated polyacrylamide gel into the electrophoresis tank, and transfer the samples to the wells.</li>
		<li>Conduct vertical electrophoresis with the following parameters: 160 V for 50 min.</li>
		<li>Stain the gel in Coomassie brilliant blue solution for 1 h, rinse in deionized water until the blue background lightens, and capture images with a camera.</li>
		<li>Perform protein blotting procedures for other gels with the following parameters: 400 mA for 20 min.</li>
		<li>Block the blotting membranes with a 5% (w/v) BSA solution for 1 h at room temperature.</li>
		<li>Wash the membranes in TBST buffer three times for 5 min each.</li>
		<li>Incubate the membranes with primary antibodies (BEV markers: LPS, OmpA, LTA; EEV markers: CD63, CD9, TSG-101, Syntenin, Integrin &#946;1; Other contamination markers: Flagellin, Calnexin) for 8-12 h at 4 &#176;C.</li>
		<li>Wash the membranes in TBST buffer three times for 10 min each.</li>
		<li>Incubate the membranes with secondary antibodies for 1 h at room temperature.</li>
		<li>Perform Step 8.5.9.</li>
		<li>Develop the blotting using a chemiluminescence apparatus.</li>
		<li>Replace PBS/BEV solution (Step 6.1.1 and Step 6.2.2) with BEVs from Escherichia coli to establish a positive control (see Table of Materials for the procedure of isolating crude E. coli-BEVs), and conduct all the above experiments for characterizing BEVs. 
		PAUSE POINT: Determine F6-F8 as the distribution of BEV-enriched fractions based on the results of the characterization described above for fecal BEVs, and utilize this narrowed range for subsequent analysis.</li>
	</ol>
	</li>
	<li>Calculate the recovery rates in terms of particles and proteins to assess the efficiency of the two DGC modes. The results below are presented as percentages.
	<ol>
		<li>Calculate the recovery rates of particles in the Top-down mode: total particles in F6-F8/particles after UC defined in Step 6.2.9.</li>
		<li>Calculate the recovery rates of particles in the Bottom-up mode: total particles in F6-F8/particles after UC defined in Step 6.2.9.</li>
		<li>Calculate the recovery rates of proteins in the Top-down mode: total protein contents in F6-F8 (&#181;g)/protein content after UC defined in Step 6.2.9 (&#181;g).</li>
		<li>Calculate the recovery rates of proteins in the Bottom-up mode: total protein contents in F6-F8 (&#181;g)/protein content after UC defined in Step 6.2.9 (&#181;g).</li>
	</ol>
	</li>
	<li>Calculate the particle/protein ratio to assess the purity traditionally defined of UC and the two DGC modes, and the results below were all presented as percentages.
	<ol>
		<li>Particle/protein ratio after UC defined in Step 6.2.9: particles after UC/protein content after UC (&#181;g).</li>
		<li>Particle/protein ratio in the Top-down mode: total particles in F6-F8/protein content in F6-F8 (&#181;g).</li>
		<li>Particle/protein ratio in the Bottom-up mode: total particles in F6-F8/protein content in F6-F8 (&#181;g).</li>
	</ol>
	</li>
	<li>Select the most suitable centrifugation mode (Top-down mode was selected in the protocol) for fecal BEV purification and future analysis.</li>
</ol>

Enhanced Bacterial Extracellular Vesicles Isolation for Microbiota Multi-Omics

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E., 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=Microbial+changes+in+stool,+saliva,+serum,+and+urine+before+and+after+anti-TNF-&#945;+therapy+in+patients+with+inflammatory+bowel+diseases.">Microbial changes in stool, saliva, serum, and urine before and after anti-TNF-&#945; therapy in patients with inflammatory bowel diseases.</a> Scientific Reports. 12 (1), 6359 (2022).</li><li>Tulkens, J., De Wever, O., Hendrix, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analyzing+bacterial+extracellular+vesicles+in+human+body+fluids+by+orthogonal+biophysical+separation+and+biochemical+characterization.">Analyzing bacterial extracellular vesicles in human body fluids by orthogonal biophysical separation and biochemical characterization.</a> Nature Protocols. 15 (1), 40-67 (2020).</li><li>Tulkens, 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=Increased+levels+of+systemic+LPS-positive+bacterial+extracellular+vesicles+in+patients+with+intestinal+barrier+dysfunction.">Increased levels of systemic LPS-positive bacterial extracellular vesicles in patients with intestinal barrier dysfunction.</a> Gut. 69 (1), 191-193 (2020).</li><li>Kang, C. S., 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=Extracellular+vesicles+derived+from+gut+microbiota,+especially+Akkermansia+muciniphila,+protect+the+progression+of+dextran+sulfate+sodium-induced+colitis.">Extracellular vesicles derived from gut microbiota, especially Akkermansia muciniphila, protect the progression of dextran sulfate sodium-induced colitis.</a> PloS one. 8 (10), 76520 (2013).</li><li>Simonsen, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+are+we+looking+at?+Extracellular+vesicles,+lipoproteins,+or+both.">What are we looking at? Extracellular vesicles, lipoproteins, or both.</a> Circulation Research. 121 (8), 920-922 (2017).</li><li>Correll, V. L., 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=Optimization+of+small+extracellular+vesicle+isolation+from+expressed+prostatic+secretions+in+urine+for+in-depth+proteomic+analysis.">Optimization of small extracellular vesicle isolation from expressed prostatic secretions in urine for in-depth proteomic analysis.</a> Journal of Extracellular Vesicles. 11 (2), 12184 (2022).</li><li>Liang, X., 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=Gut+bacterial+extracellular+vesicles:+important+players+in+regulating+intestinal+microenvironment.">Gut bacterial extracellular vesicles: important players in regulating intestinal microenvironment.</a> Gut Microbes. 14 (1), 2134689 (2022).</li><li>Alberti, G., 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=Extracellular+vesicles+derived+from+gut+microbiota+in+inflammatory+bowel+disease+and+colorectal+cancer.">Extracellular vesicles derived from gut microbiota in inflammatory bowel disease and colorectal cancer.</a> Biomedical papers of the Medical Faculty of the University Palacky, Olomouc, Czech Republic. 165 (3), 233-240 (2021).</li><li>D&#237;ez-Sainz, E., Milagro, F. I., Riezu-Boj, J. I., Lorente-Cebri&#225;n, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+gut+microbiota-derived+extracellular+vesicles+on+obesity+and+diabetes+and+their+potential+modulation+through+diet.">Effects of gut microbiota-derived extracellular vesicles on obesity and diabetes and their potential modulation through diet.</a> Journal of Physiology and Biochemistry. 78 (2), 485-499 (2022).</li><li>Lajqi, T., 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=Gut+microbiota-derived+small+extracellular+vesicles+endorse+memory-like+inflammatory+responses+in+murine+neutrophils.">Gut microbiota-derived small extracellular vesicles endorse memory-like inflammatory responses in murine neutrophils.</a> Biomedicines. 10 (2), 442 (2022).</li><li>Lee, K. E., 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=The+extracellular+vesicle+of+gut+microbial+Paenalcaligenes+hominis+is+a+risk+factor+for+vagus+nerve-mediated+cognitive+impairment.">The extracellular vesicle of gut microbial Paenalcaligenes hominis is a risk factor for vagus nerve-mediated cognitive impairment.</a> Microbiome. 8 (1), 107 (2020).</li><li>Villard, A., Boursier, J., Andriantsitohaina, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+and+eukaryotic+extracellular+vesicles+and+nonalcoholic+fatty+liver+disease:+new+players+in+the+gut-liver+axis.">Bacterial and eukaryotic extracellular vesicles and nonalcoholic fatty liver disease: new players in the gut-liver axis.</a> American Journal of Physiology-Gastrointestinal and Liver Physiology. 320 (4), G485-G495 (2021).</li><li>Wei, S., 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=Outer+membrane+vesicles+enhance+tau+phosphorylation+and+contribute+to+cognitive+impairment.">Outer membrane vesicles enhance tau phosphorylation and contribute to cognitive impairment.</a> Journal of Cellular Physiology. 235 (5), 4843-4855 (2020).</li><li>Bitto, N. J., Kaparakis-Liaskos, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+of+bacterial+membrane+vesicle+production,+purification,+quantification,+and+examination+of+their+immunogenic+functions.">Methods of bacterial membrane vesicle production, purification, quantification, and examination of their immunogenic functions.</a> Methods in Molecular Biology. 2523, 43-61 (2022).</li><li>Stentz, R., Miquel-Clop&#233;s, A., Carding, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Production,+isolation,+and+characterization+of+bioengineered+bacterial+extracellular+membrane+vesicles+derived+from+Bacteroides+thetaiotaomicron+and+their+use+in+vaccine+development.">Production, isolation, and characterization of bioengineered bacterial extracellular membrane vesicles derived from Bacteroides thetaiotaomicron and their use in vaccine development.</a> Methods in Molecular Biology. 2414, 171-190 (2022).</li><li>Zhang, Q., Jeppesen, D. K., Higginbotham, J. N., Franklin, J. L., Coffey, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comprehensive+isolation+of+extracellular+vesicles+and+nanoparticles.">Comprehensive isolation of extracellular vesicles and nanoparticles.</a> Nature Protocols. 18 (5), 1462-1487 (2023).</li><li>Iwai, K., Minamisawa, T., Suga, K., Yajima, Y., Shiba, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+human+salivary+extracellular+vesicles+by+iodixanol+density+gradient+ultracentrifugation+and+their+characterizations.">Isolation of human salivary extracellular vesicles by iodixanol density gradient ultracentrifugation and their characterizations.</a> Journal of Extracellular Vesicles. 5, 30829 (2016).</li><li>Vandeputte, D., 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=Stool+consistency+is+strongly+associated+with+gut+microbiota+richness+and+composition,+enterotypes+and+bacterial+growth+rates.">Stool consistency is strongly associated with gut microbiota richness and composition, enterotypes and bacterial growth rates.</a> Gut. 65 (1), 57-62 (2016).</li><li>Wen, M., 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=Bacterial+extracellular+vesicles:+A+position+paper+by+the+Microbial+Vesicles+Task+Force+of+the+Chinese+Society+of+Extracellular+Vesicles.">Bacterial extracellular vesicles: A position paper by the Microbial Vesicles Task Force of the Chinese Society of Extracellular Vesicles.</a> Interdisciplinary Medicine. 1, 12046 (2023).</li></ol>

The authors declare no conflicting interests.

Bacterial extracellular vesicles (BEVs) are lipid-bilayer nanoparticles secreted by bacteria, carrying a wealth of proteins, lipids, nucleic acids, and other bioactive molecules, contributing to mediating the functional effects of bacteria20. BEVs derived from the gut have been verified to be involved in the development of diseases, such as inflammatory bowel disease, Crohn&#39;s disease, and colorectal cancer, and also affect general metabolism and mediate impaired cognitive function<sup class="x...

The gut is widely recognized as the organ harboring the most abundant microbial communities in the human body, with over 90% of bacteria involved in colonization and multiplication1,2. Extensive evidence has demonstrated that the gut microbiota modulates the gut microenvironment and simultaneously interacts with dysfunction in distant organs, primarily through an impaired intestinal barrier3,4. Mounting evidence indicates a correlation between the imbalance of gut microbiota and the progression of inflammatory bowel disease (IBD)5,6, as well as cognitive disorders through the gut-brain axis5,6,7,8. Bacterial extracellular vesicles (BEVs) produced by bacteria play significant roles in these pathological processes.
BEVs are nanoscale particles encapsulating bacterial derivatives, with diameters ranging from 20 to 400 nm. They have been demonstrated to facilitate interactions between bacteria and their host organisms9,10. Despite their invisibility, these particles have garnered increasing attention from researchers due to their prospective broad applications as diagnostic biomarkers, therapeutic targets, and drug delivery vehicles11. Human feces, often used as biospecimens for studying BEVs, predominantly sourced from gut bacteria, contain a complex mixture of water, bacteria, lipids, proteins, undigested food residue, and exfoliated epithelial cells among others. The intricate fecal composition poses challenges to the isolation and purity of BEVs, thereby impeding a comprehensive, objective, and realistic analysis of BEVs. Hence, effective strategies to minimize interference from contaminating components and enhance the yield of BEVs have emerged as critical issues warranting immediate attention.
Existing isolation strategies largely rely on techniques such as ultra-high-speed centrifugation (UC), density gradient centrifugation (DGC), and size exclusion chromatography (SEC)12,13,14,15,16,17. Currently, DGC is one of the most widely applied methods in the field of BEV separation, encompassing two sedimentation-floating modes, &#34;Top-down&#34; and &#34;Bottom-up&#34;, which are determined by the initial loading position of the sample. These methodologies differentiate extracellular vesicles (EVs) from other components based on size and density disparities, yielding variable purity and recovery rates. Prior research has indicated that single-approach strategies are insufficient for adequately separating EVs from soluble proteins in body fluid samples, such as lipoprotein in blood18 and Tamm-Horsfall protein in urine19. Additionally, the size distribution of eukaryotic extracellular vesicles (EEVs) often overlaps with that of BEVs, thereby necessitating further methodological enhancements to optimize BEV yield. Consequently, advancing the study of BEVs hinges on the development of effective separation and purification methodologies. Notably, Tulkens et al15 employed an orthogonal biophysical strategy to separate fecal BEVs from EEVs, in which the centrifugation time of a Bottom-up DGC mode was up to 18 h. In contrast, this study reduced it to 7 h, greatly saving the gradient-ultracentrifugation time and simplifying the process.
In the present study, we isolated and purified fecal BEVs employing two DGC modes under optimized buffer conditions, after enriching BEVs with a range of differential centrifugation speeds, from low to extremely high velocity. Evaluations based on morphology, particle size, and concentration indicated a commendable performance by this enhanced method. This study could serve as the foundation for future research, extending its applications to a broader domain, and offering insights into the heterogeneity of BEVs within the human body. It also contributes to the standardization of BEV separation and analysis techniques.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 % (w/v) glutaraldehyde (prepared from 2.5 % stock solution in deionized water)</td>
			<td>ACMEC</td>
			<td>AP1126</td>
			<td>Morphological observation for BEVs using TEM at Step 8.3.3</td>
		</tr>
		<tr>
			<td>1 % (w/v) methylcellulose (prepared from original powder in deionized water)</td>
			<td>Sigma-Aldrich</td>
			<td>M7027</td>
			<td>Morphological observation for BEVs using TEM at Step 8.3.6</td>
		</tr>
		<tr>
			<td>1.5 % (w/v) uranyl acetate (prepared from original powder in deionized water)</td>
			<td>Polysciences</td>
			<td>21447-25</td>
			<td>Morphological observation for BEVs using TEM at Step 8.3.5</td>
		</tr>
		<tr>
			<td>1000 &#956;L, 200 &#956;L, 10 &#956;L Pipette</td>
			<td>KIRGEN</td>
			<td>KG1313, KG1212, KG1011</td>
			<td>Transfer the solution</td>
		</tr>
		<tr>
			<td>5 % (w/v) bovine serum albumin solution (prepared from the original powder in TBST buffer)</td>
			<td>Fdbio science</td>
			<td>FD0030</td>
			<td>Used in western blotting for blocking at Step 8.5.6</td>
		</tr>
		<tr>
			<td>5 &#215; loading buffer</td>
			<td>Fdbio science</td>
			<td>FD006</td>
			<td>Used in western blotting and Coomassie brilliant blue stain at Step 8.5.1</td>
		</tr>
		<tr>
			<td>75 % (v/v) alcohol</td>
			<td>LIRCON</td>
			<td>LIRCON-500 mL</td>
			<td>Surface disinfection</td>
		</tr>
		<tr>
			<td>96-well plate</td>
			<td>Rar</td>
			<td>A8096</td>
			<td>Measure the absorbance values&#160;</td>
		</tr>
		<tr>
			<td>Anti-Calnexin antibody</td>
			<td>Abcam</td>
			<td>ab92573</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-CD63 antibody</td>
			<td>Abcam</td>
			<td>ab134045</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-CD9 antibody</td>
			<td>Abcam</td>
			<td>ab236630</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-Flagellin antibody</td>
			<td>Sino Biological</td>
			<td>40067-MM06</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-Integrin beta 1 antibody</td>
			<td>Abcam</td>
			<td>ab30394</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-LPS antibody</td>
			<td>Thermo Fisher</td>
			<td>MA1-83152</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-LTA antibody</td>
			<td>Thermo Fisher&#160;</td>
			<td>MA1-7402</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-OmpA antibody</td>
			<td>CUSABIO</td>
			<td>CSB-PA359226ZA01EOD, https://www.cusabio.com/</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-Syntenin antibody</td>
			<td>Abcam</td>
			<td>ab133267</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Anti-TSG101 antibody</td>
			<td>Abcam</td>
			<td>ab125011</td>
			<td>Western blotting (Primary Antibody)</td>
		</tr>
		<tr>
			<td>Autoclave</td>
			<td>ZEALWAY</td>
			<td>GR110DP</td>
			<td>Sterilization for supplies and mediums used in the experiment</td>
		</tr>
		<tr>
			<td>Balance</td>
			<td>Mettler Toledo</td>
			<td>AL104</td>
			<td>Balance the tube sample-loaded with PBS</td>
		</tr>
		<tr>
			<td>Bicinchoninic acid assay&#160;</td>
			<td>Fdbio science</td>
			<td>FD2001</td>
			<td>Measure protein content of BEVs at Step 8.2</td>
		</tr>
		<tr>
			<td>BioRender</td>
			<td>BioRender</td>
			<td>https://app.biorender.com</td>
			<td>Make the schematic workflow of BEVs isolation and purification showed in Figure 1</td>
		</tr>
		<tr>
			<td>Biosafety cabinet</td>
			<td>Haier</td>
			<td>HR1200- II B2</td>
			<td>Peform the procedures about feces sample handling</td>
		</tr>
		<tr>
			<td>Centrifuge 5810 R; Rotor F-34-6-38</td>
			<td>Eppendorf</td>
			<td>5805000092; 5804727002, adapter: 5804774000</td>
			<td>Preprocess for BEVs (Step 3)</td>
		</tr>
		<tr>
			<td>Chemiluminescence Apparatus</td>
			<td>BIO-OI</td>
			<td>OI600SE-MF</td>
			<td>Used in western blotting for signal detection at Step 8.5.12</td>
		</tr>
		<tr>
			<td>Cytation 5</td>
			<td>BioTek</td>
			<td>F01</td>
			<td>Microplate detector for measuring the absorbance (Step 8.1) and fluorescence (Figure 6) values&#160;</td>
		</tr>
		<tr>
			<td>Dil-labled low density lipoprotein</td>
			<td>ACMEC</td>
			<td>AC12038</td>
			<td>Definition of distribution of interfering components&#160;</td>
		</tr>
		<tr>
			<td>Electrophoresis equipment</td>
			<td>Bio-rad</td>
			<td>1658033</td>
			<td>Used in western blotting for protein separation and transfer at Step 8.5.2, 8.5.3, 8.5.5</td>
		</tr>
		<tr>
			<td>Enhanced Chemiluminescence kit HRP&#160;</td>
			<td>Fdbio science</td>
			<td>FD8020</td>
			<td>Used in western blotting for signal detection at Step 8.5.12</td>
		</tr>
		<tr>
			<td>Escherichia coli&#160;</td>
			<td>American Type Culture Collection</td>
			<td>ATCC8739</td>
			<td>Isolate BEVs as a positive control. Protocol: Dissolve 25 g of the LB powder in 1 L deionized water, and autoclave. Transfer the 800 &#956;L of preserved Escherichia coli into the medium. Cultivate at 37 &#176;C in the incubator shaker. Then centrifuge at 3, 000 &#215; g for 20 min at 4 &#176;C, 12, 000 &#215; g for 30 min at 4 &#176;C, filter the supernatant through 0.22 &#956;m membrane, and perform ultra-speed centrifugation at 160, 000 &#215; g for 70 min at 4 &#176;C. Pellet defined as crude BEVs from Escherichia coli was suspended in 1.2 mL PBS (Step 3, 4).&#160;&#160;&#160;&#160;</td>
		</tr>
		<tr>
			<td>Falcon tubes 50 mL</td>
			<td>KIRGEN</td>
			<td>KG2811</td>
			<td>Preprocess for BEVs (Step 3)</td>
		</tr>
		<tr>
			<td>Feto Protein Staining Buffer</td>
			<td>Absci</td>
			<td>ab.001.50</td>
			<td>Coomassie brilliant blue staining at Step 8.5.4</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>Biosharp</td>
			<td>BS-TFP-070B</td>
			<td>Morphological observation for BEVs using TEM at Step 8.3 (Blotting the solution)</td>
		</tr>
		<tr>
			<td>Formvar/Carbon supported copper grids&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>TEM-FCF200CU50</td>
			<td>Morphological observation for BEVs using TEM at Step 8.3</td>
		</tr>
		<tr>
			<td>HEPES powder</td>
			<td>Meilunbio</td>
			<td>MB6078</td>
			<td>Prepare iodixanol buffers with different concentrations for density gradient centrifugation</td>
		</tr>
		<tr>
			<td>HRP AffiniPure Goat Anti-Mouse IgG (H+L)</td>
			<td>Fdbio science</td>
			<td>FDM007</td>
			<td>Western blotting (Secondary Antibody)</td>
		</tr>
		<tr>
			<td>HRP AffiniPure Goat Anti-Rabbit IgG (H+L)</td>
			<td>Fdbio science</td>
			<td>FDR007</td>
			<td>Western blotting (Secondary Antibody)</td>
		</tr>
		<tr>
			<td>Incubator shaker</td>
			<td>Qiangwen</td>
			<td>DHZ-L</td>
			<td>Cultivate Escherichia coli&#160;</td>
		</tr>
		<tr>
			<td>Kimwipes&#8482; Delicate Task Wipes</td>
			<td>Kimtech Science</td>
			<td>34155</td>
			<td>Wipe the inner wall of the ultracentrifuge tube at Step 4.15</td>
		</tr>
		<tr>
			<td>LB broth</td>
			<td>Hopebio</td>
			<td>HB0128</td>
			<td>Cultivate Escherichia coli&#160;</td>
		</tr>
		<tr>
			<td>Low temperature freezer (-80 &#176;C)</td>
			<td>Haier</td>
			<td>DW-86L338J</td>
			<td>Store the samples</td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Alalddin</td>
			<td>M116118</td>
			<td>Used in western blotting for activating PVDF membrane at Step 8.5.5</td>
		</tr>
		<tr>
			<td>Micro tubes 1.5 mL</td>
			<td>KIRGEN</td>
			<td>KG2211</td>
			<td>Recover fractions after density gradient centrifugation</td>
		</tr>
		<tr>
			<td>Micro tubes 2 mL</td>
			<td>KIRGEN</td>
			<td>KG2911</td>
			<td>Recover fractions after density gradient centrifugation</td>
		</tr>
		<tr>
			<td>Micro tubes 5 mL</td>
			<td>BBI</td>
			<td>F610888-0001</td>
			<td>Recover fractions after density gradient centrifugation</td>
		</tr>
		<tr>
			<td>Microplate reader&#160;</td>
			<td>Thermo Fisher&#160;</td>
			<td>Multiskan MK3</td>
			<td>Measure protein content of BEVs at Step 8.2</td>
		</tr>
		<tr>
			<td>Millipore filter 0.22 &#956;m</td>
			<td>Merck millipore</td>
			<td>SLGP033RB</td>
			<td>Filtration sterilization; Material: polyethersulfone, PES</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>GHTECH</td>
			<td>1.01307.040</td>
			<td>Density gradient centrifugation solution</td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>GHTECH</td>
			<td>1.01394.068</td>
			<td>Density gradient centrifugation solution (pH adjustment)</td>
		</tr>
		<tr>
			<td>Optima&#8482; XPN-100</td>
			<td>Beckman Coulter</td>
			<td>A94469</td>
			<td>Ultracentrifugation for BEVs isolation at Step 4, 7</td>
		</tr>
		<tr>
			<td>OptiPrep&#8482;</td>
			<td>Serumwerk Bernburg AG</td>
			<td>1893</td>
			<td>Density gradient centrifugation stock solution</td>
		</tr>
		<tr>
			<td>Orbital Shaker</td>
			<td>Youning</td>
			<td>CS-100</td>
			<td>Dissolve feces at Step 2</td>
		</tr>
		<tr>
			<td>Phosphate buffered saline</td>
			<td>Procell</td>
			<td>PB180327</td>
			<td>Dissolve feces at Step 2</td>
		</tr>
		<tr>
			<td>Pipettor</td>
			<td>Eppendorf</td>
			<td>3120000267, 3120000259</td>
			<td>Transfer the solution</td>
		</tr>
		<tr>
			<td>Plastic pasteur pipette</td>
			<td>ABCbio</td>
			<td>ABC217003-4</td>
			<td>Remove supernatant in preprocessing at Step 3.4</td>
		</tr>
		<tr>
			<td>Polyvinylidene difluoride (PVDF) membranes</td>
			<td>Millipore</td>
			<td>ISEQ00010, IPVH00010</td>
			<td>Used in western blotting for protein transfer at Step 8.5.5</td>
		</tr>
		<tr>
			<td>Prefabricated polyacrylamide gel, 4&#8211;20% 15 Wells</td>
			<td>ACE</td>
			<td>F15420Gel</td>
			<td>Used in western blotting for protein separation at Step 8.5.2, 8.5.3</td>
		</tr>
		<tr>
			<td>Primary antibody diluent</td>
			<td>Fdbio science</td>
			<td>FD0040</td>
			<td>Used in western blotting at Step 8.5.8</td>
		</tr>
		<tr>
			<td>Protein ladder</td>
			<td>Fdbio science</td>
			<td>FD0672</td>
			<td>Used in western blotting and Coomassie brilliant blue stain at Step 8.5</td>
		</tr>
		<tr>
			<td>Rapid protein blotting solution</td>
			<td>UBIO</td>
			<td>UW0500</td>
			<td>Used in western blotting for protein transfer at Step 8.5.5</td>
		</tr>
		<tr>
			<td>Rotor SW 32 Ti Swinging-Bucket Rotor</td>
			<td>Beckman Coulter</td>
			<td>369650</td>
			<td>Ultracentrifugation for BEVs isolation at Step 4, 7</td>
		</tr>
		<tr>
			<td>Syringe 20 mL, 50 mL&#160;</td>
			<td>Jetway</td>
			<td>ZSQ-20ML, YCXWJZSQ-50 mL</td>
			<td>Transfer buffers amd remove supernatant in preprocessing</td>
		</tr>
		<tr>
			<td>TBS powder</td>
			<td>Fdbio science</td>
			<td>FD1021</td>
			<td>Used in western blotting at Step 8.5</td>
		</tr>
		<tr>
			<td>Transmission electron microscope (TEM)</td>
			<td>Hitachi&#160;</td>
			<td>H-7650</td>
			<td>Morphological observation for BEVs at Step 8.3</td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Fdbio science</td>
			<td>FD0020</td>
			<td>Used in western blotting at Step 8.5</td>
		</tr>
		<tr>
			<td>Ultracentrifuge tube</td>
			<td>Beckman</td>
			<td>326823, 355642</td>
			<td>Ultracentrifugation for BEVs isolation at Step 4, 7</td>
		</tr>
		<tr>
			<td>Ultra-clean bench</td>
			<td>AIRTECH</td>
			<td>SW-CJ-2FD</td>
			<td>Peform the procedures about liquid handling</td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>Bluepard</td>
			<td>CU600</td>
			<td>Used for measuring protein content of BEVs at Step 8.2.5</td>
		</tr>
		<tr>
			<td>ZetaView</td>
			<td>Particle Metrix</td>
			<td>S/N 21-734, Software ZetaView (version 8.05.14 SP7)</td>
			<td>Nanoparticle tracking analysis (NTA) for measuring the particle size and concentrarion of BEVs at Step 8.4</td>
		</tr>
	</tbody>
</table>

isolation and purification of bacterial extracellular vesicles from human feces using density gradient centrifugation

Bacterial extracellular vesicles (BEVs) are nanovesicles derived from bacteria that play an active role in bacteria-bacteria and bacteria-host communication, transferring bioactive molecules such as proteins, lipids, and nucleic acids inherited from the parent bacteria. BEVs derived from the gut microbiota have effects within the gastrointestinal tract and can reach distant organs, resulting in significant implications for physiology and pathology. Theoretical investigations that explore the types, quantities, and roles of BEVs derived from human feces are crucial for understanding the secretion and function of BEVs from the gut microbiota. These investigations also necessitate an improvement in the current strategy for isolating and purifying BEVs.
This study optimized the isolation and purification process of BEVs by establishing two density gradient centrifugation (DGC) modes: Top-down and Bottom-up. The enriched distribution of BEVs was determined in fractions 6 to 8 (F6-F8). The effectiveness of the approach was evaluated based on particle morphology, size, concentration, and protein content. The particle and protein recovery rates were calculated, and the presence of specific markers was analyzed to compare the recovery and purity of the two DGC modes. The results indicated that the Top-down centrifugation mode had lower contamination levels and achieved a recovery rate and purity similar to that of the Bottom-up mode. A centrifugation time of 7 h was sufficient to achieve a fecal BEV concentration of 108/mg.
Apart from feces, this method could be applied to other body fluid types with proper modification according to the differences in components and viscosity. In conclusion, this detailed and reliable protocol would facilitate the standardized isolation and purification of BEVs and thus, lay a foundation for subsequent multi-omics analysis and functional experiments.

Determine the distribution of BEV-enriched fractions 
To determine the distribution of bacterial extracellular vesicles (BEVs)-enriched fractions, a blank control was established to measure the absorbance values at OD 340 nm, and the density of each fraction was calculated based on the measurements and iodixanol guidelines (Step 8.1). Table 2 presents the density results, demonstrating that fractions F4 to F9 exhibited densities within the range typically associated with extracellul...

Watch this Scientific Journal Video about Accelerating Research on Bacterial Extracellular Vesicles Separation and Heterogeneity at JoVE.com

Author Spotlight: Accelerating Research on Bacterial Extracellular Vesicles Separation and Heterogeneity 

This study describes a method to isolate and purify bacterial extracellular vesicles (BEVs) enriched from human feces via density gradient centrifugation (DGC), identifies the physical characteristics of BEVs from morphology, particle size, and concentration, and discusses the potential applications of the DGC approach in clinical and scientific research.

Isolation and Purification of Bacterial Extracellular Vesicles from Human Feces Using Density Gradient Centrifugation

Bacterial extracellular vesicles (BEVs) are nanovesicles derived from bacteria that play an active role in bacteria-bacteria and bacteria-host ...

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2.3K Views.  Southern Medical University. This study describes a method to isolate and purify bacterial extracellular vesicles (BEVs) enriched from human feces via density gradient centrifugation (DGC), identifies the physical characteristics of BEVs from morphology, particle size, and concentration, and discusses the potential applications of the DGC approach in clinical and scientific research.

Video: Author Spotlight: Accelerating Research on Bacterial Extracellular Vesicles Separation and Heterogeneity 

Isolation of CD4+ T cells from Mouse Lymph Nodes Using Miltenyi MACS Purification

Isolation of cells from the primary source is a necessary step in many more complex protocols. Miltenyi offers kits to isolate cells from several organisms including humans, non-human primates, rat and, as we describe here, mice. Magnetic bead-based cell separation allows for either positive selection (or cell depletion) as well as negative selection. Here, we demonstrate negative selection of untouched or na ve CD4+ helper T cells. Using this standard protocol we typically purify cells that are &#8805; 96% pure CD4+/CD3+. This protocol is used in conjunction with the protocol Dissection and 2-Photon Imaging of Peripheral Lymph Nodes in Mice published in issue 7 of JoVE, for purification of T cells and other cell types to adoptively transfer for imaging purposes. Although we did not demonstrate FACS analysis in this protocol video, it is highly recommended to check the overall purity of isolated cells using the appropriate antibodies via FACS. In addition, we demonstrate the non-sterile method of T cell isolation. If sterile cells are needed for your particular end-user application, be sure to do all of the demonstrated procedures in the tissue culture hood under standard sterile conditions. Thank you for watching and good luck with your own experiments!

Isolation of lymphocytes using the Miltenyi MACs kit is a reliable way to purify cells from whole lymphoid tissue homogenates. Cells purified using the Miltenyi system are typically &#8805; 96% pure. Here, we demonstrate the steps taken to isolate CD4+ T cells, one of the many kits offered by Miltenyi.

Isolation of cells from the primary source is a necessary step in many more complex protocols. Miltenyi offers kits to isolate cells from several ...

Human Pancreatic Islet Isolation: Part II: Purification and Culture of Human Islets

Management of Type 1 diabetes is burdensome, both to the individual and society, costing over 100 billion dollars annually. Despite the widespread use of glucose monitoring and new insulin formulations, many individuals still develop devastating secondary complications. Pancreatic islet transplantation can restore near normal glucose control in diabetic patients 1, without the risk of serious hypoglycemic episodes that are associated with intensive insulin therapy. Providing sufficient islet mass is important for successful islet transplantation. However, donor characteristics, organ procurement and preservation affect the isolation outcome 2. At University of Illinois at Chicago (UIC) we developed a successful isolation protocol with an improved purification gradient 3. The program started in January 2004 and more than 300 isolations were performed up to November 2008. The pancreata were sent in cold preservation solutions (UW, University of Wisconsin or HTK, Histidine-Tryptophan Ketoglutarate) 4-7 to the Cell Isolation Laboratory at UIC for islet isolation. Pancreatic islets were isolated using the UIC method, which is a modified version of the method originally described by Ricordi et al 8. As described in Part I: Digestion and Collection of Pancreatic Tissue, human pancreas was trimmed, cannulated, perfused, and digested. After collection and at least 30 minutes of incubation in UW solution, the tissue was loaded in the cell separator (COBE 2991, Cobe, Lakewood, CO) for purification 3. Following purification, islet yield (expressed as islet equivalents, IEQ), tissue volume, and purity was determined according to standard methods 9. Isolated islets were cultured in CMRL-1066 media (Mediatech, Herndon, VA), supplemented with 1.5% human albumin, 0.1% insulin-transferrin-selenium (ITS), 1 ml of Ciprofloxacin, 5 ml o f 1M HEPES, and 14.5 ml of 7.5% Sodium Bicarbonate in T175 flasks at 37&deg;C overnight culture before islets were transplanted or used for research.

Achieving high quality and appropriate quantity of human islets is one of the prominent prerequisites for successful islet transplantation. In this video, we describe step by step the procedures for human pancreatic islet isolation (part II: purification and culture of human islets) using a modified automated method.

Management of Type 1 diabetes is burdensome, both to the individual and society, costing over 100 billion dollars annually. Despite the widespread use of ...

DNA Extraction from 0.22 &mu;M Sterivex Filters and Cesium Chloride Density Gradient Centrifugation

This method is used to extract high molecular weight genomic DNA from planktonic biomass concentrated on 0.22 &mu;M Sterivex filters that have been treated with storage/lysis buffer and archived at -80&deg;C, and to purify this DNA using a cesium chloride density gradient.  The protocol begins with two one-hour incubation steps to liberate DNA from cells and remove RNA.  Next, a series of Phenol:Chloroform and Chloroform extractions are performed followed by centrifugation to remove proteins and cell membrane components, collection of the aqueous DNA extract, and several buffer exchange steps to wash and concentrate the extract.  Part Five describes the optional purification via cesium chloride density gradient.  It is recommended to work with less than 15 samples at one time to avoid confusion and cut down protocol time. The total time required for this protocol depends on the number of samples to be extracted. For 10-15 samples and assuming the proper centrifugation equipment is available, this entire protocol should take 3 days. Make sure you have the hybridization ovens set to temperature at the outset of the process.

We describe a method for extraction of high molecular weight genomic DNA from planktonic biomass concentrated on 0.22 &mu;m Sterivex filters, followed by cesium chloride density gradient centrifugation for purification.

This method is used to extract high molecular weight genomic DNA from planktonic biomass concentrated on 0.22 &mu;M Sterivex filters that have been ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Vampiric Isolation of Extracellular Fluid from Caenorhabditis elegans

The genetically tractable model organism C. elegans has provided insights into a myriad of biological questions, enabled by its short generation time, ease of growth and small size. This small size, though, has disallowed a number of technical approaches found in other model systems. For example, blood transfusions in mammalian systems and grafting techniques in plants enable asking questions of circulatory system composition and signaling. The circulatory system of the worm, the pseudocoelom, has until recently been impossible to assay directly. To answer questions of intercellular signaling and circulatory system composition C. elegans researchers have traditionally turned to genetic analysis, cell/tissue specific rescue, and mosaic analysis. These techniques provide a means to infer what is happening between cells, but are not universally applicable in identification and characterization of extracellular molecules. Here we present a newly developed technique to directly assay the pseudocoelomic fluid of C. elegans. The technique begins with either genetic or physical manipulation to increase the volume of extracellular fluid. Afterward the animals are subjected to a vampiric reverse microinjection technique using a microinjection rig that allows fine balance pressure control. After isolation of extracellular fluid, the collected fluid can be assayed by transfer into other animals or by molecular means. To demonstrate the effectiveness of this technique we present a detailed approach to assay a specific example of extracellular signaling molecules, long dsRNA during a systemic RNAi response. Although characterization of systemic RNAi is a proof of principle example, we see this technique as being adaptable to answer a variety of questions of circulatory system composition and signaling.

Vampiric Isolation of Extracellular Fluid from Caenorhabditis elegans

The model organism C. elegans uses pseudocoelomic fluid as a passive circulatory system. Direct assay of this fluid has not been previously possible. Here we present a novel technique to directly assay the extracellular space, and use systemic silencing signals during an RNAi response as a proof of principle example.

The genetically tractable model organism C. elegans has provided insights into a myriad of biological questions, enabled by its short generation time, ...

Sperm Collection of Differential Quality Using Density Gradient Centrifugation

In sexual reproduction, a male gamete or sperm cell fuses with a female gamete to bring about fertilization. However, a large number of sperm cells with fertilizing ability are required to interact with a female gamete to ensure fertilization. As such, the fertilizing ability of individual sperm cells is critical for successful reproduction. Density gradient centrifugation has been utilized for several decades as a reproducible, fast, efficient, effective and extremely adaptable method to collect only high-quality sperm to be used in assisted reproductive technology. The protocols we described herein focus on the utilization of the discontinuous Percoll density gradient centrifugation (PDGC) technique to isolate three distinct populations of rooster sperm by their quality. We were able to collect low-, medium- and high-quality sperm. We also describe reproducible protocols that entail determining fertility potential of sperm by assessing their viability, mobility and penetrability. Collection of sperm by their quality using PDGC technique would be useful to accurately and thoroughly characterize sperm with differential fertility potential.

In this paper, we aim to describe the performance of the density gradient centrifugation technique and its application in sperm physiology research.

In sexual reproduction, a male gamete or sperm cell fuses with a female gamete to bring about fertilization. However, a large number of sperm cells with ...

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 ...

Purification and Analysis of Caenorhabditis elegans Extracellular Vesicles

The secretion of small membrane-bound vesicles into the external environment is a fundamental physiological process of all cells. These extracellular vesicles (EVs) function outside the cell to regulate global physiological processes by transferring proteins, nucleic acids, metabolites, and lipids between tissues. EVs reflect the physiological state of their cells of origin. EVs are implicated to have fundamental roles in virtually every aspect of human health. Thus, EV protein and genetic cargos are being increasingly analyzed for biomarkers of health and disease. However, the EV field still lacks a tractable invertebrate model system that permits the study of EV cargo composition. C. elegans is well suited for EV research because it actively secretes EVs outside of its body into its external environment, permitting facile isolation. This article provides all the necessary information for generating, purifying, and quantifying these environmentally secreted C. elegans EVs including how to work quantitatively with very large populations of age-synchronized worms, purifying EVs, and a flow cytometry protocol that directly measures the number of intact EVs in the purified sample. Thus, the large library of genetic reagents available for C. elegans research can be tapped into for investigating the impacts of genetic pathways and physiological processes on EV cargo composition.

Purification and Analysis of Caenorhabditis elegans Extracellular Vesicles

This article presents methods for generating, purifying, and quantifying Caenorhabditis elegans extracellular vesicles.

The secretion of small membrane-bound vesicles into the external environment is a fundamental physiological process of all cells. These extracellular ...

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

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

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

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

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

Isolation and Characterization of Cyanobacterial Extracellular Vesicles

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

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