Single Step Isolation of Extracellular Vesicles from Large-Volume Samples with a Bifurcated A4F Microfluidic Device

Abstract

Extracellular vesicles (EVs) hold immense potential for various biomedical applications, including diagnostics, drug delivery, and regenerative medicine. Nevertheless, the current methodologies for isolating EVs present significant challenges, such as complexity, time consumption, and the need for bulky equipment, which hinders their clinical translation. To address these limitations, we aimed to develop an innovative microfluidic system based on cyclic olefin copolymer-off-stoichiometry thiol-ene (COC-OSTE) for the efficient isolation of EVs from large-volume samples in a continuous manner. By utilizing size and buoyancy-based separation, the technology used in this study achieved a significantly narrower size distribution compared to existing approaches from urine and cell media samples, enabling the targeting of specific EV size fractions in future applications. Our innovative COC-OSTE microfluidic device design, utilizing bifurcated asymmetric flow field-flow fractionation technology, offers a straightforward and continuous EV isolation approach for large-volume samples. Furthermore, the potential for mass manufacturing of this microfluidic device offers scalability and consistency, making it feasible to integrate EV isolation into routine clinical diagnostics and industrial processes, where high consistency and throughput are essential requirements.

Introduction

Extracellular vesicles (EVs) are cell-derived membrane-bound particles comprising two main types: exosomes (30-200 nm) and microvesicles (200-1000 nm)¹. Exosomes form through inward budding of the endosomal membrane within a multivesicular body (MVB), releasing intraluminal vesicles (ILVs) into the extracellular space upon fusion with the plasma membrane¹. In contrast, microvesicles are generated by outward budding and fission of the cell membrane². EVs play a crucial role in intercellular communication by transporting proteins, nucleic acids, lipids, and metabolites, reflecting the physiological state of the cell, including growth, angiogenesis, metastasis, proliferation, and therapy resistance³. As a result, they have emerged as promising biomarkers and therapeutic targets for diseases, including cancer, highlighting their potential in diagnostics and drug delivery systems⁴.

To fully utilize EVs in disease diagnostics and therapeutics, efficient isolation from various biofluids is crucial⁵. Common methods include ultracentrifugation (UC), density gradient centrifugation, size exclusion chromatography (SEC), filtration, and immunoisolation⁶. UC is a widely used technique but may yield particles of similar density that are not EVs and can generate EV aggregates⁷. SEC has gained popularity due to its ability to provide higher purity samples by excluding particles based on size rather than density⁸. However, careful selection of the appropriate pore size for the SEC column and optimization of chromatography conditions are essential to minimize co-isolation of unwanted particles like chylomicrons and low-density lipoproteins⁸. Despite their effectiveness, both methods are time-consuming and challenging to automate, especially for larger volume samples like cell media or urine, limiting their scalability for industrial applications⁹.

In recent years, asymmetric field flow field fractionation (A4F) has evolved as a powerful separation technique for size and buoyancy-based micro- and nanometer-sized particle separation¹⁰. The operational principle of A4F relies on a microfluidic channel endowed with a porous membrane at its base, generating a force exerted towards the membrane called cross-flow¹⁰. When combined with Brownian motion and Poiseuille flow inherent to the system, cross-flow facilitates efficient particle separation due to varying particle position within the flow dynamics¹¹. Despite the benefits, this method is limited to sample volumes within the microliter range¹² and requires an additional focusing step, extending the duration of the process¹⁰.

Over the last decade, microfluidics has gained prominence as a tool for rapid, efficient, and clinically reliable EV separation¹³. However, most microfluidic methods designed for EV separation are optimized for small-volume, high-concentration EV samples or depend on complex separation procedures¹⁴. Furthermore, within the field of microfluidics, polydimethylsiloxane (PDMS) is recognized as the golden standard material owing to its optical transparency, biocompatibility, and ease of use¹⁵. Nevertheless, its known propensity to absorb small lipophilic molecules, including EVs, can be problematic for its application in the EV field¹³.

Cyclic olefin copolymer (COC) is a frequently used material in microfluidics due to biocompatibility, small absorption of molecules, and high chemical resistance¹⁵. However, the fabrication of COC devices often involves complex processes or specialized equipment¹⁶. Alternatively, off-stoichiometry thiol-ene (OSTE) is a promising alternative to PDMS due to decreased absorption of small molecules, superior chemical stability, ease of fabrication, and scalable fabrication process^17,18. However, due to complex connections to tubing, devices can be prone to leaking¹⁹.

The aim of this study was to engineer and fabricate a microfluidic device combining OSTE and COC and bifurcated A4F principle for EV separation from large-volume samples such as urine or cell media.

Protocol

Sample collection was approved by the Latvian University Life and Medical Science Research Ethics Committee (decision N0-71-35/54)

NOTE: The materials used in this study are included in the Table of Materials file.

1. Three-dimensional (3D) printed mold fabrication

1. Design a serpentine-shaped double negative mold in CAD software with the following dimensions for the top channel: height of 0.5 mm, width of 1 mm, and length of 210 mm. For the bottom channel, use the same design with a width of 1.5 mm, height of 0.6 mm, and length of 210 mm. Dimensions such as mold width, length, height, and design isometric illustration are shown in Figure 1. Save the design as .stl file.
2. Prepare the file in Z-Suite; no support needs to be added. Print the molds using an ultraviolet liquid crystal display (UV LCD) 3D printer with tough black resin.
3. After printing, carefully remove the molds from the printer surface. Wash the molds in isopropanol (IPA) in sonication for 20 min.
4. Blow-dry the molds with N₂.
5. Expose the molds to flood exposure using an ND33 filter for 810 s, then place them into an oven at 60 °C for 48 h.

2. Preparation of the PDMS molds

1. Weight PDMS at a 10:1 w/w ratio (for each mold, use 16 g of elastomer base and 1.6 g of cross-linking agent). Mix the components in a planetary centrifugal mixer, mixing for 1 min at 500 RPM.
2. Cast the mixed PDMS into the 3D-printed molds and degas in a vacuum desiccator at -800 mbar for 30 min or until no bubbles are observed.
3. Carefully place a 100 µm thick polyvinyl chloride liner on top of the liquid PDMS.
   NOTE: Apply the liner slowly from one side to another to avoid bubble formation.
4. Insert the mold with PDMS into a Mold-in-Place Jig and tighten the jig by using a torque wrench set to 0.3 Nm.
5. Place the jig into a 60 °C oven for 3 h to cure PDMS.
6. Disassemble the jig using a hex key and remove the 3D-printed mold from the jig.
7. Carefully de-mold PDMS molds from the 3D-printed master molds, running around the edges of the mold with a scalpel, then removing the mold with tweezers.
   ​NOTE: For easier mold removal, IPA can be used; however, the molds need to be heated at 60 °C for 10 min afterward to evaporate excess IPA.

3. Preparation of OSTE-COC top channel

1. Weight OSTE at 1.09:1 w/w ratio (for 5 devices, 1.56 g of A part and 1.44 g of B part are needed). Mix the components in a planetary centrifugal mixer: mix for 5 min at 750 RPM, then degas for 5 min at 750 RPM.
2. Transfer the mixed OSTE to a centrifuge tube and degas at -800 mbar in a vacuum desiccator for 30 min.
3. Clean COC slides with 2 x 16 mini Luer connectors by sonication in IPA for 10 min. Carefully dry the slides with N₂.
4. Place the clean COC slides into the plasma asher so that the flat surface is placed upwards, and treat the surface with 800 SCCM oxygen plasma for 2 min at 700 W power with 0.133 mbar pressure.
5. Place the oxidized COC slide on the PDMS mold for the top channel so that the treated surface is in contact with the structured surface of PDMS. Place the assembly into a Mold-in-Place jig, tightening the jig with a torque wrench set to 0.3 Nm.
6. Connect the pressure system to the compressed air line at 6 bar. Take the centrifuge tube with degassed OSTE and assemble it per pressure system instructions using polytetrafluorethylene (PTFE) tubing with 0.8 mm inner diameter (ID).
7. Connect the PTFE tube to the inlet of the PDMS mold and, using 1000 mbar pressure maintained by a piezoelectric pump located in the pressure system, fill the device. Place the jig with the glass side up into the mask aligner and expose with 850 mJ dose using ND33 filter.
8. After exposure, disassemble the jig using a hex key and carefully remove the COC slide with the structured OSTE layer from the PDMS mold.
9. Bring the structured OSTE in contact with a 25 mm x 75 mm membrane with 50 nm pores and 11.8% pore density so that the channels are fully covered with the membrane.
   NOTE: It is particularly important to keep the membrane as straight as possible during the process of application.
10. Place the assembly onto a hot plate set to 60°C with the membrane side up, place a clean glass slide on top of the membrane, and apply a 1.6 kPa pressure from the top to ensure even bonding. Heat the assembly overnight.
    ​NOTE: After this step, it is especially important to evaluate bonding performance and channel deformation.

4. Preparation of OSTE-COC bottom channel and device assembly

1. Mix OSTE at 1.09:1 w/w ratio (for 5 devices 1.56 g of A part and 1.44 g of B part are needed). Mix the components in a planetary centrifugal mixer: mix for 5 minutes at 750 RPM, then degas for 5 minutes at 750 RPM.
2. Transfer the mixed OSTE to a falcon tube and degas at -800 mbar in a vacuum desiccator for 30 min.
3. Clean COC slides by sonication in IPA for 10 minutes. Carefully dry the slides with N₂.
4. Place the clean COC slides into plasma asher and treat the surface with 800 SCCM oxygen plasma for 2 min at 700 W power with 0.133 mbar pressure.
5. Place the oxidized COC slide on the PDMS mold for the bottom channel so that the treated surface is in contact with the structured surface of PDMS. Place the assembly into a Mold-in-Place jig, tightening the jig with a torque wrench set to 0.3 Nm.
6. Take the falcon tube with degassed OSTE and assemble it as per pressure system instructions using PTFE tubing with 0.8 mm ID.
7. Connect the PTFE tube to the inlet of the PDMS mold and using 1000 mbar pressure fill the device. Place the jig with the glass side up into mask aligner and expose with 1100 mJ dose using ND33 filter.
8. After exposure, disassemble the jig using a hex key and carefully remove the COC slide with the structured OSTE layer from the PDMS mold.
9. Combine the structured OSTE layer with the top channel assembly so that the design of the top and bottom layer is aligned with one another and that the OSTE bottom layer is in contact with the membrane.
10. Insert the design into a Mold-in-Place jig and, using bulldog clips, apply a 1.6 kPa pressure to the device evenly. Place the jig into a 60°C oven overnight.
    ​NOTE: Uneven placement of the clips can lead to uneven bonding.

5. Device evaluation

1. After bonding, visually evaluate the devices with a microscope for visual defects, for example, deformation of the channel or unsuccessful bonding.
2. Test the devices with no defects by passing 0.02 µm filtered deionized water through the channels with 30 mbar pressure. Observe for flow channel and Luer port leaks and water flow.
   ​NOTE: If no leakage can be detected and channel flow is uninterrupted, devices are considered usable for further experiments.

6. Device setup

1. Fill two 5 mL syringes with 2 mL of 20 nm filtered 3% H₂O₂ and fit them on a syringe pump (use the movement arrows for the needed distance, insert syringes, and fix them in place with the top bar).
   CAUTION! Always be careful when handling H₂O₂. Wear a lab coat, protective rubber gloves, and eye protection.
2. Connect 14 cm long 800 µm ID PTFE tubing to the syringes using the syringe needles and to the inlets of the device using Luer connectors.
3. Connect 20 cm long 200 µm ID polyether ether ketone (PEEK) tubing to the outlets.
4. Connect the other end of the PEEK tubing to a waste container. The setup can be seen in Figure 3.
5. Start the syringe pump at the flow speed of 500 µL/min for each inlet (on the instrument screen: Settings > System Set > Syringe (manufacturer: medical syringe, syringe: 5 mL) > Back button > Parameters (Select: infuse, Repeat: 1, Volume: 1.5 mL, Flow: 500 µL/min) > Back button > Back button). Collect the flow-through in a waste container.
6. Fill two new 5 mL syringes with 2 mL of 20 nm filtered 70% ethanol.
7. Replace the previous syringes with the ethanol-filled syringes and start the syringe pump using the same parameters described in step 6.5.
8. Use new 5 mL syringes and fill them with 5 mL of 20 nm filtered phosphate-buffered saline (PBS).
9. Replace the previous syringes with the PBS-filled syringes and flush the device with 4 mL of PBS by repeating steps 6.1. and 6.5.
   NOTE: Make sure to change the system setup parameters to Volume: 4 mL (on the instrument screen: Settings > Parameters > (Select: Volume: 4 mL).
10. Unplug the tubing from the device and close each inlet and outlet with Luer plugs.
    NOTE: Leave PBS in the device and avoid trapping any air by adding additional PBS on each inlet and outlet using a micropipette before closing.
11. Leave the device overnight in a dark place at room temperature (RT).
12. The following day, flush the PBS-filled device with 4 mL of 20 nm filtered PBS (as described in step 6.8).
13. Collect the last 1 mL of the flow-through from both outlets in a 2 mL protein low-binding tube and store at 4 °C as a blank for particles introduced by the device.
14. Remove the PBS-filled syringes and replace them with new 5 mL syringes filled with air.
15. Purge the device with 4 mL of air until all the liquid has exited the device and tubing.

7. Device testing with standardized latex beads

1. Prepare a standardized bead sample using 1.0 µm polystyrene and 0.1 µm carboxylate beads. To a 15 mL tube, add 500 µL of 1.0 µm polystyrene bead standards, 1 µL of 0.1 µm carboxylate bead standards, and 9499 µL of 20 nm filtered PBS (final concentration of 1.0 and 0.1 µm bead mix will be 5 × 108 beads/mL and 3.6 × 1010 beads/mL respectfully).
2. Vortex the bead sample for 30 s and store it at +4 °C while not in use.
3. Fill one new 5 mL syringe with 1.5 mL of the standardized bead mixture and another one with 1.5 mL of 20 nm filtered PBS.
4. Remove the previously attached syringes and attach the bead mixture syringe to the first inlet tubing and the other to the second inlet.
5. Remove the waste container and prepare at least five 0.5 mL microcentrifuge tubes for each outlet tubing.
6. Change the syringe pump system start-up parameters to Volume: 1 mL; Flow: 250 µL/min.
   NOTE: Depending on the user's needs, different flow speeds such as 250 µL/min, 200 µL/min, 150 µL/min, 100 µL/min, and 50 µL/min can be set. A preliminary study to choose the most favorable speed for the wanted outcome is recommended.
7. Start the syringe pump and collect the outflow - 5 to 6 droplets in each microcentrifuge tube.
   ​NOTE: If the same device is used multiple times, flush the device as described in step 6.8 and change all connectors, plugs, and needles. Wash the used connectors, plugs, and needles by leaving them in 20 nm filtered 70% ethanol for at least 30 min. Then, move all parts to a tube containing 20 nm filtered deionized water, shake the tube thoroughly, and transfer them to a Petri dish containing 20 mL of 20 nm filtered deionized water. Leave the Petri dish on the shaker for at least 30 min at 100 rpm, then leave the things to dry in a dry Petri dish (for at least 12 h) before reusing.

8. Device testing with urine samples

1. Prepare the device as described in steps 6.1-6.15.
2. Collect 100 mL of morning urine per donor using urine sterile containers and deliver it to the laboratory for processing within 2 hours of collection.
3. Separate each urine sample into two 50 mL tubes and centrifuge at 2'000 x g for 15 min at RT.
4. Collect the supernatant and discard the pellet.
5. Centrifuge the supernatant at 10'000 x g for 30 minutes at RT to get rid of large particles and cell debris.
6. Collect the supernatant and discard the pellet.
7. Wash the device as described in 6.6.
8. Change the 5 mL syringes to a 20 mL syringe filled with the prepared urine sample for the sample inlet and another 20 mL syringe filled with 20 mL of 20 nm filtered DEPC-PBS for the buffer inlet. Change the syringe pump settings to BDPlastipak; Syringe: cc; Volume: 20 mL; Flow: 250 µL/min (optimal flow rate), similarly as described in step 6.2. Start the syringe pump and collect the flow-through from each outlet in separate 50 mL tubes.
9. Concentrate each flow-through separately using 100,000 molecular weight cut-off concentration tubes to 100 µL each at 3,000 x g in 4 °C.
10. Transfer the concentrates to 0.5 mL microcentrifuge tubes.
11. Vortex the concentrated samples for 30 seconds, then spin for 10 seconds.
12. Aliquot the samples by transferring 25 µL of sample per tube, label them, and freeze at -80 °C for long-term storage.

9. Device testing with conditioned media

1. Prepare the device as described in steps 6.1- 6.15.
2. Culture cells in a 5% CO₂ humidified environment at 37 °C until the total count reaches 100 million according to Bajo-Santos et al.²⁰ Passage the cells to a single T175 suspension flask in 100 mL of serum-free media supplemented with 2% B-27 serum substitute supplement and culture in a 5% CO₂ humidified environment at 37 °C for additional 48 h.
3. Collect the cell suspension and centrifuge it at 300 x g for 5 min at RT to get rid of cells.
4. Collect and centrifuge the supernatant again at 3,000 x g for 30 min at +4 °C to get rid of large particles and cell debris.
5. Collect the supernatant and store it at +4°C until the separation can take place, but no longer than 3 h.
6. Change the 5 mL syringes with 20 mL syringes filled with the prepared conditioned media for the sample inlet and filled with 20 mL of 20 nm filtered PBS for the buffer inlet.
7. Set the settings in the pump station as described in step 7.8.
8. Start the syringe pump and collect the flow-through from each outlet in a separate 50 mL tube.
9. Concentrate each flow-through separately using 100,000 molecular weight cut-off concentration tubes to 150 µL each at 3,000 x g at +4 °C.
10. Transfer the concentrates to 0.5 mL microcentrifuge tubes.
11. Vortex concentrated samples for 30 s.
12. Spin concentrated samples for 10 s.
13. Aliquot the samples by transferring 25 µL of sample per tube, label them, and freeze them at -80 °C for long-term storage.

10. Isolation of EVs using ultracentrifugation

1. Centrifuge 20 mL of urine or conditioned cell media at 100,000 x g for 70 min at +4 °C using a centrifuge with a fixed-angle rotor.
2. Discard the supernatant and resuspend the pellet in 20 mL of 20 nm filtered PBS.
3. Centrifuge again, as explained in 10.1.
4. Discard the supernatant and resuspend the pellet in 12 mL of 20 nm filtered PBS.
5. Centrifuge at 100,000 x g for 70 min at +4 °C using a swing rotor.
6. Discard the supernatant.
   NOTE: Be careful when discarding the supernatant this time. It is recommended to do it with a pipette.
7. Resuspend the pellet in 100 µL of 20 nm filtered PBS.
8. Aliquot the sample and proceed to setion 12 for EV characterization. Otherwise, store at -80 °C until further use.

11. Isolation of EVs using size exclusion chromatography (SEC)

1. Concentrate 20 mL of urine or conditioned cell media to 500 µL using 100 kDa molecular weight cut-off (MWCO) centrifugal filter units at 3,000 x g at +4 °C for 15 min.
2. Transfer the concentrate onto the SEC column (35 nm load range).
3. Pour the 20 nm filtered PBS onto the column and collect 15 fractions of 0.5 mL.
4. Analyze fractions with a dynamic light scattering (DLS) system following the manufacturer's instructions.
5. Pool together the selected fractions. Choose fractions with an attenuator lower than 11 and at least 30% of total particles being bigger than 40 nm.
6. Concentrate selected fractions to 100 µL using 3 kDa molecular weight cut-off (MWCO) centrifugal filter units at 14,000 x g at +4 °C for 15 min.
7. Aliquot the sample and proceed to section 12 for EV characterization. Otherwise, store at -80 °C until further use.

12. EV characterization

1. Take the sample to be characterized from the freezer and defrost it slowly (on an ice block or at +4 °C).
2. Perform nanoparticle tracking analysis (NTA) to determine particle size distribution and amount¹³. Perform high-affinity T-cell membrane protein 4 (TIM-4) double sandwich enzyme-linked immunosorbent assay (dsELISA)¹⁴ to confirm EV presence in the samples.

13. NTA

1. Vortex the thawed EV sample for 30 s and spin for 10 s, then transfer 1 µL to 999 µL of 20 nm filtered PBS in a 2 mL microcentrifuge tube to dilute it 1000-fold. Save 1 mL of extra PBS used for dilution to check that it contains no particles. Store everything at +4 °C until measurement.
   NOTE: Filter PBS through a 0.02 nm filter prior to use.
2. Vortex the EV sample or blank for 30 s and insert it into the module using a syringe. Fill the module with approximately 0.7 mL of sample and place it in the instrument.
3. Measure the sample using the nanoparticle analyzer instrument following the manufacturer's instructions.

14. dsELISA for EV markers

1. Prepare a solution of 1 µg/mL TIM4-Fc protein. Coat the wells of a 96-well ELISA plate by transferring 100 µL of solution to each necessary well and incubate overnight at +4 °C shaking at 200 RPM.
2. The following day, prepare washing solution (1x TBS + 0.05% Tween20 + 2 mM CaCl₂).
3. Discard all the liquid in each well. Wash each well by adding 200 µL of wash solution, shaking at 200 RPM for 5 min at RT, and discarding. Repeat this step twice.
4. Prepare blocking solution (1x TBS + 0.05% Tween20 + 1% bovine serum albumin. Add 200 µL of blocking solution to each well and incubate at RT for 1 h while shaking at 200 RPM.
5. Wash each well as described in step 14.3.
6. Prepare 220 µL of each sample dilution in washing buffer (optimal EV concentration is 2.7 × 10⁵ EVs/µL). Add 100 µL of the diluted sample per well and perform two replicates. Incubate at RT for 90 min while shaking at 200 RPM.
7. Wash each well as described in step 14.3.
8. Add 100 µL of corresponding antibodies to each well. Use proper EV markers for confirming EVs and contaminants⁴. Incubate at RT for 2 h while shaking at 200 RPM. NOTE: If secondary antibodies are used, repeat steps 14.7-14.8. However, reduce the incubation period of secondary antibodies to 1 h.
9. Wash each well as described in step 14.3.
10. Add 100 µL of horseradish peroxidase substrate to each well, mix the plate for 1 min, and incubate at RT for 30 min.
11. Add 1 M H₂SO₄ to each well to stop the reaction.
12. Using a microplate reader, measure the absorbance at 450 nm.

Representative Results

We fabricated a microfluidic device using a 3D printed double negative mold (Figure 1) via soft-lithography (Figure 2A) for high throughput EV separation based on the bifurcated A4F principle (Figure 2B,C). The setup requires a pump and a flow-through station, as can be seen in Figure 3, for the isolation of EVs in an automated manner. Firstly, to evaluate the proof of concept of the devices, a mixture of polystyrene beads with diameters of 100 nm and 1000 nm was prepared to represent vesicles and fine cell debris, respectively¹⁰. Experiments were conducted with varying flow rates for the bead mix, both with and without bifurcating flow, to investigate the effect of linear velocity on separation efficiency. Across all experiments, the recovery of small beads remained consistent and above 90%¹⁰,showing the potential of the device to recover EVs.

Then, we assessed and compared the potential of the COC-OSTE device in isolating EVs from large volumes (>1 mL) from complex biofluids with minimal pre-processing. As such, urine from 10 healthy donors (Figure 4) and cell media from two different prostate cell lines (Figure 5) were used as a template to simultaneously isolate EVs following three different procedures: ultracentrifugation (UC), size exclusion chromatography (SEC), and the A4F microfluidics-based OSTE-COC device. After isolation, the total number of particles and their size distribution were assessed using nanoparticle tracking analysis (NTA). On average, the OSTE-COC device showed better total particle recovery from biofluids compared to UC, but the statistical significance was only achieved when combining the particle numbers from both ports (Figure 4A). In order to compare the device performance with other systems, R & L outlet together should be taken into consideration. As shown in Figure 4, L & R outlet together performance on recovering EVs outperforms UC and SEC. Separately, the L-port was designed to capture the small EV fraction, while the R-port was designed to collect the bigger EV fraction with other molecules of similar size. Interestingly, the recovery using the R-PORT of the OSTE-COC device was slightly higher than SEC and UC alone (Figure 4A). CD63 expression showed a similar pattern (Figure 4C). This finding indicates that the OSTE-COC device was more effective in total EV recovery. Equivalent size distribution was found between the different methodologies, except for UC, which shows a bigger particle size distribution (Figure 4B).

Comparable results were observed in cell media cultures. In both scenarios, the total particle recovery from both device ports exhibited superior performance compared to SEC or UC methodologies (as depicted in Figure 5A,B). Notably, EVs derived from PC3 cells demonstrated a distinct size distribution, with greater homogeneity in the L-PORT distribution when contrasted with other experimental groups (Figure 5C,D). Furthermore, the analysis of CD63 expression confirmed the higher EV recovery rates achieved using the COC-OSTE device (as illustrated in Figure 5E,F). A summary comparing the isolation characteristics of the different methodologies examined in this study can be found in Table 1.

Figure 1: 3DP serpentine-shaped double negative mold dimensions and isometric view. Please click here to view a larger version of this figure.

Figure 2: COC-OSTE microfluidic device. (A) Scheme of the different main steps of fabricating the OSTE-COC device. (B) Device working principle. (C) Image of the finished device. Scale bar: 15 mm. This figure has been modified with permission from Priedols et al.¹⁰ and Bajo-Santos et al.²⁰. Please click here to view a larger version of this figure.

Figure 3: Experimental configuration of the device. The syringe pump is on the left, the OSTE-COC device is in the middle, and the recovery station is on the right. This figure has been modified with permission from Priedols et al.¹⁰ and Bajo-Santos et al.²⁰. Please click here to view a larger version of this figure.

Figure 4: Urinary EV size distribution and particle recovery from 10 donors using ultracentrifugation (UC), size-exclusion chromatography (SEC), and the COC-OSTE device. (A) Particle amount recovered from each isolation method by nanoparticle tracking analysis (NTA). Data represented as mean +/- standard deviation. Statistical significance denoted by * (p<0.05). (B) Boxplots display the average particle size distribution among all urine samples assessed by NTA, with whiskers indicating the minimum and maximum values. P-values were derived from comparisons to UC, with **** indicating high statistical significance (p<0.0001). (C) The median and range of the average CD63 amount, assessed using double-sandwich enzyme-linked immunosorbent assay (dsELISA), for each isolation method, were calculated across all samples. L-port: Left port; R-port: Right port. This figure has been modified with permission from Bajo-Santos et al.²⁰. Please click here to view a larger version of this figure.

Figure 5: Characterization of EVs isolated from PC3 and LNCaP cells using different methods. (A) Particle amount recovered from PC3 media using nanoparticle tracking analysis (NTA) represented by mean+/- standard deviation. (B) Particle amount recovered from LNCaP cell media using NTA represented as mean +/- standard deviation. (C) Median particle size distribution of EVs from PC3 cultures, along with the range, was determined by NTA. (D) Median size distribution of EVs isolated from LNCaP cultures with range. (E) Median and range of CD63 expression for each isolation method for PC3 cell line-derived EV, using double-sandwich enzyme-linked immunosorbent assay (dsELISA). (F) Median and range of LNCaP-derived EV CD63 expression by dsELISA for each isolation method. UC: Ultracentrifugation; SEC: Size-exclusion chromatography; L-port: Left port; R-port: Right port. This figure has been modified with permission from Bajo-Santos et al.²⁰. Please click here to view a larger version of this figure.

	UC	SEC	COC-OSTE
Processing time/sample	++/+++	+++	+
Throughput	++	+	+++
Cost/sample	+	+++	++
Purity	+	+++	++
Automatization	++	+	+++
EV Yield	++	++	+++
Size selection	+	++	++

Table 1. Comparison of isolation characteristics of EVs with the three methods UC SEC, and the COC-OSTE device. UC: Ultracentrifugation; SEC: Size-exclusion chromatography. COC-OSTE: cyclic olefin copolymer-off-stoichiometry thiol-ene. +: low; ++: medium; +++: high. * Device dependent.

Discussion

The presented microfluidic device offers a promising method for the isolation and extraction of EVs from biological fluids, addressing some of the critical limitations of existing gold standard methods such as UC and SEC¹². UC and SEC are known to be labor-intensive, time-consuming, and suffer from low yield, making them less suitable for high-throughput applications where large quantities of EVs are needed^21,22. In contrast, the microfluidic device described can continuously process a substantial volume of up to 20 mL of biological fluid with minimal user input, making it a potential game-changer for industrial or clinical settings. One of the key advantages of this microfluidic device is its ability to standardize EV isolation and improve reproducibility compared to traditional methods that involve manual steps. By reducing the variability of EV isolation device manufacturing through mass manufacturing, the device's performance can be better controlled and consistent, which is vital for applications like EV-based therapeutics and diagnostics. Furthermore, the device's compatibility with large-volume manufacturing via reaction injection molding of suitable substrates further enhances its potential for scalability and broader adoption.

To ensure reproducibility and consistent performance, an extensive and well-defined protocol is necessary, both to produce the chip and for its usage in experiments. The chip is designed with industrialization in mind. However, during the device fabrication in the laboratory environment, attention must be paid to avoiding the formation of bubbles, ensuring precise and robust bonding, and conducting thorough testing at higher flow speeds than those expected in real experiments. This precaution is especially important for COC-OSTE devices, as they can be prone to leaking due to unsuccessful bonding, especially at higher flow rates. Additionally, since OSTE is a photosensitive material, the device fabrication should be performed in yellow light to avoid unnecessary exposure to UV. As this technology is still in its initial stages, researchers should experiment with different flow speeds to identify the optimal setting for their specific application since standardized protocols for this novel method are yet to be established, and liquid viscosity can clearly affect EV isolation efficacy based on our results. By following these guidelines and addressing the challenges of standardization, the potential of this microfluidic device for EV isolation can be fully realized across various research areas and applications.

The method of EV isolation using the microfluidic device allows for extensive modification and troubleshooting to optimize performance. To improve particle separation, researchers can explore longer channels, different membrane porosities and pore densities, and altered dimensions. If there are a lot of large particles in the small particle outlet, testing a higher buffer inlet speed can be beneficial, while if there are many small particles in the large particle outlet, experimenting with a longer channel or a membrane with larger pores is recommended. Addressing particle absorption can involve adjusting flow speeds, surface modifications, and OSTE composition or changing the UV exposure time. For devices experiencing instability and leakage, decreasing flow speeds and ensuring proper gluing are crucial steps. To minimize EV damage, researchers can reduce flow speeds, wash with PBS more thoroughly after device sterilization, and consider using smaller membrane pores or alternative mold design techniques and materials. Through these adjustments, the microfluidic device can be fine-tuned, unleashing its potential for diverse applications in EV research and related fields. Additionally, to optimize the device's performance, upstream sample characterization, including density and viscosity measurements, could be incorporated to adjust flow parameters. Moreover, particular emphasis must be placed in the nature of the sample, given the established non-sterility of biofluids²³. Consequently, the sterilization of samples before their administration in clinical applications should be duly considered.

Despite these challenges, the microfluidic device holds immense potential in various research areas. Its ability to continuously isolate EVs from large volumes of highly heterogeneous samples opens the door for automation, facilitating high-throughput applications and enabling more in-depth analysis of EVs from diverse sources. Integrating this microfluidic module into different devices, such as hollow fiber cell bioreactor cartridges or high-resolution flow cytometers, could make EV applications more industry-friendly and drive innovation in fields like drug delivery, diagnostics, and cosmetics⁴. Overall, this microfluidic device represents a significant step forward in the field of EV research, offering an efficient and standardized method for EV isolation with promising applications in a wide range of research an industrial settings.

Materials

Name	Company	Catalog Number	Comments
0.1 µm carboxylate FluoSpheres	Invitrogen	#F8803	Stock concentration: 3.6 x 1013 beads/mL (LOT dependent)
0.5 mL microcentrifuge tubes	Starstedt	72.704
1 mL Luer cone syringe single use without needle	RAYS	TUB1ML
1.0 µm polystyrene FluoSpheres	Invitrogen	#F13083	Stock concentration: 1 x 1010 beads/mL (LOT dependent)
10 mL Serological pipettes	Sarstedt	86.1254.001
15 mL (100k) Amicon Ultra centrifugal filters	Merck Millipore	UFC910024
2.0 mL Protein LoBind tubes	Eppendorf	30108132
20 mL syringes	BD PlastikPak	10569215
250 µm ID polyether ether ketone tubing	Darwin Microfluidics	CIL-1581
3 kDa MWCO centrifugal filter units	Merck Millipore,	UFC200324
5 mL Medical Syringe without Needle	Anhui Hongyu Wuzhou Medical	159646
50 mL conical tubes	Sarstedt	62.547.254
70 Ti fixed angle ultracentrifuge rotor	Beckman Coulter	337922
800 µm ID polytetrafluoroethylene tubing	Darwin Microfluidics	LVF-KTU-15
96 well microplate, f-bottom, med. binding	Greiner Bio-One	655001	ELISA plate
B-27 Supplement (50x), serum free	Thermo Fisher Scientific	17504044
Bovine serum albumin	SigmaAldrich	A7906-100G
COC Topas microscopy slide platform	Microfluidic Chipshop	10000002
COC Topas microscopy slide platform 2 x 16 Mini Luer	Microfluidic Chipshop	10000387
Elveflow OB1 pressure controller	Elvesys Group
Luer connectors	Darwin Microfluidics	CS-10000095
Mask aligner Suss MA/BA6	SUSS MicroTec Group
Mixer Thinky ARE-250	Thinky Corporation
NanoSight NS300	Malvern Panalytical	NS300	nanoparticle analyzer
Optical microscope Nikon Eclipse LV150N	Nikon Metrology NV
OSTE 322 Crystal Clear	Mercene Labs
PBS TABLETS.Ca/Mg free. Fisher Bioreagents. 100 g	Fisher Scientific	BP2944-100
PC membrane (50 nm pore diameter, 11.8% density)	it4ip S.A., Louvain-La Neuve, Belgium
Petri dishes, sterile	Sarstedt	82.1472.001
Plasma Asher GIGAbatch 360 M	PVA TePla America, LLC
qEVoriginal/35 nm column	Izon	SP5	SEC column
QSIL 216 Silicone Elastomer Kit	PP&S
Resin Tough Black	Zortrax
SW40 Ti swing ultracentrifuge rotor	Beckman Coulter	331301
Syringe pump	DK Infusetek	ISPLab002
T175 suspension flask	Sarstedt	83.3912.502
TIM4-Fc protein	Adipogen LifeSciences	AG-40B-0180B-3010
TMB (3,3',5,5'-tetramethylbenzidine)	SigmaAldrich	T0440-100ML	Horseradish peroxidase substrate
Tween20	SigmaAldrich	P1379-100ML
Ultracentrifuge Optima L100XP	Beckman Coulter
Ultrasonic cleaning unit P 60 H	Elma Schmidbauer GmbH
Universal Microplate Spectrophotometer	Bio-Tek instruments	71777-1
Urine collection cup, 150mL, sterile	APTACA	2120_SG
Whatman Anotop 25 Syringe Filter	SigmaAldrich	68092002
Zetasizer Nano ZS	Malvern Panalytical		dynamic light scattering (DLS) system
Zortrax Inkspire	Zortrax