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Design of an IDS Fusion Protein Containing TAT

(1) The TAT peptide fragment was synthesized by a biotechnology company and then PCR amplified to obtain TAT fragments containing homologous recombination sequences at both ends.

(2) Using existing LAMP2A and IDS plasmids as templates, we amplified fragments containing homologous recombination sequences. The resulting homologous recombination fragments were verified by agarose gel electrophoresis.

(3) The pLVX-N1-ACGFP vector was subjected to double digestion with the restriction enzymes EcoR I and Xba I, followed by identification of the digested products using nucleic acid electrophoresis. Linearized plasmid vector was obtained through recovery and purification using a DNA recovery kit.

Plasmid Ligation by Homologous Recombination

• 1.Dilute the homologous recombination fragments obtained from the previous steps using ddH₂O according to the calculated ratios.


• 2.Prepare a recombinant reaction mixture on ice using the diluted TAT fragment, LAMP2A fragment, IDS fragment, and linearized vector fragment with the 2× ClonExpress Mix provided by the ClonExpress® Ultra kit. For the multi-fragment homologous recombination reaction, the optimal amounts for each fragment should follow this formula: Optimal amount of cloning vector = [0.02 × number of base pairs in the cloning vector] ng (0.03 pmol); Optimal amount for each fragment = [0.02 × number of base pairs in each fragment] ng (0.03 pmol).


• 3.Gently mix the solution using a pipette (do not vortex) and briefly centrifuge to collect the reaction mixture at the bottom of the tube.


• 4.Incubate in a water bath at 50°C for 30 minutes, then immediately transfer to ice to cool.

Construction of plasmids

(1) Linearization of plasmid template: the pLVX-Puro vector was double digested with restriction enzymes EcoR I and Xba I, and the digestion product was identified by nucleic acid electrophoresis. The linear electrophoresis fragment was recovered and purified by DNA recovery and purification kit to obtain the linearized plasmid vector.

(2) The fragment containing homologous recombination sequence was amplified by using the target plasmid as template.

(3) Ligation:The linearized vector and the IDS fragment were ligated using a homologous recombination enzyme. The mixture was incubated at 50°C and then cooled on ice.

(4) Transformation: A vial of DH5α competent cells was thawed on ice. We added 10 µL of the recombinant product to the competent cells and gently flicked the tube to mix. The cells were incubated on ice for 30 minutes, followed by a heat shock at 42°C for 90 seconds. After heat shock, the cells were immediately placed on ice to cool for 3–5 minutes. We then added LB liquid medium (without antibiotics) and cultured the cells at 37°C with shaking at 220 rpm for 1 hour. After incubation, the cells were centrifuged at 5,000 rpm for 5 minutes, and the supernatant was discarded. The remaining cells were resuspended in the residual medium and spread evenly on plates containing ampicillin for selection. The plates were inverted and incubated at 37°C for 12–16 hours.

(5) Colony Identification:After overnight incubation, several monoclonal colonies formed on the transformation plates. We picked several clones from the plates and inoculated them onto blank plates containing ampicillin for storage. Colony PCR was performed using the universal primers CMV-F and PGK-R (Table 2.2). Positive colonies with PCR product sizes around 1700 bp were selected for Sanger sequencing.

(6) Expansion and Storage of Bacterial Strains
The bacterial strains with correct sequencing results were expanded by picking colonies from the storage plates and transferring them into LB liquid medium containing antibiotics. The cultures were incubated overnight at 37°C with shaking at 220 rpm. From each of the two recombinant plasmid cultures, 500 µL of the bacterial liquid was aliquoted into cryovials, and 500 µL of 60% glycerol was added. The contents of the cryovials were mixed thoroughly, sealed, and stored at -80°C.

Lentiviral Packaging

(1) Observe the number of HEK293T cells under an inverted microscope. Transfection can be performed when the cell density in the field of view reaches approximately 70% confluency.

(2) One hour prior to transfection, discard the original culture medium and add 5 mL of DMEM containing 5% FBS without antibiotics. Place the culture flask in the incubator to prepare.

(3) Take two 15 mL centrifuge tubes and add 250 µL of Opti-MEM medium to each. To one tube, add 30 µL of PEI (1 mg/mL), mix gently, and let it sit for 5 minutes to prepare the PEI dilution. In the other tube, add the three-plasmid system, mix gently, and let it sit for 5 minutes to prepare the plasmid DNA dilution.

(4) Add the PEI dilution to the plasmid DNA dilution and gently mix by pipetting. The three-plasmid lentiviral packaging system is outlined in the following table:

After incubating the infection reaction mixture at room temperature for 15 minutes, add it to the HEK293T cells and gently swirl to evenly distribute the mixture in the culture flask. Place the culture flask in the incubator for 24 hours.

(6) Discard the transfection reaction mixture and add 10 mL of complete medium to continue culturing for another 48 hours.

(7) Collect the supernatant from the culture flask, centrifuge at 1000 rpm for 5 minutes, and then filter the supernatant using a 0.22 µm filter membrane to obtain the viral solution.

Lentiviral Transfection

• 1.Seed the cells to be transfected into a 6-well plate. Under an inverted microscope, observe the cell density and proceed with transfection when the density is around 40%.


• 2.Discard the medium from the culture flask and add 2 mL of viral solution. Transfect for 4–6 hours.


• 3.Add 2 mL of complete medium and continue culturing for 48 hours.


• 4.Discard the mixture of viral solution and medium, and add 2 mL of complete medium to continue culturing for another 24 hours.


• 5.Discard the medium and add complete medium containing 3 µg/mL puromycin for cell selection.


• 6.Continue the selection process for one week, changing the medium daily to remove floating apoptotic cells. When the cell density becomes too high, perform cell passage until no dead cells remain in the culture medium.


• 7.Obtain a stable pLVX-IDS-expressing cell line through long-term selection, and subsequently perform cell cryopreservation.

Exosome Extraction by Differential Centrifugation

(1) Seed the stable transfected cell line into a 10 cm culture dish and grow in DMEM complete medium.

(2) When the cell density reaches approximately 60–70%, discard the complete medium and add serum-free DMEM medium to cover the bottom of the dish. Continue incubation in the incubator for 48 hours.

(3) Transfer the medium to a centrifuge tube and centrifuge at 4°C, first at 300 g for 10 minutes, then at 2000 g for 20 minutes, and finally at 10,000 g for 30 minutes. After this, transfer the supernatant to an ultracentrifuge tube and equilibrate. Centrifuge at 105,400 g at 4°C for 120 minutes.

(4) Discard the supernatant and resuspend the pellet in pre-cooled sterile PBS solution to prepare the exosome suspension.

(5) The exosome suspension can be stored at -80°C for future use. Through the differential centrifugation method, we obtained exosomes produced by the stable transfected cell line, providing experimental materials for subsequent cell and animal studies.

After incubating the infection reaction mixture at room temperature for 15 minutes, add it to the HEK293T cells and gently swirl to evenly distribute the mixture in the culture flask. Place the culture flask in the incubator for 24 hours.

Characterization of Exosomes by Transmission Electron Microscopy (TEM)

The purified exosomes were fixed using a solution of 4% paraformaldehyde and 4% glutaraldehyde, and then placed on carbon-coated copper grids. The grids were soaked in 2% phosphotungstic acid solution, and data was collected using an electron microscope at a voltage of 80 kV.

Extraction of Total Protein from Cells and Exosomes

(1) When the cells reach 80% confluency, discard the culture medium and wash the cells with pre-chilled PBS solution, repeating this 1–2 times.

(2) Place the culture flask on ice and add an appropriate amount of RIPA lysis buffer containing protease inhibitors. Lyse the cells for 30 minutes while gently shaking the flask.

(3) Use a cell scraper to gently detach the cells from the bottom of the flask and collect the cell lysate. Centrifuge at 14,000 rpm for 15 minutes at 4°C, and the supernatant obtained is the total protein extract from the cells.

(4) Remove the exosome suspension from the -80°C freezer, thaw it at room temperature, and add an appropriate amount of RIPA lysis buffer containing protease inhibitors. Lyse for 30 minutes while gently shaking the centrifuge tube.

(5) Centrifuge at 14,000 rpm for 15 minutes at 4°C, and the supernatant obtained is the total protein extract from the exosomes.

Protein Concentration Determination by BCA Assay

(1) Prepare a 0.5 mg/mL BSA protein standard solution according to the BCA assay kit instructions. Dilute the standard solution with PBS to concentrations of 0.025, 0.05, 0.1, 0.2, 0.3, 0.4, and 0.5 mg/mL, and transfer 20 µL of each into 0.5 mL EP tubes.

(2) Dilute the total protein extract with PBS at a 5:1 ratio and transfer 20 µL to a 0.5 mL EP tube.

(3) Prepare the BCA working reagent based on the amounts of the standard and sample solutions, and add it to the standard and sample solutions to create the reaction mixture. Mix by vortexing and briefly centrifuge to collect the mixture at the bottom of the tube.

(4) Place the EP tubes in a 60°C dry oven for 10 minutes, then transfer the reaction liquid to a 96-well plate and measure the absorbance at 562 nm using an enzyme-linked immunosorbent assay (ELISA) reader.

(5) Plot a standard curve based on the measurement results and calculate the protein concentration of the samples.

Western Blot Detection of IDS Protein Levels

(1) Adjust the sample concentrations to be equal using PBS, then add 5× loading buffer, mix gently, and heat in a water bath at boiling temperature for 10 minutes.

(2) Prepare the SDS-PAGE gel according to the molecular weight of the target protein and configure the SDS-PAGE electrophoresis buffer.

(3) Load the protein samples and markers, and run the electrophoresis according to the following program:

(4) Soak the PVDF membrane in methanol for activation, then transfer the proteins at a constant current of 250 mA for 120 minutes at 4°C.

(5) Soak the transferred PVDF membrane in a 3% BSA solution and incubate on a shaker at room temperature for 90 minutes to block.

(6) Cut the PVDF membrane according to the different target protein molecular weights and immerse it in the corresponding primary antibody dilution. Incubate overnight on a shaker.

(7) Recover the antibody and wash the PVDF membrane with TBST buffer for 5 minutes, three times.

(8) After washing, add the corresponding secondary antibody at a dilution of 1:5000 and incubate on a shaker at room temperature for 1 hour.

(9) Discard the secondary antibody and wash the PVDF membrane with TBST buffer for 5 minutes, three times.

(10) Use Pro-light HRP chemiluminescent detection reagent for visualization and scan the image using a gel imaging system.

Fluorescent Labeling of Exosomes

(1) Dissolve DiI dye in DMSO to prepare a dye dilution with a concentration of 10 mg/mL.

(2) Add the dye dilution to the exosome suspension to achieve a final concentration of 10 μM, and incubate in the dark at room temperature for 30 minutes.

(3) Centrifuge at 10,000 g for 30 minutes at 4°C, transferring the supernatant to a 10 kDa ultrafiltration tube.

(4) Centrifuge again at 10,000 g for 5 minutes at 4°C, discard the filtrate, and wash the exosomes with sterile PBS.

(5) Resuspend the pellet in sterile PBS to create a labeled exosome suspension.

Detection of Exosome Uptake by Flow Cytometry

(1) Seed U87MG cells at a density of 1×10^5 in a 12-well plate, setting up three groups with three replicates each.

(2) Once the cell density reaches 70%, add complete medium containing HEK293T sEVs, IDS sEVs, and TAT-IDS sEVs, then incubate the 12-well plate in a CO2 incubator for 2 hours.

(3) Discard the supernatant and wash the cells with PBS.

(4) Add trypsin to digest the cells, creating a cell suspension. Centrifuge at 1,000 rpm for 5 minutes at 4°C, discard the supernatant, and resuspend the pellet in PBS to prepare a cell suspension.

(5) Analyze the exosome uptake in U87MG cells using flow cytometry.

Detection of Exosome Uptake by Fluorescence Imaging

(1) Prepare 18 mm cover slips by soaking them in alcohol for 24 hours, then expose them to UV light for 2 hours.

(2) Place the cover slips in a 12-well plate and seed U87MG cells at a density of 1×10^5, setting up three groups with three replicates each.

(3) Once the cell density reaches 70%, add IDS sEVs and IDS Exosignal sEVs, and incubate the plate in a CO2 incubator for 2 hours.

(4) Discard the supernatant and wash the cells with PBS.

(5) Add 4% paraformaldehyde to fix the cells for 15 minutes at room temperature in the dark, followed by the addition of 10 mM glycine to neutralize the fixative.

(6) Stain the cells with DAPI for 10 minutes, then discard the staining solution and wash the cells with PBS.

(7) Transfer the cover slips to glass slides, adding mounting medium to secure the cover slips.

(8) Use a laser scanning confocal microscope to capture images of the slides.

Establishing an In Vitro Blood-Brain Barrier Model

(1) Remove Corning® Matrigel® from -20°C storage and thaw it on ice. Dilute with pre-cooled (4°C) serum-free DMEM medium. Uniformly coat the diluted Matrigel in the upper chamber of a 12-well co-culture plate and the bottom of the Transwell insert.

(2) Place the 12-well plate in an incubator at 37°C for 1 hour to solidify. Afterward, aspirate the supernatant.

(3) Seed HCMEC/D3 cells at a density of 5×10^4 cells in the upper chamber, adding 400 μL of DMEM medium containing 10% FBS to the upper chamber and 1950 μL of the same medium to the lower chamber.

(4) Continue incubating the 12-well plate in the cell culture incubator, changing the medium every two days. Monitor cell growth under an inverted microscope during this period.

(5) Discard all media from both the upper and lower chambers of the co-culture plate, then add 500 μL of a serum-free 10 μg/mL RB-Dextran solution to the Transwell upper chamber. Add 1 mL of serum-free medium to the lower chamber.

(6) Incubate the 12-well plate in the incubator at 37°C for 30 minutes.

(7) Transfer 200 μL of the medium from the lower chamber to a black transparent-bottom 96-well cell culture plate, and measure the leakage of the RB-Dextran solution using a fluorescence microplate reader (excitation wavelength 544 nm, emission wavelength 599 nm).

(8) Collect and analyze experimental data continuously for 8 days.

(9) Seed U87MG cells at a density of 5×10^4 cells in the lower chamber, adding 1950 μL of DMEM medium containing 10% FBS. After 1 day of culture, the BBB required for the experiment can be obtained.

Detection of Exosome Uptake in the In Vitro BBB Model by Flow Cytometry

(1) Retrieve the completed in vitro blood-brain barrier model. Discard the medium from both the upper and lower chambers, then add 400 μL of complete medium containing IDS sEVs and IDS Exosignal sEVs to the upper chamber, and 1950 μL of complete medium to the lower chamber. Incubate the 12-well plate in the cell culture incubator for 24 hours.

(2) Discard the culture medium and wash the cells in the lower chamber with PBS.

(3) Add trypsin to the lower chamber to digest the cells, creating a cell suspension. Centrifuge at 4°C and 1000 rpm for 5 minutes, discard the supernatant, and resuspend the pellet in PBS to create a cell suspension.

(4) Use flow cytometry to detect the uptake of exosomes by U87MG cells in the lower chamber to evaluate the BBB permeability efficiency of different exosomes.

Detection of Exosome Uptake in the In Vitro BBB Model by Fluorescence Imaging

(1) Prepare 18 mm cover slips and immerse them in alcohol for 24 hours, then expose them to UV light for 2 hours.

(2) Place the cover slips in a 12-well plate used to construct the in vitro blood-brain barrier model. Discard the medium from both the upper and lower chambers, then add 400 μL of complete medium containing IDS sEVs and IDS Exosignal sEVs to the upper chamber, and 1950 μL of complete medium to the lower chamber. Incubate the 12-well plate in the cell culture incubator for 24 hours.

(3) Discard the culture medium and wash the cells in the co-culture chamber with PBS.

(4) Under light-protected conditions at room temperature, add 4% paraformaldehyde to fix for 15 minutes. Then add 10 mM glycine.

(5) Add DAPI dye to the lower chamber for 10 minutes for staining. Discard the staining solution and wash the cells with PBS.

(6) Transfer the cover slips to slides, adding a mounting medium to fix the cover slips.

(7) Use a laser confocal microscope to image the slides.