Porous monolithic polymer for separation and purification of biological particles, method for producing porous monolithic polymer for separation and purification of biological particles, and method for separating and purifying biological particles comprising phospholipids using porous monolithic polymer for separation and purification of biological particles

Abstract

[Abstract] [Problem] To provide a porous monolithic polymer for separation and purification of biological particles, a method for producing the same, and a method for separating and purifying biological particles comprising phospholipids using porous monolithic polymer for separation and purification of biological particles, which can be used for efficient and rapid purification of EVs. [Solution] The present invention relates to a porous monolithic polymer for separation and purification of biological particles, said polymer being hybridized with an olefinic thermoplastic resin and an inorganic oxide, a method for producing the same, and a method for separating and purifying biological particles with phospholipids on the surface thereof by means of said porous monolithic polymer for separation and purification of biological particles. [Selected Figure] Fig. 1a

Description

Porous monolithic polymer for separating and purifying biological particles, method for producing porous monolithic polymer for separating and purifying biological particles, and method for separating and purifying biological particles composed of phospholipids using porous monolithic polymer for separating and purifying biological particles

[0001] The present invention relates to a porous monolithic polymer for separating and purifying biological particles, which is a mixture of a polymer and an inorganic oxide, particularly TiO2, a method for producing said polymer, and a method for separating and purifying biological particles composed of phospholipids.

[0002] Extracellular vesicles (EVs) are nanosized, membranous particles released by all types of cells and present in various bodily fluids, such as blood, urine, and saliva. In particular, small EVs (SEVs), particles with a diameter of approximately 100 nm, contain various biomolecules, including proteins, lipids, and nucleic acids, and reflect the physiological and pathological states of donor cells. This characteristic not only makes SEVs highly desirable as noninvasive biomarkers in bodily fluids, but also draws attention in the drug discovery industry as a promising tool for drug delivery due to their ability to cross biological barriers and deliver cargo to target cells. Numerous studies have demonstrated the potential of SEVs carrying therapeutic cargos to treat various diseases, including cancer, neurodegenerative diseases, and infectious diseases.

[0003] Despite this, researchers have yet to achieve an optimal approach for collecting SEVs. Ultracentrifugation (UC) is the gold-standard method for SEV purification, which can purify SEVs from cellular debris and contaminating proteins based on their density or size. However, UC requires complex procedures and long processing times, resulting in low recovery rates and limited throughput. Alternatively, immunoaffinity based on membrane markers can be used to collect highly purified SEVs. However, biological ligands are expensive and can only be manipulated in limited quantities. This makes immunoaffinity methods infeasible for processing large amounts of samples. Another drawback is the heterogeneity of SEV molecular populations, which poses a risk of overlooking specific SEV populations bearing target molecules. Other methods, such as polymer precipitation, size-exclusion chromatography, microfluidic chips, and flow field-flow fractionation, have been developed, but they suffer from protein contamination, complex procedures, limited small-scale experiments, and the need for sophisticated equipment. Therefore, to accelerate clinical applications and the development of the pharmaceutical industry, an efficient and rapid purification method for SEVs that can be generally applied to a wide range of biological samples is needed.

[0004] For example, the invention described in Patent Document 1 proposes a granular porous body to be used in a liquid chromatography column for separating biomolecules such as peptides, proteins, or nucleic acids. However, the invention of Patent Document 1 is limited to small-scale implementation because it takes time and effort to prepare the porous body, it also takes time to separate and purify, and there is a limit to the amount of sample that can be loaded at one time.

[0005] The invention described in Patent Document 2 proposes a method for separating and recovering extracellular vesicles from a sample in the presence of a polymer. However, in the conventional process for separating and purifying extracellular vesicles, the method merely involves adding a water-soluble polymer to the sample to improve the accuracy of separation.

[0006] JP 2016-70937 WO 2020 / 080387

[0007] The present invention aims to provide a porous monolithic polymer for the separation and purification of biological particles, which can be used to efficiently and quickly purify EV, a method for producing the same, and a method for the separation and purification of biological particles composed of phospholipids using the monolithic polymer for the separation and purification of biological particles. Here, a monolith refers to a porous mass having a continuous skeleton and voids, and is a functional material that can be used as a column or a catalyst immobilization support due to its structural characteristics.

[0008] The present inventors have conducted extensive research to address the aforementioned problems of the prior art and have found that a spongy monolithic polymer (SPM), a mixture of an olefin-based thermoplastic resin and an inorganic oxide, provides a simple, rapid, and highly efficient method for purifying extracellular vesicles (EVs). Surprisingly, this method can purify 130.1 times smaller EVs from the same volume of cell culture medium within one hour, with minimal steps, compared to ultracentrifugation. Furthermore, the flexibility of the material's shape and size allows it to be applied to small clinical samples and large volumes of bovine serum. The inorganic oxide-hybridized SPM of the present invention can serve as a universal platform for collecting and purifying SEVs, significantly contributing to drug discovery research and biomarker discovery using EVs.

[0009] The invention according to claim 1 is a porous monolithic polymer for separating and purifying biological particles, which is a hybrid of an olefin thermoplastic resin and an inorganic oxide.

[0010] The invention according to claim 2 may be the porous monolithic polymer for separating and purifying biological microparticles according to claim 1, wherein the thermoplastic resin is at least one selected from the group consisting of poly(ethylene-co-glycidyl methacrylate) (PEGM), ethylene-vinyl acetate copolymer, high-density polyethylene, low-density polyethylene, olefin-based elastomers, and polyamide-based elastomers.

[0011] The invention according to claim 3 may be the porous monolithic polymer for separating and purifying biological microparticles according to claim 1, wherein the inorganic oxide is one or more selected from the group consisting of titanium oxide, magnesium oxide, aluminum oxide, silicon dioxide, and calcium phosphate.

[0012] The invention according to claim 4 relates to the porous monolithic polymer for separating and purifying biological microparticles according to claim 1, characterized in that the porosity is 50 to 90%, the pore diameter is 1 μm to 200 μm, and the melting point is 60°C to 180°C.

[0013] The invention according to claim 5 is a method for producing a porous monolithic polymer for separating and purifying biological microparticles according to claim 1, which comprises melting the thermoplastic resin, kneading the thermoplastic resin with a pore-forming agent and a pore-forming aid together with the inorganic oxide, and molding the resin, followed by eluting the pore-forming agent with an aqueous solvent.

[0014] The invention according to claim 6 may be the manufacturing method according to claim 5, wherein the pore-forming agent is an organic or inorganic substance that does not react with the thermoplastic resin and the inorganic oxide during kneading.

[0015] The invention according to claim 7 may be the manufacturing method according to claim 6, wherein the pore-forming agent is one or more selected from the group consisting of pentaerythritol, corn starch, anhydrous sodium sulfate, starches such as wheat starch and potato starch, artificial sweeteners such as aspartame and saccharin, polysaccharides such as hemicellulose, acid-soluble inorganic compounds such as calcium carbonate and magnesium carbonate, and table salt.

[0016] The invention according to claim 8 may be the manufacturing method according to claim 5, wherein the diameter of the particles of the pore-forming agent is 1 μm to 200 μm.

[0017] The invention according to claim 9 may be the manufacturing method according to claim 5, characterized in that the pore-aiding agent is a monomer or polymer of a polyhydric alcohol.

[0018] The invention according to claim 10 may be the manufacturing method according to claim 9, wherein the pore-forming aid is at least one selected from the group consisting of poly(oxyethylene-oxypropylene))triol, polyalkylene glycols such as polyethylene glycol, and diethylene glycol.

[0019] The invention according to claim 11 relates to a method for separating and purifying biological particles having phospholipids on their surfaces by using the porous monolithic polymer for separating and purifying biological particles according to claim 1.

[0020] The invention according to claim 12 may be the method according to claim 11, wherein the biological microparticles are extracellular vesicles.

[0021] According to the first aspect of the present invention, there is an effect that a porous monolithic polymer, which is a mixture of a thermoplastic resin and an inorganic oxide, can be used for separating and purifying biological particles.

[0022] According to the second aspect of the present invention, a porous monolithic polymer for separating and purifying biological particles can be obtained from readily available materials.

[0023] According to the third aspect of the present invention, a porous monolithic polymer for separating and purifying biological particles can be obtained from readily available materials.

[0024] According to the fourth aspect of the present invention, a porous monolithic polymer for separating and purifying biological particles can be obtained from readily available materials, which serves as a novel and excellent medium for use in separating and purifying biological particles.

[0025] According to the fifth aspect of the present invention, it is possible to produce a porous monolithic polymer in which the pore size and porosity are adjusted as desired.

[0026] According to the sixth aspect of the present invention, when producing a porous monolithic polymer, it is possible to select a pore-forming agent from a wide range of options.

[0027] According to the seventh aspect of the present invention, it is possible to produce a porous monolithic polymer having pores by using readily available materials.

[0028] According to the eighth aspect of the present invention, a porous monolithic polymer having pores of 1 μm to 200 μm can be produced, which has the effect of facilitating the capture of biological microparticles.

[0029] According to the ninth aspect of the present invention, there is an effect that a porous monolithic polymer can be produced using a readily available pore-assisting agent.

[0030] According to the tenth aspect of the present invention, there is an effect that a porous monolithic polymer can be produced using a readily available pore-assisting agent.

[0031] According to the eleventh aspect of the present invention, the porous monolithic polymer for separating and purifying biological particles of the present invention adsorbs the phospholipids of the biological particles, thereby providing the effect of facilitating the separation and purification of the biological particles.

[0032] According to the invention of claim 12, the porous monolithic polymer for separating and purifying biological microparticles of the present invention adsorbs phospholipids of extracellular vesicles, thereby providing the effect of facilitating the separation and purification of extracellular vesicles.

[0033] 1 shows SEM images of an SPM and a TiO2-SPM according to one embodiment of the present invention. 2 shows the pore characteristics of a TiO2-SPM according to one embodiment of the present invention measured by a mercury porosimeter. 3 shows the LNP adsorption capacity of a TiO2-SPM column according to one embodiment of the present invention. 4 shows the number of LNPs in each fraction separated and purified by a TiO2-SPM column according to one embodiment of the present invention. 5 shows the particle size profile of LNPs and the elution rate of TiO2-SPM in a loaded sample according to one embodiment of the present invention. 6 shows the lipid components of prepared LNPs. 7 shows the particle size profile of LNPs and the elution rate of TiO2-SPM in a loaded sample according to one embodiment of the present invention. 8 shows the dynamic light scattering distribution curve of nanoparticles in a loaded sample. 9 shows the particle size profile of LNPs and the elution rate of TiO2-SPM in a loaded sample according to one embodiment of the present invention. 10 shows the size distribution of LNPs in the elution fractions determined by nanotracking analysis. This figure shows the number of NPs in the elution fraction from a protein-immobilized TiO2-SPM according to one embodiment of the present invention. Proteins were immobilized to suppress nonspecific adsorption. Each centrifugation was performed at 1000 × g for 1 minute. This figure shows the number of NPs in the elution fraction at different loading centrifugation speeds using a protein-immobilized TiO2-SPM according to one embodiment of the present invention. This figure shows the number of NPs collected from culture supernatant by SEV purification according to one embodiment of the present invention. (Right) This figure shows the number of NPs collected by UC from 38.5 mL of culture supernatant and the number of NPs collected by TiO2-BSA-SPM from 0.4 mL of culture supernatant. (Left) Equivalent values ​​per mL of culture supernatant. Particle size profiling of NPs collected by TiO2-BSA-SPM and UC. This figure shows transmission electron microscope images of NPs collected by TiO2-BSA-SPM. Figure 1 shows single particle analysis by imaging flow cytometry of EVs collected by TiO2-BSA-SPM and stained for lipid membranes. Figure 2 shows single particle analysis of particles after Triton-X treatment. Figure 3 shows overlapping proteins quantified in NPs collected by TiO2-BSA-SPM and UC.Figure 1 shows the top 5 cellular components from GO enrichment analysis of only proteins identified in NPs collected with TiO2-BSA-SPM. Figure 2 shows the top 5 cellular components from GO enrichment analysis of only proteins identified in NPs collected with UC. Figure 3 shows the characteristics of EV-depleted FBS produced with syringe barrel TiO2-BSA-SPM. a) Particle size profiling of NPs in FBS and EV-depleted FBS prepared with TiO2-BSA-SPM and UC. Figure 4 shows the number of NPs in EV-depleted FBS. Images of HEK293T cells cultured with EV-depleted FBS prepared with TiO2-BSA-SPM (right) and UC (left). Figure 1 shows the number of viable cells cultured in Dulbecco's modified Eagle's medium (Thermo Fisher Scientific) containing 10% EV-depleted FBS (viable cells were counted using an automated cell counter (LUNA-FL™, Logos Biosystems)). Figure 1 shows the gating strategy for single particle analysis of SEV collected by TiO2-BSA-SPM. Gating to distinguish flow control beads. "Speed ​​Beads" is the region representing flow control beads, and "Region 1" is the region representing objects derived from the sample. The vertical axis represents side scatter intensity, and the horizontal axis represents the bright-field area of ​​the objects. Figure 1 shows the gating strategy for single particle analysis of SEV collected by TiO2-BSA-SPM. Gating to remove aggregates from particles detected in Region 1. The vertical axis plots the area of ​​the side scatter image, and the horizontal axis plots the side scatter intensity. Figure 1 shows the characteristics of NPs collected from serum samples of healthy donors by TiO2-BSA-SPM. a) Size distribution of NPs collected from serum samples. b) Characteristics of NPs collected from serum samples of healthy donors by TiO2-BSA-SPM. c) Overlap of proteins quantified in serum samples and NPs. Proteomic analysis was performed three times using independent experiments with technical replicates. c) Characteristics of NPs collected from serum samples of healthy donors by TiO2-BSA-SPM. c) Top five cellular components from GO enrichment analysis of only proteins identified in NPs.

[0034] Hereinafter, embodiments of the porous monolithic polymer for separating and purifying biological microparticles according to the present invention, its manufacturing method, and a method for separating and purifying biological microparticles composed of phospholipids using the porous monolithic polymer for separating and purifying biological microparticles will be described with reference to the drawings.

[0035] The porous monolithic polymer for separating and purifying biological particles of the present invention is a kneaded product of an olefin-based thermoplastic resin and an inorganic oxide. By kneading the thermoplastic resin and the inorganic oxide, the inorganic oxide is hybridized with the thermoplastic resin.

[0036] Examples of thermoplastic resins used in the present invention include flexible and inexpensive polymers such as poly(ethylene-co-glycidyl methacrylate) (PEGM), ethylene-vinyl acetate copolymer, high-density polyethylene, low-density polyethylene, olefin-based elastomers, and polyamide-based elastomers. Two or more of these thermoplastic resins can also be used in combination. The resulting sponge-like monolithic polymers (SPMs) can be used as separation media of various shapes and sizes.

[0037] Suitable examples of inorganic oxides used in the present invention include metal oxides such as titanium oxide, aluminum oxide, and magnesium oxide, as well as inorganic oxides such as silica (silicon dioxide), particularly fumed silica, and calcium phosphate, particularly tricalcium phosphate. Two or more inorganic oxides can also be used in combination.

[0038] The sponge-like monolithic polymer (SPM) of the present invention has pores with a pore size of 1 μm to 200 μm and a porosity of 50% to 90%. Pore sizes smaller than 1 μm induce clogging, while pore sizes larger than 200 μm result in biological particles not being captured and being lost. Porosity smaller than 50% increases the time and effort required for separation and purification, while porosity larger than 90% makes the SPM brittle and unsuitable for separation and purification. More preferably, the SPM has pores with a pore size of 5 μm to 10 μm and a porosity of 70% to 80%. This pore size is much larger than the diameter of extracellular vesicles (EVs), particularly small particles (SEVs), which are approximately 100 nm in diameter, and therefore provides a flow path suitable for low-pressure chromatographic separation of SEVs without causing clogging.

[0039] The TiO2-hybridized SPM of the present invention selectively captures phosphate groups, which are abundant on the lipid bilayer surface of SEV, via bidentate bonds. This bidentate bond is pH-responsive and reversible, capturing SEV under neutral conditions and releasing it under basic conditions (pH 10-12). Hybridizing silica to the SPM of the present invention selectively captures basic amino acid residues of SEV. Furthermore, hybridizing calcium phosphate to the SPM allows for the capture of phosphate and hydroxyl groups.

[0040] The SPM of the present invention may be prepared by kneading a pore-forming agent and a pore-aiding agent together with the thermoplastic resin when mixing the inorganic oxide. 15 to 35 wt% of the thermoplastic resin melted at 60 to 180°C is uniformly mixed with 1 to 20 wt% of the inorganic oxide, 50 to 80 wt% of the pore-forming agent, and 7 to 15 wt% of the pore-aiding agent, and the mixture is molded. The pore-forming agent is then eluted with an aqueous solvent to obtain the SPM. The thermoplastic resin loses fluidity at temperatures below 80°C and is unable to maintain its shape at temperatures above 180°C. Furthermore, if the thermoplastic resin is less than 15 wt%, it is compressed by the liquid pressure during liquid delivery, significantly reducing liquid permeability. If the amount is more than 35 wt%, the porosity decreases, resulting in poor liquid permeability, and the surface area within the sponge is reduced, resulting in poor SEV capture performance. If the inorganic oxide content is less than 1 wt%, the amount of biological particles that can be captured will be too small; if it is more than 20 wt%, the efficiency of hybridization with the thermoplastic resin will be reduced, and during the process of eluting the pore-forming agent with an aqueous solvent after molding, much of the inorganic oxide will fall off from the thermoplastic resin, resulting in waste. If the pore-forming agent content is less than 50 wt%, the liquid permeability will deteriorate due to increased density, and if it is more than 80 wt%, the resin will be destroyed. If the pore-forming agent content is less than 7 wt%, the liquid permeability between the pores will be poor, and if it is more than 15 wt%, the surface area inside the sponge will be too small. These weight ratios are determined by balancing the ability of the SPM of the present invention to separate and purify biological particles.

[0041] The pore-forming agent is adjusted to a particle size of 1 to 200 μm. This is to adjust the pore size of the SPM to 1 to 200 μm, and more preferably a particle size of 1 to 10 μm. A particulate hair-like substance that does not dissolve in the thermoplastic resin is used. Preferred materials include starches such as pentaerythritol, corn starch, wheat starch, and potato starch, sugar, and polysaccharides such as hemicellulose, as well as tetramethylolmethane (chemical name: 2,2-dihydroxymethyl-1,3-propanediol) represented by the following formula 1 (Chemical Formula 1).

[0042] The pore aid is used to shorten the water dissolution time of the pore aid. It is preferable to use an agent that has a high boiling point, can improve the fluidity of the resin with a small amount, and has a small heat loss. Suitable agents include poly(oxyethylene-oxypropylene) triol, polyalkylene glycols such as polyethylene glycol, and diethylene glycol.

[0043] The separation and purification of biological particles of the present invention is carried out by filling a column with this SPM and loading a sample. The column used here is not particularly limited, and may be a spin column or a syringe barrel column. Spin columns are suitable for centrifugation, while syringe barrel types are suitable for separation by pressure.

[0044] The bioparticles having phospholipids adsorbed by the hybridized inorganic oxide enter the pores of the SPM, thereby enabling the separation and purification of the bioparticles according to the present invention.

[0045] Our SEV purification platform was developed by hybridizing TiO2 particles with a sponge-like polymer (SPM). The TiO2-SPM was molded into a solid-phase extraction column, which is effective for collecting SEVs from various biological samples via bidentate binding to the embedded TiO2. The large pores of the polymer's sponge-like structure allow for simple flow-through processing by centrifugation or 1 minute of manual pressure. This method, including pretreatment, can be completed within 1 hour and recovers 130.7 times more SEVs than UC from the same volume of cell culture supernatant. Bottom-up proteome analysis also demonstrated that SEVs collected by TiO2-SPM are more comprehensive than UC. This method is applicable to clinical samples, allowing efficient collection of SEVs from 100 μL plasma samples. Furthermore, by modifying the format to a syringe barrel, the adsorption capacity and sample loading volume were increased, enabling complete removal of EVs from 10 mL of FBS (fetal bovine serum). The TiO2-SPM of the present invention can serve as a universal platform for collecting and purifying SEVs, and can make a significant contribution to drug discovery research and biomarker discovery using SEVs.

[0046] The monolithic polymer for biological separation of the present invention will be explained below using examples, but the present invention is not limited to these examples.

[0047] Example 1 Preparation of TiO2-SPM 35 wt% poly(ethylene-co-glycidyl methacrylate) (PEGM) (containing 12% glycidyl methacrylate), 52 wt% pore-forming agent (pentaerythritol) (with a particle size of approximately 10 μm), 7 wt% pore-forming agent (poly(oxyethylene-oxypropylene)) triol), and 20 wt% TiO2 particles were melted at 150°C and uniformly kneaded. The resulting material was extruded into pillars at 130°C. The pillars were immediately cooled in water to obtain rods. The stick-shaped material was then washed with methanol and water using ultrasound to remove soluble compounds.

[0048] The resulting TiO2-hybridized sponge-like monolithic polymer (TiO2-SPM) had a porosity of approximately 75% and a cross-sectional diameter of 8.0 mm. Furthermore, the morphology of the TiO2-SPM was observed using a field-emission scanning electron microscope (JSM6700-M, JEOL, Tokyo, Japan; hereafter referred to as SEM). The SEM image of the prepared TiO2-SPM showed that the TiO2 particles were properly hybridized with the SPM by being exposed on the surface through kneading (Fig. 1). The median pore size of the TiO2-SPM, characterized by mercury porosimetry, was 5.21 μm. The pores were much larger than those of the SEV (Fig. 1b).

[0049] Column Preparation: Next, the TiO2-SPM rods were sliced ​​into 1-cm-thick circles to obtain TiO2-SPM disks. To prepare TiO2-SPM-packed solid-phase extraction (SPE) columns, two types of empty polypropylene cartridges were used based on the intended application and sample volume: spin columns (2 cm × 7.0 mm i.d.) and syringe barrel columns (5 cm × 7.0 mm i.d.). The spin column type is suitable for centrifugation, while the syringe barrel type is suitable for manual SPE. Multiple TiO2-SPM disks can be packed into the syringe barrel type to increase the adsorption capacity of EVs (up to five disks). The diameter of the TiO2-SPM disks (8.0 mm) was larger than the inner diameter of the empty cartridge. Nevertheless, the elasticity of the sponge-like monolith material made packing easy. The packing procedure was simple. The TiO2-SPM disk was immersed in methanol to completely wet it. The disk was then placed parallel to the top of the cartridge and pressed straight down to the bottom using a stainless steel rod. The disk had to be pressed carefully to avoid distortion or wrinkling within the cartridge during the process. The prepared column was then washed with a 1:1 methanol / water solution, five times the volume of the packed TiO2-SPM, to remove the pore template and ensure uniform packing.

[0050] Separation Behavior of Liposomal Nanoparticles on TiO2-SPM To evaluate the adsorption capacity of TiO2-SPM, liposomal nanoparticle (LNP) solutions (0.4 mL) with different concentrations were prepared and loaded onto a TiO2-SPM packed into a spin column. Due to the high permeability resulting from the spongy morphology, the sample loading process was rapidly completed by centrifugation at 1,000 × g for 1 min. After collecting the flow-through fraction, the spin column was washed with phosphate-buffered saline (PBS) (0.4 mL × 5, wash fraction). The amount of LNP adsorbed onto the TiO2-SPM column was estimated by subtracting the sum of the flow-through and wash fractions from the initial amount of LNP loaded (Fig. 1c). The results showed that the number of adsorbed LNPs increased with the amount of LNP loaded, and the maximum adsorption capacity was determined to be 1.17 × 1010 particles. Next, we attempted to elute the adsorbed particles from the TiO2-SPM. Previous reports have shown that high pH conditions can effectively disrupt the bismuth bonds of TiO particles with the phosphate groups on the surface of SEV (a) F. Gao, F. Jiao, C. Xia, Y. Zhao, W. Ying, Y. Xie, X. Guan, M. Tao, Y. Zhang, W. Qin, X. Qian, Chem. Sci. 2019, 10, 1579; b) X. Xiang, F. Guan, F. Jia, H. Li, W. Zhang, Y. Zhang, W. Qin, Anal. Methods 2021, 13, 1591). Therefore, 10% NH3·HO was used as the elution solvent. The collected elution fractions were rapidly replaced with PBS by ultrafiltration (30 kDa MWCO, Amicon®) to protect the lipid bilayer from high pH conditions. As shown in Figure 1d, the adsorbed LNPs were rapidly eluted under high pH conditions, and the elution fraction contained 9.07 × 109 LNPs (84.6% of the adsorbed LNPs). The average diameter of the LNPs in the elution fraction was 97 nm, similar to the size of the previously loaded LNPs (Figure 2c). This adsorption ability was not observed with conventional SPM, indicating effective bidentate binding between the phosphate groups on the LNP surface and the hybridized TiO2. The strong affinity between TiO2-SPM and LNPs was confirmed.

[0051] Example 2 Purification of SEVs from Biological Samples SEVs were purified and collected from the supernatant of cell culture medium of the human embryonic kidney cell line HEK293T (RIKEN BRC) by ultracentrifugation using a TiO2-SPM column. Cells were cultured in Dulbecco's modified Eagle's medium (DMEM, Thermo Fisher Scientific) containing 10% fetal bovine serum (FBS, Thermo Fisher Scientific) at 37°C and 5% CO2 until they reached approximately 70-80% confluence. They were then cultured in EV-depleted medium for 48 hours. Supernatants were prepared from cells at approximately 70-80% confluence and centrifuged at 400 x g for 10 minutes and 2000 x g for 10 minutes at 4°C to remove cell debris and large EVs, respectively. 0.4 mL of the prepared supernatant was loaded onto a TiO2-SPM spin column, and the column was treated as in Example 1. Nanoparticle tracking analysis (NTA) results showed that 1.45 × 108 nanoparticles (NPs) were observed in the elution fraction, indicating successful recovery of SEVs by TiO2-SPM (Fig. 2a, left bar). However, the number of NPs collected from the culture supernatant was much lower than the expected number of LNPs.

[0052] The surface of SEV contains membrane proteins such as CD9 and CD81, which may nonspecifically adsorb to the hydrophobic surface of TiO2-SPM (E.G. Evtuushenko, D.V. Bagrov, V.N. Lazarev, M.a. Livshits, E. Khomyakova, PlosOne2021, 15, e0243738 ., E. Kanao, S. Wada, H. Nishida, T. Kubo, T. Tanigawa, k. Imami, a. Shimoda, k. Umezaki, Y. Sasaki, k. Akiyoshi, J. Adachi, k. Otsuka, Y. Ishihama, Anal. Chem. 2022, 94, 18025.). In addition, in the culture supernatant of a contaminated system, due to competing interactions with other substances, hybridized TiO2 may have a longer adsorption time to SEV than LNP.

[0053] It has also been reported that the incubation time required to confirm sufficient binding between urinary SEV and TiO2 particles was increased by five times (F. Gao, F. Jiao, C. Xia, Y. Zhao, W. Ying, Y. Xie, X. Guan, M. Tao, Y. Zhang, W. Qin, X. Qian, Chem. Sci. 2019, 10, 1579;).

[0054] Example 3: Protein Immobilization To improve the efficiency of NP collection from culture supernatants, we immobilized a protein (bovine serum albumin) on the TiO2-SPM surface to improve the surface hydrophilicity and slow the centrifugation speed. Proteins could be easily immobilized to TiO2-SPM via the surface epoxy groups by incubating at 37°C. 7.34 × 108 nanoparticles were observed in the elution fraction of BSA-immobilized TiO2-SPM (TiO2-BSA-SPM), indicating that the improved hydrophilicity significantly reduced nonspecific adsorption (Figure 3a). The centrifugation speed during sample loading was optimized at 300 × g, and 1.17 × 109 nanoparticles were collected (Figure 3b). The centrifugation process was also completed within 1 minute, but the loaded supernatant did not pass below 300 × g. Surprisingly, the number of nanoparticles recovered from 0.4 mL of culture supernatant using TiO2-BSA-SPM was greater than that recovered from 38.5 mL by ultracentrifugation (Fig. 4a, left). Considering the sample volume loaded, the NP collection efficiency of TiO2-BSA-SPM could be equivalent to 130.7 times that of UC (Fig. 4a, right).

[0055] Here, we investigated the characteristics of NPs in the elution fraction of TiO2-BSA-SPM. The size of the collected NPs was determined by NTA to be approximately the same as that of UC, a result confirmed by observation with a H-7650 transmission S4 electron microscope (Hitachi, Tokyo, Japan; hereafter referred to as "Hitachi") and digital images captured using an XR-41 CCCD camera system (Advanced Microscopy Techniques, Woburn, USA) (Figures 4b and 4c). Furthermore, NPs collected by TiO2-BSA-SPM were fluorescently labeled with ExoSparkler@ (Exosome Membrane Labeling Kit-Red, Dojindo, Kumamoto, Japan), which selectively stains lipid bilayers. Bright-field images of the particles were captured using a 40x magnification lens. The flow rate was set to the slowest setting, and the lasers corresponding to fluorescence and side scattering were used at maximum power. Gating was performed to analyze single NPs (Figures 6a and 6b). Specifically, flow control beads were distinguished based on side-scattered light intensity and bright-field image size, while other objects were considered particles originating from the sample. Aggregates were also excluded based on scattered light intensity and image size, and nanoparticles were analyzed for dye-derived fluorescence and scattered light. Dye-stained nanoparticles were detected in samples collected by the TiO2-BSA-SPM (Figure 4d). Furthermore, the addition of Triton-X significantly reduced the number of particles with detected dye fluorescence (Figure 4e). This suggests that the addition of surfactant disrupted the particles. These results clearly demonstrate that the nanoparticles collected by the TiO2-BSA-SPM contain lipid bilayers.

[0056] Example 4: LC-MS / MS Analysis Proteome analysis was also performed by LC-MS / MS to investigate the protein composition of NPs collected with TiO2-BSA-SPM and UC. To confirm reproducibility, three replicates were prepared for each method, and only proteins quantified in two or three replicates were recognized as identified proteins. The sample amount for LC-MS / MS measurements was measured using a NanoDrop (Thermo Fisher Scientific) at absorbance 205 nm with an extinction coefficient of 31 and normalized to 0.5 μg per measurement. As a result, 1,174 proteins were identified from nanoparticles collected with TiO2-BSA-SPM, including 94% of the top 100 exosome marker proteins registered in Exocarta (http: / / exocarta.org / index.html). In NPs collected with TiO2-BSA-SPM and UC, 944 proteins were commonly identified, including membrane proteins such as CD9, CD63, and CD81, as well as the typical exosome marker protein, Syntenin-1 (Fig. 4f). Gene Ontology (GO) annotations obtained from the UniProt database using the David 6.8 database revealed that 230 proteins identified exclusively in NPs collected with TiO2-BSA-SPM were assigned the keyword "extracellular exosome" as a cellular component, whereas 32 proteins in UC were not assigned this keyword (Fig. 4f, g, h). These results suggest that TiO2-BSA-SPM can collect more diverse and pure SEV populations than UC.

[0057] To test this hypothesis, we examined the up-regulated proteins in SEVs collected by TiO2-SPM and UC. Fold change (FC) and p-values ​​were calculated using label-free quantification (LFQ) intensities with Welch's t-test. An FC of >1 and p-value of <0.05 were set as the criteria for selecting up-regulated proteins. GO analysis identified both groups of proteins as cellular components, assigned by the keyword "extracellular exosome." These results demonstrate that the average protein composition of collected SEVs differs significantly between the two methods, supporting the possibility that some SEVs collected by TiO2-BSA-SPM may be missed by UC. In summary, TiO2-BSA-SPM is a significantly superior SEV purification method to UC in terms of both processing time and collection efficiency.

[0058] Supplementary Explanation of the Examples Preparation of Fluorescent Liposomal Nanoparticles Lipid stock solutions were mixed and homogenized in chloroform at a molar ratio of 49.8% 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC, Tokyo Chemical Industry Co., Ltd., Tokyo, Japan), 14.9% 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE, Tokyo Chemical Industry Co., Ltd.) and 14.9% cholesterol (Tokyo Chemical Industry Co., Ltd.), 19.9% ​​1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-L-serine (POPS, Tokyo Chemical Industry Co., Ltd.), and 0.5% 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-yl) (ammonium salt) (NBD-PE, Tokyo Chemical Industry Co., Ltd.).

[0059] <Isolation of SEVs from HEK293 Cells by Ultracentrifugation> The human embryonic kidney cell line HEK293T [obtained from Riken BRC] was cultured in Dulbecco's modified Eagle's medium (DMEM, Thermo Fisher Scientific) containing 10% fetal bovine serum (FBS, Thermo Fisher Scientific) at 37°C and 5% CO2. Cells were grown until approximately 70-80% confluent and then cultured in EV-depleted medium for 48 hours. SEVs were collected from the resulting culture supernatant by ultracentrifugation to reduce size-related heterogeneity. The supernatant was centrifuged at 300 × g for 10 minutes, 2,000 × g for 10 minutes, and 10,000 × g for 30 minutes. The temperature was maintained at 4°C throughout the centrifugation process. After filtration through a 0.22 μm membrane filter, the SEVs were ultracentrifuged at 120,000 g for 100 minutes at 4°C. The SEV pellet was washed with PBS by recentrifugation under the same conditions. Cell morphology was observed using an all-in-one fluorescence microscope (BZ-X810, Keyence, Osaka, Japan).

[0060] <Transmission Electron Microscopy> SEV samples were placed on carbon-coated grids for 10 minutes, then washed with 2% uranyl acetate solution and negatively stained for 1 minute. The grid was then dried after removing the stain with filter paper. SEV samples were observed using a H-7650 S4 transmission electron microscope (Hitachi, Tokyo, Japan). Digital images were captured using an XR-41 CCCD camera system (Advanced Microscopy Techniques, Woburn, USA).

[0061] Nanoparticle Tracking Analysis (NTA) The concentration and size distribution of EVs were measured based on Brownian motion using a NanoSight NS300 nanoparticle characterization system (Malvern Panalytical, Tokyo, Japan). Five 60-second shots were used per acquisition. The average of three captures for each biological replicate was used to determine the mode of size distribution and nanoparticle concentration.

[0062] <Single particle analysis of SEVs using imaging flow cytometry> Measurements were performed using an AMNIS ImageStreamX Mark II flow cytometer (Luminex, TX, USA). Calibration was performed before each sample measurement. The flow rate was calibrated from the average velocity of SpeedBeads (Luminex, TX, USA) flowing through the channel. The side-scattered light intensity was calibrated using the default intensity of the 785 nm laser irradiated on the SpeedBeads. Each laser intensity was calibrated by measuring the scattered light intensity when irradiated on the flowing SpeedBeads. For SEV particle measurements, signals were detected at Ch4 in bright field, Ch3 for MemDye-Red on the SEV membrane, and Ch6 for side scattering (SSC). Laser power was maximized (561 nm: 200 mW, 642 nm: 150 mW, SSC: 70 mW). All measurements were performed at low flow rates and 40x magnification. SEV were measured as fluorescence-corrected samples. SEV were corrected using a TiO2-SPM, and the solvent was changed to PBS using a 10 kDa ultrafiltration membrane (Merck, Darmstadt, Germany). SEV solution (90 μL) was mixed with water (10 μL) and ExoSperkler MemDye solution (2 μL) and incubated at 37°C for 30 minutes. To verify the presence of lipid bilayer particles, a control sample was prepared by adding 10 μL of 1% Triton X-100 aqueous solution to 90 μL of SEV solution. The mixture was left at room temperature for 1 hour, and then reacted with 2 μL of ExoSperkler Mem Dye solution. Each solution was placed in a filter tube and washed with PBS to remove unreacted dye compounds. Analysis of the fluorescence intensity of each object was performed using IDEAS (Luminex, TX, USA).

[0063] Protein Digestion and Purification of Peptide Samples for LC-MS / MS Analysis: SEV was dried and digested using a trypsin digestion protocol utilizing phase-transfer surfactant (PTS) as previously described [T. Masuda, M. Tomita, Y. Ishihama, J. Proteome Res. 2008, 7, 731.]. Proteins were reduced with 10 mM dithiothreitol (DTT) (FUJIFILM Wako Pure Chemical Corporation, Osaka, Japan) at 37°C for 30 minutes, followed by alkylation with 50 mM 2-iodoacetamide (IAA) (FUJIFILM Wako Pure Chemical Corporation) for 30 minutes at room temperature in the dark. Samples were diluted to 2 M urea with 50 mM ammonium bicarbonate. Proteins were digested with 1 μg of lysyl endopeptidase (LysC) (FUJIFILM Wako Pure Chemical Corporation) and 1 μg of trypsin (Promega, Tokyo, Japan) overnight at 37°C in a shaking incubator. The resulting peptides were acidified with 0.5% trifluoroacetic acid (TFA, final concentration) and fractionated on StageTips containing SDB-XC (top) and SCX (bottom) Empore disc membranes (GL Sciences, Tokyo, Japan). Peptides were washed with 0.1% TFA in 5% ACN and 0.1% TFA in 80% ACN. The peptides were then eluted from the tip with 4% TFA in 30% ACN containing 500 mM ammonium acetate and 30% ACN containing 500 mM ammonium acetate. The sample solution was evaporated in a SpeedVac (Thermo Fisher Scientific), and the residue was resuspended in 0.5% TFA in 5% ACN. Finally, the peptides were desalted again using a StageTip with an SDB-XC Empore disc membrane and suspended in loading buffer (0.5% TFA in 5% ACN) for subsequent LC-MS / MS analysis. After digestion, the peptide concentration was measured using a NanoDrop (ThermoFisher Scientific); the absorbance was measured at 205 nm with an extinction coefficient of 31 [R. K. Scopes, Anal. Biochem. 1974, 59, 277.].Peptides were separated on a pull-in house needle column packed with ReproSil-Pur 120C18-AQ3-μm RP material (Dr Maisch, Ammerbuch, Germany).

[0064] <LC-MS / MS Analysis> NanoLC-MS / MS analysis was performed using a Q-Exactive (Thermo Fisher Scientific) connected to an UltiMate 3000 pump (Thermo Fisher Scientific) and an HTC-PAL autosampler (CTC Analytics). Peptides were separated on a House needle column (length 150 mm, internal diameter 100 μm, needle opening 6 μm) packed with ReproSil-Pur 120 C18-AQ 3-μm RP material (Dr Maisch, Ammerbuch, Germany) [Y. Ishihama, J. Rappsilber, J. S. Andersen, M. Mann, J. Chromatogr. Asia 2002, 979, 233.]. Samples were applied by full-loop injection of 5 μL, with a flow rate of 500 nL / min. Separation was performed using a four-step linear gradient: 4-10% ACN for 5 min, 10-40% MeCN for 60 min, 40-99% MeCN for 10 min, and 99% MeCN for 10 min with 0.5% TFA. The electrospray voltage was set to 2.4 kV in positive mode. Full MS scans were acquired over a mass range of 350-1500 m / z with a resolution of 70,000, an automatic gain control (AGC) target of 3e6, and a maximum injection time of 100 mS. MS / MS scans were performed using the Top 10 method with a resolution of 17,500, an AGC target of 1e5, a maximum injection time of 100 mS, and an isolation window of 2.0 Th. Precursor ions were fragmented by higher-energy collisional dissociation with a normalized collision energy of 27%.

[0065] Database Searching: For all experiments, raw MS data files were analyzed using MaxQuant / v2.0.3.0 [J. Cox, M. Mann, Nat. Biotechnol. 2008, 26, 1367.]. Peptides and proteins were identified by automated database searching using Andromeda against the human SwissProt database (version 2023-02, 20,588 protein entries) with a precursor mass tolerance of 20 ppm for the initial search, 4.5 ppm for the main search, and a fragment ion mass tolerance of 20 ppm. The enzyme was set as trypsin / P with two missed cleavages. Cysteine ​​carbamidomethylation was set as a fixed modification. Protein N-terminal methionine oxidation and acetylation were set as variable modifications. Search results were filtered for peptide spectrum match (PSM) and FDR <1% at the protein level. To mitigate the issue of missing values, the Match-between-run algorithm (MBR) was utilized via the Identification subtab of the Global Parameters tab in MaxQuant. Default settings for MBR were used (0.7-minute match window and 20-minute alignment time). Proteins identified only by site, potential contaminants, and those with reversed sequences were removed for data analysis. Missing or zero values ​​were replaced with the minimum value to assess relative abundance.

[0066] Notably, TiO2-BSA-SPM is highly suitable for clinical samples. We investigated the particle size and proteome profile of NPs collected using TiO2-BSA-SPM from 10 μL serum samples from healthy donors (CosmoBio; Human Serum (Pool) Human, KOJ-12181201, BJ11104a). The average size of collected nanoparticles was found to be 117.8 ± 7.1 nm (Figure 7a). Furthermore, compared with serum samples, 355 proteins were identified exclusively in NPs collected with TiO2-BSA-SPM, including typical exosome markers (Figure 7b). According to GO annotation, a protein group was assigned to the keyword "extracellular exosome" as a cellular component (Figure 7c). The protein group was also assigned to the keyword "blood microparticles." This keyword refers to phospholipid microvesicles derived from any of several cell types, such as platelets, blood cells, and endothelial cells. However, due to the characteristics of the TiO2-BSA-SPM method, particularly the lipid bilayer membrane, we were unable to separate SEVs from these particles. This result highlights the effectiveness and wide applicability of the TiO2-SPM method for SEV collection. Finally, we applied the TiO2-BSA-SPM method to remove EVs from fetal bovine serum (FBS), demonstrating its high throughput and flexibility. FBS contains essential proteins, lipids, and carbohydrates required for many cell cultures and is widely used as a growth factor for in vitro cell culture. However, bovine-derived EVs are present in FBS and can increase the background EV content in assay systems. UC has commonly been used for EV removal, but it requires more than 16 hours due to its low throughput. To accelerate EV research, a technology for removing EVs from large amounts of FBS is needed. Here, a syringe barrel-type solid-phase extraction cartridge was prepared as a spin column type that processes only 0.5 μL of sample. The syringe barrel type can store up to five TiO2-BSA-SPM disks, increasing the adsorption capacity. Furthermore, by placing an additional syringe on top as a sample reservoir, up to 25 mL of sample can be processed at one time.The loading process was performed by manual pressure, and the large pores of the TiO2-BSA-SPM enabled the processing of viscous FBS. Figure 2 shows the results of EV removal from 10 mL of FBS using TiO2-BSA-SPM. TiO2-BSA-SPM treatment significantly reduced the number of nanoparticles in the FBS (Fig. 5a). In particular, when four or more TiO2-BSA-SPM disks were used, a higher EV removal efficiency was achieved than with UC at 16 h (Fig. 5b). To examine the effects on cells, HEK293T cells were cultured using EV-depleted FBS prepared with TiO2-BSA-SPM. The cell counts over time were similar between cells cultured with UC and EV-depleted FBS prepared with TiO2-BSA-SPM, indicating that this method did not damage the cells (Fig. 5c, d). In short, it was found that TiO2-BSA-SPM can provide a simple method for removing EVs from large volumes.

[0067] The TiO2-SPM of the present invention will become a universal platform for collecting and purifying SEVs, and will make a significant contribution to drug discovery research and biomarker discovery using SEVs.

Claims

1. A porous monolithic polymer for the separation and purification of biological particles, hybridized with an olefin thermoplastic resin and an inorganic oxide.

2. The porous monolithic polymer for separating and purifying biological microparticles according to claim 1, wherein the thermoplastic resin is at least one selected from the group consisting of poly(ethylene-co-glycidyl methacrylate) (PEGM), ethylene-vinyl acetate copolymer, high density polyethylene, low density polyethylene, olefin-based elastomers, and polyamide-based elastomers.

3. The porous monolithic polymer for separating and purifying biological microparticles according to claim 1, wherein the inorganic oxide is one or more selected from the group consisting of titanium oxide, magnesium oxide, aluminum oxide, silicon dioxide, and calcium phosphate.

4. The porous monolithic polymer for separating and purifying biological particles according to claim 1, characterized in that the porosity is 50 to 90%, the pore size is 1 to 200 μm, and the melting point is 60° C. to 180° C.

5. A method for producing a porous monolithic polymer for the separation and purification of biological microparticles as described in claim 1, which comprises melting the thermoplastic resin, kneading the thermoplastic resin with a pore forming agent and a pore aid together with the inorganic oxide, and, after molding, dissolving the pore forming agent in an aqueous solvent.

6. The method according to claim 5, wherein the pore forming agent is an organic or inorganic substance that does not react with the thermoplastic resin and the inorganic oxide during kneading.

7. The method of claim 6, wherein the pore-forming agent is one or more selected from the group consisting of pentaerythritol, corn starch, medieval anhydrous sodium sulfate, starches such as wheat starch and potato starch, artificial sweeteners such as aspartame and saccharin, polysaccharides such as hemicellulose, acid-soluble inorganic compounds such as calcium carbonate and magnesium carbonate, and table salt.

8. The method according to claim 5, wherein the diameter of the particles of said pore forming agent is from 1 to 200 μm.

9. The method according to claim 5, wherein the forming aid is a monomer or polymer of a polyhydric alcohol.

10. The method according to claim 9, wherein the pore-forming aid is at least one member selected from the group consisting of (poly(oxyethylene-oxypropylene))triol, polyalkylene glycols such as polyethylene glycol, and diethylene glycol.

11. A method for separating and purifying biological microparticles having phospholipids on their surfaces, using the porous monolithic polymer for separating and purifying biological microparticles described in claim 1.

12. The method of claim 11, wherein the biological microparticle is an extracellular vesicle.

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