Method for preparing lipid nanoparticles

Abstract

The present invention relates to a method for preparing lipid nanoparticles, the method comprising a step of mixing an organic solution containing a lipid component with an aqueous solution containing a nucleic acid component, thereby forming lipid nanoparticles, wherein the aqueous solution is an aqueous solution having a neutral pH, changes such as the change in the size of lipid nanoparticles due to a change in the pH of a solvent during the preparation of lipid nanoparticles is suppressed, thereby enabling the size of nanoparticles to be kept small, and nanoparticles can be prepared containing, as a component of lipid nanoparticles, a material that is unstable under acidic pH conditions.

Description

Method for manufacturing lipid nanoparticles

[0001] The present invention relates to a method for manufacturing lipid nanoparticles, and more specifically, to a method for manufacturing lipid nanoparticles that minimizes in vivo side effects and can effectively deliver pharmacologically active substances to target cells, wherein, when mixing an organic solution containing lipid components and an aqueous solution containing nucleic acid components, the aqueous solution used for manufacturing has a neutral pH.

[0002]

[0003] Even for the same drug, efficacy varies widely depending on the delivery method. A drug delivery system (DDS) refers to the administration route and form of the drug that efficiently delivers the required amount of medication while minimizing side effects and maximizing efficacy. In the pharmaceutical industry, drug delivery systems can be considered a high-value-added core technology with a high probability of success, capable of generating economic benefits comparable to new drug development.

[0004] Recently, among various drug delivery systems, the development of gene therapies utilizing gene drug delivery systems is gradually expanding. To successfully and safely perform gene therapy, it is crucial to deliver genes or gene regulatory factors to the desired tissues. Representative examples of such genes or gene regulatory factors include mRNA, siRNA, and miRNA. Since nucleic acids such as mRNA perform the function of expressing specific proteins, they can play a role in supplementing proteins deficient due to genetic factors. Furthermore, using mRNA that expresses cancer markers or viral surface proteins enables the development of anticancer drugs and vaccines that activate the body's immune response. Additionally, nucleic acids such as siRNA and miRNA are substances capable of inhibiting the expression of specific proteins within the body, and are gaining attention as important tools for the treatment of cancer, genetic diseases, infectious diseases, and autoimmune diseases. However, since it is difficult to deliver these nucleic acids directly into cells and they are easily degraded by enzymes in the blood, extensive research is being conducted to overcome these limitations.

[0005] Non-viral gene carriers, such as cationic liposomes and polymers, are attracting attention for effectively delivering these substances. The improved stability profile and ease of fabrication and manipulation of polymer carriers have accelerated research on the design and synthesis of non-toxic and biodegradable polymer carriers for effective and safe gene delivery. Poly(L-lysine), polyethyleneimine, starburst, polyamidoamine dendrimers, and cationic liposomes have been widely studied as non-viral gene carriers because they can self-assemble spontaneously and compress plasmid DNA (pDNA) into structures small enough to enter cells via endocytosis.

[0006] In particular, lipid nanoparticles (LNPs) have recently been actively researched as non-viral gene delivery vehicles (Non-Patent Literature 1-9). Since LNPs utilize substances present in the body such as phospholipids and cholesterol, they have high bioavailability and affinity, enable drug release and control, and are particulate drug delivery vehicles with high stability against degradation by enzymes.

[0007]

[0008] Meanwhile, mRNA-based vaccines and therapeutics exert their effects by administering mRNA encoding a specific antigen or a protein exhibiting a therapeutic effect into the body, thereby causing it to be expressed as that antigen or protein. Regarding mRNA vaccines, extensive research has been conducted on cancer vaccines using mRNA encoding cancer antigens, and they have garnered even more attention since the efficacy of COVID-19 vaccines was proven in 2020. LNP forms are generally used as carriers to deliver mRNA vaccines and therapeutics into the body (U.S. Patent Publication 2020-0405844 A).

[0009] The efficacy of LNPs used for delivering oligos such as mRNA, siRNA, and miRNA is largely determined by the characteristics of ionized lipids, which are one of the components constituting the particles; however, in the early stages of LNP development, positively charged lipids were widely used rather than ionized lipids. This was due to considerations regarding the ease of nanoparticle manufacturing and in vitro efficacy. Since positively charged lipids constantly carry a positive charge, they facilitate the binding of negatively charged oligos through electrostatic attraction, thereby forming nanoparticles. Furthermore, because the lipid membranes constituting cell membranes and endosomes carry a negative charge, it was known that this promotes the fusion of positively charged lipid nanoparticles with the cell membrane, resulting in high efficiency in delivering oligos into the cell (U.S. Patent Publication 2018-0221510 A, 2018-0369384 A). However, lipid nanoparticles manufactured using positively charged lipids also have an overall positive charge, so a major disadvantage has been highlighted: they cause various side effects due to unnecessary interactions with unspecified cells while circulating through the bloodstream. Ionized lipids have been used to overcome these disadvantages of positively charged lipids.

[0010] The ionizable lipids constituting LNPs are characterized by being non-charged at neutral pH but positively charged under acidic conditions. Since LNPs maintain an electrically neutral state while circulating within the body, they can overcome the disadvantages associated with nanoparticles composed of positively charged lipids. Conversely, when LNPs enter the cell's endosomes, the ionizable lipids transition to a positively charged state due to the acidic pH environment inside the endosomes. The transition of nanoparticles to a positively charged state by ionizable lipids within the endosome is of significant importance to the efficiency of the nanoparticles. It is known that a fusion process between the lipid components constituting the nanoparticles and the endosome lipid membrane is necessary to deliver the oligos contained within the nanoparticles to the cytoplasm by passing through the endosome lipid membrane. Since endosome lipid membranes typically carry a negative charge, this fusion is facilitated by electrostatic attraction with the positively charged nanoparticles. To this end, it is desirable for ionized lipids to be designed with a chemical structure having a pKa value that is non-charged at neutral pH and positively charged at acidic pH. Accordingly, the pKa of suitable ionized lipids is known to be 5 to 7, but this range of pKa is only a necessary condition for the efficient development of ionized lipids, not a sufficient condition.

[0011] As previously described, the fact that ionized lipids have a pKa value of 5 to 7 has a significant impact on the manufacturing process of LNPs in addition to biological aspects. The manufacturing process of LNPs is basically a method of preparing nanoparticles composed of lipid components in such a way that nucleic acids are incorporated into the interior of the particles. Conventional LNPs are manufactured in the following manner. First, lipids and nucleic acids, which are the main components constituting the LNP, are prepared in separate solutions. The lipid components are prepared in an organic solvent such as ethanol, and the nucleic acids are prepared in an aqueous solution. When these two solutions are mixed rapidly and uniformly, the lipid components form nanometer-sized particles in the mixture of the organic solvent and the aqueous solution. At this time, the ionized lipids bind to the nucleic acids, thereby encapsulating the nucleic acids inside the particles.

[0012] The encapsulation efficiency of nucleic acids is determined by the binding of ionized lipids and nucleic acids through mutual electrostatic attraction during the mixing process. Nucleic acids are composed of a phosphate backbone and constantly carry a negative charge. Therefore, the ionized lipids must be transferred to a positive charge during the mixing process, which is possible by maintaining the pH of the aqueous solution containing the nucleic acids under acidic conditions. This is because when an acidic aqueous solution with a pH lower than the pKa of the ionized lipids is mixed with an ethanol solution of the lipid components, the ionized lipids become protonated and take on a positive charge, making it easier for them to bind with the negative charge of the oligos.

[0013] When prepared using this method, the produced LNP exists in a mixed solution of ethanol and aqueous solution under acidic pH conditions. Therefore, as the final step in LNP preparation, the LNP in the mixed solution is dialyzed in a buffer solution at neutral pH; this process removes the ethanol while simultaneously changing the acidic pH to neutral, thereby completing the preparation of the LNP.

[0014] However, the method of preparing such lipid nanoparticles, namely the method of preparing nanoparticles under acidic pH conditions and then replacing them with a solvent under neutral pH conditions, has several disadvantages. First, this method makes the final physical properties of the nanoparticles non-uniform. Prior art (Non-patent Literature 11) describes that when particles formed under acidic pH conditions are changed to neutral pH conditions, the charge of the ionized lipids changes from positive to neutral, causing aggregation between particles. In this process, additional structural variations of the particles are induced, leading to an increase in particle size and the production of particles of generally non-uniform size.

[0015] Secondly, another problem with nanoparticle manufacturing methods that generate lipid nanoparticles under acidic pH conditions is that it is difficult to use chemically unstable materials as components when manufacturing nanoparticles under acidic pH conditions. When intending to use such materials, a method can be employed in which the pH is rapidly changed to neutral conditions immediately after manufacturing LNPs at acidic pH. However, this method makes it difficult to ensure the structural stability of the material under acidic pH conditions and not only requires an additional mixing process to control the pH of the solution during the LNP manufacturing process, but the rapid change in solution pH also acts as a factor that is disadvantageous to maintaining particle uniformity.

[0016] Accordingly, the inventors made diligent efforts to solve the above problem and confirmed that when LNP is prepared using ionized lipids with a pKa of 7 or less, LNP can be stably prepared in an aqueous solution of neutral pH without using acidic pH conditions, thereby completing the present invention.

[0017] In addition, although the pKa of the ionized lipids constituting LNP is typically distributed between 5 and 7, the fact that nanoparticles can be produced using ionized lipids with pKa < 7 in a neutral pH aqueous solution makes it obvious that LNPs can also be produced in a neutral pH aqueous solution when using ionized lipids with pKa > 7 or higher.

[0018]

[0019] [Prior Art Literature]

[0020] [Patent Literature]

[0021] US Patent Publication US 2020-0405844 A1

[0022] US Patent Publication US 2018-0221510 A1

[0023] US Patent Publication US 2018-0369384 A1

[0024] Republic of Korea Registered Patent No. 10-2516680

[0025]

[0026] [Non-patent literature]

[0027] Moss, K.H., Popova, P., Hadrup, SR, Astakhova, K. & Taskova, M. Lipid Nanoparticles for Delivery of Therapeutic RNA Oligonucleotides. Mol Pharm 16, 2265-2277 (2019)

[0028] Kulkarni, J. A., Witzigmann, D., Chen, S., Cullis, P. R. & van der Meel, R. Lipid Nanoparticle Technology for Clinical Translation of siRNA Therapeutics. ACC Chem Res 52, 2435-2444 (2019)

[0029] Buck, J., Grossen, P., Cullis, P.R., Huwyler, J. & Witzigmann, D. Lipid-Based DNA Therapeutics: Hallmarks of Non-Viral Gene Delivery. ACS Nano 13, 3754-3782 (2019)

[0030] Akinc, A. et al. The Onpattro story and the clinical translation of nanomedicines containing nucleic acid-based drugs. Nat Nanotechnol 14, 1084-1087 (2019)

[0031] Springer, A.D. & Dowdy, S.F. GalNAc-siRNA Conjugates: Leading the Way for Delivery of RNAi Therapeutics. Nucleic Acid Ther 28, 109-118 (2018)

[0032] Kulkarni, J.A., Cullis, P.R. & van der Meel, R. Lipid Nanoparticles Enabling Gene Therapies: From Concepts to Clinical Utility. Nucleic Acid Ther 28, 146-157 (2018)

[0033] Rietwyk, S. & Peer, D. Next-Generation Lipids in RNA Interference Therapeutics. ACS Nano 11, 7572-7586 (2017)

[0034] Fang, Y. et al. Cleavable PEGylation: a strategy for overcoming the "PEG dilemma" in efficient drug delivery. Drug Deliv 24, 22-32 (2017)

[0035] Cullis, P. R. & Hope, M. J. Lipid Nanoparticle Systems for Enabling Gene Therapies. Mol Ther 25, 1467-1475 (2017)

[0036] Suzuki, Y. & Ishihara, H. Difference in the lipid nanoparticle technology employed in three approved siRNA (Patisiran) and mRNA (COVID-19 vaccine) drugs. Drug Metab Pharmacokinet 41, 100424 (2021)

[0037] Kulkarni, J. A. et al. On the Formation and Morphology of Lipid Nanoparticles Containing Ionizable Cationic Lipids and siRNA. ACS Nano 12, 4787-4795 (2018)

[0038]

[0039] The objective of the present invention is to provide a method for manufacturing lipid nanoparticles that produce efficacy by delivering pharmacologically active substances, such as nucleic acids, into the cytoplasm.

[0040] Another objective of the present invention is to provide lipid nanoparticles produced by the above method and a drug delivery composition containing the same.

[0041]

[0042] To achieve the above objective, the present invention provides a method for manufacturing lipid nanoparticles comprising: (a) a step of mixing an organic solution containing a lipid component and an aqueous solution containing a nucleic acid component to form lipid nanoparticles; and (b) a step of removing the solvent from the mixed solution, wherein the aqueous solution is a neutral pH aqueous solution.

[0043]

[0044] The method for manufacturing lipid nanoparticles according to the present invention can maintain the size of the nanoparticles small by suppressing changes such as changes in the size of lipid nanoparticles due to changes in the pH of the solvent that occur during the LNP manufacturing process.

[0045] Nanoparticles can be manufactured by including substances unstable under acidic pH conditions as components of lipid nanoparticles.

[0046]

[0047] Figure 1 shows A) size, B) RNA encapsulation rate, C) TNS assay graph, and D) pKa measurements of nanoparticles prepared under acidic pH conditions (L1), neutral pH conditions (L2), and distilled water conditions (L3), respectively.

[0048] Figure 2 shows A) particle size, B) RNA encapsulation rate, C) TNS assay graph, and D) pKa measurements when nanoparticles were prepared using MC3 ionized lipid or ALC-0315 ionized lipid under acidic pH conditions (L1, L4), neutral pH conditions (L2, L5), and distilled water conditions (L3, L6), respectively.

[0049] Figure 3 is a graph showing the mRNA expression rates of nanoparticles prepared using SM-102 ionized lipid (A) or MC3 ionized lipid (B) under acidic pH conditions (L1), neutral pH conditions (L2), and distilled water conditions (L3), respectively.

[0050] Figure 4 shows data obtained by preparing nanoparticles containing PEG-lipids that decompose under acidic pH conditions under acidic pH conditions (L1), neutral pH conditions (L2), and distilled water conditions (L3), respectively, and measuring A) particle size, B) RNA encapsulation rate, and C) hEPO protein expression amount.

[0051] Figure 5 shows A) RNA encapsulation rate and B) particle size graphs of nanoparticles prepared using SM-102 ionized lipids under various buffer solution conditions, and C) RNA encapsulation rate and D) particle size graphs of nanoparticles prepared using MC3 ionized lipids.

[0052] Figure 6 shows A) RNA encapsulation rate and B) particle size data measured by preparing nanoparticles under various buffer solution conditions with different concentrations.

[0053] Figure 7 shows the A) size and B) RNA encapsulation rate of nanoparticles prepared using an RNA aqueous solution prepared in distilled water, with NP ratios of 9 (L1), 7 (L2), and 5 (L3), respectively.

[0054]

[0055] Specific details for implementing the invention

[0056] Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by a skilled expert in the art to which this invention pertains. In general, the nomenclature used herein is well known and commonly used in the art.

[0057]

[0058] According to the present invention, in the process of manufacturing lipid nanoparticles capable of minimizing in vivo side effects and effectively delivering pharmacologically active substances to target cells, when adjusting the mixing conditions of an organic solution containing lipid components and an aqueous solution containing nucleic acid components, it was confirmed that when manufacturing LNPs using ionized lipids with a pKa of 7 or less, LNPs can be stably manufactured in an aqueous solution of neutral pH without using acidic pH conditions.

[0059]

[0060] Accordingly, in one aspect, the present invention relates to a method for preparing lipid nanoparticles comprising: (a) a step of mixing an organic solution containing a lipid component and an aqueous solution containing a nucleic acid component to form lipid nanoparticles; and (b) a step of removing the solvent from the mixed solution, wherein the aqueous solution is a neutral pH aqueous solution.

[0061]

[0062] In the present invention, neutrality means having neither positive nor negative charge in electrical characteristics, and neutral pH means intermediate between acidity and basicity. Accordingly, neutral pH is pH 6.5 to 8.0, preferably pH 6.8 to 7.5, most preferably pH 7, and includes all pH values ​​within the above range.

[0063] In the present invention, the neutral pH aqueous solution is preferably distilled water.

[0064] In the present invention, the neutral pH aqueous solution may further comprise an amine compound, wherein the amine compound is TRIS (tris-hydroxymethyl-aminomethane), imidazole, HEPES (Hydroxyethyl piperazine ethane sulfonic acid), MOPSO (3-Morpholino-2-hydroxypropanesulfonic acid), MOPS (3-(N-Morpholino)propanesulfonic acid), PIPES (1,4-Piperazinediethanesulfonic acid), BES (N,N-Bis(2-hydroxyethyl)-2-aminoethanesulfonic acid), DIPSO (3-(N,N-Bis[2-hydroxyethyl]amino)-2-hydroxypropanesulfonic acid), HEPPSO (N-(2-Hydroxyethyl)piperazine-N′-2-hydroxypropanesulphonic acid), POPSO (Piperazine-N,N′-bis(2-hydroxypropanesulfonic acid), EPPS (4-(2-Hydroxyethyl)-1-piperazinepropanesulfonic acid), ACES (N-(2-Acetamido)-2-aminoethanesulfonic acid), TES (2-[(2-Hydroxy-1,1-bis(hydroxymethyl)ethyl)amino]ethanesulfonic acid acid), TAPSO (2-Hydroxy-3-[tris(hydroxymethyl)methylamino]-1-propanesulfonic acid), glycylglycine (Glycylglycine, Diglycine, Gly-Gly), Tricine (N-[Tris(hydroxymethyl)methyl]glycine) and Bicine (N,It can be selected from the group consisting of N-Bis(2-hydroxyethyl)glycine, preferably TRIS, imidazole, or HEPES, but is not limited thereto.

[0065] In the present invention, the volume ratio of the organic solution and the buffer aqueous solution may be 2:1 to 1:20, preferably 1:1 to 1:10, and more preferably 1:2 to 1:5. If the volume ratio of the organic solution and the buffer aqueous solution is greater than the above range, the solubility of lipid components in the mixed solution is high, making it difficult to form nanoparticles; conversely, if it is smaller than the above range, there is a problem of non-uniform lipid precipitates being generated.

[0066] In the present invention, the lipid component may be an ionized lipid, a PEG-lipid, or a mixture thereof.

[0067] In the present invention, the ionized lipid is a compound composed of a head portion containing an amine group and a hydrophobic lipid portion, and includes, for example, the following substances. (6Z, 9Z, 28Z, 31Z)-hephtatriaconta 6,9,28,31-tetraene-19-yl4-(dimethylamino)butanoate (DLin-MC3-DMA), [(4-hydroxybutyl)azandyl]di(hexane-6,1-diyl)bis(2-hexyldecanoate)(ALC-0315), 8-[(2-hydroxyethyl)[6-oxo-6-(undecyloxy)hexyl]amino]-octanic acid (SM-102), 1-linoleoyl-2-linoleyloxy-3-dimethylaminopropane (DLin-2-DMAP), 1,2-dilinoleylcarbamoyloxy-3-dimethylaminopropane (DLin-C-DAP), 1,2-Dilinoleoyl-3-Dimethylaminopropane (DLin-DAP), 1,2-Dilinoleyloxy-N,N-Dimethylaminopropane (DLin-DMA), 2,2-Dilinoleyl-4-Dimethylaminomethyl-[1,3]-Dioxolane (DLin-K-DMA), 2,2-Dilinoleyl-4-(2-Dimethylaminoethyl)-[1,3]-Dioxolane (DLin-KC2-DMA), 1,2-Dioleoyl-3-Dimethylammonium Propane (DODAP), N,N-Dimethyl-(2,3-Dioleyloxy)Propyamine (DODMA), DioctadeciramideGlycyrrhoxyspermin (DOGS), SperminCholesterylCarbamate (GL-67);Bis-Guanidinium-Spermidine-Cholesterol (BGTC), 3β-(N-(N',N'-dimethylaminoethane)-carbamoyl) cholesterol (DC-Chol), 1,1'-(2-(4-(2-((2-(bis(2-hydroxydecyl)amino)ethyl)(2-hydroxydecyl)amino)ethyl)piperazine-1-yl)ethylazandyl)dododecane-2-ol (C12-200), Nt-butyl-N'-tetradecylamino-propionamidine (diC14-amidine);Dimethyl dioctadecirinemonium bromide (DDAB), N-(1,2-dimyristyl oxyprop-3-yl)-N,N-dimethyl-N-hydroxyethylammonium bromide (DMRIE), N,N-dioleyl-N,N-dimethylammonium chloride (DODAC), dioleyl oxypropyl-3-dimethylhydroxyethylammonium bromide (DORIE), N-(1-(2,3-dioleyl oxyl)propyl)-N-2-(spermine carbokixamide)ethyl)-N,N-dimethylammonium trifluoroacetate (DOSPA), 1,2-dioleoyl trimethylammonium propane chloride (DOTAP), N-(1-(2,3-dioleyl oxy)propyl)-N,N,N-trimethylammonium chloride (DOTMA) and One or more may be selected from the group consisting of aminopropyl-dimethyl-bis(dodecyloxy)-propane aluminum bromide (GAP-DLRIE).

[0068] In the present invention, the ionized lipid may additionally include sterol lipids and neutral lipids.

[0069] Among the components of the lipid nanoparticles according to the present invention, sterol lipids provide structural rigidity to the lipid filling within the lipid nanoparticles and are dispersed in the core and surface of the nanoparticles to improve the stability of the nanoparticles, and cholesterol derivatives including cholesterol and cholesteryl esters can be used.

[0070] Among the components of the lipid nanoparticles according to the present invention, the neutral lipid plays a role in protecting the core formed by the interaction of ionized lipids and drugs within the lipid nanoparticles by surrounding it, and binds to the lipid bilayer of target cells to facilitate the passage through the cell membrane and escape from the endosome during intracellular delivery of the drug. Among the components of the lipid nanoparticles according to the present invention, the neutral lipid may be used without limitation as long as it is a phospholipid or sphingolipid capable of promoting the fusion of lipid particles, and preferably DOPE (dioleoylphosphatidylethanolamine), DSPC (distearoylphosphatidylcholine), POPC (palmitoyloleoylphosphatidylcholine), EPC (egg phosphatidylcholine), DOPC (dioleoylphosphatidylcholine), DPPC (dipalmitoylphosphatidylcholine), DOPG (dioleoylphosphatidylglycerol), DPPG (dipalmitoylphosphatidylglycerol), DSPE (distearoylphosphatidylethanolamine), PE (Phosphatidylethanolamine), DPPE (dipalmitoylphosphatidylethanolamine), DOPE (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine), POPE It may be (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine), POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine), DOPS (1,2-dioleoyl-sn-glycero-3-[phospho-L-serine]), ceramide, or sphingomyelin.

[0071] In this specification, “PEG-lipid” refers to a form in which PEG (polyethyleneglycol) and lipid are conjugated, and is used interchangeably with “lipid-PEG,” “PEG-lipid,” or “lipid-PEG,” and means a lipid in which polyethyleneglycol (PEG), a hydrophilic polymer, is attached to one end. The PEG-lipid contributes to the stability of the nanoparticles in serum within the lipid nanoparticles and prevents aggregation between nanoparticles. Additionally, the PEG-lipid protects the nucleic acid from degrading enzymes during in vivo delivery, thereby enhancing the in vivo stability of the nucleic acid and can increase the half-life of the drug encapsulated within the nanoparticles.

[0072] In the above PEG-lipid, PEG can be directly conjugated to the lipid or linked to the lipid through a linker moiety. Any linker moiety suitable for binding PEG to the lipid may be used, and includes, for example, ester-free linker moiety and ester-containing linker moiety. The above ester-free linker moiety includes, but is not limited to, amido (-C(O)NH-), amino (-NR-), carbonyl (-C(O)-), carbamate (-NHC(O)O-), urea (-NHC(O)NH-), disulfide (-SS-), ether (-O-), succinyl (-(O)CCH2CH2C(O)-), succinamidyl (-NHC(O)CH2CH2C(O)NH-), and hydrazone (-C(O)NHN=R-, -C=NNH-), as well as combinations thereof (e.g., a linker containing both a carbamate linker moiety and an amido linker moiety). The above ester-containing linker moiety includes, for example, esters (-C(O)O-), carbonates (-OC(O)O-), succinoyl, phosphate esters (-O-(O)POH-O-), sulfonate esters and combinations thereof, but is not limited thereto.

[0073] In the present invention, the lipid within the PEG-lipid can be used without limitation as long as it is a lipid capable of binding with polyethylene glycol, and phospholipids and / or cholesterol, which are other components of the lipid nanoparticles, can also be used. Specifically, the lipid within the PEG-lipid may be dimyristoylglycerol (DMG), ceramide, succinoyl-diacylglycerol (s-DAG), distearoylphosphatidylcholine (DSPC), distearoylphosphatidylethanolamine (DSPE), or cholesterol, but is not limited thereto. In addition, the above lipid may be a lipid comprising a structure in which two types of lipids selected from C3-C30 single and branched chain forms of alkyl, alkenyl, and alkynyl hydrocarbons are connected to glycerol by a bond of ether (-O-) or ester (-C(O)O-, -OC(O)-).

[0074]

[0075] The PEG in the above PEG-lipid is a hydrophilic polymer that has the ability to inhibit the adsorption of plasma proteins, thereby increasing the circulation time of lipid nanoparticles in the body and preventing aggregation between nanoparticles. In addition, PEG-bound lipids exhibit a stealth function in vivo, which can prevent the degradation of nanoparticles.

[0076] The above PEG may be a functionalized PEG in which a functional group is attached to the side that is not bound to lipids. The functional groups available for use may be one or more selected from the group consisting of succinyl, carboxylic acid, maleimide, amine, biotin, and cyanur.

[0077] The above PEG may have a ligand bound to the side that is not bound to lipids. The ligands available for use at this time may be one or more selected from the group consisting of N-acetyl-D-galactosamine (GalNAc), N-acetyl-D-galactose, D-galactose, N-acetyl-D-glucosamine, N-acetyl-D-glucosamine, D-glucose, D-mannose, L-fucose, carbohydrate derivatives, folate, transferrin, RGD peptide, cyclic RGD peptide, TAT peptide, R9 peptide, CADY peptide, HA2 peptide, monoclonal antibody, antigen-binding fragment or antibody fragment, single-chain variable fragment (scFv), and aptamer. Here, the antigen-binding fragment of an antibody or antibody fragment refers to a fragment possessing antigen-binding function, and includes Fab, F(ab'), F(ab')2, and scFv, etc.

[0078] The PEG-lipid used in the present invention may be used alone, or mixed with a PEG-lipid that includes a functional group or ligand at the PEG terminal that is not bound to the lipid.

[0079] It was confirmed that by the method for manufacturing lipid nanoparticles according to the present invention, changes such as changes in the size of lipid nanoparticles due to changes in the pH of the solvent occurring during the LNP manufacturing process can be suppressed, thereby enabling the production of nanoparticles with a small size, and that nanoparticles can be manufactured by including a substance unstable under acidic pH conditions as a component of the lipid nanoparticles.

[0080] Accordingly, the present invention relates to lipid nanoparticles produced by the above-described manufacturing method in another aspect.

[0081] In the present invention, the lipid nanoparticles may include nucleic acids and ionized lipids.

[0082] In the present invention, the size of the particle may be 40 to 200 nm, preferably 60 to 150 nm, and the RNA encapsulation rate may be 50 to 100%, preferably 80 to 100%.

[0083] If the particle size is smaller than the range described above, it is easily removed from the body through methods such as excretion; however, if the particle size is larger than the range described above, there is a disadvantage in that movement between tissues within the body is not smooth. Since a higher RNA encapsulation rate minimizes raw material loss during the nanoparticle manufacturing process, it is desirable to maintain a high RNA encapsulation rate.

[0084]

[0085] The present invention will be described in more detail below through examples. These examples are intended solely to illustrate the present invention, and it will be obvious to those skilled in the art that the scope of the present invention is not to be interpreted as being limited by these examples.

[0086]

[0087] Example 1: Effect of buffer solution pH on the physical properties of nanoparticles during lipid nanoparticle preparation

[0088] Example 1-1: Nanoparticle Preparation Conditions and Particle Size

[0089] A conventional method for preparing lipid nanoparticles involves rapidly mixing an ethanol solution containing lipid components with a buffer solution containing RNA. The lipid components included in the ethanol solution are ionized lipids, phospholipids, cholesterol, and PEG-lipids. In this example, the same materials and compositional ratios used by Moderna in the manufacture of the Covid-19 vaccine were used for evaluation. Specifically, SM-102 was used as the ionized lipid, and DSPC (distearoylphosphatidylcholine, a phospholipid), cholesterol, and PEG-DMG (1,2-dimyristoyl-3-PEG-glycerol) were used in a molar ratio of 50:10:38.5:1.5, respectively. The lipid concentration of the solution was prepared to be 4.5 mg / mL.

[0090] Meanwhile, three types of RNA-containing solutions were prepared and evaluated: acidic pH buffer solution, neutral pH buffer solution, and distilled water. For the acidic pH buffer solution, a 50 mM citric acid buffer solution with a pH of 4 was used, and for the neutral pH buffer solution, a 50 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) buffer solution with a pH of 7.4 was used. RNA was used at a concentration of 0.04 mg / mL in all solutions.

[0091] Lipid nanoparticles were prepared by mixing an ethanol solution and an aqueous solution in a 3:1 volume ratio using a micromixer chip or a T-line chip, with the flow rate of the mixed solution set to 4 mL / min. The mixture was collected and dialyzed for 24 hours in a pH 7.5 PBS buffer solution. Lipid nanoparticles L1, L2, and L3, prepared in acidic pH buffer solution, neutral pH buffer solution, and distilled water, respectively, were prepared, and their particle sizes were measured using a Dynamic Light Scattering (DLS) device (Fig. 1A). It was confirmed that the nanoparticles prepared in neutral pH buffer solution or distilled water were relatively smaller in size compared to the nanoparticles prepared under acidic pH conditions.

[0092]

[0093] Example 1-2: Measurement of RNA encapsulation

[0094] To verify the RNA encapsulation rate of lipid nanoparticles prepared using ionized lipids according to the present invention, the Quant-it RiboGreen RNA assay kit (Thermo Fisher Scientific) was used. RNA standard solutions of various concentrations were prepared in 1X TE buffer solution to fall within the range of 0 to 0.1 μM. Two types of lipid nanoparticle samples were prepared by diluting the lipid nanoparticle solutions so that the concentration of RNA contained in the lipid nanoparticles fell within the range of the RNA standard solutions. One of the lipid nanoparticle samples was diluted in 1X TE buffer solution, and the other was diluted in 2% TritonX-100 and 1X TE buffer solution. After mixing the RiboGreen reagent with the RNA standard solution and the two types of lipid nanoparticle samples according to the method described in the RiboGreen Assay kit manual, fluorescence was measured at 480 nm for excitation and 520 nm for emission. The respective concentrations can be determined by substituting the fluorescence values ​​measured from the lipid nanoparticle samples into the calibration curve of the fluorescence values ​​measured from the RNA standard solutions. The RNA concentration measured in the lipid nanoparticle sample containing TritonX-100 is the total RNA concentration (C_tot), and the RNA concentration measured in the sample not containing TritonX-100 is the RNA concentration that was not encapsulated in the lipid nanoparticle (C_out). Therefore, the proportion of RNA encapsulated in the nanoparticle can be calculated as follows.

[0095]

[0096] [Mathematical Formula 1]

[0097] Encapsulation Rate (%) = { 1 - C_out / C_tot} x 100

[0098]

[0099] The RNA encapsulation rate calculated according to Equation 1 is as shown in Fig. 1B. It was confirmed that the RNA encapsulation rate of nanoparticles prepared using a neutral pH buffer solution or distilled water is similar to that of nanoparticles prepared using an acidic pH buffer solution.

[0100]

[0101] Examples 1-3: pKa measurement of nanoparticles

[0102] The pKa of the nanoparticles prepared under each condition was measured. TNS (6-p-Toluidino-2-naphthalenesulfonic acid), a fluorescent substance, was used to measure the pKa. Although TNS is not fluorescent in an aqueous solution, it becomes fluorescent when adsorbed onto the lipid components of lipid nanoparticles. When lipid nanoparticles containing ionized lipids are introduced into various types of buffer solutions with different pH values, the surface charge of the nanoparticles remains neutral in buffer solutions with a pH higher than the pKa of the ionized lipids, thus preventing the adsorption of TNS. However, in buffer solutions with a pH lower than the pKa of the ionized lipids, the surface of the nanoparticles becomes positively charged, inducing the adsorption of TNS and allowing for the measurement of fluorescence. By utilizing this property, the pKa of the ionized lipids constituting the nanoparticles can be measured. To this end, buffer solutions were prepared in pH increments of 0.5 from pH 2 to pH 12. The composition of the buffer solution was based on 10 mM HEPES, 10 mM MES, 10 mM Ammonium acetate, and 130 mM NaCl, with the pH adjusted. The solution volume was set to 100 μl in a 96-well Black plate, and 100 μM lipid nanoparticles were mixed with the buffer solution based on the ionized lipid concentration. A sample without added lipid nanoparticles was also prepared. After maintaining at room temperature for 10 minutes, TNS was added to each well to achieve a concentration of 1 μM, and the mixture was stirred using an orbital shaker for 2 minutes. The fluorescence of the samples was measured at Ex321 nm / Em429 nm using a multiplate reader, and each fluorescence measurement was corrected to a value excluding lipid nanoparticles.

[0103] In the graph (Fig. 1C) showing the fluorescence values ​​in each pH buffer solution, the pKa value was determined as the pH value at which half of the maximum fluorescence value is reached. The pKa values ​​of the nanoparticles prepared using acidic pH buffer solution, neutral pH buffer solution, and distilled water were all measured to be similar to the pKa of SM-102 ionized lipids described in the literature (Non-patent Literature 10).

[0104] The results of Example 1 are summarized in Table 1. It can be seen that compared to the case where an acidic pH buffer solution, which is a conventional manufacturing condition, was used, the particle size is relatively smaller, while other physical properties such as RNA encapsulation rate and pKa remain unchanged.

[0105]

[0106] Example 2: Preparation and evaluation of nanoparticles in neutral pH buffer solution using various ionized lipids

[0107] In Example 1, the effect of RNA buffer solution conditions on the physical properties of nanoparticles was evaluated when preparing nanoparticles using SM-102 ionized lipid. It is predicted that these preparation conditions and nanoparticle characteristics can be applied to all cases where the pKa of the ionized lipids is similar. To confirm whether this can be applied not only to the specific ionized lipid SM-102 but also to similar ionized lipids, two additional types of ionized lipids were used for evaluation. MC3 and ALC-0315 are ionized lipids used in the preparation of lipid nanoparticle therapeutics containing siRNA and a COVID-19 virus vaccine, respectively, and both therapeutics are commercially available treatments.

[0108] Six types of lipid nanoparticles were prepared using the ionized lipid MC3 and ALC-0315 according to the method described in Example 1. MC3 was used for L1-L3, and ALC-0315 was used for L4-L6. When preparing nanoparticles using each ionized lipid, an acidic pH buffer solution (citrate pH 4), a neutral pH buffer solution (Hepes pH 7.4), and distilled water were used, respectively, in the same manner as in Example 1.

[0109] The size of the prepared nanoparticles was measured using the method described in Example 1-1 (Fig. 2A). As with the nanoparticles prepared using SM-102, the size of the nanoparticles prepared using an acidic pH solution was relatively large, while the size of the nanoparticles prepared using a neutral pH buffer solution or distilled water was relatively small. This phenomenon was observed identically in the ionized lipid MC3 and ALC-0315.

[0110] In addition, the RNA content encapsulated in the manufactured nanoparticles was measured using the method described in Example 1-2 (Fig. 2B). As a result of the measurement, it was confirmed that all nanoparticles maintained similar levels.

[0111] The pKa of each nanoparticle was measured using the method described in Examples 1-3 (Figs. 2C, 2D). It was confirmed that the pKa of the nanoparticles prepared using MC3 and ALC-0315 ionized lipids was measured as a constant value regardless of the pH of the RNA buffer solution. It can be seen that the measured values ​​are similar to the pKa of MC3 and ALC-0315 reported in the literature (Non-patent Literature 10).

[0112] The results of Example 2 are summarized in Table 2. It can be seen that compared to the case where an acidic pH buffer solution, which is a conventional manufacturing condition, was used, the particle size was relatively smaller, while other physical properties such as RNA encapsulation rate and pKa remained unchanged.

[0113]

[0114] As can be seen from Examples 1 and 2, it is possible to prepare nanoparticles using a neutral pH buffer solution or distilled water as the RNA solution, regardless of the type of ionized lipid. In addition, while the size of the nanoparticles prepared using a neutral pH buffer solution or distilled water is relatively small, the RNA encapsulation rate and the pKa of the nanoparticles remain unchanged.

[0115] In particular, it is important to note that, contrary to what is previously known, it was confirmed that nanoparticles can be manufactured under pH conditions higher than the pKa of the ionized lipids. It is known that when nanoparticles encapsulate nucleic acids such as RNA, the negative charge of the RNA and the positive charge of the ionized lipids combine through electrostatic attraction to form nanoparticles. Therefore, since ionized lipids with a pKa generally in the range of 5 to 7 are typically used for nanoparticle manufacturing, nanoparticles are usually manufactured under pH conditions lower than the pKa of the ionized lipids to ensure that these lipids acquire a positive charge during the manufacturing process. This is because under pH conditions lower than the pKa, the ionized lipids undergo protonation and acquire a positive charge. However, contrary to this conventional wisdom, Examples 1 and 2 demonstrate that nanoparticles can be manufactured under pH conditions higher than the pKa of the ionized lipids, and that the RNA encapsulation rate and the pKa of the nanoparticles can be maintained identically.

[0116]

[0117] Example 3: Expression efficiency of nanoparticles prepared under conditions of neutral pH buffer solution or distilled water

[0118] To measure the in vitro efficacy of lipid nanoparticles according to the present invention, lipid nanoparticles containing SM-102 or MC3 ionized lipids were prepared by the method described in Example 1-1. mRNA expressing luciferase was prepared in an acidic pH buffer solution (citrate pH 4), a neutral pH buffer solution (Hepes pH 7.4), and distilled water, and nanoparticles were prepared by mixing them with a lipid solution containing SM-102 or MC3 ionized lipids. HepG2 and HEK293 cell lines were treated with these lipid nanoparticles at mRNA concentrations of 6.25, 12.5, 25, and 50 ng for 24 hours, and then luciferase activity was measured (Fig. 3). Measurements at each concentration were performed in three replicates, and the average value was calculated and expressed as a relative value. As a result of the measurements, it was confirmed that luciferase activity was measured in proportion to the RNA concentration treated in the cells, and it was confirmed that the in vitro expression efficiency of the nanoparticles did not differ significantly whether they were prepared using a neutral pH buffer solution or distilled water.

[0119]

[0120] Example 4: Preparation of lipid nanoparticles containing PEG-lipids that decompose under acidic pH conditions

[0121] When manufacturing conventional lipid nanoparticles, a PEG-lipid content of about 0.5 to 2 mol% is used. PEG facilitates the formation of nanoparticles because it inhibits inter-particle interactions on the particle surface. If PEG-lipids are not used when manufacturing lipid nanoparticles, the nanoparticles themselves cannot be formed due to particle aggregation, so their use is unavoidable. However, since PEG also inhibits nanoparticles from interacting with cells, the efficiency of nanoparticles decreases if the PEG-lipid content is high. Therefore, the range of PEG-lipid content used in the manufacture of conventional LNPs is limited to 0.5 to 2 mol%.

[0122] In contrast, prior art (Patent Literature 4) describes lipid nanoparticles containing a high PEG-lipid content. This technology is characterized by blocking non-specific interactions with other biomaterials by increasing the PEG-lipid content, enabling delivery to specific target cells solely through ligands. In particular, unlike conventional lipid nanoparticles, efficiency improves as the PEG-lipid content increases. This is possible because the nanoparticles utilize PEG-lipids designed to degrade under acidic pH conditions inside the endosome once they enter the cell's endosome. The PEG-lipids used in this technology contain degradable functional groups at the PEG-lipid linkage that degrade under acidic pH conditions. Therefore, the structure involves the removal of PEG from the particle surface and subsequent fusion with the endosome lipid membrane as the nanoparticles enter the endosome.

[0123] Since these nanoparticles utilize PEG-lipids (PEG-lipids containing degradable functional groups) that degrade under acidic pH conditions, an additional process is required during their preparation. Specifically, the conventional LNP manufacturing process uses an acidic pH solution; however, because PEG-lipids are highly likely to degrade in acidic solutions, the pH of the mixed solution must be rapidly changed to a neutral pH immediately after mixing RNA and lipids under acidic conditions to minimize this risk, just as in the existing method. Nevertheless, the possibility of PEG-lipids degrading under acidic pH conditions still persists. Furthermore, the particle size may increase unevenly during the process of rapidly changing from an acidic pH to a neutral pH.

[0124] These disadvantages in the manufacturing process can be resolved by applying the method of preparation using a neutral pH buffer solution or distilled water as described in the present invention. To verify this, nanoparticles presented in the relevant prior art were prepared using the components described in the literature. An ethanol solution with a concentration of 4.5 mg / mL was prepared containing SM-102, DSPC, cholesterol, PEG-lipids containing degradable functional groups, and GalNAc-PEG-lipids in a molar ratio of 40:10:39:10:1. GalNAc (N-acetyl-D-galactosamine) is a ligand commonly used for delivery to liver cells. Additionally, an aqueous RNA solution was prepared in an acidic pH buffer solution (citrate pH 4), a neutral pH buffer solution (HEPES pH 7.4), and distilled water, respectively, containing hEPO (human erythropoietin) mRNA at a concentration of 0.04 mg / mL. Nanoparticles were produced by mixing an ethanol solution containing lipid components with different RNA aqueous solutions in the manner described in Example 1-1 and left to stand at room temperature for 30 minutes. The preparation of nanoparticles was completed by performing dialysis in a 1X PBS buffer solution to replace the solvent.

[0125] The physical properties of the nanoparticles prepared under each aqueous solution condition were measured in the manner described in Example 1, including particle size and RNA encapsulation rate (Figs. 4A, 4B). As a result of the measurements, it was found that the nanoparticles (L1) prepared under acidic pH conditions showed a significant increase in size compared to other nanoparticles prepared under neutral pH conditions. This is because, immediately after particle formation, the acidic pH of the solution was not neutralized to a neutral pH, causing the PEG lipids distributed on the surface of the particles to decompose, resulting in the nanoparticles clumping together and precipitating. On the other hand, it was found that this phenomenon was not observed in the nanoparticles (L2) prepared in a neutral pH buffer solution and the nanoparticles (L3) prepared in distilled water. Additionally, it was confirmed that the RNA encapsulation rate was very low for L1, whereas L2 and L3 showed very high efficiency.

[0126] To verify the efficiency of each nanoparticle, a dose of 0.5 mpk was injected into the tail vein of mice. After 6 hours, serum was separated and blood hEPO concentrations were measured using an hEPO ELISA kit (Fig. 4C). While almost no protein was detected in L1, normal protein expression was observed in mice administered L2 and L3.

[0127] Through this embodiment, it can be seen that when lipid nanoparticles are manufactured using a material that is unstable under acidic pH conditions, the problem can be solved by applying the manufacturing method under neutral pH conditions presented in this invention.

[0128]

[0129] Example 5: Preparation of lipid nanoparticles in various neutral pH buffer solutions

[0130] Through the present invention, it was confirmed that lipid nanoparticles using ionized lipids can be manufactured under neutral pH conditions, and it was verified whether these manufacturing conditions are reproducible when using various buffer solutions. To this end, in addition to the citrate pH 4, distilled water, and Hepes pH 7 solution used in the previous example, Imidazole, Tris, MOPS (3-(N-morpholino)propanesulfonic acid), PIPES (piperazine-N,N′-bis(2-ethanesulfonic acid), TAPSO (3-[[1,3-dihydroxy-2-(hydroxymethyl)propan-2-yl]amino]-2-hydroxypropane-1-sulfonic acid), Carbonate, Phosphate, and Citrate were used as the pH 7.4 buffer solution, and Tricine (N-[1,3-Dihydroxy-2-(hydroxymethyl)propan-2-yl]glycine) and 1X PBS (Phosphate Buffered Saline) were used as the pH 7.4 buffer solution. Furthermore, nanoparticles were prepared in a 130 mM NaCl solution. Except for distilled water, 1X PBS, and 130 mM NaCl solution, the concentrations of the buffer solutions used in this experiment were The concentration was fixed at 10 mM, and the RNA concentration in each aqueous solution was set to 0.07 mg / mL. SM-102 was used as the ionized lipid, and all preparation conditions, except for the buffer solution, were carried out as described in Example 1. The RNA encapsulation rate and particle size of the nanoparticles prepared in this manner were measured, respectively (Figs. 5A, 5B).

[0131] Among the buffer solutions used, the RNA encapsulation rate of nanoparticles prepared using pH 7 HEPES, Imidazole, Tris, MOPS, PIPES, and TAPSO buffer solutions and pH 7.4 Tricine buffer solution was measured to be similar to that of nanoparticles (L1) prepared under acidic pH conditions. On the other hand, nanoparticles prepared using pH 7 carbonate, phosphate, and citrate buffer solutions and pH 7.4 1X PBS buffer solution showed relatively low RNA encapsulation rates. Thus, even when using buffer solutions with neutral pH, the prepared nanoparticles can be classified into two groups in terms of RNA encapsulation rate, and the difference between these two groups is determined by the chemical structure of the material used to prepare the buffer solution. That is, the group with a high RNA encapsulation rate was prepared using a buffer solution containing an amine group, while the group with a low RNA encapsulation rate is characterized by the use of buffer solutions of organic and inorganic acid series that do not contain an amine group. In addition, in the group with a high RNA encapsulation rate, the particle size is relatively smaller compared to the nanoparticles (L1) prepared under acidic pH conditions, whereas no distinct trend is observed in the group with a low RNA encapsulation rate.

[0132] In addition, to determine whether the salt contained in distilled water affects the RNA encapsulation rate when nanoparticles are prepared using distilled water, nanoparticles (L12) were prepared using an aqueous solution of 130 mM NaCl. The RNA encapsulation rate of L12 was measured to be 19%, which is a very low level compared to the RNA encapsulation rate of 92% of nanoparticles (L2) prepared using distilled water. Considering that RNA is encapsulated inside the nanoparticles due to the electrostatic attraction between RNA and ionized lipids when nanoparticles are prepared, it appears that the NaCl ions present in the aqueous solution play a role in inhibiting this binding.

[0133] To determine whether MC3 ionized lipids exhibit a similar trend to SM-102 ionized lipids, nanoparticles were prepared using MC3 in the same manner in acidic pH citric acid buffer solution, pH 7 Imidazole, HEPES, MOPS, PIPES, TAPSO, and Carbonate buffer solutions, and pH 7.4 Tricine buffer solution. The RNA encapsulation rate and particle size of the nanoparticles prepared in this manner were measured (Figs. 5C, 5D). Similar to the case of SM-102 ionized lipids, MC3 ionized lipids also showed a high RNA encapsulation rate in neutral pH buffer solutions prepared with amine-containing substances, but a very low encapsulation rate in buffer solutions prepared with the organic acid Carbonate.

[0134]

[0135] Example 6: Evaluation of buffer solution concentration

[0136] In Example 5, it was confirmed that the RNA encapsulation rate of nanoparticles can be inhibited by NaCl contained in the aqueous solution used for nanoparticle preparation. Since a certain amount of salt is included even when preparing buffer solutions, an experiment was conducted to determine whether the concentration of the buffer solution used for nanoparticle preparation affects the RNA encapsulation rate. Acidic pH buffer solutions and neutral pH buffer solutions were used at concentrations of 10 mM and 50 mM, respectively. For the neutral pH buffer solutions, buffer solutions prepared with amine-containing substances such as Tris, Imidazole, and HEPES were used, while phosphate and citrate buffer solutions were used as amine-free inorganic or organic acid-based buffer solutions. Nanoparticles were prepared using SM-102 ionized lipids in the same manner as described in Example 5, and the RNA encapsulation rate and particle size were measured, respectively (Figs. 6A, 6B).

[0137] Measurement results confirmed that the concentration of the buffer solution used does not affect the RNA encapsulation rate. It was found that when using buffer solutions prepared with amine-containing substances among acidic and neutral pH buffer solutions, both 10 mM and 50 mM showed high RNA encapsulation rates. Furthermore, it appears that even when using phosphate and citrate buffer solutions, which have low RNA encapsulation rates, the concentration of the buffer solution does not significantly change the RNA encapsulation rate.

[0138]

[0139] Example 7: Mixing ratio of lipids and RNA

[0140] The RNA encapsulation rate is influenced by the relative amounts of RNA and ionized lipids used when manufacturing nanoparticles. Since ionized lipids carry a positive charge and RNA carries a negative charge, nanoparticles are typically manufactured using an equivalent amount of ionized lipids that is several times the total charge of RNA. At this time, the relative molar ratio of ionized lipids to RNA charge is called the NP ratio. The higher the NP ratio—that is, the more ionized lipids are used in excess relative to RNA charge—the higher the RNA encapsulation rate tends to be. Conversely, the lower the NP ratio—that is, the higher the relative amount of RNA—the lower the RNA encapsulation rate tends to be.

[0141] To verify whether these characteristics are reproduced in nanoparticles prepared using a neutral pH aqueous solution, nanoparticles were prepared using various NP ratios. The RNA aqueous solution was prepared using distilled water, and solutions with RNA concentrations of 0.044 mg / mL, 0.057 mg / mL, and 0.08 mg / mL were used. The lipid solution was prepared so that the lipid concentration in ethanol was 4.5 mg / mL, and nanoparticles were prepared using the method described in Example 1. The NP ratios of the nanoparticles prepared in this way correspond to 9, 7, and 5, respectively. The particle size and RNA encapsulation rate of the prepared nanoparticles were measured (Fig. 7).

[0142] As a result of measurement, it can be confirmed that when the NP ratio decreases, the particle size relatively increases and the RNA encapsulation rate tends to decrease.

[0143]

[0144] Foregoing, specific parts of the present invention have been described in detail. It will be apparent to those skilled in the art that such specific descriptions are merely preferred embodiments and do not limit the scope of the invention. Accordingly, the actual scope of the invention is defined by the appended claims and their equivalents.

Claims

1. A method for preparing lipid nanoparticles comprising the following steps: (a) a step of forming lipid nanoparticles by mixing an organic solution containing lipid components and an aqueous solution containing nucleic acid components; and (b) Step of removing the solvent from the mixed solution, Here, the above aqueous solution is characterized as being a neutral pH aqueous solution.

2. A method for manufacturing lipid nanoparticles according to claim 1, characterized in that the neutral pH aqueous solution is distilled water.

3. A method for producing lipid nanoparticles according to claim 1, characterized in that the neutral pH aqueous solution is a buffer solution prepared from an amine compound. 4.The method of claim 3, wherein the amine compound is TRIS (tris-hydroxymethyl-aminomethane), imidazole, HEPES (Hydroxyethyl piperazine Ethane Sulfonic acid), MOPSO (3-Morpholino-2-hydroxypropanesulfonic acid), MOPS (3-(N-Morpholino)propanesulfonic acid), PIPES (1,4-Piperazinediethanesulfonic acid), BES (N,N-Bis(2-hydroxyethyl)-2-aminoethanesulfonic acid), DIPSO (3-(N,N-Bis[2-hydroxyethyl]amino)-2-hydroxypropanesulfonic acid), HEPPSO (N-(2-Hydroxyethyl)piperazine-N′-2-hydroxypropanesulphonic acid) acid), POPSO (Piperazine-N,N′-bis(2-hydroxypropanesulfonic acid), EPPS A method for preparing lipid nanoparticles characterized by being selected from the group consisting of (4-(2-Hydroxyethyl)-1-piperazinepropanesulfonic acid), ACES (N-(2-Acetamido)-2-aminoethanesulfonic acid), TES (2-[(2-Hydroxy-1,1-bis(hydroxymethyl)ethyl)amino]ethanesulfonic acid), TAPSO (2-Hydroxy-3-[tris(hydroxymethyl)methylamino]-1-propanesulfonic acid), glycylglycine (Glycylglycine, Diglycine, Gly-Gly), tricin (N-[Tris(hydroxymethyl)methyl]glycine), and bicin (N,N-Bis(2-hydroxyethyl)glycine).

5. A method for manufacturing lipid nanoparticles according to claim 1, characterized in that the volume ratio of the organic solution to the aqueous solution is 2:1 to 1:

20.

6. A method for manufacturing lipid nanoparticles according to claim 1, characterized in that the lipid component is an ionized lipid, a PEG-lipid, or a mixture thereof.

7. In claim 6, the ionized lipid is an ionized lipid composed of a head portion containing an amine group and a hydrophobic lipid portion. (6Z, 9Z, 28Z, 31Z)-hephtatriaconta 6,9,28,31-tetraene-19-yl4-(dimethylamino)butanoate (DLin-MC3-DMA), [(4-hydroxybutyl)azandyl]di(hexane-6,1-diyl)bis(2-hexyldecanoate)(ALC-0315), 8-[(2-hydroxyethyl)[6-oxo-6-(undecyloxy)hexyl]amino]-octanic acid (SM-102), 1-linoleoyl-2-linoleyloxy-3-dimethylaminopropane (DLin-2-DMAP), 1,2-dilinoleylcarbamoyloxy-3-dimethylaminopropane (DLin-C-DAP), 1,2-Dilinoleoyl-3-Dimethylaminopropane (DLin-DAP), 1,2-Dilinoleyloxy-N,N-Dimethylaminopropane (DLin-DMA), 2,2-Dilinoleyl-4-Dimethylaminomethyl-[1,3]-Dioxolane (DLin-K-DMA), 2,2-Dilinoleyl-4-(2-Dimethylaminoethyl)-[1,3]-Dioxolane (DLin-KC2-DMA), 1,2-Dioleoyl-3-Dimethylammonium Propane (DODAP), N,N-Dimethyl-(2,3-Dioleyloxy)Propyamine (DODMA), DioctadeciramideGlycyrrhoxyspermin (DOGS), SperminCholesterylCarbamate (GL-67);Bis-Guanidinium-Spermidine-Cholesterol (BGTC), 3β-(N-(N',N'-dimethylaminoethane)-carbamoyl) cholesterol (DC-Chol), 1,1'-(2-(4-(2-((2-(bis(2-hydroxydecyl)amino)ethyl)(2-hydroxydecyl)amino)ethyl)piperazine-1-yl)ethylazandyl)dododecane-2-ol (C12-200), Nt-butyl-N'-tetradecylamino-propionamidine (diC14-amidine);Dimethyl dioctadecirinemonium bromide (DDAB), N-(1,2-dimyristyl oxyprop-3-yl)-N,N-dimethyl-N-hydroxyethylammonium bromide (DMRIE), N,N-dioleyl-N,N-dimethylammonium chloride (DODAC), dioleyl oxypropyl-3-dimethylhydroxyethylammonium bromide (DORIE), N-(1-(2,3-dioleyl oxyl)propyl)-N-2-(spermine carbokixamide)ethyl)-N,N-dimethylammonium trifluoroacetate (DOSPA), 1,2-dioleoyl trimethylammonium propane chloride (DOTAP), N-(1-(2,3-dioleyl oxy)propyl)-N,N,N-trimethylammonium chloride (DOTMA) and A method for preparing lipid nanoparticles characterized by selecting one or more from the group consisting of aminopropyl-dimethyl-bis(dodecyloxy)-propane aluminum bromide (GAP-DLRIE).

8. A method for manufacturing lipid nanoparticles according to claim 6, wherein the ionized lipid further comprises sterol lipids and neutral lipids.

9. A method for manufacturing lipid nanoparticles according to claim 8, characterized in that the sterol lipid is cholesterol or a cholesteryl ester.

10. A method for manufacturing lipid nanoparticles according to claim 8, characterized in that the neutral lipid is a phospholipid or a sphingolipid.

11. The method of item 10, wherein the phospholipids are DOPE (dioleoylphosphatidylethanolamine), DSPC (distearoylphosphatidylcholine), POPC (palmitoyloleoylphosphatidylcholine), EPC (egg phosphatidylcholine), DOPC (dioleoylphosphatidylcholine), DPPC (dipalmitoylphosphatidylcholine), DOPG (dioleoylphosphatidylglycerol), DPPG (dipalmitoylphosphatidylglycerol), DSPE (distearoylphosphatidylethanolamine), PE (Phosphatidylethanolamine), DPPE (dipalmitoylphosphatidylethanolamine), DOPE (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine), POPE (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine), POPC A method for preparing lipid nanoparticles characterized by being selected from the group consisting of (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine), DOPS (1,2-dioleoyl-sn-glycero-3-[phospho-L-serine]), ceramide, sphingomyelin, and mixtures thereof.

12. A method for manufacturing lipid nanoparticles according to claim 6, characterized in that the lipid is directly conjugated to the PEG or connected to the lipid through a linker moiety.

13. A method for producing lipid nanoparticles according to claim 12, characterized in that the linker moiety is an ester-free linker moiety or an ester-containing linker moiety.

14. A method for preparing lipid nanoparticles according to claim 13, wherein the ester-free linker moiety is selected from the group consisting of amido (-C(O)NH-), amino (-NR-), carbonyl (-C(O)-), carbamate (-NHC(O)O-), urea (-NHC(O)NH-), disulfide (-SS-), ether (-O-), succinyl (-(O)CCH2CH2C(O)-), succinamidyl (-NHC(O)CH2CH2C(O)NH-), hydrazone (-C(O)NHN=R-, -C=NNH-) and combinations thereof.

15. A method for preparing lipid nanoparticles according to claim 13, wherein the ester-containing linker moiety is selected from the group consisting of ester (-C(O)O-), carbonate (-OC(O)O-), succinoyl, phosphate ester (-O-(O)POH-O-), sulfonate ester, and combinations thereof.

16. A method for manufacturing lipid nanoparticles according to claim 12, characterized in that the lipid is a lipid capable of being combined with polyethylene glycol.

17. In Paragraph 16, A method for preparing lipid nanoparticles characterized in that the above lipid comprises a structure in which two types of lipids selected from C3-C30 single and branched chain forms of alkyl, alkenyl, and alkynyl hydrocarbons are connected to glycerol by ether (-O-) or ester (-C(O)O-, -OC(O)-) bonds.

18. Lipid nanoparticles produced by the method of any one of claims 1 to 17.

19. In paragraph 18, the lipid nanoparticles are lipid nanoparticles comprising nucleic acids and ionized lipids.

20. Lipid nanoparticles according to claim 18, characterized in that the particle size is 40 to 200 nm and the RNA encapsulation rate is 80 to 100%.

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