Microfluidic apparatus for producing lipid nanoparticles of various sizes, and manufacturing method using the same

Abstract

The present invention relates to a microfluidic apparatus for producing nucleic acid-deliverable lipid nanoparticles, and a method for producing lipid nanoparticles using the same. Using the microfluidic apparatus, lipid nanoparticles of a desired size can be produced by adjusting the molar ratio between compositions and the Reynolds number.

Description

[Technical Field]

[0001] The present invention relates to a microfluidic apparatus for producing lipid nanoparticles for nucleic acid delivery, and a method for producing lipid nanoparticles of various sizes using the same. Specifically, the present invention relates to a method for producing lipid nanoparticles of a specific size by adjusting the composition ratio and Reynolds number of the sterols and constituent components constituting the lipid nanoparticles, and to a microfluidic apparatus capable of efficient mixing for the production of lipid nanoparticles, enabling efficient mass production of lipid nanoparticles of various sizes. [Background technology]

[0002] Recently, the market for gene therapy drugs has been growing. Gene therapy drugs are medications that treat and prevent diseases by delivering genes to the lesion and producing proteins from those genes. A variety of gene therapy drugs with diverse mechanisms of action have been approved by the FDA or are currently undergoing clinical trials, ranging from COVID-19 vaccines from Pfizer and Moderna to Qalsody, a treatment for amyotrophic lateral sclerosis (ALS) from Biogen and IONS that utilizes RNA interference (RNAi).

[0003] For such gene therapies, it is crucial to accurately deliver genes that are easily damaged in the human body to the lesion site, and various studies are being conducted to improve the efficiency of delivery within the human body. Gene therapies using various types of carriers (vectors) are being studied, and since carriers (vectors) alone are still highly likely to be damaged in the human body, lipid nanoparticles (LNPs) are also being actively researched to safely protect and deliver them.

[0004] Lipid nanoparticles (LNPs) are nanoparticles composed of lipids such as phospholipids and cholesterol. They play a role in protecting carriers from damage within the human body and neutralizing their charge, thereby increasing their efficiency in permeating cell membranes. They are also effective in drug delivery using modalities other than genes, and many pharmaceutical companies are conducting research on new lipid nanoparticles. However, synthesizing and manufacturing such lipid nanoparticles presents a problem: it is difficult to control the spontaneous aggregation and mutual bonding of phospholipids and cholesterol, which can lead to the formation of nanoparticles of various sizes, such as micelles and liposomes.

[0005] Within lipid nanoparticles, cholesterol is known to have diverse effects on phospholipid-based biological membranes. Generally, the higher the cholesterol content in the biological membrane, the lower the membrane's fluidity and the stronger the bonds between phospholipids tend to be. Researchers have confirmed that lipid nanoparticles with a high cholesterol content, intended for the delivery of small molecules, exhibit improved drug loading efficiency and particle stability. For these reasons, lipid nanoparticles for nucleic acid delivery generally contain cholesterol as a component. However, as mentioned above, the inclusion of cholesterol makes it difficult to manufacture uniform lipid nanoparticles, and controlling their size while encapsulating gene therapy drugs at a high level is extremely challenging.

[0006] The COVID-19 vaccine is currently the most commercially available lipid nanoparticle vaccine. The messenger RNA (mRNA) used in it generally has a length of several thousand bases. The length of mRNA commonly used as a therapeutic agent is known to be 1,000 to 5,000 nt. In contrast, antisense oligonucleotides (ASOs), which have a length of several tens of bases, are a different concept of gene therapy substance that uses RNA expression inhibition, and development of therapeutic agents using lipid nanoparticles for these is also progressing. In the technology of utilizing lipid nanoparticles as nucleic acid delivery devices, different levels of leachate occur depending on the length of the nucleic acid base sequence used, and therefore the cholesterol composition ratio must be adjusted according to the length of the nucleic acid being carried. That is, in the case of nucleic acids with relatively long base sequences (e.g., mRNA), the ratio of equal amounts of phospholipid to cholesterol, which is the composition ratio of typical lipid nanoparticles, means an unnecessarily excessive cholesterol content. On the other hand, nucleic acids with relatively short base sequences (e.g., oligonucleotides) experience relatively high leachate in lipid nanoparticles, so a higher cholesterol content is required.

[0007] The inventors conducted various studies on the production of lipid nanoparticles with different composition ratios. As a result, they devised a method for producing lipid nanoparticles in which the molar ratio and Reynolds number of the lipid nanoparticle composition are adjusted, and a method for producing these nanoparticles with high efficiency using a separately designed, expandable microfluidic apparatus. By elucidating the effects of this method, the inventors completed the present invention. [Prior art documents] [Patent Documents]

[0008] [Patent Document 1] U.S. Patent No. 8058069 [Non-patent literature]

[0009] [Non-Patent Document 1] Camilla, H., A. et al. The role of lipid components in lipid nanoparticles for vaccines and gene therapy. Adv. Drug Deliv. Rev., 188, 114416 (2022) [Non-Patent Document 2] Cheng, Q. et al. Selective organ targeting (SORT) nanoparticles for tissue-specific mRNA delivery and CRISPR-Cas gene editing. Nat. Nanotech., 15(4), 313-320 (2020) [Overview of the Initiative] [Problems that the invention aims to solve]

[0010] As mentioned earlier, lipid nanoparticles are formed from a variety of lipids and include a variety of nanoparticle by-products, which presents a problem in that it is very difficult to control the size of the synthesized particles.

[0011] The present invention aims to provide a method for controlling the particle size, which varies depending on the molar ratio between lipid nanoparticle compositions and the Reynolds number, for lipid nanoparticles intended for nucleic acid delivery.

[0012] The present invention aims to provide a manufacturing method that improves the efficiency of lipid nanoparticle production by carrying out the above manufacturing method on a separately devised microfluidic apparatus. [Means for solving the problem]

[0013] The present invention relates to a microfluidic device comprising an inlet channel, a mixing channel, and an outlet channel, (a) The step of injecting a lipid mixture into the inflow channel, (b) Injecting nucleic acid into an inflow channel different from the aforementioned inflow channel; (c) Mixing a lipid mixture and nucleic acid in the aforementioned mixing channel to produce lipid nanoparticles, and providing a method for producing lipid nanoparticles using a microfluidic device.

[0014] The mixing channel may include microposts.

[0015] In the microfluidic device, there may be two or more inflow channels, and the flows of fluids flowing into different inflow channels among the inflow channels may be in different directions from each other.

[0016] The lipid mixture can include at least one selected from the group consisting of ionizable lipid, helper lipid, PEGylated lipid, and sterol, but is not limited thereto.

[0017] The ionizable lipid can include at least one selected from the group consisting of 9-heptadecanoyl 8-{ (2-hydroxyethyl)[6-oxo-6-(undecyloxy)hexyl]amino}octanoic acid (SM-102), 1,2-dilinoleyloxy-N,N-dimethyl-3-aminopropane (DLinDMA), 2,2-dilinoleyl-4-dimethylaminoethyl-[1,3]-dioxolane (DLin-KC2-DMA), (6Z,9Z,28Z,31Z)-heptatriaconta-6,9,28,31-tetraene-19-yl 4-(dimethylamino)butanoic acid (DLin-MC3-DMA), 6-((2-hexyldecanoyl)oxy)-N-(6-((2-hexyldecanoyl)oxy)hexyl)-N-(4-hydroxybutyl)hexane-1-aminium (ALC-0315), 1,2-dioleyloxy-3-dimethylaminopropane (DODMA), 1,2-dioleyl-3dimethylammonium-propane (DODAP), etc., but is not limited thereto.

[0018] In one embodiment, the ionizable lipid may be 9-heptadecanoyl 8-{(2-hydroxyethyl)[6-oxo-6-(undecyloxy)hexyl]amino}octanoic acid (SM-102).

[0019] The helper lipid may be at least one selected from the group consisting of distearoyl phosphatidylcholine (DSPC), palmitoyl oleoyl phosphatidylcholine (POPC), 1,2-dioleoyl-sn-glycero-3-phosphate (18:1PA), dipalmitoyl phosphatidylcholine (DPPC), dioleoyl phosphatidylcholine (DOPC), dioleoyl phosphatidylglycerol (DOPG), dipalmitoyl phosphatidylglycerol (DPPG), dioleoyl phosphatidylethanolamine (DOPE), palmitoyl oleoyl phosphatidylethanolamine (POPE), dioleoyl-phosphatidylethanolamine 4-(N-maleimidomethyl)-cyclohexane-1-carboxylate (DOPE-mal), dipalmitoyl phosphatidylethanolamine (DPPE), 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), sphingomyelin (SM), and dioleoyl phosphatidylglycerol (DOPG), etc., but is not limited thereto.

[0020] In one embodiment, the helper lipid may be distearoyl phosphatidylcholine (DSPC) or palmitoyl oleoyl phosphatidylcholine (POPC).

[0021] The PEGylated lipid may be at least one selected from the group consisting of 1,2-dimyristoyl-rac-glycerol methoxypolyethylene glycol (DMG-PEG), polyethylene glycol-diacylglycerol (PEG-DAG), polyethylene glycol-dialkyloxypropyl (PEG-DAA), polyethylene glycol-dimethacrylate (PEG-DMA), distearoyl-rac-glycerol methoxypolyethylene glycol (DSG-PEG), polyethylene glycol-ceramide (PEG-Ceramide), polyethylene glycol-phospholipid (PEG-phospholipid), polyethylene glycol-phosphatidylethanolamine (PEG-PE), polyethylene glycol-succinoyl 1,2-diacylglycerol (PEG-S-DAG), and polyethylene glycol-dialkyloxypropylcarbamate (PEG-dialkyloxypropylcarbamate), but is not limited thereto.

[0022] In one embodiment, the PEGylated lipid may be 1,2-dimyristoyl-rac-glycerol methoxypolyethylene glycol (DMG-PEG2k).

[0023] The aforementioned sterols include cholesterol, ergosterol, campesterol, stigmasterol, sitosterol, fucosterol, betulin, lupeol, ursolic acid, oleanolic acid, brassicasterol, 9,11-dehydroergosterol, daucosterol, β-sitosterol acetate, and β-sitosterol amino acid conjugates. It may be, but is not limited to, at least one selected from the group consisting of conjugates, 20-hydroxycholesterol, 22-hydroxycholesterol, 24-hydroxycholesterol, 25-hydroxycholesterol, and 27-hydroxycholesterol.

[0024] In one embodiment, the sterol may be cholesterol.

[0025] The nucleic acid may be at least one selected from the group consisting of mRNA, siRNA, aiRNA, miRNA, dsRNA, shRNA, lncRNA, saRNA, rRNA, tRNA, piRNA, circRNA, RNA, DNA, cDNA, plasmid, aptamer, oligonucleotide, ribozyme, PNA, and DNAzyme, but is not limited thereto.

[0026] In one embodiment, the nucleic acid may be mRNA or oligonucleotide.

[0027] The polydispersity index (PDI) of the lipid nanoparticles may be 0.3 or less, preferably 0.2 or less.

[0028] The nucleic acid encapsulation rate of the lipid nanoparticles may be 50% or more, preferably 70% or more.

[0029] The nucleic acid content of the lipid nanoparticles may be 15% or more, preferably 20% or more.

[0030] The size of the lipid nanoparticles can be adjusted by the Reynolds number with respect to the fluid flow in the mixing channel, and the Reynolds number may be 10 to 150, preferably 25 to 100.

[0031] The size of the lipid nanoparticles may be 10 nm to 200 nm, preferably 20 nm to 150 nm, and most preferably 30 nm to 90 nm.

[0032] The size of the lipid nanoparticles can be adjusted by the molar ratio of the lipid mixture to the sterol. The molar ratio of the lipid mixture to the sterol may be 1:0.1 or higher, preferably 1:0.3 or higher.

[0033] The present invention provides lipid nanoparticles obtained by the method for producing lipid nanoparticles described above.

[0034] The present invention includes an inflow channel, a mixing channel, and an outflow channel. The aforementioned inflow channels are two or more, The aforementioned mixed channel includes a micropost, The present invention provides a microfluidic apparatus for producing lipid nanoparticles, characterized in that fluids flowing into different inflow channels flow in different directions.

[0035] A fluid containing a lipid mixture and nucleic acids can flow into the microfluidic device through the inflow channel, and the lipid mixture may, but is not limited to, include at least one selected from the group consisting of ionizable lipids, helper lipids, PEGylated lipids, and sterols.

[0036] The lipid mixture can flow into one inflow channel, while the nucleic acids can flow into the other inflow channel.

[0037] The lipid mixture can flow in the same direction as the fluid flow, while the nucleic acids can flow in a direction different from the fluid flow, preferably in a direction perpendicular to the fluid flow.

[0038] The aforementioned micropost may or may not be connected to the wall constituting the mixed channel.

[0039] The microposts may be arranged in one or more columns in the direction of fluid flow, and in a single column, one or more microposts may be arranged in a direction different from the direction of fluid flow.

[0040] The plurality of microposts do not necessarily have to be arranged in a line in the direction of fluid flow; they may be arranged so as to partially or completely cover the gaps between microposts arranged in adjacent rows when viewed from the direction of fluid flow.

[0041] The size of the lipid nanoparticles can be adjusted by the Reynolds number with respect to the fluid flow in the mixing channel, and the Reynolds number may be 10 to 150, preferably 25 to 100.

[0042] The size of the lipid nanoparticles may be adjusted by the molar ratio of the lipid mixture to the sterol. The molar ratio of the lipid mixture to the sterol may be 1:0.1 or higher, preferably 1:0.3 or higher.

[0043] The polydispersity index (PDI) of the lipid nanoparticles may be 0.3 or less, preferably 0.2 or less.

[0044] The nucleic acid encapsulation rate of the lipid nanoparticles may be 50% or more, preferably 70% or more.

[0045] The nucleic acid content of the lipid nanoparticles may be 15% or more, preferably 20% or more. [Effects of the Invention]

[0046] The present invention provides an optimal method for producing lipid nanoparticles by controlling the particle size of the lipid nanoparticles through adjustment of the composition ratio of raw materials containing cholesterol. The present invention's method for producing lipid nanoparticles can be used in the production of pharmaceuticals containing lipid nanoparticles.

[0047] Furthermore, by using the lipid nanoparticle manufacturing method of the present invention in the manufacturing process of pharmaceuticals that use nucleic acids as a modality, messenger ribonucleic acid can be encapsulated in lipid nanoparticles of various sizes at high concentrations, thereby improving therapeutic efficiency. [Brief explanation of the drawing]

[0048] [Figure 1] This is a schematic diagram illustrating the synthesis process of lipid nanoparticles (LNPs) encapsulating nucleic acids using the micromixing channel apparatus of the present invention, and the morphology of endogenous nucleic acid LNPs. [Figure 2] This is a schematic diagram showing the lengths of different parts of the apparatus for the structure and design of the apparatus of the present invention. [Figure 3] This shows the fluid flow using the apparatus of the present invention and the fluid flow around the micropost. [Figure 4] The results of measuring the mixing efficiency based on the number of rows of microposts in the apparatus of the present invention are shown. [Figure 5]The results of measuring the mixing efficiency based on the number of microposts present in a single row in the apparatus of the present invention are shown. [Figure 6] This result shows a Dean vortex formed by the width spacing of microposts arranged in the mixing area within the apparatus of the present invention. [Figure 7] These results show the Dean number and mixing efficiency based on the width spacing of the microposts arranged in the mixing area of ​​the apparatus of the present invention. [Figure 8] The results of measuring the mixing efficiency with respect to the change in Reynolds number in the apparatus of the present invention are shown. [Figure 9] The results of measuring the mixing efficiency due to changes in channel height in the apparatus of the present invention are shown. [Figure 10] The results of measuring the mixing efficiency based on the injection ratio of aqueous solution and lipid mixture in the apparatus of the present invention are shown. [Figure 11] This section describes a method for calculating the residence time of a fluid. [Figure 12] The fluid flow within the apparatus according to Manufacturing Example 1 and Manufacturing Example 2 of the present invention is shown. [Figure 13] The results of measuring the mixing efficiency in the apparatus of the present invention, based on changes in the structure's starting interval, the spacing of microposts, and the vertical length of microposts, are shown. [Figure 14] The results of measuring the mixing efficiency due to changes in the blockage rate in the apparatus of the present invention are shown. [Figure 15] This result shows the mixing efficiency within the apparatus based on the front and rear width spacing provided by the microposts in the apparatus of the present invention. [Figure 16] This result shows the shear ratio based on the Reynolds number for different width intervals using microposts within the apparatus of the present invention. [Figure 17] This result shows the size-specific design conditions for the device of the present invention. [Figure 18] This result shows the mixing efficiency of the apparatus of the present invention based on the Reynolds number for different sizes. [Figure 19] This shows the flow velocity based on the Reynolds number for different sizes of the apparatus according to the present invention. [Figure 20]This result shows the shear rate based on the Reynolds number for different sizes of the apparatus of the present invention. [Figure 21] This graph compares the Reynolds number, LNP size, and polydispersity index (PDI) when the composition ratio (molar ratio) of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k is 50:10:38.5:1.5. [Figure 22] and [Figure 23] Figure 21 shows the DLS results for LNP size as a function of Reynolds number. [Figure 24] This graph compares the Reynolds number, LNP size, and polydispersity index (PDI) when the composition ratio (molar ratio) of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k is 28:5.7:65.5:0.8. [Figure 25] and [Figure 26] Figure 24 shows the DLS results for LNP size as a function of Reynolds number. [Figure 27] This graph compares the Reynolds number, LNP size, and polydispersity index (PDI) when the composition ratio (molar ratio) of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k is 20:4:75.5:0.5. [Figure 28] and [Figure 29] Figure 27 shows the DLS results for LNP size as a function of the Reynolds number. [Figure 30] This graph compares the size of LNPs and PDI based on the molar ratio of cholesterol. [Figure 31] and [Figure 32] Figure 30 shows the DLS results for LNP size based on cholesterol molar ratio, and is a number distribution diagram. [Figure 33] and [Figure 34]Figure 30 shows the DLS results for LNP size based on cholesterol molar ratio, and is a volume distribution diagram. [Figure 35] and [Figure 36] Figure 30 shows the DLS results for LNP size based on cholesterol molar ratio, and is an intensity distribution diagram. [Figure 37] This graph compares the size of LNPs and the polydispersity index (PDI) at Reynolds numbers 50 and 100 when the composition ratio (molar ratio) of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k is 48.5:9.5:37.5:4.5. [Figure 38] Figure 37 shows the DLS results for LNP size as a function of Reynolds number. [Figure 39] This graph compares the size and PDI of bare LNPs and mCherry LNPs (which lack mRNA) under synthesis conditions for a size of 30 nm. [Figure 40] This graph compares the size and PDI of bare LNPs and mCherry LNPs (which lack mRNA) under synthesis conditions for a size of 60 nm. [Figure 41] This graph compares the size and PDI of bare LNPs and mCherry LNPs (which lack mRNA) under synthesis conditions for a size of 90 nm. [Figure 42] and [Figure 43] Figures 39, 40, and 41 show the DLS results for the sizes of bare LNPs and mCherry LNPs without mRNA. [Figure 44] and [Figure 45] This graph compares the size and PDI of bare LNPs (without mRNA) and Cas9 LNPs under synthesis conditions for a size of 30 nm. [Figure 46] This graph compares the size and PDI of bare LNPs (LNPs without mRNA) and Cas9 LNPs under synthesis conditions for a size of 60 nm. [Figure 47]This graph compares the size and PDI of bare LNPs (without mRNA) and Cas9 LNPs under synthesis conditions for a size of 90 nm. [Figure 48] and [Figure 49] Figures 44, 45, 46, and 47 show the DLS results for the sizes of bare LNPs and Cas9 LNPs without mRNA. [Figure 50] This graph compares the encapsulation efficiency (EE) of mCherry LNPs under synthesis conditions of sizes 30nm, 60nm, and 90nm. [Figure 51] This graph compares the encapsulation efficiency (EE) of Cas9 LNPs under synthesis conditions of sizes 30nm, 60nm, and 90nm. [Figure 52] This graph compares the size and PDI of bare LNPs (LNPs without mRNA) and oligo LNPs (LNPs without mRNA) under synthesis conditions for a size of 30 nm. [Figure 53] This graph compares the size and PDI of bare LNPs (LNPs without mRNA) and oligo LNPs (LNPs without mRNA) under synthesis conditions for a size of 60 nm. [Figure 54] This graph compares the size and PDI of bare LNPs (LNPs without mRNA) and oligo LNPs (LNPs without mRNA) under synthesis conditions for a size of 90 nm. [Figure 55] and [Figure 56] Figures 52, 53, and 54 show the DLS results for the size of bare LNPs and oligo LNPs without mRNA. [Figure 57] This graph compares the encapsulation efficiency (EE) of Oligo LNPs under synthesis conditions of sizes 30nm, 60nm, and 90nm. [Figure 58] These are fluorescence images showing the degree of mCherry protein expression after processing human glioblastoma U87MG with mCherry LNP (200 ng of mCherry mRNA) under synthesis conditions of sizes 30 nm, 60 nm, and 90 nm. [Figure 59] Figure 58 is a graph showing the quantification of the fluorescence of the mCherry protein in the U87MG fluorescence image. [Figure 60] This is a fluorescence image showing the degree of mCherry protein expression in the human hepatocellular carcinoma cell line HepG2, under synthesis conditions of 60 nm and 90 nm sizes, using mCherry mRNA processing amounts (500, 200, and 20 ng). [Figure 61] Figure 60 is a graph showing the quantification of the fluorescence of the mCherry protein in the HepG2 fluorescence image. [Figure 62] This is a fluorescence image showing the degree of mCherry protein expression in the human mononuclear cell line THP-1, under synthesis conditions of 60 nm and 90 nm sizes, using mCherry mRNA processing amounts (500, 200, and 20 ng). [Figure 63] Figure 62 is a graph showing the quantification of the fluorescence of the mCherry protein in the THP-1 fluorescence image. [Figure 64] and [Figure 65] These are the results regarding the storage stability of 60 nm mCherry LNPs after synthesis, in terms of size, polydispersity index (PDI), mCherry mRNA encapsulation rate (EE), and endogenous rate (LE). [Figure 66] and [Figure 67] These are the results regarding the storage stability of 90nm mCherry LNPs after synthesis, in terms of size, polydispersity index (PDI), mCherry mRNA encapsulation rate (EE), and endogenous rate (LE). [Figure 68] and [Figure 69] These are the results regarding the storage stability of 30nm Cas9 LNPs after synthesis, in terms of size, polydispersity index (PDI), Cas9 mRNA encapsulation rate (EE), and endogenous rate (LE). [Figure 70] and [Figure 71] These are the results regarding the storage stability of 90nm Cas9 LNPs after synthesis, in terms of size, polydispersity index (PDI), Cas9 mRNA encapsulation rate (EE), and endogenous rate (LE). [Figure 72] and [Figure 73] These are the results regarding the storage stability of 50nm oligo LNPs (target size: 30nm) after synthesis, specifically in terms of size, polydispersity index (PDI), oligonucleotide encapsulation rate (EE), and endogenous content (LE). [Figure 74] and [Figure 75] These are the results regarding the storage stability of 100 nm oligo LNP (target size: 60 nm) after synthesis, specifically in terms of size, polydispersity index (PDI), oligonucleotide encapsulation rate (EE), and endogenous content (LE). [Figure 76] and [Figure 77] These are the results regarding the storage stability of 130nm oligo LNP (target size: 90nm) after synthesis, specifically in terms of size, polydispersity index (PDI), oligonucleotide encapsulation rate (EE), and endogenous content (LE). [Modes for carrying out the invention]

[0049] Hereinafter, embodiments and examples of the present application will be described in detail with reference to the attached drawings, so that they can be easily implemented by a person with ordinary skill in the art to which the present invention pertains. However, the present application can be realized in various forms and is not limited to the embodiments and examples described herein.

[0050] In the entirety of the specification of this application, when a part "includes" a certain component, this means that, unless otherwise stated, it may include other components rather than excluding them.

[0051] As used herein, the term "lipid nanoparticles" refers to four types of nanoparticles: ionizable lipids, helper lipids, PEGylated lipids, and sterols, where helper lipids may consist of at least one lipid. It also refers to nano-sized particles capable of encapsulating nucleic acids.

[0052] As used herein, the terms “apparatus” or “microfluidic apparatus” mean an apparatus that includes microchannels, etc., provided on a substrate made of a variety of materials including plastics containing organic polymers, glass, metal, or silicone, through which fluids can flow.

[0053] In the apparatus of the present invention, the inlet channel, mixing channel, and outlet channel can be arranged sequentially in the direction of fluid flow.

[0054] The inflow channel may include a channel through which nucleic acids flow and a channel through which a lipid mixture flows, and the nucleic acids and the lipid mixture may each flow in through different inflow channels. Here, the lipid mixture may flow in the same direction as the fluid flow direction in the mixing channel, and the nucleic acids may flow in the same direction as the fluid flow direction in the mixing channel. Preferably, the lipid mixture may flow in the same direction as the fluid flow direction, and the nucleic acids may flow in a direction perpendicular to the fluid flow direction.

[0055] The inflow channel through which the lipid mixture flows may be one or more, and the inflow channel through which the nucleic acid flows may be one or more. Preferably, the apparatus of the present invention may include one inflow channel through which the lipid mixture flows and two inflow channels through which the nucleic acid flows.

[0056] In the aforementioned mixing channel, nucleic acids and lipid mixtures can be mixed with each other.

[0057] The particle size can be adjusted by changing the Reynolds number with respect to the fluid flow in the mixing channel. The Reynolds number with respect to the fluid flow in the mixing channel may be 10 to 150, preferably 25 to 100.

[0058] The height of the channel may be 200 to 800 μm.

[0059] As used herein, the term “fluid flow” refers to the direction in which fluid flows from the inlet channel to the outlet channel within the apparatus. Preferably, the lipid mixture flows in through an inlet channel positioned in the same direction as the flow direction in the mixing channel, and the nucleic acids flow in through an inlet channel positioned perpendicular to the flow direction in the mixing channel.

[0060] Generally, the flow velocity within a device can be expressed by the Reynolds number (Re), a dimensionless number that represents the ratio of inertial force to viscous force, and its calculation formula is as follows:

number

[0061] At this time, D h ρ represents the hydraulic diameter within the microfluidic apparatus, while ρ, μ, ν, and Q represent the fluid density, kinematic viscosity, velocity, and flow rate, respectively. Therefore, the flow rate increases proportionally with increasing Reynolds number. When the Reynolds number exceeds 500, turbulence-like flow is formed within the apparatus, preventing the generation of controllable vortex patterns. The increased shear force between the solvent and the substance due to the high flow rate can cause instability in the raw material drug and the resulting nanoparticles. When the Reynolds number is 300 or higher, uncontrollable chaotic flow begins to occur within the apparatus, making quality control for the production of uniform nanoparticles difficult. Conversely, when the Reynolds number is low (10 or less), effective mixing of lipid mixtures and nucleic acids becomes difficult due to diffusion, making it difficult to achieve superior productivity compared to existing nanoparticle synthesis methods.

[0062] The apparatus of the present invention can adjust particle size by changing the Reynolds number.

[0063] As used herein, the term "micropost" refers to a structure that obstructs the linear flow of a fluid within a mixing channel. The microposts of the present invention are structures that allow a fluid flowing into a device to form a microvortex and mix efficiently, and can include all forms, preferably columnar. Microposts can be designed to separate, meander, or merge fluid flows that collide with them. Microposts can be designed to maintain the main fluid flow and prevent stagnation, for example, by colliding with the fluid at a right angle to the fluid with respect to the direction of fluid flow.

[0064] The micropost may or may not be connected to the wall forming the mixed channel, and may be at least one selected from the group consisting of a polygonal pyramid, a frustum of a polygon, a polygonal prism, and variations thereof, and may preferably be a quadrangular prism.

[0065] The microposts can be arranged in one or more columns in the direction of fluid flow, preferably in six or more columns in the direction of fluid flow. In addition, one or more microposts can be arranged in a direction different from the direction of fluid flow within a single column, preferably 1 to 6 microposts can be arranged in a direction different from the direction of fluid flow within a single column, more preferably 1 to 6 microposts can be arranged in a direction perpendicular to the direction of fluid flow within a single column, and most preferably 1 to 2 microposts can be arranged in a direction perpendicular to the direction of fluid flow within a single column.

[0066] The plurality of microposts do not necessarily have to be arranged in a line in the direction of fluid flow; the plurality of microposts may be arranged so as to partially or completely cover the gaps between microposts arranged in adjacent lines when viewed from the direction of fluid flow.

[0067] In the present invention, microposts may be arranged alternately with microposts in adjacent rows. In the present invention, "arranged alternately" means that the multiple rows of microposts are not arranged in a straight line or parallel to each other in a single row in the direction of fluid flow, and microposts arranged in one row may be arranged to partially or completely cover the gaps between microposts in adjacent rows when viewed from the direction of fluid flow.

[0068] In the present invention, the microposts have a height of 200 to 800 μm, and the microposts may or may not be connected to the walls constituting the mixing channel.

[0069] The term "mixing efficiency" as used herein eff This value expresses the ratio as a percentage, using the fact that a lipid mixture and nucleic acids, each having a mass fraction of 1, are mixed with each other in a channel and eventually converge to 0.5. The formula for calculating this is as follows:

number

[0070] This time, Mix eff This represents the mass fraction, and the closer the mass fraction is to 0.5, the higher the synthesis efficiency proportionally.

[0071] The aforementioned efflux channel may be a channel through which the generated lipid nanoparticles leach out.

[0072] As used herein, the term "inlet volume flow rate ratio" refers to the ratio of the inlet volume flow rate of an aqueous solution to the inlet volume flow rate of a fluid containing a lipid mixture.

[0073] As used herein, the term "flow blockage ratio" is the ratio of the width of microposts to the width of the overall channel in the first column along the direction of fluid flow in a mixing channel.

[0074] The term "encapsulation efficiency (EE)" as used herein is calculated using a formula obtained by detecting the total mRNA and unencapsulated mRNA (free mRNA) based on fluorescence intensity in each mRNA LNP.

number

[0075] The term "Total mRNA" refers to the total amount of mRNA contained within the final product of the lipid nanoparticle manufacturing process, while "Free mRNA" refers to the amount of mRNA present outside the lipid nanoparticles in the final product of the lipid nanoparticle manufacturing process.

[0076] The term "loading efficiency (LE)" as used herein is calculated using a formula obtained by detecting the total mRNA and unencapsulated mRNA (free mRNA) based on fluorescence intensity in each mRNA LNP.

number

[0077] The aforementioned "feeding mRNA" refers to the total amount of mRNA introduced into the lipid nanoparticle manufacturing process.

[0078] The present invention will be described in more detail below through examples, but these examples are for illustrative purposes only and are not intended to limit the scope of the present invention. [Examples]

[0079] [Example 1] Design of equipment for lipid nanoparticle production A microfluidic device for producing lipid nanoparticles (LNPs) was designed (Figures 1, 2, and 3). The microfluidic device included three inlet channels and one outlet channel. Of the three inlet channels, the central channel was injected with a lipid mixture, and the two channels on either side were injected with aqueous solutions containing nucleic acids. A micropost structure was also introduced inside the device to effectively mix the lipid mixture and nucleic acids.

[0080] [Example 2] The effect of a single micropost structure To confirm the mixing efficiency of the lipid mixture and nucleic acid aqueous solution based on the presence, number, and arrangement of microposts placed inside the device, the distribution of the lipid mixture within the microfluidic device was analyzed using hydrodynamic techniques.

[0081] As shown in Figure 3, we confirmed that the lipid mixture and the nucleic acid aqueous solution are efficiently mixed by microvortexes generated when the lipid mixture flows through the micropost structure.

[0082] In this invention, the inlet volume flow rate ratio refers to the ratio of the inlet flow rate of an aqueous solution fluid to the inlet flow rate of a fluid containing a lipid mixture, and the flow blockage ratio is the ratio of the width of the microposts to the width of the overall channel in the first column along the fluid flow direction in the mixing channel.

[0083] The results were confirmed under the following conditions: channel height of 200 μm, inlet volume flow rate ratio of 1:5.5 (lipid mixture:aqueous solution), Reynolds number (Re) of 50, flow blockage ratio of 0.5, and rectangular micropost size of 1000 μm × 400 μm. The results are shown in Figures 4 and 5. The number of microposts was designed to be evenly distributed across the channel width while maintaining the micropost size and keeping the fluid-passage width between microposts constant.

[0084] As shown in Figures 4 and 5, we confirmed that the mixing efficiency improved by approximately 1.5 to 4.0 times when microposts were present compared to when they were absent, depending on the number of microposts present.

[0085] [Table 1]

[0086] As shown in Figure 4 and Table 1, the mixing efficiency increases as the number of columns increases, and it can be seen that when there are 6 or more columns, an excellent mixing efficiency of 98% or more is observed.

[0087] [Table 2]

[0088] As shown in Figure 5 and Table 2, the mixing efficiency actually decreases as the number of microposts located within a channel in a single column increases, and the best mixing efficiency is observed when the number is 1 to 2.

[0089] Therefore, the mixing efficiency changes depending on the number and arrangement of microposts placed inside the device, and it can be seen that the highest mixing efficiency is obtained when the single micropost structure is arranged alternately, as shown in Figures 6 and 7. The microposts were evenly distributed with respect to the channel width, and the width spacing through which the fluid passes between the placed microposts was kept constant.

[0090] [Example 3] Optimization of the usable flow velocity range and flow velocity ratio range within the device. To optimize LNP synthesis by controlling the flow rates of lipid mixtures and aqueous solutions within the apparatus, as well as their injection ratios, mixing efficiency was analyzed under various Reynolds number (Re) and flow rate ratio conditions.

[0091] To investigate the usable flow velocity range, the mixing efficiency was analyzed in the apparatus when the Reynolds number was varied from 12.5 to 200 with a channel height of 200 μm and an injection ratio of 1:5.5. The results are shown in Figure 8 and Table 3.

[0092] [Table 3]

[0093] As shown in Table 3, we confirmed that mixing efficiencies of 90% or higher were obtained when the Reynolds number was between 12.5 and 200, and that mixing efficiencies of 98% were obtained when the Reynolds number was 50 and 100.

[0094] Furthermore, the mixing efficiency was analyzed when the channel height was varied from 200 to 800 μm at an injection ratio of 1:5.5 and a Reynolds number of 50, and the results are shown in Figure 9 and Table 4.

[0095] [Table 4]

[0096] As shown in Table 4, we confirmed that the mixing efficiency was 95% or higher when the channel height was 200 to 800 μm.

[0097] Furthermore, the mixing efficiency was analyzed when the injection ratio was varied from 1:2.5 to 1:8.5 in the apparatus with a channel height of 200 μm and a Reynolds number (Re) of 50, and the results are shown in Figure 10 and Table 5.

[0098] [Table 5]

[0099] As shown in Table 5, we confirmed that when the injection ratio was between 1:2.5 and 1:8.5, the mixing efficiency was 97% or higher.

[0100] [Example 4] Research on apparatus height for increasing mixing efficiency To maximize the mixing efficiency based on the height of the mixing channel in the device, the mixing efficiency was analyzed under different height conditions.

[0101] When raw materials are injected at a relatively high flow rate (Re>50) during the nanoparticle synthesis process using the apparatus, strong shear stress is generated from the walls due to the narrow width of the channel.

[0102] Organic substances in fluids have been reported to aggregate and deform above certain thresholds. In particular, in the case of proteins, various studies indicate that shear stress from fluid flow generated in external equipment induces protein deformation, including aggregation, bending, and degradation. Such deformation of precursor proteins alters the structure, size, and function of the manufactured nanoparticles. Generally, the range in which deformation is induced varies depending on the structure of the equipment generating shear stress, and the form and size of the materials used. However, for proteins with a size of 1 nm to 10 nm (mass range of 20 kDa to 300 kDa), such as insulin, enzymes, and immunoglobulins, a shear stress of 1000 dyne / cm² is considered to induce deformation. 2It has been reported that shear flow can deform the structure of proteins within a certain range (Bekard, I., et al. The Effects of Shear Flow on Protein Structure and Function. Biopolymers, 95, 11, 733-745 (2011)).

[0103] Therefore, in the apparatus of the present invention, shear stress τ generated via the fluid flow w To confirm whether or not the precursor material and nanoparticles were deformed, the shear stress generated according to each channel height was first calculated as follows.

number

[0104] In this case, Q is the flow rate, and μ, h, and w represent the kinematic viscosity coefficient of the fluid, the height of the channel cross-section, and the lateral length of the channel cross-section, respectively.

[0105] [Table 6]

[0106] As shown in Table 6, in the apparatus of the present invention, the shear force increases geometrically as the channel height decreases, but the protein structure deformation is 1000 dyne / cm. 2 We confirmed that it did not exceed the specified range.

[0107] However, if the channel height is 100 μm, increasing the Reynolds number, which corresponds to the flow velocity, to 300 will result in a flow rate of 1000 dyne / cm². 2 This range is reached. Furthermore, considering that the size of the nanoparticles to be manufactured is in the range of 10 nm to 100 nm, which is larger than the size of the protein, it can be seen that when the channel height is 200 μm or more, the precursor material and nanoparticles do not deform due to shear stress.

[0108] Also, as shown in Table 7 below, when the height of the mixing channel of the device increases, the volume increases, and the residence time τ of the precursor to be mixed for the same flow rate remains in the channel. res and the mixing time τ required for the synthesis of the precursor mix increases. The specific calculation method of the residence time for the substance to be mixed to remain in the channel is shown in Fig. 11. The mixing time τ required for the synthesis of the substance mix can be calculated by the more dominant physical phenomenon between convection and diffusion in the device, and can be calculated as follows using the Peclet number (Pe), which is a dimensionless number indicating the ratio of the convection time to the diffusion time. [Number]

[0109] In the device of the present invention, the Peclet number (Pe) has a value of 100 to 5000. Based on this, it can be seen that mixing by convection is dominant. Therefore, the mixing time τ by convection in the channel mix、convection is calculated as follows (Valencia, P. et al. Single-Step Assembly of Homogenous Lipid-Polymeric and Lipid-Quantum Dot Nanoparticles Enabled by Microfluidic Rapid Mixing. ACS Nano 4, 3, 1671-1679 (2010), and Rhee, M. et al. Drop Mixing in a Microchannel for Lab-on-a-Chip Platforms. Langmuir, 24 (2), 590-601 (2008)). [Number]

[0110] At this time, D represents the diffusion coefficient, and L and w represent the length of the channel and the horizontal length of the channel cross-section, respectively.

[0111] [Table 7]

[0112] In this case, basically, for the substance to be sufficiently mixed within the channel, the residence time τ is necessary. res is the mixing time τ mix It must be longer than the residence time, but as shown in Table 7, the residence time is longer than the mixing time at all channel heights, so we can confirm that sufficient mixing time has been ensured at the given channel height.

[0113] Furthermore, the increase in mixing time due to the increase in channel height is calculated to be 53 ms at the largest height of 800 μm, as shown in Table 7. This is because the aggregation time τ is the time it takes for the injected substances to aggregate and come together. agg This is the time at which the range begins to fall within 50-100 ms, which can be considered the maximum height for applying the apparatus and manufacturing method of the present invention (see Rohit, K et al. Microfluidic Platform for Controlled Synthesis of Polymeric Nanoparticles. Nano Letters, 8, 9, 2906-12, (2008), and Johnson, BK et al. Mechanism for Rapid Self-Assembly of Block Copolymer Nanoparticles. Physical Review Letters, 91, 11, 118302-1-4 (2003)).

[0114] Therefore, as shown in Table 7, the optimal height of the mixing channel that can prevent aggregation of nanoparticles and the generation of non-uniform nanoparticles within the apparatus to the greatest extent possible is 200 to 800 μm.

[0115] [Example 5] Fluid flow within the device Based on channel height and micropost conditions, the apparatus of the present invention was designed as two manufacturing examples, as shown in Table 8 below.

[0116] [Table 8]

[0117] The above-mentioned Production Examples 1 and 2 were prepared, and in order to confirm the mixing flow pattern of the lipid mixture and aqueous solution, the distribution of the substances was visualized using ink and observed under a microscope.

[0118] Fluids containing lipid mixtures were visualized using 6% ethanol ink, and fluids containing hydrophilic substances were visualized using physiological saline. The results are shown in Figure 12. As shown in Figure 12, it can be seen that the lipid mixture and aqueous solution are efficiently mixed by microvortexes generated when the lipid mixture flows through the micropost structure.

[0119] [Example 6] Optimization of the structural design within the device To optimize the channel size and micropost mixing efficiency within the device, the mixing efficiency was verified under various conditions within the device structure, including 1) the starting interval of microposts, 2) the spacing between microposts, 3) the vertical length of microposts, and 4) the flow blocking ratio.

[0120] The mixing efficiency for each variable was confirmed under fixed channel height of 0.2 mm, injection ratio of 1:5.5, and Reynolds number of 50. The results are shown in Figures 13 and 14.

[0121] [Table 9]

[0122] As shown in Figure 13 and Table 9, we confirmed that the fluid flow morphology through the microposts designed within the channel remained largely unchanged regardless of the post spacing and longitudinal length conditions, maintaining high mixing efficiency (>0.95) under all conditions.

[0123] [Table 10]

[0124] On the other hand, as shown in Figure 14 and Table 10, in the case of the occlusion ratio, which is the ratio of the lateral spacing of microposts in the channel, it was confirmed that the mixing efficiency increases as the value increases, and when the occlusion ratio is 0.5, the mixing efficiency is maintained at a constant level of 0.98 or higher.

[0125] Furthermore, Figure 15 shows the mixing efficiency based on the width spacing before and after the fluid passage. Figure 16 shows the shear ratio when the front and rear width spacings are the same. The narrower the width spacing through which the fluid passes, the higher the shear ratio. Since the shear ratio is proportional to the shear stress, this can be said to result in higher shear stress.

[0126] In other words, as the flow occlusion rate increases, the channel width also narrows, and consequently, the shear stress increases proportionally. High shear stress, when it exceeds a certain value as previously calculated, can cause aggregation and deformation of organic matter present in the fluid (Bekard, I., et al. The Effects of Shear Flow on Protein Structure and Function. Biopolymers, 95, 11, 733-745 (2011)). When the flow velocity is increased to Re300 under conditions where the occlusion rate exceeds 0.65, the shear stress becomes 1000 dyne / cm². 2 When considering the above increase, the channel flow occlusion rate may be 0.2 to 0.8, and preferably in the range of 0.35 to 0.65, it can be seen that high mixing efficiency is not greatly affected by shear stress.

[0127] [Example 7] Design of equipment for mass production of lipid nanoparticles In this invention, we designed an apparatus capable of mass-producing nanoparticles containing LNPs. As shown in Figure 17, with the microfluidic apparatus in Figures 1 and 2 set as the base ×1, the mixing efficiency, flow velocity, and shear rate based on the Reynolds number were analyzed using hydrodynamic techniques when the two-dimensional size of the apparatus, i.e., the starting interval of microposts, the spacing between microposts, the vertical length of microposts, and the width spacing before and after the fluid flow, were increased by 2x (×2), 3x (×3), and 4x (×4). When the apparatus was enlarged, the channel height was maintained at 200 μm and the flow blockage ratio at 0.5.

[0128] As shown in Figure 18, the standard microfluidic apparatus in Figure 2 can be confirmed to have a mixing efficiency of 0.95 or higher when the Reynolds number is between 50 and 300. When the size of the apparatus is doubled (×2), the mixing efficiency decreases to 0.84 when the Reynolds number is 50. However, when the Reynolds number is 100 or higher, it was confirmed that the mixing efficiency remains above 0.95 up to a Reynolds number of 300. When the size of the apparatus is tripled (×3), it was confirmed that the mixing efficiency remains above 0.95 when the Reynolds number is between 150 and 300. When the size of the apparatus is quadrupled (×4), it was confirmed that the mixing efficiency remains above 0.95 when the Reynolds number is between 200 and 300.

[0129] As shown in Figure 19, it can be confirmed that to maintain the same Reynolds number as the size of the apparatus increases, only the flow velocity of the apparatus needs to be increased. Since an increase in flow velocity means an increase in the amount of nanoparticles that can be synthesized per unit time, it can be seen that nanoparticles can be mass-produced by two-dimensionally expanding the microfluidic apparatus shown in Figures 1 and 2.

[0130] As shown in Figure 20, it can be seen that the shear ratio decreases as the size of the apparatus increases. In other words, since the shear ratio is proportional to the shear stress, it can be said that the shear stress decreases as the size of the apparatus increases. This indicates that increasing the size of the apparatus does not lead to the aggregation and deformation of organic substances (in the case of LNPs, nucleic acids) present in the fluid.

[0131] The microfluidic apparatus of the present invention can be shown to be capable of mass production of LNPs containing nucleic acids without causing aggregation and deformation of organic substances present in the fluid, simply by increasing its size in two dimensions.

[0132] [Example 8] Optimization of lipid nanoparticle synthesis In this invention, as shown in Figure 1, lipid nanoparticles (LNPs) were synthesized using a microfluidic apparatus. Of the three inflow channels, the central channel was injected with a lipid mixture, and the two channels on either side were injected with aqueous solutions. The lipid mixture and aqueous solution were efficiently mixed by microposts inside the apparatus, and the synthesized LNPs could finally be obtained through the outflow channel of the micromixing channel apparatus.

[0133] As a lipid mixture for LNP synthesis, ionizable lipids, helper lipids, PEGylated lipids, and sterols can be used. Specifically, the present invention uses SM-102 [9-heptadecanyl 8-{(2-hydroxyethyl)[6-oxo-6-(undecyloxy)hexyl]amino}octanoate], DSPC (distearoylphosphatidylcholine), POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine), DMG-PEG2k [1,2-dimyristoyl-rac-glycerol-methoxypolyethylene glycol-2000], and cholesterol.

[0134] Furthermore, an aqueous solution with a pH of 3.0 to 6.0 is required for LNP synthesis, and in this invention, a citrate buffer with a pH of 3.0 was used.

[0135] 8-1 [Synthesis of lipid nanoparticles by flow rate] Lipid mixtures were prepared using SM-102, DSPC / POPC, cholesterol, and DMS-PEG2k, with composition ratios set to 50:10:38.5:1.5 (molar ratio), 28:5.7:65.5:0.8, or 20:4:75.5:0.5, respectively. LNP synthesis was performed while changing the Reynolds number to 12.5, 25, 50, 100, and 200 to compare LNP size based on flow rate. The three ratios of the lipid mixtures were set to increase the molar ratio of cholesterol in the lipid mixture, based on the commonly used ratio of 50:10:38.5:1.5.

[0136] The synthesized LNPs were analyzed using dynamic light scattering (DLS) with a Zetasizer Pro Blue instrument to determine their hydrodynamic size relative to a number-weighted size distribution, and their degree of uniformity was indicated by the polydispersity index (PDI).

[0137] [Table 11]

[0138] LNPs were synthesized by fixing the molar ratio of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k to 50:10:38.5:1.5 and changing the Reynolds number to 12.5, 25, 50, 100, and 200. As shown in Table 11 and Figures 21, 22, and 23, the sizes of the LNPs were measured to be 83, 66, 64, 67, and 58 nm, respectively, and the polydispersity index (PDI) of all LNPs was measured to be 0.15 or less, confirming that very uniform LNPs were synthesized overall. The PDI value considered stable for LNPs may vary depending on the circumstances, but the FDA guidelines recommend that the PDI of LNPs used as drugs be 0.3 or less.

[0139] [Table 12]

[0140] The molar ratio of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k was fixed at 28:5.7:65.5:0.8 (a condition in which only the ratio of cholesterol was increased compared to Table 1), and LNPs were synthesized by changing the Reynolds number to 12.5, 25, 50, 100, and 200. As shown in Table 12 and Figures 24, 25, and 26, the sizes of the LNPs were measured to be 95, 94, 102, 84, and 95 nm, respectively, and the PDI of all LNPs was measured to be 0.1 or less, confirming that very uniform LNPs were synthesized.

[0141] [Table 13]

[0142] The molar ratio of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k was fixed at 20:4:75.5:0.5 (a condition in which only the ratio of cholesterol was increased compared to Table 12), and LNPs were synthesized by changing the Reynolds number to 12.5, 25, 50, 100, and 200. As shown in Table 13 and Figures 27, 28, and 29, the sizes of the LNPs were measured to be 94, 83, 93, 76, and 86 nm, respectively, and the PDI of all LNPs was measured to be 0.1 or less, confirming that very uniform LNPs were synthesized.

[0143] Specifically, when LNPs were synthesized using the microfluidic apparatus shown in Figure 1, it was confirmed that highly uniform LNPs (with a PDI value not higher than 0.1, except for Reynolds number 12.5 in Table 11) were synthesized regardless of flow rate, across a variety of lipid composition ratios. However, it was confirmed that there was no significant change in LNP size with respect to flow rate.

[0144] 8-2 [Synthesis of lipid nanoparticles based on the composition ratio of lipid mixtures] To alter the composition ratio of a lipid mixture consisting of SM-102, DSPC / POPC, cholesterol, and DMG-PEG2k, LNPs were synthesized by changing the molar ratio of cholesterol in the lipid mixture to 38.5, 55.5, 65.5, 75.5, 86.0, and 98.45. During this process, the composition ratios of the lipid mixture (ionized lipids, helper lipids, and PEGylated lipids) excluding cholesterol were kept constant.

[0145] [Table 14]

[0146] Table 14 shows the composition ratios of the overall lipid mixture obtained by changing the molar ratio of cholesterol. Using the composition ratio of each lipid mixture, LNPs were synthesized using a microfluidic apparatus as shown in Figure 1. Based on the results of Example 8-1, a Reynolds number of 50 was selected, which resulted in the smallest PDI, i.e., the synthesis of the most uniform LNPs.

[0147] [Table 15]

[0148] As shown in Table 14 above, Table 15 shows the LNP size and PDI results when LNPs were synthesized by adjusting the composition ratio of the lipid mixture during LNP synthesis. As shown in Figure 30 and Table 15, the sizes of the synthesized LNPs with cholesterol molar ratios of 38.5, 55.5, 65.5, 75.5, 86.0, and 98.45 in the lipid mixture were measured to be 64, 84, 102, 93, 103, and 105 nm.

[0149] The uniformity of LNPs synthesized via microfluidic apparatus was confirmed by PDI. PDI was measured to be 0.2 or less up to a cholesterol molar ratio of 86.0, confirming highly uniform synthesis. The uniformity of the synthesized LNPs can also be confirmed by the narrow appearance of a single peak in the number-weighted size distribution results (Figures 31 and 32), the volume-weighted size distribution results (Figures 33 and 34), and the intensity-weighted size distribution results (Figures 35 and 36).

[0150] When using the aforementioned microfluidic apparatus, it can be seen that LNPs are synthesized with uniform size even when the molar ratio of cholesterol in a commonly used lipid mixture composition is 86.0, which is more than twice the molar ratio of 38.5, meaning that 80% of the total lipid mixture composition is cholesterol. Furthermore, when using the aforementioned microfluidic apparatus, it can be confirmed that LNPs are synthesized even in an extreme lipid mixture composition where the molar ratio of cholesterol is 98.45. However, in this case, the PDI was 0.274, indicating inferior monodispersity compared to other conditions. The reason for the slightly higher PDI is that the excess cholesterol caused LNP aggregation.

[0151] Furthermore, under conditions where the cholesterol molar ratio was 60 or less, the LNP size tended to increase as the proportion of cholesterol increased, but no change in LNP size was observed under conditions where the cholesterol molar ratio was 65.5.

[0152] 8-3 [Optimization of lipid nanoparticle synthesis by size] As can be seen from the results of Examples 8-1 and 8-2, the size of LNPs synthesized using the microfluidic apparatus is 60 nm to 100 nm. To synthesize LNPs smaller than 50 nm, the molar ratio of PEGylated lipids in the lipid mixture was increased. At this time, the composition ratio of the lipid mixture SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k was set to 48.5:9.5:37.5:4.5, and the Reynolds numbers were set to 50 and 100 to confirm whether a change in LNP size occurs with respect to the Reynolds number.

[0153] [Table 16]

[0154] As shown in Table 16, Figure 37, and Figure 38, when the Reynolds number is 50 and 100, the LNPs are synthesized at 35 and 36 nm, respectively, and the PDI is around 0.2, confirming that they were synthesized relatively uniformly.

[0155] [Table 17]

[0156] When summarizing the results of the final synthesized LNPs, the conditions for Reynolds number and lipid mixture composition ratios required to synthesize highly uniform LNPs of specific sizes of 30 nm, 60 nm, and 90 nm using the microfluidic apparatus shown in Figure 1 are shown in Table 17.

[0157] Generally, the average diameter of LNPs used as mRNA vaccines falls within the 80-100 nm range. Smaller LNPs (~60 nm) impart high curvature stress to the particles, making them difficult to manufacture and resulting in lower particle stability and poor drug loading efficiency (intrinsicity). Conversely, smaller LNP sizes tend to increase the particle's surface area, leading to improved drug delivery efficiency to target organs.

[0158] Furthermore, the size of LNPs also affects the target properties of the particles for various organs in the body. Generally, it is known that for organs such as the liver, which contain many microvessels, the target properties of the particles for those organs decrease as the size of the LNPs increases. In addition, there are research results showing that small LNPs less than 60 nm (especially in the 30 nm range) have excellent target properties for dendritic cells in lymph nodes and many other immune cells.

[0159] For the reasons mentioned above, it is crucial to set an appropriate particle size according to the intended use of LNPs and to have a manufacturing method to achieve this. The present invention provides a method for producing LNPs of a suitable size by adjusting the Reynolds number and particle composition ratio using a microfluidic apparatus.

[0160] [Example 9] Optimization of the synthesis of mRNA-encapsulated lipid nanoparticles of different sizes. Lipid nanoparticles (mRNA LNPs) encapsulating messenger ribonucleic acid (mRNA) were synthesized using the same microfluidic apparatus as in Example 8. The synthesis was carried out in the same manner as in Example 8, and the mRNA was prepared in a pH 3.0 citrate buffer so that the aqueous solution contained mRNA in an N / P ratio of 6.

[0161] mRNA LNPs were synthesized using the size-specific LNP synthesis conditions set out in Table 17 of Example 8-3. The mRNAs used were mCherry mRNA with a length of 996 nucleotides and Cas9 mRNA with a length of 4521 nucleotides. mCherry LNPs and Cas9 LNPs were synthesized in different sizes using two mRNAs with a length difference.

[0162] [Table 18]

[0163] As shown in Table 18 and Figures 39, 40, and 41, the sizes of mCherry LNPs synthesized targeting 30, 60, and 90 nm were measured to be 40, 57, and 89 nm, respectively. The PDI values ​​at this time were 0.105, 0.054, and 0.052, respectively, indicating very uniform synthesis. Furthermore, the similarity in size distribution between mCherry LNPs and bare LNPs (LNPs without mRNA) can be confirmed. The uniformity of mCherry LNPs by size can also be confirmed from the number-weighted size distribution results in Figures 42 and 43.

[0164] [Table 19]

[0165] As shown in Table 19 and Figures 44, 45, 46, and 47, the sizes of mCherry LNPs synthesized targeting 30, 60, and 90 nm were measured to be 37, 57, and 90 nm, respectively. The PDI values ​​at these times were 0.190, 0.036, and 0.014, respectively. It was found that the Cas9 LNPs targeting 60 nm and 90 nm were synthesized very uniformly, and the Cas9 LNPs targeting 30 nm also showed that nanoparticle uniformity was still ensured, as their PDI was less than 0.2. Furthermore, the similarity in the size distribution between Cas9 LNP and Bare LNP can be confirmed. The uniformity of Cas9 LNPs by size can also be confirmed from the number-weighted size distribution results in Figures 48 and 49.

[0166] Nucleic acid encapsulation efficiency (EE) is calculated using the total mRNA detected based on fluorescence intensity in each mRNA LNP, and the value obtained by detecting unencapsulated mRNA (free mRNA), using the following formula:

number

number

[0167] In short, nucleic acid encapsulation rate (EE) is an indicator of the quality (mRNA content) of the final product, while nucleic acid endogenous rate (LE) is an indicator of the efficiency (or yield) of mRNA LNPs being loaded from the raw material.

[0168] As shown in Table 18 and Figure 50, the nucleic acid encapsulation rates for 30, 60, and 90 nm mCherry LNPs were 84%, 95%, and 100%, respectively. As shown in Table 19 and Figure 51, the nucleic acid encapsulation rates for 30, 60, and 90 nm Cas9 LNPs were 74%, 98%, and 100%, respectively. Regardless of the mRNA type, the nucleic acid encapsulation rate for 60 nm or 90 nm mRNA LNPs was over 95%, which means that the lipid mixture and mRNA are mixed very efficiently by the microfluidic device. The nucleic acid encapsulation rate for 30 nm mCherry LNPs, which have a small space for encapsulating nucleic acids, was 84%, and the nucleic acid encapsulation rate for 30 nm Cas9 LNPs was 74%, which are still relatively high. This means that the lipid mixture and mRNA are mixed efficiently by the microfluidic device. A high nucleic acid encapsulation rate can be said to mean that the probability of impurity problems caused by unencapsulated mRNA is very low.

[0169] As can be seen from Table 18, the nucleic acid endogenous concentrations of mCherry LNPs at 30, 60, and 90 nm are 38%, 39%, and 44%, respectively. As can be seen from Table 19, the nucleic acid endogenous concentrations of Cas9 LNPs at 30, 60, and 90 nm are 44%, 29%, and 37%, respectively. The fact that nucleic acid endogenous concentrations of over 30% are observed regardless of target size or mRNA type means that the lipid mixture and mRNA are efficiently mixed by the microfluidic device. Furthermore, a high nucleic acid endogenous concentration means that there is less raw material loss.

[0170] In other words, the microfluidic apparatus of the present invention is capable of synthesizing mRNA LNPs of a specific size with high uniformity, regardless of mRNA length, and can be said to be an excellent synthesis platform capable of developing highly efficient LNP-based therapeutic drugs or vaccines that uniformly possess stability and functionality against a variety of diseases.

[0171] [Example 10] Optimization of the synthesis of size-specific lipid nanoparticles encapsulated with oligonucleotides. Oligonucleotide-encapsulated lipid nanoparticles (Oligo LNPs) were synthesized using the same microfluidic apparatus as in Example 8 and Example 9. The synthesis was carried out in the same manner as in Example 8, using the conditions shown in Table 17. Oligonucleotides were prepared in an aqueous solution in a citrate buffer at pH 3.0 so that the N / P ratio was 6.

[0172] Oligo LNPs were synthesized using the size-specific LNP synthesis conditions set out in Example 8-3. The oligonucleotide used in this case was DNA composed of only 20 (20mer) thymine (T) molecules.

[0173] [Table 20]

[0174] As shown in Table 20 and FIGS. 52, 53 and 54, the sizes of the Oligo LNPs synthesized targeting 30, 60, 90 nm were measured to be 50, 99, 132 nm respectively, and it was confirmed that they were synthesized to a size 20 nm to 40 nm larger than the target size. The PDI at this time was 0.029, 0.053, 0.036 respectively. Although synthesized to be larger than the target size, it can be seen that all sizes were synthesized very uniformly even when the oligonucleotide was encapsulated in the LNP. Also, the uniformity of the Oligo LNPs by size can be confirmed from the number-weighted size distribution results in FIGS. 55 and 56.

[0175] After detecting the total oligonucleotide and the free oligonucleotide based on the fluorescence intensity in Olilgo LNP, the nucleic acid encapsulation rate and the nucleic acid incorporation rate of the oligonucleotide were calculated using the obtained values. The calculation formula for the nucleic acid encapsulation rate is

Number

Number

[0176] As shown in Table 20 and FIG. 57, the nucleic acid encapsulation rates of the 50, 100, 130 nm Oligo LNPs are 90, 89, 91% respectively, and their nucleic acid incorporation rates are 53, 48, 47% respectively. Through the nucleic acid encapsulation rate of 90%, it can be seen that the probability of occurrence of impurity problems due to the unencapsulated oligonucleotide is low, and through the nucleic acid incorporation rate of 45% or more, it can be seen that the raw material loss is small. That is, it can be confirmed that the lipid mixture and the oligonucleotide are efficiently mixed by the microfluidic device.

[0177] Unlike mRNA LNPs, the synthesis of Oligo LNPs larger than the target size may be due to the different nucleic acids that are endogenous substances. Oligonucleotides are composed of 20 nucleotides, while mRNA is composed of 1000 to 5000 nucleotides, showing a length difference of 50 to 250 times. Comparing the nucleic acid encapsulation rates among target sizes of 30, 60, and 90 nm in Table 18, Table 19, and Table 20, it can be seen that Oligo LNPs are up to 15%, 20%, and 10% higher than mRNA LNPs. This seems to indicate that the size increases as the amount of the shorter oligo nucleotides encapsulated in LNPs increases.

[0178] That is, the microfluidic device in the present invention is a device capable of synthesizing Oligo LNPs of a specific size with high uniformity, and can be said to be an excellent synthetic platform for developing highly efficient therapeutic agents or vaccines based on LNPs with uniform stability and functionality against various diseases.

[0179] [Example 11] Confirmation of intracellular protein expression by size - specific mCherry LNP treatment To verify the LNP - based mRNA delivery ability in cells, the degree of protein expression in cells treated with mRNA LNPs was confirmed by fluorescence images using a confocal microscope. As cells for confirming the protein expression of mRNA, U87MG, a human glioblastoma, HepG2, a human hepatoma cell line, and THP - 1, a human monocytic cell line, were used. For mRNA, mCherry mRNA that can emit fluorescence was used to confirm the presence or absence of protein expression of intracellular mRNA. mCherry LNPs were synthesized according to the target size (30, 60, 90 nm) as in Example 9.

[0180] In human glioblastoma U87MG cells, the degree of mCherry protein expression was observed based on mCherry LNP size. U87MG cells were treated with 200 ng of mRNA of mCherry LNPs of different sizes (30, 60, and 90 nm) and then cultured for 24 hours under 5% CO2 conditions at 37°C. After removing any remaining mCherry LNPs from the culture medium, the cells were cultured for another 24 hours. The culture medium was then changed to confirm mCherry protein expression, and the cell nucleus and cytoskeleton were stained. After staining, the U87MG cells were fixed by treating them with 1 mL of 4% paraformaldehyde solution for 15 minutes. The fluorescence intensity of mCherry protein expressed in each fixed cell was recorded using a confocal microscope with a filter containing an excitation wavelength of 587 nm and an emission wavelength of 610 nm.

[0181] As shown in Figure 58, the expression of mCherry protein in U87MG cells can be confirmed by fluorescence imaging. The expression of mCherry protein in the intracellular cytoplasm was confirmed by the overlap of the fluorescence staining the cytoskeleton (green) and the fluorescence of mCherry (red). When U87MG cells were treated with three different sizes of mCherry LNPs at equal doses of 200 ng, it was confirmed that mCherry protein expression increased with increasing LNP size. As shown in the quantitative results in Figure 59, mCherry protein expression was 4 times and 14 times higher when treated with 60 nm and 90 nm mCherry LNPs than when treated with the smallest size 30 nm mCherry LNPs, respectively. When treated with the largest size 90 nm mCherry LNPs, mCherry protein expression was 3 times higher than when treated with the intermediate size 60 nm mCherry LNPs.

[0182] In the human liver cancer cell line HepG2, the degree of protein expression at different mRNA LNP concentrations was used to confirm the degree of mCherry LNP. Two conditions were prepared by synthesizing mCherry LNPs of sizes 60 and 90 nm. HepG2 cells were treated with 20, 200, and 500 ng of mCherry LNP at 60 and 90 nm, respectively, and then cultured for 24 hours under conditions of 5% CO2 and 37°C. After removing any remaining mCherry LNP from the culture medium, the cells were cultured for another 24 hours. After changing the culture medium to confirm mCherry protein expression, the cell nucleus and cytoskeleton were stained. After staining, the HepG2 cells were fixed by treating them with 1 mL of 4% paraformaldehyde solution for 15 minutes. The fluorescence intensity of the mCherry protein expressed in each fixed cell was imaged using a confocal microscope with a filter containing an excitation wavelength of 587 nm and an emission wavelength of 610 nm.

[0183] The degree of mCherry protein expression in HepG2 cells after treatment with mCherry mRNA at different concentrations was confirmed from the fluorescence images in Figure 60. When mCherry LNPs were treated in HepG2 cells, the expression of mCherry protein in the cytoplasmic region of the cell was confirmed by the overlap of the fluorescence staining the cytoskeleton (green) and the fluorescence of mCherry (red). When mCherry LNPs of different sizes were treated in HepG2 cells, it was confirmed that the amount of mCherry protein expressed in HepG2 cells increased proportionally to the amount of treated mCherry mRNA (20, 200, 500 ng), regardless of the size of the mCherry LNP (60 nm or 90 nm). Furthermore, as can be seen from the quantitative results in Figure 61, it was confirmed that when the same amount of mCherry mRNA was treated, the mCherry protein expression rate was higher with larger LNP sizes.

[0184] In the human mononuclear cell line THP-1, the degree of protein expression at different mRNA LNP concentrations was examined. mCherry LNP was synthesized under two conditions: 60 nm and 90 nm. THP-1 cells were treated with 20, 200, and 500 ng of mCherry LNP at 60 nm and 90 nm, respectively, and then cultured for 24 hours under 5% CO2 conditions at 37°C. After removing any remaining mCherry LNP from the culture medium, the cells were cultured for another 24 hours. The culture medium was then changed to confirm mCherry protein expression, and the cell nucleus and cytoskeleton were stained. After staining, the THP-1 cells were fixed by treating them with 1 mL of 4% paraformaldehyde solution for 15 minutes. The fluorescence intensity of the mCherry protein expressed in each fixed cell was recorded via a confocal microscope using a filter containing an excitation wavelength of 587 nm and an emission wavelength of 610 nm.

[0185] As shown in Figure 62, the degree of mCherry protein expression in THP-1 cells after treatment with mCherry mRNA at different concentrations was confirmed from fluorescence images. The expression of mCherry protein in the cytoplasmic region of THP-1 cells after treatment with mCherry LNP was confirmed by the overlap of the fluorescence staining the cytoskeleton (green) and the fluorescence of mCherry (red).

[0186] When 60nm mCherry LNPs were treated with THP-1, we confirmed that the amount of mCherry protein expressed within THP-1 increased proportionally to the amount of treated mCherry mRNA (20, 200, 500 ng). However, when 90nm mCherry LNPs were treated with THP-1, we confirmed that the amount of treated mCherry mRNA (20, 200, 500 ng) did not correlate with the amount of mCherry protein expressed within THP-1.

[0187] Furthermore, when comparing the degree of mCherry expression based on mCherry LNP size (60 nm or 90 nm) after processing with the same amount of mCherry mRNA, the fluorescence image in Figure 62 and the quantitative results in Figure 63 confirmed that 60 nm mCherry LNPs showed even higher mCherry expression than 90 nm mCherry LNPs.

[0188] In this invention, we confirmed that in both epithelial cells (HepG2) and epithelial-like cells (U87MG), larger mCherry LNP size is associated with higher mCherry protein expression. This may be because, while the PEGylated lipid content (mol%) increases as the LNP size decreases, the increased PEG content in the LNP delays intracellular uptake or endosomal escape of the LNP.

[0189] Furthermore, in human immune cells such as THP-1, unlike epithelial (or epithelial-like) cells, we confirmed that mCherry protein expression was even higher when treated with relatively smaller 60nm mCherry LNPs, rather than the larger 90nm mCherry LNPs. This result suggests that there is an LNP size that allows for efficient nucleic acid delivery depending on the type of cell being targeted.

[0190] [Example 12] Verification of the stability of endogenous LNPs in nucleic acids To verify the stability of nucleic acid-endogenous LNPs, the size-specific LNP synthesis conditions set out in Table 17 of Example 8-3 were applied using the microfluidic apparatus used in Examples 8, 9, and 10. Nucleic acid-endogenous LNPs were synthesized by preparing an aqueous solution containing nucleic acid (mRNA or oligonucleotide) in a pH 3.0 citrate buffer with an N / P ratio of 6.

[0191] To confirm the storage stability of mCherry LNPs and Cas9 LNPs, two sizes were synthesized and compared for each mRNA type (mCherry mRNA and Cas9 mRNA). mCherry LNPs were synthesized at 60 and 90 nm, and Cas9 LNPs at 30 and 90 nm. The synthesized mCherry LNPs and Cas9 LNPs were stored in PBS containing 1% trehalose, and their storage stability for 0, 7, 14, and 28 days under refrigerated conditions at 4°C was confirmed using DLS results (average size, size at the highest peak in the number-weighted size distribution, percentage at the highest peak in the volume-weighted size distribution, PDI), nucleic acid inclusion rate, and nucleic acid endogenous rate.

[0192] As shown in Figures 64 and 65, the average size and the highest peak size in the number-weighted size distribution of 60 nm mCherry LNPs remained largely unchanged for 28 days under refrigerated conditions at 4°C. Therefore, the highest peak in the volume-weighted size distribution, which represents the fraction of the major size within the nanoparticle size distribution, remained largely unchanged at around 20% in volume%, and the PDI, which indicates uniformity, also remained within 0.1, confirming that it was very uniform even after 28 days. The nucleic acid encapsulation rate decreased from 100% on day 0 to 82% on day 7, and further decreased to 73% on day 28. The nucleic acid inclusion rate remained around 20% for 28 days.

[0193] As shown in Figures 66 and 67, 90 nm mCherry LNPs showed similar results in size to 60 nm mCherry LNPs under refrigerated conditions at 4°C. The average size and the highest peak size in the number-weighted size distribution remained largely unchanged for 28 days. Therefore, the volume-weighted size distribution, which represents the fraction of the major size within the nanoparticle size distribution, also remained largely unchanged at 15%-20%, and the PDI, indicating uniformity, remained within 0.1, confirming very high uniformity even after 28 days. The nucleic acid encapsulation rate showed a different pattern from 60 nm mCherry LNPs, maintaining around 80% for 28 days, while the nucleic acid inclusion rate remained around 40% for 28 days.

[0194] As shown in Figures 68 and 69, the average size and the highest peak size in the number-weighted size distribution of 30 nm Cas9 LNPs remained largely unchanged for 28 days under refrigerated conditions at 4°C. Therefore, the volume-weighted size distribution, which represents the fraction of the major size within the nanoparticle size distribution, also maintained a volume of around 15%. The PDI, indicating uniformity, was measured at 0.22. Subsequently, the PDI remained in the 0.2 range without significant change for 28 days, confirming the maintenance of uniformity. Generally, a PDI of 0.7 or higher is considered indicative of poor nanoparticle uniformity; therefore, a PDI of 0.2 indicates that the nanoparticles are still uniform. The nucleic acid inclusion rate remained at 70% to 80% for 28 days despite the small size of the LNPs, and the nucleic acid inclusion rate remained at around 40% for 28 days.

[0195] As can be seen from Figures 70 and 71, the average size and the highest peak size in the number-weighted size distribution of 90 nm Cas9 LNPs remained largely unchanged for 28 days under refrigerated conditions at 4°C. Therefore, the volume-weighted size distribution, which represents the fraction of the major size within the nanoparticle size distribution, also remained largely unchanged at 15% to 20%, and the PDI, which indicates uniformity, remained around 0.1, confirming that it was very uniform even after 28 days. The nucleic acid encapsulation rate was confirmed to remain around 80% for 28 days, and the nucleic acid inclusion rate remained around 30% for 28 days.

[0196] When mRNA LNPs are synthesized using the microfluidic apparatus of the present invention, size stability and LNP uniformity are maintained for 28 days under refrigerated conditions, regardless of mRNA length or LNP size. However, the nucleic acid encapsulation rate decreases in one of the four types of mRNA LNPs (a 60nm mCherry LNP), indicating that optimization of storage conditions is necessary.

[0197] Oligo LNPs were synthesized in three sizes using the size-specific LNP synthesis conditions set out in Table 17 of Example 8-3. Similar to Example 10, the synthesized Oligo LNPs were larger than the target size. The sizes of Oligo LNPs synthesized targeting 30, 60, and 90 nm were measured to be 50, 100, and 130 nm, respectively, resulting in sizes 20 nm to 40 nm larger than the target size. The synthesized Oligo LNPs were stored in PBS containing 1% trehalose, and their storage stability for 0, 7, 14, and 28 days under refrigerated conditions at 4°C was confirmed by DLS results (average size, size at the highest peak in the number-weighted size distribution, percentage at the highest peak in the volume-weighted size distribution (volume%), PDI), nucleic acid inclusion rate, and nucleic acid endogenous rate.

[0198] As shown in FIGS. 72 and 73, it was confirmed that the 50-nm Oligo LNP (target size: 30 nm) maintained the size at the highest peak of the average size and the number-weighted size distribution without significant change for 28 days under refrigerated conditions at 4°C. Therefore, the percentage (volume%) at the highest peak of the volume-weighted size distribution, which represents the fraction of the main size in the size distribution within the nanoparticles, also maintained 20%, and the PDI indicating uniformity also remained around 0.1 and was very uniform even after 28 days. The nucleic acid encapsulation rate was confirmed to maintain 80% or more for 28 days, and the nucleic acid incorporation rate maintained around 50% for 28 days.

[0199] As shown in FIGS. 74 and 75, it was confirmed that the 100-nm Oligo LNP (target size: 60 nm) maintained the size at the highest peak of the average size and the number-weighted size distribution without significant change for 28 days under refrigerated conditions at 4°C. Therefore, the percentage (volume%) at the highest peak of the volume-weighted size distribution, which represents the fraction of the main size in the size distribution within the nanoparticles, also maintained 20%, and the PDI indicating uniformity also remained around 0.1 and was very uniform even after 28 days. The nucleic acid encapsulation rate was confirmed to maintain 80% or more for 28 days, and the nucleic acid incorporation rate maintained around 50% for 28 days.

[0200] As shown in Figures 76 and 77, we confirmed that the average size and the highest peak size in the number-weighted size distribution of 130 nm Oligo LNP (target size: 90 nm) remained largely unchanged for 28 days under refrigerated conditions at 4°C. Therefore, the volume-weighted size distribution, which represents the fraction of the major size within the nanoparticle size distribution, also remained at 15% to 20%, and the PDI, which indicates uniformity, remained around 0.1, confirming that it was very uniform even after 28 days. The nucleic acid encapsulation rate decreased slightly from 89% on day 0 to 82% on day 7, but was confirmed to remain at 80% on day 28. The nucleic acid inclusion rate remained at around 50% for 28 days.

[0201] When oligo LNPs are synthesized using the microfluidic apparatus of the present invention, size stability and uniformity of the LNPs are maintained for 28 days under refrigerated conditions, regardless of LNP size. Significant changes in nucleic acid encapsulation rate and nucleic acid endogenous rate were also confirmed.

[0202] Based on the results of the above-described examples, synthesizing size-specific LNPs using a microfluidic apparatus under three optimized conditions allows for the highly uniform synthesis of LNPs of desired sizes with a high mixing efficiency of 0.95 or higher, enabling the encapsulation of nucleic acids of various lengths with high nucleic acid encapsulation and intrinsic nucleic acid rates. Therefore, the optimized LNP synthesis technology in the present invention is considered to enable the selection and development of LNPs of suitable sizes with high uniformity according to the diverse indications to be treated or prevented.

Claims

1. Including inflow channels, mixing channels and outflow channels, The aforementioned inflow channels are two or more, The aforementioned mixed channel includes a micropost, The fluids flowing into different inflow channels are made to flow in different directions. The lipid mixture flows into one inflow channel, and the nucleic acids flow into the other inflow channel. The lipid mixture comprises sterols and further comprises at least one selected from the group consisting of ionized lipids, helper lipids, and PEG-modified lipids. The microposts are arranged in one or more rows in the direction of fluid flow, and in one row, one or more microposts are arranged in a direction different from the direction of fluid flow. The aforementioned plurality of microposts are not arranged in a single line in the direction of fluid flow, but are arranged so as to partially or completely cover the gaps between microposts arranged in adjacent rows when viewed from the direction of fluid flow. The Reynolds number for the fluid flow in the mixing channel is 25 to 100. A microfluidic apparatus for producing lipid nanoparticles, wherein the molar ratio of sterols to the lipid mixture is 0.3 to 0.

86.

2. The microfluidic apparatus for producing lipid nanoparticles according to claim 1, characterized in that the lipid mixture flows in in the same direction as the fluid flow, and the nucleic acids flow in in a direction different from the fluid flow.

3. The microfluidic apparatus for producing lipid nanoparticles according to claim 2, characterized in that the lipid mixture flows in in the same direction as the fluid flow, and the nucleic acids flow in in a direction perpendicular to the fluid flow.

4. The microfluidic apparatus for producing lipid nanoparticles according to claim 1, characterized in that the microposts are connected to or not connected to the walls constituting the mixing channel.

5. The microfluidic apparatus for producing lipid nanoparticles according to claim 1, characterized in that the size of the lipid nanoparticles is adjusted by the molar ratio of the lipid mixture to the sterol.

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