Microfluidic device for preparing lipid nanoparticles of various sizes, and preparation method using same

Abstract

The present invention relates to a microfluidic device for preparing lipid nanoparticles that can deliver nucleic acids, and a lipid nanoparticle preparation method using same. By using the microfluidic device, lipid nanoparticles of a desired size can be prepared by adjusting the molar ratio between compositions and the Reynolds number.

Description

Microfluidic device for manufacturing lipid nanoparticles of various sizes and manufacturing method using the same

[0001] The present invention relates to a microfluidic device 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 controlling the composition ratio of sterols and constituent components constituting the lipid nanoparticles and the Reynolds number, and to a microfluidic device capable of efficient mixing in the production of lipid nanoparticles, thereby enabling efficient mass production of lipid nanoparticles of various sizes.

[0002]

[0003] The gene therapy market has recently been growing. Gene therapy is a medicine that delivers genes to the affected area, treating or preventing disease through the production of proteins produced from those genes. From Pfizer and Moderna's COVID-19 vaccines to Biogen and Ionis' RNA interference (RNAi)-based treatment for Lou Gehrig's disease, Qalsody, gene therapies with various mechanisms are currently undergoing FDA approval or clinical trials.

[0004] For these gene therapies, it's crucial to accurately deliver genes susceptible to destruction within the human body to the lesion site. Various studies are being conducted to improve delivery efficiency within the body. Gene therapies using various vectors are being studied. Furthermore, since vectors alone are still highly susceptible to destruction within the body, lipid nanoparticles (LNPs) are being actively researched to safely protect and deliver these genes.

[0005] Lipid nanoparticles (LNPs) are nanoparticles composed of lipids such as phospholipids and cholesterol. They protect carriers from destruction in the human body and neutralize the carrier's charge to increase cell membrane penetration efficiency. They are also effective for drug delivery in modalities other than genes, and many pharmaceutical companies are conducting research on new lipid nanoparticles. However, the synthesis and manufacturing of these lipid nanoparticles poses a challenge. This is because it is difficult to control the spontaneous aggregation and mutual association of phospholipids and cholesterol, which can lead to the formation of nanoparticles of various sizes, such as micelles and liposomes.

[0006] Cholesterol in lipid nanoparticles is known to have various effects on phospholipid-based membranes. In general, the higher the cholesterol content in a membrane, the lower the membrane fluidity and the stronger the bonds between phospholipids. Researchers have confirmed that lipid nanoparticles with a high cholesterol content for small molecule drug delivery have improved drug loading efficiency and particle stability. For this reason, lipid nanoparticles for nucleic acid delivery have typically incorporated cholesterol as a component. However, as mentioned above, the inclusion of cholesterol complicates the manufacture of uniform lipid nanoparticles, and controlling their size while encapsulating a high level of gene therapy is quite challenging.

[0007] The COVID-19 vaccine is currently the most commercially available lipid nanoparticle vaccine. The messenger RNA (mRNA) used in this vaccine typically has a sequence length of several thousand bases. The mRNA typically used as a therapeutic is known to be 1,000 to 5,000 nt. On the other hand, antisense oligonucleotides (ASOs), which are tens of bases long, are a different type of gene therapy that utilizes RNA expression inhibition. These are also being developed as therapeutics using lipid nanoparticles. In technologies utilizing lipid nanoparticles as nucleic acid delivery vehicles, varying levels of leakage occur depending on the length of the nucleic acid sequence used. Therefore, the cholesterol composition ratio needs to be adjusted to match the length of the loaded nucleic acid. In other words, for nucleic acids with relatively long sequences (e.g., mRNA), the typical lipid nanoparticle composition of equal amounts of phospholipids and cholesterol results in an unnecessarily excessive cholesterol content. On the other hand, nucleic acids with relatively short base sequences (e.g., oligonucleotides) exhibit relatively high efflux from lipid nanoparticles, and therefore require higher cholesterol contents.

[0008] The present inventors have conducted various studies on the production of lipid nanoparticles having different composition ratios, and as a result, have devised a method for producing lipid nanoparticles having controlled molar ratios of lipid nanoparticle compositions and Reynolds numbers, and a method for producing the same with high efficiency in a separately designed expandable microfluidic device, and have elucidated the effects thereof, thereby completing the present invention.

[0009]

[0010] (Prior art literature)

[0011] (Patent Document)

[0012] (Patent Document 1) U.S. Registered Patent US8058069B2

[0013]

[0014] (Non-patent literature)

[0015] (Non-patent Document 1) Camilla, H., A. et al. The role of lipid components in lipid nanoparticles for vaccines and gene therapy. Adv. Drug Delivery. Rev., 188, 114416 (2022).

[0016] (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).

[0017]

[0018] As mentioned above, lipid nanoparticles have the problem that it is very difficult to control the particle size due to the various nanoparticle byproducts formed from various lipids.

[0019] The present invention aims to provide a molar ratio between the compositions of lipid nanoparticles for the purpose of nucleic acid delivery, and to provide a manufacturing method for controlling the particle size that varies according to the molar ratio and the Reynolds number.

[0020] The purpose of the present invention is to provide a manufacturing method that improves the efficiency of lipid nanoparticle production by performing the above manufacturing method on a separately designed microfluidic device.

[0021]

[0022] The present invention relates to a microfluidic device comprising an inlet channel, a mixing channel, and an outlet channel.

[0023] (a) a step of injecting a lipid mixture into an inlet channel;

[0024] (b) a step of injecting nucleic acid into an inlet channel other than the above inlet channel, and

[0025] (c) A method for producing lipid nanoparticles using a microfluidic device is provided, comprising a step of producing lipid nanoparticles by mixing a lipid mixture and nucleic acid in the above mixing channel.

[0026] The above mixing channel may include micropillars.

[0027] In the above microfluidic device, there may be two or more inlet channels, and the flow of fluids flowing into different inlet channels among the inlet channels may be in different directions.

[0028] The lipid mixture may include, but is not limited to, one or more selected from the group consisting of ionizable lipid, helper lipid, PEGylated lipid, and sterol.

[0029] The above ionized lipids are 9-heptadecanyl 8-{(2-hydroxyethyl)[6-oxo-6-(undecyclooxy)hexyl]amino}octanoate (SM-102), 1,2-dirinoleyloxy-N,N-dimethyl-3-aminopropane (DLinDMA), 2,2-dirinoleyl-4-dimethylaminoethyl-[1,3]-dioxolane (DLin-KC2-DMA), (6Z,9Z,28Z,31Z)-heptatriaconta-6,9,28,31-tetraen-19-yl 4-(dimethylamino)butanoate (DLin-MC3-DMA), It may include at least one selected from the group consisting of 6-((2-hexyldecanoyl)oxy)-N-(6-((2-hexyldecanoyl)oxy)hexyl)-N-(4-hydroxybutyl)hexan-1-aminium (ALC-0315), 1,2-dioleyloxy-3-dimethylaminopropane (DODMA), 1,2-dioleyl-3-dimethylammonium-propane (DODAP), etc., but is not limited thereto.

[0030] In one embodiment, the ionizable lipid may be 9-heptadecanyl 8-{(2-hydroxyethyl)[6-oxo-6-(undecyclooxy)hexyl]amino}octanoate (SM-102).

[0031] The above helper lipids are distearoylphosphatidylcholine (DSPC), palmitoyloleoylphosphatidylcholine (POPC), 1,2-dioleyl-sn-glycero-3-phosphate (18:1 PA), dipalmitoylphosphatidylcholine (DPPC), dioleoylphosphatidylcholine (DOPC), dioleoylphosphatidylglycerol (DOPG), dipalmitoylphosphatidylglycerol (DPPG), dioleoylphosphatidylethanolamine (DOPE), palmitoyloleoylphosphatidylethanolamine (POPE), dioleoyl-phosphatidylethanolamine 4-(N-maleimidomethyl)-cyclohexane-l-carboxylate (DOPE-mal), dipalmitoylphosphatidylethanolamine (DPPE), It may be at least one selected from the group consisting of, but is not limited to, 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), sphingomyelin (SM), and dioleoylphosphatidylglycerol (DOPG).

[0032] In one embodiment, the helper lipid may be distearoylphosphatidylcholine (DSPC) or palmitoyloleoylphosphatidylcholine (POPC).

[0033] The above 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-dialkyoxypropylcarbamate (PEG-dialkyoxypropylcarbamate), but is not limited thereto.

[0034] In one embodiment, the pegylated lipid may be 1,2-dimyristoyl-rac-glycerol methoxypolyethylene glycol (DMG-PEG2k).

[0035] The above sterols are cholesterol, ergosterol, campesterol, stigmasterol, sitosterol, fucosterol, betulin, lupeol, ursolic acid, oleanolic acid, brassicasterol, 9, 11-dehydroergosterol, daucosterol, β-sitosterol-acetate, β-sitosterol-amino acid conjugates, 20-hydroxycholesterol, It may be at least one selected from the group consisting of 22-hydroxycholesterol, 24-hydroxycholesterol, 25-hydroxycholesterol, 27-hydroxycholesterol, etc., but is not limited thereto.

[0036] In one embodiment, the sterol may be cholesterol.

[0037] 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.

[0038] In one embodiment, the nucleic acid may be mRNA or an oligonucleotide.

[0039] The polydispersity index (PDI) of the above lipid nanoparticles may be 0.3 or less, and preferably 0.2 or less.

[0040] The nucleic acid encapsulation rate of the above lipid nanoparticles may be 50% or more, and preferably 70% or more.

[0041] The nucleic acid content of the above lipid nanoparticles may be 15% or more, and preferably 20% or more.

[0042] The size of the lipid nanoparticles can be controlled according to the Reynolds number for the flow of fluid in the mixing channel, and the Reynolds number can be 10 to 150, preferably 25 to 100.

[0043] 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.

[0044] The size of the lipid nanoparticles can be controlled depending on the molar ratio of the lipid mixture and the sterol. The molar ratio of the lipid mixture and the sterol may be 1:0.1 or more, and preferably 1:0.3 or more.

[0045] The present invention provides lipid nanoparticles obtained by the method for producing the above lipid nanoparticles.

[0046] The present invention includes an inlet channel, a mixing channel and an outlet channel,

[0047] There are two or more of the above inflow channels,

[0048] The above mixing channel includes micropillars,

[0049] A microfluidic device for manufacturing lipid nanoparticles is provided, characterized in that fluids flowing into different inflow channels flow in different directions.

[0050] A fluid including a lipid mixture and nucleic acid can be introduced into the microfluidic device through the above inflow channel, and 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.

[0051] The above lipid mixture can be introduced into one inlet channel, and the nucleic acid can be introduced into another inlet channel.

[0052] The lipid mixture may be introduced in the same direction as the flow of the fluid, and the nucleic acids may be introduced in a different direction from the flow of the fluid, preferably, the nucleic acids may be introduced in a direction perpendicular to the flow of the fluid.

[0053] The above micro-pillars may or may not be connected to the walls forming the mixing channel.

[0054] The above micro-columns can be arranged in one or more rows (columns) in the direction of the fluid flow, and in one column, one or more micro-columns can be arranged in a direction different from the direction of the fluid flow.

[0055] The above-described plurality of micro-pillars may not be arranged in a row in the direction of fluid flow, and may be arranged so as to partially or completely cover the gaps between micro-pillars arranged in adjacent rows when viewed in the direction of fluid flow.

[0056] The size of the lipid nanoparticles can be controlled according to the Reynolds number for the flow of fluid in the mixing channel, and the Reynolds number can be 10 to 150, preferably 25 to 100.

[0057] The size of the lipid nanoparticles can be controlled by the molar ratio of the lipid mixture and the sterol. The molar ratio of the lipid mixture and the sterol can be 1:0.1 or more, and preferably 1:0.3 or more.

[0058] The polydispersity index (PDI) of the above lipid nanoparticles may be 0.3 or less, and preferably 0.2 or less.

[0059] The nucleic acid encapsulation rate of the above lipid nanoparticles may be 50% or more, and preferably 70% or more.

[0060] The nucleic acid content of the above lipid nanoparticles may be 15% or more, and preferably 20% or more.

[0061]

[0062] The lipid nanoparticle manufacturing method of the present invention provides an optimal lipid nanoparticle manufacturing method by controlling the particle size of lipid nanoparticles according to the composition ratio of raw materials including cholesterol. The lipid nanoparticle manufacturing method of the present invention can be used in the production of pharmaceuticals containing lipid nanoparticles.

[0063] In addition, by using the lipid nanoparticle manufacturing method of the present invention in the manufacturing process of a pharmaceutical product that uses nucleic acid as a modality, messenger ribonucleic acid can be encapsulated in lipid nanoparticles of various sizes at a high concentration, thereby increasing therapeutic efficiency.

[0064]

[0065] Figure 1 is a schematic diagram of the synthesis process of lipid nanoparticles (LNPs) containing nucleic acid using the micro mixing channel device of the present invention and the shape of nucleic acid-containing LNPs.

[0066] Figure 2 is a schematic diagram showing the length of each part of the device for the structure and design of the device of the present invention.

[0067] Figure 3 shows the flow of fluid through the device of the present invention and the flow of fluid around the micro-pillars.

[0068] Figure 4 shows the results of measuring the mixing efficiency according to the number of micro-columns in the device of the present invention.

[0069] Figure 5 shows the results of measuring mixing efficiency according to the number of micro-pillars present in one column in the device of the present invention.

[0070] Figure 6 is a result showing a Dean vortex formed by the width spacing of micro-pillars arranged in the mixing zone of the device of the present invention.

[0071] Figure 7 shows the results of the number of Deans and mixing efficiency according to the width spacing of the micro-pillars arranged in the mixing zone of the device of the present invention.

[0072] Figure 8 shows the results of measuring mixing efficiency according to changes in Reynolds number in the device of the present invention.

[0073] Figure 9 shows the results of measuring mixing efficiency according to changes in channel height in the device of the present invention.

[0074] Figure 10 shows the results of measuring the mixing efficiency according to the injection ratio of the aqueous solution and lipid mixture in the device of the present invention.

[0075] Figure 11 shows a calculation method for the residence time of a fluid.

[0076] Figure 12 shows the fluid flow within the device according to Manufacturing Examples 1 and 2 of the present invention.

[0077] Figure 13 shows the results of measuring different mixing efficiencies depending on changes in the structure start interval, micro-column interval, and micro-column vertical length in the device of the present invention.

[0078] Figure 14 shows the results of measuring mixing efficiency according to changes in blockage rate in the device of the present invention.

[0079] Figure 15 is a result showing the mixing efficiency within the device according to the front and rear width gap by the micro pillars in the device of the present invention.

[0080] Figure 16 shows the results of shear rate according to Reynolds number at each width interval by micro-pillars in the device of the present invention.

[0081] Figure 17 shows the results of design conditions by size for the device of the present invention.

[0082] Figure 18 shows the results of mixing efficiency according to the Reynolds number according to the device size of the present invention.

[0083] Figure 19 shows the results of flow velocity according to Reynolds number according to device size of the present invention.

[0084] Figure 20 shows the results of shear rate according to Reynolds number according to device size of the present invention.

[0085] Figure 21 is a graph comparing the size of LNP and polydispersity index (PDI) with Reynolds number when the composition ratio (molar ratio) of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k is 50:10:38.5:1.5.

[0086] Figures 22 and 23 are DLS results for the size of LNP according to the Reynolds number of Figure 21.

[0087] Figure 24 is a graph comparing the size of LNP and polydispersity index (PDI) with Reynolds number when the composition ratio (molar ratio) of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k is 28:5.7:65.5:0.8.

[0088] Figures 25 and 26 are DLS results for the size of LNP according to the Reynolds number of Figure 24.

[0089] Figure 27 is a graph comparing the size and polydispersity index (PDI) of LNPs with the Reynolds number when the composition ratio (molar ratio) of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k is 20:4:75.5:0.5.

[0090] Figures 28 and 29 are DLS results for the size of LNP according to the Reynolds number of Figure 27.

[0091] Figure 30 is a graph comparing the size and PDI of LNP according to the cholesterol molar ratio.

[0092] Figures 31 and 32 are number distributions of the DLS results for the size of LNP according to the cholesterol molar ratio of Figure 30.

[0093] Figures 33 and 34 are volume distributions of the DLS results for the size of LNPs according to the cholesterol molar ratio of Figure 30.

[0094] Figures 35 and 36 are intensity distributions of the DLS results for the size of LNP according to the cholesterol molar ratio of Figure 30.

[0095] Figure 37 is a graph comparing the size and polydispersity index (PDI) of LNPs at Reynolds numbers of 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.

[0096] Figure 38 is a DLS result for the size of LNP according to the Reynolds number of Figure 37.

[0097] Figure 39 is a graph comparing the size and PDI of bare LNPs without mRNA and mCherry LNPs under synthetic conditions with a size of 30 nm.

[0098] Figure 40 is a graph comparing the size and PDI of bare LNPs without mRNA and mCherry LNPs under synthetic conditions with a size of 60 nm.

[0099] Figure 41 is a graph comparing the size and PDI of bare LNPs without mRNA and mCherry LNPs under synthetic conditions with a size of 90 nm.

[0100] Figures 42 and 43 are DLS results for the sizes of Bare LNPs and mCherry LNPs without mRNA of Figures 39, 40, and 41.

[0101] Figures 44 and 45 are graphs comparing the size and PDI of bare LNPs without mRNA and Cas9 LNPs under synthetic conditions with a size of 30 nm.

[0102] Figure 46 is a graph comparing the size and PDI of bare LNPs without mRNA and Cas9 LNPs under synthetic conditions with a size of 60 nm.

[0103] Figure 47 is a graph comparing the size and PDI of bare LNPs without mRNA and Cas9 LNPs under synthetic conditions with a size of 90 nm.

[0104] Figures 48 and 49 are DLS results for the sizes of bare LNPs and Cas9 LNPs without mRNA of Figures 44, 45, 46, and 47.

[0105] Figure 50 is a graph comparing the encapsulation efficiency (EE) of mCherry LNPs under synthetic conditions with sizes of 30 nm, 60 nm, and 90 nm.

[0106] Figure 51 is a graph comparing the encapsulation efficiency (EE) of Cas9 LNPs under synthetic conditions with sizes of 30 nm, 60 nm, and 90 nm.

[0107] Figure 52 is a graph comparing the size and PDI of bare LNPs without mRNA and oligo LNPs under synthetic conditions with a size of 30 nm.

[0108] Figure 53 is a graph comparing the size and PDI of bare LNPs without mRNA and oligo LNPs under synthetic conditions with a size of 60 nm.

[0109] Figure 54 is a graph comparing the size and PDI of bare LNPs and oligo LNPs without mRNA under synthetic conditions with a size of 90 nm.

[0110] Figures 55 and 56 are DLS results for the sizes of Bare LNPs and Oligo LNPs without mRNA of Figures 52, 53, and 54.

[0111] Figure 57 is a graph comparing the encapsulation efficiency (EE) of Oligo LNPs under synthetic conditions with sizes of 30 nm, 60 nm, and 90 nm.

[0112] Figure 58 is a fluorescence image showing the level of mCherry protein expression after treatment with mCherry LNPs (200 ng of mCherry mRNA) of synthetic conditions with sizes of 30 nm, 60 nm, and 90 nm in human glioblastoma U87MG.

[0113] Figure 59 is a graph quantifying the fluorescence of mCherry protein in the U87MG fluorescence image of Figure 58.

[0114] Figure 60 is a fluorescence image showing the degree of mCherry protein expression according to the amount of mCherry mRNA processed (500, 200, 20 ng) under synthetic conditions with sizes of 60 nm and 90 nm in the human liver cancer cell line HepG2.

[0115] Figure 61 is a graph quantifying the fluorescence of mCherry protein in the HepG2 fluorescence image of Figure 60.

[0116] Figure 62 is a fluorescence image showing the degree of mCherry protein expression according to the amount of mCherry mRNA processed (500, 200, 20 ng) under synthetic conditions with sizes of 60 nm and 90 nm in the human monocyte cell line THP-1.

[0117] Figure 63 is a graph quantifying the fluorescence of mCherry protein in the THP-1 fluorescence image of Figure 62.

[0118] Figures 64 and 65 show the results for size, polydispersity index (PDI), mCherry mRNA encapsulation ratio (EE), and endogenous content (LE) for storage stability after synthesis of 60 nm mCherry LNPs.

[0119] Figures 66 and 67 show the results for size, polydispersity index (PDI), mCherry mRNA encapsulation ratio (EE), and endogenous content (LE) for storage stability after synthesis of 90 nm mCherry LNPs.

[0120] Figures 68 and 69 show the results for size, polydispersity index (PDI), Cas9 mRNA encapsulation rate (EE), and endogenous content (LE) for storage stability after synthesis of 30 nm Cas9 LNPs.

[0121] Figures 70 and 71 show the results for size, polydispersity index (PDI), Cas9 mRNA encapsulation rate (EE), and endogenous content (LE) for storage stability after 90 nm Cas9 LNP synthesis.

[0122] Figures 72 and 73 show the results for size, polydispersity index (PDI), oligonucleotide encapsulation ratio (EE), and endogenous content (LE) for storage stability after synthesis of 50 nm Oligo LNP (target size: 30 nm).

[0123] Figures 74 and 75 show the results for size, polydispersity index (PDI), oligonucleotide encapsulation ratio (EE), and endogenous content (LE) for storage stability after synthesis of 100 nm Oligo LNP (target size: 60 nm).

[0124] Figures 76 and 77 show the results for size, polydispersity index (PDI), oligonucleotide encapsulation ratio (EE), and endogenous content (LE) for storage stability after synthesis of 130 nm Oligo LNP (target size: 90 nm).

[0125]

[0126] Hereinafter, with reference to the attached drawings, embodiments and examples of the present invention will be described in detail so that those skilled in the art can easily implement the present invention. However, the present invention may be implemented in various forms and is not limited to the embodiments and examples described herein.

[0127] Throughout this specification, whenever a part is said to "include" a component, this means that it may include other components, but not to the exclusion of other components, unless otherwise stated.

[0128] The term "lipid nanoparticle" as used herein is composed of four types: ionizable lipid, helper lipid, PEGylated lipid, and sterol. The helper lipid may be composed of more than one type of lipid. It also refers to a nano-sized particle capable of encapsulating nucleic acids therein.

[0129] The term "device" or "microfluidic device" as used herein means a device that includes microchannels or the like that allow fluid to flow on a substrate made of various materials including plastic, glass, metal or silicon that includes an organic polymer material.

[0130] In the device of the present invention, the inlet channel, the mixing channel, and the outlet channel can be arranged sequentially in the direction of the flow of the fluid.

[0131] The above inflow channel may include a channel for introducing nucleic acids and a channel for introducing a lipid mixture, wherein the nucleic acids and the lipid mixture may each be introduced through different inflow channels. Here, the lipid mixture may be introduced in the mixing channel in the same direction as the fluid flow direction, and the nucleic acids may be introduced in a direction different from the fluid flow direction. Preferably, the lipid mixture may be introduced in the same direction as the fluid flow direction, and the nucleic acids may be introduced in a direction orthogonal to the fluid flow direction.

[0132] The inlet channel through which the lipid mixture is introduced may be one or more, and the inlet channel through which the nucleic acid is introduced may be one or more. Preferably, the device of the present invention may include one inlet channel through which the lipid mixture is introduced and two inlet channels through which the nucleic acid is introduced.

[0133] In the above mixing channel, nucleic acid and lipid mixture can be mixed with each other.

[0134] The particle size can be controlled by changing the Reynolds velocity for the fluid flow in the mixing channel. In addition, the Reynolds velocity for the fluid flow in the mixing channel can be 10 to 150, and preferably 25 to 100.

[0135] The height of the above channel may be 200 to 800 μm.

[0136] The term "fluid flow" as used herein refers to the direction in which fluid flows from an inlet channel to an outlet channel within the device. Preferably, the lipid mixture is introduced through an inlet channel arranged in the same direction as the flow direction in the mixing channel, and the nucleic acids are introduced through an inlet channel arranged orthogonal to the flow direction in the mixing channel.

[0137] In general, the flow velocity within the device can be expressed by the Reynolds number (Re), a dimensionless number that represents the ratio of inertial force and viscous force, and the calculation formula is as follows.

[0138]

[0139] At this time, D h is the hydraulic diameter within the microfluidic device, and ρ, μ, υ, Q represent the density, kinematic viscosity, velocity, and flow rate of the fluid, respectively. Therefore, as the Reynolds number increases, the flow rate increases proportionally. When the Reynolds number exceeds 500, turbulence-like flow is formed within the device, which does not generate a controllable vortex pattern, and the increased shear force between the solvent and the substance due to the high flow rate may cause instability of the raw drug substance and the produced nanoparticles. When the Reynolds number is 300 or higher, uncontrollable chaotic flow begins to occur within the device, making quality control for the production of uniform nanoparticles difficult. On the other hand, when the Reynolds number is low, such as 10 or less, it relies on diffusion, making it difficult to achieve effective mixing of the lipid mixture and nucleic acid, making it difficult to achieve superior productivity compared to existing nanoparticle synthesis methods.

[0140] The device of the present invention can control the size of particles by changing the Reynolds number.

[0141] The term "micropost" as used herein refers to a structure that disrupts the straight flow of fluid within a mixing channel. The micropost of the present invention is a structure that allows fluids introduced into the device to form microvortices and be efficiently mixed, and may include any shape, and is preferably a columnar shape. The micropost may be designed to separate or bend the flow of fluids that collide with the post, or to merge with other flows. The micropost may be designed to maintain the main flow of the fluid and prevent it from stagnating, and for example, may collide with the fluid at a right angle to the direction of flow of the fluid.

[0142] The above micro-columns may or may not be connected to the walls forming the mixing channel, and may be at least one selected from the group consisting of polypyramids, polypyramidal truncations, polygonal columns, and modified shapes thereof, and preferably may be square columns.

[0143] The above micro-columns may be arranged in one or more columns in the direction of fluid flow, and preferably in six or more columns in the direction of fluid flow. In addition, one or more micro-columns may be arranged in a direction different from the direction of fluid flow in one column, and preferably, one to six micro-columns may be arranged in a direction different from the direction of fluid flow in one column, more preferably, one to six micro-columns may be arranged in a direction perpendicular to the direction of fluid flow in one column, and most preferably, one to two micro-columns may be arranged in a direction perpendicular to the direction of fluid flow in one column.

[0144] The plurality of micro-columns may not be arranged in a row in the direction of the fluid flow, and the plurality of micro-columns may be arranged so as to partially or completely cover the gaps between micro-columns arranged in adjacent rows when viewed in the direction of the fluid flow.

[0145] In the present invention, the micro-columns may be arranged in an alternating manner with respect to the micro-columns in adjacent rows. In the present invention, "arranged in an alternating manner" means that the micro-columns in the plurality of rows are not arranged in a straight line or in a single row parallel to each other in the direction of fluid flow, and the micro-columns arranged in one row may be arranged so as to partially or completely cover the gaps between the micro-columns arranged in adjacent rows when viewed in the direction of fluid flow.

[0146] In the present invention, the micro-columns may have a height of 200 to 800 μm, and the micro-columns may or may not be connected to the walls forming the mixing channel.

[0147] The term "mixing efficiency" as used herein )" is a value expressed as a percentage using the fact that the lipid mixture and nucleic acid, each having a mass fraction of 1, are mixed together in the channel and ultimately converge to 0.5, and the calculation formula is as follows.

[0148]

[0149] At this time is the mass fraction, and the closer the mass fraction is to 0.5, the higher the synthesis efficiency proportionally.

[0150] The above-mentioned outflow channel may be a channel through which the generated lipid nanoparticles outflow.

[0151] The term "inlet volume flow rate ratio" as used herein means the ratio of the inlet volume flow rate of an aqueous fluid to the inlet volume flow rate of a fluid containing a lipid mixture, respectively.

[0152] The term "flow blockage ratio" as used herein is the ratio of the width of the micro-pillars to the width of the entire channel in the first row along the direction of fluid flow in the mixing channel.

[0153] The term "encapsulation efficiency (EE)" used in this specification is calculated using the values ​​obtained by detecting total mRNA and free mRNA based on fluorescence intensity in each mRNA LNP. ) is the value calculated according to the following.

[0154] The above “Total mRNA” refers to the total amount of mRNA contained within the final product of the lipid nanoparticle manufacturing process, and the above “Free mRNA” refers to the amount of mRNA present outside the lipid nanoparticle among the final product of the lipid nanoparticle manufacturing process.

[0155] The term "nucleic acid loading efficiency (LE)" used in this specification is calculated using the values ​​obtained by detecting total mRNA and free mRNA based on fluorescence intensity in each mRNA LNP. ) is calculated according to the above. The above “feeding mRNA” refers to the total amount of mRNA input into the lipid nanoparticle manufacturing process.

[0156] The present invention will be described in more detail through the following examples; however, the following examples are for illustrative purposes only and are not intended to limit the scope of the present invention.

[0157]

[0158] [Example 1]

[0159] Device design for manufacturing lipid nanoparticles

[0160] A microfluidic device for manufacturing lipid nanoparticles (LNPs) was designed (Figs. 1, 2, and 3). The microfluidic device includes three inlet channels and one outlet channel. A lipid mixture was injected into the central channel of the three inlet channels, and an aqueous solution containing nucleic acid was injected into the two channels on either side. In addition, a micro-pillar structure was introduced into the device to effectively mix the lipid mixture and nucleic acid.

[0161]

[0162] [Example 2]

[0163] Effect of a single micro-column structure

[0164] To determine the mixing efficiency of lipid mixtures and nucleic acid solutions according to the presence, number, and arrangement of micropillars placed inside the device, the distribution of lipid mixtures inside the microfluidic device was analyzed using fluid dynamics techniques.

[0165] As shown in Fig. 3, it was confirmed that the lipid mixture and the nucleic acid solution were efficiently mixed by microvortex generated when the lipid mixture flowed through the micro-column structure.

[0166] In the present invention, the inlet volume flow rate ratio refers to the ratio of the inlet volume flow rate of the aqueous fluid to the inlet volume flow rate of the fluid containing the lipid mixture, and the flow blockage ratio refers to the ratio of the width of the micro-columns to the width of the entire channel in the first row along the direction of fluid flow in the mixing channel.

[0167] The results were confirmed under the conditions of a channel height of 200 μm, an inlet volume flow rate ratio of 1:5.5 (lipid mixture:aqueous solution), a Reynolds number (Re) of 50, a flow blockage ratio of 0.5, and a rectangular micro-pillar size of 1000 μm X 400 μm, and the results are shown in Figs. 4 and 5. The number of micro-pillars was designed to be evenly distributed across the channel width while maintaining the size of the micro-pillars and maintaining a constant width gap between the micro-pillars through which the fluid passes.

[0168] As shown in FIGS. 4 and 5, it was confirmed that the mixing efficiency was improved by approximately 1.5 to 4.0 times depending on the number of micro-pillars distributed in the case with micro-pillars compared to the case without micro-pillars.

[0169]

[0170]

[0171]

[0172] As shown in Fig. 4 and Table 1 above, as the number of columns increases, the mixing efficiency increases, and when the number is 6 or more, an excellent mixing efficiency of 98% or more is shown.

[0173]

[0174]

[0175]

[0176] As shown in Fig. 5 and Table 2 above, as the number of micro-pillars located within a channel in one column increases, the mixing efficiency actually decreases, and the best mixing efficiency is shown when the number is 1-2.

[0177] Therefore, the mixing efficiency varies depending on the number and arrangement of the micro-pillars arranged within the device, and in particular, as shown in FIGS. 6 and 7, the highest mixing efficiency is achieved when a single micro-pillar structure is arranged in an interleaved manner. The micro-pillars are evenly arranged with respect to the channel width, and the width gap between the arranged micro-pillars through which the fluid passes is maintained constant.

[0178]

[0179] [Example 3]

[0180] Optimization of the available flow rate range and flow rate ratio range within the device

[0181] To optimize LNP synthesis according to the flow rates and injection ratios of lipid mixture and aqueous solution within the device, the mixing efficiency was analyzed under various Reynolds number (Re) and flow rate ratio conditions.

[0182] To investigate the available flow rate range, the mixing efficiency was analyzed when the Reynolds number was varied from 12.5 to 200 at a channel height of 200 μm and an injection ratio of 1:5.5 in the device, and the results are shown in Fig. 8 and Table 3.

[0183]

[0184]

[0185]

[0186] As shown in Table 3 above, a mixing efficiency of 90% or more was obtained when the Reynolds number was 12.5 to 200, and a mixing efficiency of 98% was confirmed when the Reynolds number was 50 and 100.

[0187] In addition, the mixing efficiency was analyzed when the channel height was changed 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 Fig. 9 and Table 4.

[0188]

[0189]

[0190]

[0191] As shown in Table 4 above, it was confirmed that the mixing efficiency was 95% or more when the channel height was 200 to 800 μm.

[0192] In addition, the mixing efficiency was analyzed when the channel height in the device was 200 μm, the Reynolds velocity (Re) was 50, and the injection ratio was changed from 1:2.5 to 1:8.5, and the results are shown in Fig. 10 and Table 5.

[0193]

[0194]

[0195]

[0196] As shown in Table 5 above, it was confirmed that the mixing efficiency was 97% or higher when the injection ratio was 1:2.5 to 1:8.5.

[0197]

[0198] [Example 4]

[0199] Study on the height of the device to increase mixing efficiency

[0200] To maximize the mixing efficiency according to the height of the mixing channel of the device, the mixing efficiency was analyzed under different height conditions.

[0201] In the process of synthesizing nanoparticles using the device, when raw materials are injected at a relatively high flow rate (Re>50), strong shear stress is generated from the wall due to the narrow width within the channel.

[0202] Organic substances in fluids have been reported to undergo aggregation and deformation when exceeding a certain value. In particular, in the case of proteins, several literatures have shown that protein deformation, including aggregation, folding, and degradation, is induced by the shear stress of fluid flow generated in external equipment. This deformation of the precursor protein changes the structure, size, and even the function of the manufactured nanoparticles. In general, the range of deformation induced varies depending on the structure of the equipment generating the shear stress and the form and size of the material used. However, for proteins with a size of 1 nm - 10 nm (mass range of 20 kDa - 300 kDa), such as insulin, enzymes, and immunoglobulins, the shear stress is 1000 dyne / cm. 2 It has been reported that shear flow can modify the structure of proteins in a range of directions. (Bekard, I., et al. The Effects of Shear Flow on Protein Structure and Function. Biopolymers, 95, 11, 733-745 (2011))

[0203] Therefore, the shear stress generated through the fluid flow in the device of the present invention ) to determine whether the precursor material and nanoparticles were deformed, the shear stress generated according to each channel height was first calculated as follows.

[0204]

[0205] Here, Q is the flow rate, represent the dynamic viscosity coefficient of the fluid, the height of the channel cross-section, and the horizontal length of the channel cross-section, respectively.

[0206]

[0207]

[0208]

[0209] As shown in Table 6 above, as the height of the channel in the device of the present invention decreases, the shear force increases exponentially, but the protein structure deformation is 1000 dyne / cm. 2 It was confirmed that the range was not exceeded.

[0210] However, if the channel height is 100 μm, the Reynolds number corresponding to the flow velocity is increased to 300, which is 1000 dyne / cm. 2 In addition, 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 the precursor and nanoparticles are not deformed due to shear stress when the channel height is 200 μm or more.

[0211] In addition, as shown in Table 7 below, as the height of the mixing channel of the device increases, the volume increases, and for the same flow rate, the residence time ( ) and the mixing time required for the precursors to be synthesized ( ) increases. The specific calculation method for the residence time of the mixed substances staying in the channel is shown in Fig. 11. The mixing time required for the substances to be synthesized ( ) can be calculated through the more dominant physical phenomenon among convection and diffusion within the device, and can be calculated as follows through the Peclet number (Pe), a dimensionless number representing the ratio of convection time to diffusion time.

[0212]

[0213] In the device of the present invention, the Peclet number (Pe) is 100 to 5000, and based on this, it can be seen that mixing by convection is dominant, so the mixing time by convection in the channel ( ) was calculated as follows. (Velencia, 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))

[0214]

[0215] Here, 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.

[0216]

[0217]

[0218]

[0219] At this time, basically the residence time ( ) is the mixing time ( ) so that the materials can be sufficiently mixed within the channel. As shown in Table 7 above, since the residence time is longer than the mixing time at all channel heights, it can be confirmed that sufficient mixing time is secured at a given channel height.

[0220] In addition, the increase in mixing time according to the increase in channel height is calculated to be 53 ms at the highest height of 800 μm as shown in Table 7 above, which is the coagulation time ( ) can be seen as the maximum height for applying the device and manufacturing method of the present invention as the time when it starts to enter the range of 50 - 100 ms. (Refer to 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))

[0221] Therefore, as shown in Table 7 above, it can be seen that the optimal mixing channel height that can prevent aggregation and non-uniform nanoparticle generation of the obtained nanoparticles within the device is 200 to 800 μm.

[0222]

[0223] [Example 5]

[0224] Fluid flow within the device

[0225] Depending on the channel height and micro-pillar conditions, the device of the present invention was designed in two manufacturing examples as shown in Table 8 below.

[0226]

[0227]

[0228]

[0229] To manufacture the above Manufacturing Examples 1 and 2 and confirm the mixing flow pattern of the lipid mixture and the aqueous solution, the distribution of the material was visualized through ink and observed under a microscope.

[0230] The fluid containing the lipid mixture was visualized using 6% ink in ethanol, and the fluid containing the hydrophilic substance was visualized using saline solution, and the results are shown in Fig. 12. As shown in Fig. 12, it can be seen that the lipid mixture and the aqueous solution are efficiently mixed by the microvortex generated when the lipid mixture flows through the micro-column structure.

[0231]

[0232] [Example 6]

[0233] Optimization of device-in-device structural design

[0234] In order to optimize the mixing efficiency according to the size of the designed channel and micro-pillars within the device, the mixing efficiency was confirmed under various conditions including 1) the starting spacing of micro-pillars, 2) the spacing between micro-pillars, 3) the vertical length of micro-pillars, and 4) the flow blockage ratio within the structure of the device.

[0235] The mixing efficiency for each variable was confirmed under the conditions of a fixed channel height of 0.2 mm, an injection ratio of 1:5.5, and a Reynolds number of 50, and the results are shown in Figs. 13 and 14.

[0236]

[0237]

[0238]

[0239] As shown in Fig. 13 and Table 9, it was confirmed that the flow pattern of the fluid passing through the designed micro-pillars in the channel did not change significantly regardless of the spacing and vertical length conditions of the pillars, and that a high mixing efficiency (>0.95) was maintained under all conditions.

[0240]

[0241]

[0242]

[0243] On the other hand, as shown in Fig. 14 and Table 10, in the case of the blockage rate, which is the horizontal spacing ratio of the micro-pillars in the channel, the mixing efficiency increases as the value increases, and it was confirmed that the mixing efficiency was maintained at a constant level of 0.98 or higher when the blockage rate was 0.5.

[0244] In addition, the mixing efficiency according to the width gap between the front and rear passages through which the fluid passes is shown in Figure 15. The shear rate when the width gap between the front and rear passages is the same is shown in Figure 16. The narrower the width gap through which the fluid passes, the higher the shear rate. Since the shear rate is proportional to the shear stress, it can be said that the shear stress increases.

[0245] That is, when the flow blockage ratio increases, the width of the channel also narrows, and accordingly, the shear stress increases proportionally. As previously calculated, high shear stress can cause aggregation and deformation of organic substances existing in the fluid when it exceeds a certain value (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 Re 300 under the condition that the blockage ratio exceeds 0.65, the shear stress increases to 1000 dyne / cm 2 Considering the increasing shear stress, it can be seen that the flow blockage ratio of the channel can be 0.2 to 0.8, and preferably, when it is in the range of 0.35 to 0.65, it has a high mixing efficiency without being greatly affected by the shear stress.

[0246]

[0247] [Example 7]

[0248] Device design for mass production of lipid nanoparticles

[0249] In the present invention, a device capable of mass-producing nanoparticles including LNPs was designed. As shown in Fig. 17, when the microfluidic device of Figs. 1 and 2 was used as a reference (X1), the two-dimensional size of the device, i.e., the starting spacing of micropillars, the spacing between micropillars, the vertical length of micropillars, and the width spacing between the front and back of the fluid flow, was expanded by two times (X2), three times (X3), and four times (X4), and the mixing efficiency, flow velocity, and shear rate according to the Reynolds number were analyzed using fluid dynamics techniques. When the device was expanded, the channel height was maintained at 200 μm, and the flow blockage ratio was maintained at 0.5.

[0250] As shown in Fig. 18, it can be confirmed that the microfluidic device of Fig. 2, which is the reference, has a mixing efficiency of 0.95 or higher when the Reynolds number is 50 to 300. When the size of the device is doubled (X2), the mixing efficiency decreases to 0.84 when the Reynolds number is 50. However, when the Reynolds number is 100 or more, it is confirmed that the mixing efficiency is 0.95 or higher up to Reynolds number 300. When the size of the device is tripled (X3), it is confirmed that the mixing efficiency is 0.95 or higher when the Reynolds number is 150 to 300. When the size of the device is quadrupled (X4), it is confirmed that the mixing efficiency is 0.95 or higher when the Reynolds number is 200 to 300.

[0251] As shown in Figure 19, it can be confirmed that as the device size increases, only the device flow rate needs to increase to maintain the same Reynolds number. Since an increase in flow rate means an increase in the amount of nanoparticles that can be synthesized per hour, it can be seen that mass production of nanoparticles is possible through two-dimensional expansion of the microfluidic device of Figures 1 and 2.

[0252] As shown in Figure 20, it can be seen that the shear rate decreases as the device size increases. That is, since the shear rate is proportional to the shear stress, it can be said that the shear stress decreases as the device size increases. It can be seen that increasing the device size does not lead to the aggregation and deformation of organic substances (in the case of LNP, nucleic acids) present in the fluid.

[0253] It can be seen that the microfluidic device of the present invention is a device capable of mass-producing LNPs containing nucleic acids without causing aggregation or deformation of organic substances present in the fluid through two-dimensional expansion of the size.

[0254]

[0255] [Example 8]

[0256] Optimization of the synthesis of lipid nanoparticles

[0257] In the present invention, lipid nanoparticles (LNPs) were synthesized using a microfluidic device, as illustrated in Figure 1. A lipid mixture was injected into the central channel of the three inlet channels, and an aqueous solution was injected into the two channels on either side. The lipid mixture and aqueous solution were efficiently mixed by micro-pillars within the device, and ultimately, synthesized LNPs were obtained through the outlet channels of the micro-mixing channel device.

[0258] Ionizable lipids, helper lipids, PEGylated lipids, and sterols can be used as lipid mixtures for LNP synthesis. Specifically, in the present invention, SM-102 (9-heptadecanyl 8-{(2-hydroxyethyl)[6-oxo-6-(undecyloxy)hexyl]amino}octanoate), DSPC (Distearoylphosphatidylcholine), POPC (Palmitoyloleoylphosphatidylcholine, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine), DMG-PEG2k (1,2-Dimyristoyl-rac-glycero-3-methoxypolyethylene glycol-2000), and cholesterol were used.

[0259] In addition, a buffer solution of pH 3.0 to 6.0 is required as an aqueous solution for LNP synthesis, and in the present invention, a citrate buffer solution of pH 3.0 was used.

[0260]

[0261] 8-1 [Synthesis of Lipid Nanoparticles According to Flow Velocity]

[0262] The lipid mixture used SM-102, DSPC / POPC, cholesterol, and DMG-PEG2k, and the composition ratio was 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 the size of LNPs according to flow rate. The three ratios of the lipid mixture were set to increase the composition molar ratio of cholesterol in the lipid mixture based on the commonly used ratio of 50:10:38.5:1.5.

[0263] The synthesized LNPs were characterized by hydrodynamic size based on number-weighted size distribution and polydispersity index (PDI) using a Zetasizer Pro Blue instrument using dynamic light scattering (DLS). The degree of uniformity of the LNPs was indicated by the polydispersity index (PDI).

[0264]

[0265]

[0266]

[0267] 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 can be seen in Table 11 and Figs. 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 lower, confirming that very uniform LNPs were synthesized overall. The PDI value considered as a stable LNP may vary depending on each situation, but the FDA guidelines recommend that the PDI of LNPs used as drugs be 0.3 or lower.

[0268]

[0269]

[0270]

[0271] LNPs were synthesized by fixing the molar ratio of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k to 28:5.7:65.5:0.8 (a condition in which only the ratio of cholesterol was increased compared to Table 1) and changing the Reynolds number to 12.5, 25, 50, 100, and 200. As can be seen in Table 12 and Figures 24, 25, and 26, the sizes of 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 highly uniform LNPs were synthesized.

[0272]

[0273]

[0274]

[0275] LNPs were synthesized by fixing the molar ratio of SM-102:DSPC / POPC:Cholesterol:DMG-PEG2k to 20:4:75.5:0.5 (a condition in which only the ratio of cholesterol was increased compared to Table 12) and changing the Reynolds number to 12.5, 25, 50, 100, and 200. As can be seen in Table 13 and Figs. 27, 28, and 29, the sizes of 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 highly uniform LNPs were synthesized.

[0276] That is, when LNPs were synthesized using the microfluidic device shown in Fig. 1, it was confirmed that very uniform LNPs (except for the Reynolds flow rate of 12.5 in Table 11, with a PDI value not higher than 0.1) were synthesized regardless of the flow rate at various lipid composition ratios. However, it was confirmed that there was no significant change in the LNP size depending on the flow rate.

[0277]

[0278] 8-2 [Synthesis of Lipid Nanoparticles According to Lipid Mixture Composition Ratio]

[0279] To change the composition ratio of the lipid mixture consisting of SM-102, DSPC / POPC, cholesterol, and DMG-PEG2k, LNPs were synthesized with the composition molar ratios of cholesterol in the lipid mixture being 38.5, 55.5, 65.5, 75.5, 86.0, and 98.45. At this time, the composition ratio among the lipid mixtures (ionized lipid, helper lipid, and PEGylated lipid) excluding cholesterol was kept constant.

[0280]

[0281]

[0282]

[0283] The composition ratio of the entire lipid mixture according to the change in the cholesterol composition molar ratio is shown in Table 14. Using the composition ratio of each lipid mixture, LNPs were synthesized using a microfluidic device as shown in Fig. 1. At this time, the Reynolds number was selected as 50, which is the Reynolds number at which the PDI is the smallest, i.e., the most uniform LNPs are synthesized, based on the results of Example 8-1.

[0284]

[0285]

[0286]

[0287] As shown in Table 14 above, the LNP size and PDI results when LNPs were synthesized by controlling the composition ratio of the lipid mixture during LNP synthesis are shown in Table 15. As can be seen in Fig. 30 and Table 15, the sizes of the synthesized LNPs according to the molar ratios of cholesterol in the lipid mixture of 38.5, 55.5, 65.5, 75.5, 86.0, and 98.45 were measured as 64, 84, 102, 93, 103, and 105 nm.

[0288] The uniformity of LNPs synthesized using the microfluidic device was confirmed through PDI. PDI was measured to be less than 0.2 up to the condition of cholesterol molar ratio 86.0, confirming that the synthesis was very uniform. The uniformity of the synthesized LNPs can also be confirmed by the narrow single peak in the number-weighted size distribution results in FIGS. 31 and 32, the volume-weighted size distribution results in FIGS. 33 and 34, and the intensity-weighted size distribution results in FIGS. 35 and 36.

[0289] When using the above microfluidic device, it was confirmed that LNPs were synthesized uniformly in size even when the cholesterol molar ratio was 86.0, which is more than twice the cholesterol molar ratio of 38.5 in a commonly used lipid mixture composition, that is, when 80% of the total lipid mixture composition was cholesterol. In addition, when using the above microfluidic device, it was confirmed that LNPs were synthesized even with an extreme lipid mixture composition with a cholesterol molar ratio of 98.45. However, in this case, it was confirmed that the PDI was 0.274, indicating lower monodispersity compared to other conditions. The reason for the slightly higher PDI is that the LNPs clumped together (aggregation) due to the excess cholesterol.

[0290] Additionally, under conditions where the cholesterol molar ratio was 60 or less, the LNP size tended to increase as the cholesterol ratio increased, but no change in the LNP size was observed from the condition where the cholesterol molar ratio was 65.5.

[0291]

[0292] 8-3 [Optimization of Synthesis of Size-Specific Lipid Nanoparticles]

[0293] As can be seen from the results of Examples 8-1 and 8-2 above, the size of the LNP synthesized using the microfluidic device is 60 nm to 100 nm. In order to synthesize LNPs of 50 nm or less, LNPs were synthesized by increasing the molar ratio of PEGylated lipids in the lipid mixture. 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 in order to confirm whether the LNP size changes according to the Reynolds number, the Reynolds number was 50 and 100.

[0294]

[0295]

[0296]

[0297] As can be seen in Table 16 above and Figures 37 and 38, when the Reynolds number is 50 and 100, LNPs were synthesized to be 35 and 36 nm, respectively, and it can be confirmed that the PDI was synthesized relatively uniformly around 0.2.

[0298]

[0299]

[0300]

[0301] When the final synthesized LNP results were summarized, the Reynolds number and lipid mixture composition conditions for synthesizing very uniform LNPs with specific sizes of 30 nm, 60 nm, and 90 nm using the microfluidic device shown in Fig. 1 were shown in Table 17.

[0302] The average diameter of LNPs typically used as mRNA vaccines falls within the 80–100 nm range. LNPs smaller than this (~60 nm) are difficult to manufacture due to the high curvature stress they impose on the particles, which tends to reduce particle stability and impair drug loading (inclusion rate). Conversely, smaller LNPs tend to have higher surface area, leading to increased drug delivery efficiency to target organs.

[0303] Furthermore, the size of LNPs also influences their targeting properties for each organ. In general, organs such as the liver, which contain numerous microvessels, are known to exhibit decreased targeting properties for those organs as LNP size increases. Furthermore, studies have shown that small LNPs less than 60 nm (especially those in the 30 nm range) exhibit superior targeting properties for dendritic cells in lymph nodes and numerous other immune cells.

[0304] For the above reasons, it is very important to set an appropriate particle size according to the intended use of LNP and to have a manufacturing method for implementing the same. The present invention provides a method for manufacturing LNPs of a size suitable for the intended use by controlling the Reynolds flux and the composition ratio of particles using a microfluidic device.

[0305]

[0306] [Example 9]

[0307] Optimization of the synthesis of size-specific lipid nanoparticles encapsulated with mRNA

[0308] Lipid nanoparticles (mRNA LNPs) encapsulating messenger ribonucleic acid (mRNA) were synthesized using the same microfluidic device used in Example 8. The synthesis was performed using the same method as in Example 8, and mRNA was prepared in a citrate buffer solution at pH 3.0 so that the aqueous solution contained mRNA at an N / P ratio of 6.

[0309] mRNA LNPs were synthesized by applying the size-specific LNP synthesis conditions set in Table 17 of Example 8-3 above. At this time, the mRNAs used were mCherry mRNA with a length of 996 nucleotides and Cas9 mRNA with a length of 4521 nucleotides, and mCherry LNPs and Cas9 LNPs were synthesized by size using two mRNAs with length differences.

[0310]

[0311]

[0312]

[0313] As shown in Table 18 and FIGS. 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, and the PDI at this time was 0.105, 0.054, and 0.052, respectively, indicating that they were synthesized very uniformly. In addition, the similarity in size distribution between mCherry LNPs and LNPs without mRNA (Bare LNP) can be confirmed. The uniformity of mCherry LNPs by size can also be confirmed through the number-weighted size distribution results in FIGS. 42 and 43.

[0314]

[0315]

[0316]

[0317] As shown in Table 19 and FIGS. 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, and the PDI at this time was 0.190, 0.036, and 0.014, respectively. It can be seen that the Cas9 LNPs targeting 60 nm and 90 nm were synthesized very uniformly, and the Cas9 LNP targeting 30 nm also had a PDI of 0.2 or less, indicating that the uniformity of the nanoparticles was still secured. In addition, the similarity in the size distributions of Cas9 LNP and Bare LNP can also be confirmed. The uniformity of Cas9 LNPs by size can also be confirmed through the number-weighted size distribution results in FIGS. 48 and 49.

[0318] The nucleic acid encapsulation efficiency (EE) was calculated using the values ​​obtained by detecting total mRNA and free mRNA based on the fluorescence intensity in each mRNA LNP. ) is the value calculated according to the equation. Similarly, the nucleic acid loading efficiency (LE) is calculated using the values ​​obtained by detecting total mRNA and free mRNA based on the fluorescence intensity in each mRNA LNP. ) is calculated according to the ratio of nucleic acid inclusion rate and nucleic acid endogenous rate. The difference between the ratio of nucleic acid inclusion rate and nucleic acid endogenous rate is that the denominator is different, and there is a difference in whether it is the total mRNA detected or the feeding mRNA.

[0319] In other words, in summary, the nucleic acid encapsulation rate (EE) is an indicator of the quality (mRNA content) of the final product, and the nucleic acid endogenous rate (LE) is an indicator of the efficiency (or yield) with which mRNA from raw materials is loaded into LNPs.

[0320] As shown in Table 18 and Figure 50, the nucleic acid encapsulation rates of 30, 60, and 90 nm mCherry LNPs are 84, 95, and 100%, respectively, and as shown in Table 19 and Figure 51, the nucleic acid encapsulation rates of 30, 60, and 90 nm Cas9 LNPs are 74, 98, and 100%, respectively. Regardless of the type of mRNA, the nucleic acid encapsulation rate of 60 nm or 90 nm mRNA LNPs is over 95%, which means that the lipid mixture and mRNA are mixed very efficiently by the microfluidic device. The nucleic acid encapsulation rate of 30 nm mCherry LNP, which is an LNP with a small space for nucleic acid encapsulation, is still high at 84%, and the nucleic acid encapsulation rate of 30 nm Cas9 LNP is still high at 74%. This means that the lipid mixture and mRNA are efficiently mixed by the microfluidic device. A high nucleic acid encapsulation rate may mean that the probability of encountering impurity problems due to unencapsulated mRNA is very low.

[0321] As shown in Table 18, the nucleic acid internalization rates of mCherry LNPs of 30, 60, and 90 nm were 38, 39, and 44%, respectively, and as shown in Table 19, the nucleic acid internalization rates of Cas9 LNPs of 30, 60, and 90 nm were 44, 29, and 37%, respectively. The high nucleic acid internalization rate of over 30% regardless of target size or mRNA type also indicates that the lipid mixture and mRNA were efficiently mixed by the microfluidic device. In addition, the high nucleic acid internalization rate means less raw material loss.

[0322] That is, the microfluidic device of the present invention is a device that can synthesize mRNA LNPs of a specific size with high uniformity regardless of the mRNA length, and can be said to be an excellent synthesis platform that can develop high-efficiency LNP-based therapeutic agents or vaccines with both stability and functionality for various diseases.

[0323]

[0324] [Example 10]

[0325] Optimization of the Synthesis of Size-Specific Oligonucleotide-Encapsulated Lipid Nanoparticles

[0326] Using the same microfluidic device used in Examples 8 and 9, lipid nanoparticles (Oligo LNP) encapsulating oligonucleotides were synthesized. Synthesis was performed in the same manner as in Example 8 using the conditions in Table 17 above, and the oligonucleotides were prepared in a citrate buffer solution at pH 3.0 so that the aqueous solution contained the oligonucleotides at an N / P ratio of 6.

[0327] Each Oligo LNP was synthesized under the size-specific LNP synthesis conditions set in Example 8-3. At this time, the oligonucleotide used was DNA composed of only 20 (20 mer) thymine (T).

[0328]

[0329]

[0330]

[0331] As shown in Table 20 and FIGS. 52, 53, and 54, the sizes of the Oligo LNPs synthesized with targets of 30, 60, and 90 nm were measured to be 50, 99, and 132 nm, respectively, confirming that they were synthesized in a size that was 20 nm to 40 nm larger than the target size. The PDI at this time was 0.029, 0.053, and 0.036, respectively. Although they were synthesized to be larger than the target size, it can be seen that the oligonucleotides were synthesized very uniformly when encapsulated in the LNPs at all sizes. In addition, the uniformity of the Oligo LNPs by size can also be confirmed through the number-weighted size distribution results in FIGS. 55 and 56.

[0332] Based on the fluorescence intensity in Olilgo LNP, the total oligonucleotide and the free oligonucleotide are detected, and the obtained values ​​are used to calculate the nucleic acid encapsulation rate and nucleic acid internalization rate of the oligonucleotide. The nucleic acid encapsulation rate calculation formula is And the calculation formula for nucleic acid intrinsic rate is am.

[0333] As shown in Table 20 and Figure 57, the nucleic acid encapsulation rates of the 50, 100, and 130 nm Oligo LNPs are 90, 89, and 91%, respectively, and the nucleic acid internalization rates are 53, 48, and 47%, respectively. The nucleic acid encapsulation rate of 90% indicates a low probability of impurity problems due to unencapsulated oligonucleotides, and the nucleic acid internalization rate of over 45% indicates a low loss of raw materials. In other words, it can be confirmed that the lipid mixture and oligonucleotides are efficiently mixed by the microfluidic device.

[0334] Unlike mRNA LNPs, the fact that Oligo LNPs synthesize larger than the target size may be due to the difference in the nucleic acids they contain. Oligonucleotides consist of 20 nucleotides, while mRNAs consist of 1,000 to 5,000 nucleotides, resulting in a 50- to 250-fold difference in length. Comparing the nucleic acid inclusion rates for target sizes of 30, 60, and 90 nm in Tables 18, 19, and 20 above, Oligo LNPs are up to 15%, 20%, and 10% higher than mRNA LNPs. This increase in size appears to be due to the increased amount of shorter oligonucleotides incorporated into the LNPs.

[0335] That is, the microfluidic device of 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 synthesis platform capable of developing highly efficient LNP-based therapeutic agents or vaccines with both stability and functionality for various diseases.

[0336]

[0337] [Example 11]

[0338] Confirmation of intracellular protein expression following size-specific mCherry LNP treatment

[0339] To verify the intracellular mRNA delivery capability of LNPs, protein expression levels in cells treated with mRNA LNPs were determined using fluorescence imaging using a confocal microscope. Human glioblastoma U87MG, human hepatoma HepG2, and human monocyte cell line THP-1 were used to determine mRNA protein expression. To determine intracellular mRNA protein expression, mCherry mRNA, which can emit fluorescence, was used. mCherry LNPs were synthesized according to target sizes (30, 60, and 90 nm) in the same manner as in Example 9.

[0340] In human glioblastoma U87MG cells, the expression level of mCherry protein according to the size of mCherry LNP was observed. U87MG cells were treated with 200 ng of mRNA of each size (30, 60, and 90 nm) of mCherry LNP and then cultured for 24 h in a 5% CO2, 37 oC conditions. After removing the mCherry LNP remaining in the culture medium, the cells were cultured for an additional 24 hours, and the culture medium was replaced to confirm mCherry protein expression. The nucleus and cytoskeleton of the cells were stained. After staining, U87MG cells were fixed by treating with 1 mL of 4% paraformaldehyde solution for 15 minutes. The fluorescence intensity of the mCherry protein expressed in each fixed cell was captured using a confocal microscope using a filter containing an excitation wavelength of 587 nm and an emission wavelength of 610 nm.

[0341] As shown in Figure 58, the expression of mCherry protein in U87MG cells was confirmed by fluorescence imaging. The expression of mCherry protein in the cytoplasmic region of the cells was confirmed by the overlap of the fluorescence of the cytoskeleton (green) and the fluorescence of mCherry (red). When U87MG cells were treated with the same amount of 200 ng of three different sized mCherry LNPs, it was confirmed that mCherry protein expression increased as the LNP size increased. As shown in the quantitative results in Figure 59, the mCherry protein expression was 4-fold and 14-fold higher when treated with 60 nm and 90 nm sized mCherry LNPs, respectively, than when treated with the smallest 30 nm sized mCherry LNP, and the mCherry protein expression was 3-fold higher when treated with the largest 90 nm sized mCherry LNP than when treated with the middle sized 60 nm mCherry LNP.

[0342] HepG2 cells, a human liver cancer cell line, were used to confirm the level of protein expression according to the concentration of mRNA LNP. mCherry LNP was synthesized in two sizes of 60 and 90 nm. After treating HepG2 cells with 20, 200, and 500 ng of each 60 and 90 nm mCherry LNP, they were cultured for 24 hours in a 5% CO2, 37 o C conditions. After removing the mCherry LNP remaining in the culture medium, the cells were cultured for an additional 24 hours, and the culture medium was replaced to confirm mCherry protein expression. The nucleus and cytoskeleton of the cells were stained. After staining, HepG2 cells were fixed by treating with 1 mL of 4% paraformaldehyde solution for 15 minutes. The fluorescence intensity of the mCherry protein expressed in each fixed cell was captured using a confocal microscope using a filter containing an excitation wavelength of 587 nm and an emission wavelength of 610 nm.

[0343] The degree of mCherry protein expression according to the concentration-specific mCherry mRNA treatment in HepG2 cells was confirmed through the fluorescence images in Fig. 60. When mCherry LNPs were treated to HepG2 cells, mCherry protein expression in the cytoplasmic region was confirmed through the overlap of the fluorescence (green) staining the cytoskeleton and the fluorescence of mCherry (red). When mCherry LNPs of various sizes were treated to HepG2 cells, it was confirmed that as the amount of mCherry mRNA treated (20, 200, 500 ng) increased, the amount of mCherry protein expressed in HepG2 increased proportionally, regardless of the size of mCherry LNP (60 nm or 90 nm). In addition, as can be seen in the quantitative results in Fig. 61, when the same amount of mCherry mRNA was treated, the mCherry protein expression rate increased as the LNP size increased.

[0344] Human monocyte cell line THP-1 cells were used to confirm the level of protein expression according to the concentration of mRNA LNP, and mCherry LNP was synthesized in two sizes of 60 and 90 nm. THP-1 cells were treated with 20, 200, and 500 ng of each 60 and 90 nm mCherry LNP, and then cultured for 24 hours in a 5% CO2, 37 oC conditions. After removing the mCherry LNP remaining in the culture medium, the cells were cultured for an additional 24 hours, and the culture medium was replaced to confirm mCherry protein expression. The nucleus and cytoskeleton of the cells were stained. After staining, THP-1 cells were fixed by treating with 1 mL of 4% paraformaldehyde solution for 15 minutes. The fluorescence intensity of the mCherry protein expressed in each fixed cell was captured using a confocal microscope using a filter containing an excitation wavelength of 587 nm and an emission wavelength of 610 nm.

[0345] As shown in Figure 62, the degree of mCherry protein expression in THP-1 cells according to mCherry mRNA treatment at different concentrations was confirmed through images acquired by fluorescence imaging. When mCherry LNP was treated to THP-1 cells, mCherry protein expression in the intracellular cytoplasmic region was confirmed through the overlap of the fluorescence (green) staining the cytoskeleton and the fluorescence of mCherry (red).

[0346] When THP-1 was treated with 60 nm mCherry LNP, it was confirmed that the amount of mCherry protein expressed in THP-1 increased proportionally as the amount of treated mCherry mRNA (20, 200, 500 ng) increased. However, when THP-1 was treated with 90 nm mCherry LNP, it was confirmed that the amount of treated mCherry mRNA (20, 200, 500 ng) and the amount of mCherry protein expressed in THP-1 were not proportional.

[0347] In addition, when comparing the level of mCherry expression according to the size of mCherry LNP (60 nm or 90 nm) when treating the same amount of mCherry mRNA, it was confirmed through the fluorescence image of Fig. 62 and the quantitative results of Fig. 63 that 60 nm mCherry LNP showed higher mCherry expression than 90 nm mCherry LNP.

[0348] In the present invention, we confirmed that the larger the mCherry LNP size, the higher the mCherry protein expression in both HepG2 epithelial cells and U87MG epithelial-like cells. This is because the smaller the LNP size, the higher the PEGylated lipid content (mol%), which may be due to the delay in cellular uptake or endosomal escape of the LNP due to the increase in PEG contained in the LNP.

[0349] In addition, in human immune cells, THP-1, unlike epithelial (like) cells, we confirmed that mCherry protein expression was higher when treated with relatively small 60 nm mCherry LNPs rather than large 90 nm mCherry LNPs. This result suggests that there is an LNP size that can efficiently deliver nucleic acids depending on the cell type to be targeted.

[0350]

[0351] [Example 12]

[0352] Stability verification of nucleic acid-containing LNPs

[0353] In order to verify the stability of nucleic acid-containing LNPs, the size-specific LNP synthesis conditions set in Table 17 of Example 8-3 were applied using the microfluidic device used in Examples 8, 9, and 10, and nucleic acid-containing LNPs were synthesized by preparing a citrate buffer solution at pH 3.0 so that the aqueous solution contained nucleic acid (mRNA or oligonucleotide) at an N / P ratio of 6.

[0354] To confirm the storage stability of mCherry LNP and Cas9 LNP, two sizes were synthesized by mRNA type (mCherry mRNA and Cas9 mRNA) and compared. mCherry LNPs were synthesized in sizes of 60 and 90 nm, and Cas9 LNPs were synthesized in sizes of 30 and 90 nm. The synthesized mCherry LNP and Cas9 LNP were stored in PBS containing 1% trehalose, and the storage stability for 0, 7, 14, and 28 days under 4℃ refrigerated conditions was confirmed through DLS results (average size, size at the highest peak of the number-weighted size distribution, percentage at the highest peak of the volume-weighted size distribution, PDI), nucleic acid inclusion rate, and nucleic acid internalization rate.

[0355] As shown in Figures 64 and 65, the 60 nm mCherry LNPs maintained the average size and the highest peak size of the number-weighted size distribution for 28 days under 4℃ refrigeration conditions. Accordingly, the percentage (volume %) of the highest peak of the volume-weighted size distribution, which indicates the major size fraction in the size distribution within the nanoparticles, did not change significantly within about 20%, and the PDI, which indicates uniformity, was also maintained 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 to 73% on day 28. The nucleic acid internalization rate was maintained at about 20% for 28 days.

[0356] As shown in Figures 66 and 67, 90 nm mCherry LNPs showed similar size results to 60 nm mCherry LNPs under 4℃ refrigerated conditions. It was confirmed that the average size and the highest peak size of the number-weighted size distribution were maintained without significant change for 28 days. Accordingly, the percentage at the highest peak of the volume-weighted size distribution, which indicates the fraction of the major size in the size distribution within the nanoparticles, also remained at 15% to 20% and did not change significantly. The PDI, which indicates uniformity, was also maintained within 0.1, confirming that it was very uniform even after 28 days. The nucleic acid encapsulation rate showed a different pattern from that of 60 nm mCherry LNPs, and was confirmed to be maintained at approximately 80% for 28 days, and the nucleic acid internalization rate was maintained at approximately 40% for 28 days.

[0357] As shown in Figures 68 and 69, the 30 nm Cas9 LNPs maintained the average size and the highest peak size of the number-weighted size distribution without significant change for 28 days under 4℃ refrigerated conditions. Accordingly, the percentage (volume %) at the highest peak of the volume-weighted size distribution, which indicates the fraction of the major size in the size distribution within the nanoparticles, also remained around 15%. The PDI, which indicates uniformity, was measured to be 0.22. It can be confirmed that the uniformity was also maintained by maintaining the 0.2 range without significant change in the PDI for up to 28 days. Since a PDI of 0.7 or higher is generally considered to be poor quality nanoparticles, a PDI corresponding to 0.2 still means that they are uniform. It was confirmed that the nucleic acid encapsulation rate was maintained at 70% to 80% for 28 days despite the small size of the LNP, and the nucleic acid internalization rate was maintained at approximately 40% for 28 days.

[0358] As shown in Figures 70 and 71, the 90 nm Cas9 LNPs maintained the average size and the highest peak size of the number-weighted size distribution without significant change for 28 days under 4℃ refrigeration conditions. Accordingly, the percentage (volume %) at the highest peak of the volume-weighted size distribution, which indicates the fraction of the major size in the size distribution within the nanoparticles, also remained at 15% to 20% and did not significantly change, and the PDI, which indicates uniformity, was also maintained at around 0.1, confirming that it was very uniform even after 28 days. The nucleic acid encapsulation rate was confirmed to be maintained at around 80% for 28 days, and the nucleic acid internalization rate was maintained at around 30% for 28 days.

[0359] When mRNA LNPs were synthesized using the microfluidic device of the present invention, size stability and LNP uniformity were maintained for 28 days under refrigerated conditions, regardless of mRNA length or LNP size. Nucleic acid encapsulation rates decreased for one of the four mRNA LNP types (60 nm mCherry LNP), suggesting the need for optimization of storage conditions.

[0360] Oligo LNPs were synthesized for three sizes using the size-specific LNP synthesis conditions set in Table 17 of Example 8-3. As in Example 10, they were synthesized to be larger than the target size. The sizes of the Oligo LNPs synthesized with targets of 30, 60, and 90 nm were measured to be 50, 100, and 130 nm, respectively, which means that they were synthesized to be 20 nm to 40 nm larger than the target size. The synthesized Oligo LNPs were stored in PBS containing 1% trehalose, and the storage stability for 0, 7, 14, and 28 days under 4℃ refrigerated conditions was confirmed through DLS results (average size, size at the highest peak of the number-weighted size distribution, percentage (volume %) at the highest peak of the volume-weighted size distribution, PDI), nucleic acid encapsulation rate, and nucleic acid internalization rate.

[0361] As shown in Figures 72 and 73, the 50 nm Oligo LNP (target size: 30 nm) was confirmed to maintain the average size and the highest peak size of the number-weighted size distribution without significant changes for 28 days under 4℃ refrigeration conditions. Accordingly, the percentage (volume %) at the highest peak of the volume-weighted size distribution, which indicates the fraction of the major size in the size distribution within the nanoparticles, also maintained 20%, and the PDI, which indicates uniformity, was also maintained at around 0.1, confirming that it was very uniform even after 28 days. The nucleic acid encapsulation rate was confirmed to be maintained at 80% or more for 28 days, and the nucleic acid internalization rate was maintained at around 50% for 28 days.

[0362] As shown in Figures 74 and 75, the 100 nm Oligo LNP (target size: 60 nm) was confirmed to maintain the average size and the highest peak size of the number-weighted size distribution without significant changes for 28 days under 4℃ refrigeration conditions. Accordingly, the percentage (volume %) at the highest peak of the volume-weighted size distribution, which indicates the fraction of the major size in the size distribution within the nanoparticles, also maintained 20%, and the PDI, which indicates uniformity, was also maintained at around 0.1, confirming that it was very uniform even after 28 days. The nucleic acid encapsulation rate was confirmed to be maintained at 80% or more for 28 days, and the nucleic acid internalization rate was maintained at around 50% for 28 days.

[0363] As shown in Figures 76 and 77, it was confirmed that the 130 nm Oligo LNP (target size: 90 nm) maintained the average size and the highest peak size of the number-weighted size distribution without significant changes for 28 days under 4℃ refrigerated conditions. Accordingly, the percentage (volume %) at the highest peak of the volume-weighted size distribution, which indicates the fraction of the major size in the size distribution within the nanoparticles, also maintained 15% to 20%, and the PDI, which indicates uniformity, was also maintained at 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 be maintained at 80% on day 28, and the nucleic acid internalization rate was maintained at around 50% for 28 days.

[0364] When oligo LNPs were synthesized using the microfluidic device of the present invention, size stability and uniformity of LNPs were maintained for 28 days under refrigerated conditions, regardless of LNP size. It was also confirmed that there was no significant change in nucleic acid encapsulation rate or nucleic acid internalization rate.

[0365] Based on the above-described exemplary results, by synthesizing LNPs of various sizes under three optimized conditions using a microfluidic device, LNPs of desired sizes can be synthesized very uniformly with a high mixing efficiency of 0.95 or higher, and nucleic acids of various lengths can be made to have high nucleic acid encapsulation and nucleic acid internalization rates. Therefore, the optimized LNP synthesis technology of the present invention is expected to enable the selection and development of LNPs of appropriate sizes with high uniformity according to various indications for treatment or prevention.

Claims

1. Including inflow channel, mixing channel and outflow channel, There are two or more of the above inflow channels, The above mixing channel comprises micro-pillars, The fluids entering different inlet channels flow in different directions, The lipid mixture flows into one inlet channel, and the nucleic acids flow into the other inlet channel. The lipid mixture comprises a sterol and further comprises at least one selected from the group consisting of an ionized lipid, a helper lipid and a pegylated lipid, The above micro-columns are arranged in one or more rows in the direction of the fluid flow, In one column, micro-columns are arranged one or more in a direction different from the direction of fluid flow. The above-mentioned plurality of micro-pillars are not arranged in a row in the direction of the fluid flow, but are arranged so as to partially or completely cover the gaps between micro-pillars arranged in adjacent rows when viewed in the direction of the fluid flow. The Reynolds number for the flow of fluid in the above mixing channel is 25 to 100, A microfluidic device for producing lipid nanoparticles, wherein the molar ratio of sterol to the lipid mixture is 0.3 to 0.

86.

2. A microfluidic device for manufacturing lipid nanoparticles, characterized in that in the first paragraph, the lipid mixture flows in the same direction as the flow of the fluid, and the nucleic acid flows in a direction different from the flow of the fluid.

3. A microfluidic device for manufacturing lipid nanoparticles, characterized in that in the second paragraph, the lipid mixture is introduced in the same direction as the flow of the fluid, and the nucleic acid is introduced in a direction orthogonal to the flow of the fluid.

4. A microfluidic device for manufacturing lipid nanoparticles, characterized in that in paragraph 1, the micro-pillars are connected or not connected to the walls forming the mixing channel.

5. A microfluidic device for manufacturing lipid nanoparticles, characterized in that the size of the lipid nanoparticles in paragraph 1 is controlled by the molar ratio of the lipid mixture and the sterol.

Images