Lipid nanoparticle encapsulation of large sized RNA

Abstract

To provide a method of producing lipid-encapsulated RNA nanoparticles.SOLUTION: A production method according to the present invention includes the steps of: flowing an aqueous solution comprising an RNA through a first tube having a first inner diameter (ID), the RNA including about 6,000 to about 13,000 nucleotides; flowing an ethanol solution including lipids through a second tube having a second inner diameter (ID), at a flow rate of about 0.2 to about 1 time relative to the aqueous solution through the first tube, the lipids including a cationic lipid; and mixing the ethanol solution with the aqueous solution, the first ID and second ID and flow rates through the first tube and second tube are selected to produce a shear force sufficiently low to preserve integrity of the RNA, the mixing producing an output solution flowing in the first tube including a turbulent flow of the RNA and the lipids in about 10% to 75% ethanol, and the lipid-encapsulated RNA nanoparticles having a bilayer structure.SELECTED DRAWING: Figure 1

Description

[Background technology]

[0001] Lipids are used as materials for ribonucleic acid (RNA) due to their ability to form lipid nanoparticles that encapsulate RNA for delivery to target cells upon parenteral administration (Zimmermann, 2006, Nature, doi:10.1038 / nature04688).

[0002] Various methods for producing lipid-encapsulated RNA nanoparticles are known. For example, WO2001 / 005373 discloses a technique for preparing lipid-encapsulated RNA nanoparticles using a static mixer that creates a turbulent environment, followed by an ethanol injection process in which the vesicle formers are combined with therapeutic molecules. US2004 / 0142025 discloses a technique for forming lipid-encapsulated RNA nanoparticles using non-turbulent mixing and a series of continuous stepwise dilutions. US6,843,942 discloses a non-turbulent mixing method for forming particles by spraying lipids through an orifice in an organic solution pipe onto nucleic acid in an aqueous solution flowing through the orifice. US9,005,654 discloses encapsulating siRNA in lipid nanoparticles (LNPs) using turbulent mixing, whereby lipids and RNA flowing in opposite directions enter a T-shaped mixing chamber at approximately the same speed from opposite arms to produce a 45-60% ethanol solution containing vesicles, which is then collected and further diluted (direct dilution method).US9,404,127 discloses that the majority of LNPs produced by the direct dilution method have a nonlamellar morphology, i.e., a non-bilayer structure.

[0003] The application of conventional methods to form lipid-encapsulated RNA nanoparticles from large RNAs presents challenges that present difficulties. For example, one such problem arises from the forces that arise in RNA during LNP formation, causing the structural integrity of RNA to be compromised. Therefore, there is a need for improved processes and devices for formulating lipid-encapsulated RNA nanoparticles in the context of large RNA sequences. Such methods must be suitable for scale-up while targeting desired particle size and polydispersity. The embodiments herein address one or more of these problems and other issues recognized by those skilled in the art. Summary of the Invention

[0004] In some aspects, embodiments herein provide a method for preparing a nucleic acid sequence comprising: a) flowing an aqueous solution containing RNA through a first tube having an inner diameter (ID) of about 0.01 inches to about 0.08 inches, wherein the pH of the aqueous solution ranges from about 3.0 to about 4.5, with an optional NaCl concentration of up to about 300 mM, and the RNA comprises about 6,000 to about 13,000 nucleotides; and b) flowing the first tube through an ID of about 0.01 inches to about 0.08 inches at a flow rate that is about 0.2 to about 1 times the flow rate of the aqueous solution through the first tube. a) flowing an ethanol solution containing lipids into a second tube having an ID of 0.04 inches, wherein the lipids comprise cationic lipids; and b) mixing the ethanol solution with the aqueous solution, wherein the mixing produces an output solution flowing through the first tube containing approximately 10% to 75% ethanol (v / v) and containing a turbulent flow of the RNA and the lipids, wherein the lipid-encapsulated RNA nanoparticles have a bilayer structure.

[0005] In some aspects, embodiments herein provide a method for producing lipid-encapsulated RNA nanoparticles, comprising: a) flowing an aqueous solution containing RNA through a first tube having a first inner diameter (ID), wherein the RNA comprises about 6,000 to about 13,000 nucleotides; b) flowing an ethanol solution containing lipids through a second tube having a second inner diameter (ID) at a flow rate that is about 0.2 to about 1 times the flow rate of the aqueous solution through the first tube, wherein the lipids comprise cationic lipids; and c) mixing the ethanol solution with the aqueous solution, wherein the flow rates through the first and second IDs and the first and second tubes are selected to produce shear forces that are low enough to protect the integrity of the RNA, and wherein the mixing produces an output solution flowing through the first tube that is about 10% to 75% ethanol (v / v) and comprises a turbulent flow of the RNA and the lipids, and wherein the lipid-encapsulated RNA nanoparticles have a bilayer structure. [Brief explanation of the drawings]

[0006] [Figure 1]

[0023] Figure 1 shows a flow chart diagram for one embodiment of a process for producing lipid nanoparticles. Lipids are dissolved in ethanol, and RNA is dissolved in an acidic aqueous solution (e.g., citrate buffer), which are both filter-sterilized. The solutions are mixed to form particles using the process described herein, and PDI and particle size (PS) are analyzed. The particles are concentrated and purified by tangential flow filtration (TFF) to remove ethanol and unbound RNA, and PDI and PS are again monitored. The particle concentration is then adjusted according to the measured total RNA concentration. The particles are filter-sterilized, filled, polished, and frozen. [Figure 2]This figure shows an apparatus for producing lipid-encapsulated RNA nanoparticles. An aqueous solution containing RNA is pumped through tubing by an HPLC pump, and an organic solution containing lipids is pumped through separate tubing by an HPLC pump. The organic solution can be pumped into the aqueous solution at a 90-degree angle within the mixing zone. The outlet tubing transports the mixed lipid-RNA solution, connected to an outlet by polypropylene tubing, which meets the dilution buffer at a 45-degree angle within the dilution zone. The tubing, which meets the dilution buffer at a 45-degree angle within the dilution zone, is serially diluted and can include one, two, three, or four dilution zones, each at a 45-degree angle. After the dilution process, the diluted particles can be collected in a stainless steel-lined container maintained at 15-20°C. The particles can be further processed by tangential flow filtration using peristaltic, diaphragm, or centrifugal pumps. [Figure 3] A more detailed view of the mixing module is shown. Nucleic acids in a buffer solution are transported through the input arm of a first stainless steel tube. Lipids in ethanol (or other suitable organic solvent / solvent mixture) are transported through a second stainless steel tube attached perpendicular to the first tube. Holes in the wall of the first tube allow liquid to be transported from the second tube to the inside of the first tube. The resulting lipid-encapsulated RNA nanoparticles exit through the output arm of the first tube. [Figure 4] 1 shows the adjusted mean fluorescence intensity (MFI) for various dose levels of lipid nanoparticle formulations, corresponding to the amount of anti-COVID19 spike protein antibodies generated in response to lipid nanoparticle administration in mice, as described in Example 13. DETAILED DESCRIPTION OF THE INVENTION

[0007] It is understood that various configurations of the subject technology will be readily apparent to those skilled in the art from this disclosure, and various configurations of the subject technology have been shown and described by way of example. As will be understood, the subject technology is capable of other and different configurations, and its several details are capable of modification in various other respects, all without departing from the scope of the subject technology. Accordingly, the summary, drawings, and detailed description are to be regarded as illustrative in nature, and not as restrictive.

[0008] Embodiments herein provide a process for encapsulating large RNAs (e.g., self-replicating RNAs) in lipid nanoparticles. For example, large mRNAs can have sizes on the order of 6,000 to approximately 15,000 nucleotides. As disclosed herein, these large nucleotides were found to be incompatible with typical LNP formation processes. Using the LNP process typically used to prepare mRNA-LNP compositions, we attempted to create LNPs encapsulating self-replicating RNAs. The following procedure is typical for RNAs containing approximately 1,000 to approximately 5,000 nucleotides. Bulk formulations are prepared by mixing an ethanolic solution of lipids with an aqueous solution of an RNA drug substance or other small mRNA, as outlined below: Lipid excipients (cationic lipids, phospholipids, cholesterol (Chol), and PEG-lipid conjugates) are dissolved in ethanol and filtered through a 0.2 μm polyethersulfone (PES) filter. An aqueous solution of mRNA is prepared in citrate buffer (pH 4.0) and then filtered through a 0.2 μm PES filter. The mRNA solution was then mixed with an ethanolic solution in a stainless steel mixing module. The nanoparticles thus formed were stabilized by serial dilution with phosphate buffer (pH 6.0) followed by HEPES buffer (pH 8.0). The nanoparticle formulation was then subjected to ultrafiltration and diafiltration (UF / DF) by tangential flow filtration (TFF) using a modified PES hollow fiber membrane (100 kDa MWCO (molecular weight cut-off)) and HEPES buffer (pH 8.0). After UF / DF, the formulation was filtered through a 0.2 μm PES filter and stored at 2–8°C until filling. Next, in-process mRNA concentration analysis was performed. The concentration of the formulation was adjusted to the final target mRNA concentration (0.2 mg / mL) and then filtered through a 0.2 μm PES sterilizing grade filter. After sterile filtration, the bulk product was aseptically filled into glass bottles, stoppered, capped, and frozen at -70±10°C.

[0009] Using this process, the resulting LNPs containing larger RNAs (greater than approximately 6,000 nucleotides) had poor size, dispersion, and encapsulation efficiency. Furthermore, a significant portion of these larger RNA structures was found to be degraded during the formulation process, which was hypothesized to be due to the shear forces generated by the specific process pressure and flow rate used in this method. It was hypothesized that such shear forces could be controlled by varying the flow rate and tubing size. However, mixing conditions and operating pressure alone were not considered the only variables necessarily resulting in the requisite LNP-wrapped RNA. Adjusting mixing conditions to reduce shear forces could result in conditions that were not sufficient for LNP formulation. Therefore, the challenge presented by large RNAs was considered complex, as other parameters were considered to be factors in successfully formulating LNP-wrapped large RNAs, including, but not limited to, buffer and salt concentrations, RNA and lipid concentrations, system pH, and overall backpressure.

[0010] Embodiments herein provide a workable solution for forming LNP-encapsulated large RNA. Among the advantages of the methods disclosed herein are: (1) the large variability in composition allowed by different lipid components, such as a range of phospholipid / helper lipid concentrations, cationic lipid concentrations and cholesterol, and RNA size; (2) transferability between modules with different scalability for small-, medium-, and large-scale production; and (3) the ability to scale up production while maintaining reduced batch volumes.

[0011] In embodiments, the method comprises the steps of: a) flowing an aqueous solution containing RNA through a first tube having an inner diameter (ID) of about 0.01 inches to about 0.08 inches, wherein the pH of the aqueous solution ranges from about 3.0 to about 4.5, with an optional NaCl concentration of up to about 300 mM, and the RNA comprises about 6,000 to about 13,000 nucleotides; and b) flowing the aqueous solution through an ID of about 0.01 inches to about 0.04 inches at a flow rate that is about 0.2 to about 1 times the flow rate of the aqueous solution through the first tube. a) flowing an ethanol solution containing lipids into a second tube having a first end D, wherein the lipids include cationic lipids; and b) mixing the ethanol solution with the aqueous solution, wherein the mixing produces an output solution flowing through the first tube containing approximately 10% to 75% ethanol (v / v) and containing a turbulent flow of the RNA and the lipids, and wherein the lipid-encapsulated RNA nanoparticles have a bilayer structure.

[0012] In one embodiment, a method for producing lipid-encapsulated RNA nanoparticles is provided, comprising the steps of: a) flowing an aqueous solution containing RNA through a first tube having a first inner diameter (ID), the RNA comprising about 6,000 to about 13,000 nucleotides; b) flowing an ethanol solution containing lipids through a second tube having a second inner diameter (ID) at a flow rate that is about 0.2 to about 1 times the flow rate of the aqueous solution through the first tube, the lipids comprising cationic lipids; and c) mixing the ethanol solution with the aqueous solution, wherein the flow rates through the first and second IDs and the first and second tubes are selected to produce shear forces low enough to protect the integrity of the RNA, and the mixing produces an output solution flowing through the first tube that is about 10% to 75% ethanol (v / v) and comprises a turbulent flow of the RNA and the lipids, and wherein the lipid-encapsulated RNA nanoparticles have a bilayer structure.

[0013] In an embodiment, the mixing comprises flowing the ethanol solution and the aqueous solution through a mixing module comprising a second tube perpendicularly connected to the first tube.

[0014] In an embodiment, the mixing comprises passing the ethanol solution and the aqueous solution through a multi-inlet vortex mixer.

[0015] In embodiments, the concentration of RNA in the aqueous solution ranges from about 85 micrograms / mL to about 2100 micrograms / mL, in embodiments, the range is from about 85 to about 200 micrograms / mL, or from about 200 to about 500, or from about 500 to about 800, or from about 800 to about 1000, or from about 1000 to about 1500, or from about 1500 to about 2100 micrograms / mL, including subranges and fractions thereof.

[0016] In embodiments, the concentration of lipid in the ethanol solution ranges from about 5.0 mg / mL to about 125 mg / mL, and in embodiments, the range is from about 5.0 to about 30 mg / mL, or from about 30 to about 60 mg / mL, or from about 60 to about 90 mg / mL, or from about 90 to about 125 mg / mL, including subranges and fractions thereof.

[0017] In embodiments, the aqueous solution is pumped through a first tube by a first pump and the ethanol solution is pumped through a second tube by a second pump at a back pressure of about 200 psi or less, hi some embodiments, the back pressure is about 195 psi or less, or about 190 psi or less, or about 180 psi or less.

[0018] In embodiments, the first tube has an ID in the range of about 0.01 inches to about 0.08 inches, and the second tube has an ID in the range of about 0.01 inches to about 0.04 inches. In embodiments, the first tube has an ID in the range of about 0.02 inches to about 0.03 inches, and the second tube has an ID in the range of about 0.01 inches to about 0.02 inches. In embodiments, the first tube has an ID of about 0.02 inches, and the second tube has an ID of about 0.01 inches. In embodiments, the first tube has an ID of about 0.03 inches, and the second tube has an ID of about 0.01 inches. Such measurements include these subranges and fractions thereof.

[0019] In embodiments, the aqueous solution is pumped at a flow rate ranging from about 40 mL / min to about 375 mL / min, or from about 40 to about 80 mL / min, or from about 80 to about 120 mL / min, or from about 120 to about 160 mL / min, or from about 160 to about 200 mL / min, or from about 200 to about 240 mL / min, or from about 240 to about 280 mL / min, or from about 280 to about 320 mL / min, or from about 320 to about 375 mL / min, including subranges and fractions thereof.

[0020] In embodiments, the ethanol solution is pumped at a flow rate ranging from about 10 mL / min to about 75 mL / min, hi some embodiments, the flow rate ranges from about 10 to about 30 mL / min, or from about 30 to about 50, or from about 50 to about 75 mL / min, including subranges and fractions thereof.

[0021] In embodiments, the aqueous solution, the ethanol solution, and the output solution are maintained at a temperature ranging from about 10°C to about 25°C.

[0022] In embodiments, the method may further include pumping a first dilution buffer and introducing the dilution buffer into the output solution, thereby mixing the dilution buffer with the output solution to create the first diluted output solution.

[0023] In embodiments, the method may further include pumping a second dilution buffer into the first diluted output solution to form a final diluted output solution, wherein there is a delay between pumping the first dilution buffer and pumping the second dilution buffer.

[0024] In embodiments, the delay is from about 0.1 to about 30 seconds, and the delay is created by the length of the tubing. In some embodiments, there is no delay. In some embodiments, the delay is from 0.1 to about 5 seconds, or from about 5 to about 10 seconds, or from about 10 to about 15 seconds, or from about 15 to about 20 seconds, or from about 20 seconds to about 30 seconds, including subranges and fractions thereof.

[0025] In embodiments, the first dilution buffer comprises a buffer having a pH of about 5.5 to about 7.0, and optionally a sodium chloride concentration of up to about 100 mM. For example, the first dilution buffer may comprise up to about 20 mM Tris buffer, or 40 mM to 90 mM phosphate buffer, or 20 mM to 50 mM HEPES buffer, or 45 mM pH 6.5 phosphate buffer.

[0026] In embodiments, the first dilution buffer may optionally include a sodium chloride concentration of up to about 50 mM.

[0027] In embodiments, the second dilution buffer may comprise a buffering agent having a pH of about 7.4 to 8.0, and optionally, a sodium chloride concentration of up to about 100 mM.

[0028] In embodiments, the second dilution buffer may optionally include a sodium chloride concentration of up to about 50 mM.

[0029] In embodiments, the second buffer contains up to about 15% w / v sucrose. In some embodiments, sucrose is present at about 12% w / v, or up to about 10%, or up to about 8%, or up to about 5%, or up to about 1%. In some embodiments, sucrose is absent.

[0030] In embodiments, the second buffer contains up to about 0.5% w / v antioxidant. In some embodiments, antioxidant is absent. In some embodiments, antioxidant is present at up to about 0.4% w / v, or about 0.3%, or about 0.2%, or about 0.1% w / v.

[0031] In embodiments, the second buffer comprises up to 20 mM of a chelating agent.

[0032] In embodiments, the first diluted output solution comprises about 1.0% to about 10.0% ethanol. In some embodiments, the first diluted output solution comprises about 2% to about 8% ethanol, or about 3% to about 7% ethanol, including subranges and fractions thereof.

[0033] In embodiments, the first dilution buffer is pumped at a flow rate of about 80 mL / min to about 900 mL / min. In some embodiments, the flow rate is about 80 mL / min to about 150 mL / min, or about 150 to about 200 mL / min, or about 200 to about 250 mL / min, or about 250 to about 300 mL / min, or about 300 to about 400 mL / min, or about 400 to about 500 mL / min, or about 500 to about 600 mL / min, or about 600 to about 700 mL / min, or about 700 to about 900 mL / min, including subranges and fractions thereof.

[0034] In embodiments, the second dilution buffer is pumped at a flow rate of about 240 mL / min to about 5400 mL / min. In some embodiments, the flow rate is about 240 to about 500 mL / min, or about 500 to about 1000 mL / min, or about 1000 to about 1500 mL / min, or about 1500 to about 2000 mL / min, or about 2000 to about 2500 mL / min, or about 2500 to about 3000 mL / min, or about 3000 to about 3500 mL / min, or about 3500 to about 4000 mL / min, or about 4000 to about 4500 mL / min, or about 4500 to about 5000 mL / min, or about 5000 to about 5500 mL / min, including subranges and fractions thereof.

[0035] In embodiments, the output solution has a total flow rate in the range of about 120 mL / min to about 300 mL / min.

[0036] In embodiments, the cationic lipid has the structure of Formula I: [ka] or a pharmaceutically acceptable salt or solvate thereof, wherein: R 5 and R 6 are each independently a straight-chain or branched C1-C 31 Alkyl, C2-C 31 Alkenyl, or C2-C 31 selected from the group consisting of alkynyl, and cholesteryl; L 5 and L 6 are each independently a straight chain C1-C 20 Alkyl and C2-C 20 alkenyl, X 5 is -C(O)O- or -OC(O)-, X 6 is -C(O)O- or -OC(O)-, X 7 is S or O, L 7 is absent or lower alkyl, R 4 is a straight-chain or branched C1-C6 alkyl, and R 7 and R 8 are each independently selected from the group consisting of hydrogen and straight-chain or branched C1-C6 alkyl.

[0037] In embodiments, the lipid-encapsulated RNA nanoparticles have an average particle size ranging from about 50 nm to about 120 nm, hi some embodiments, the average particle size is about 60 to about 120 nm, about 70 to about 120 nm, or about 70 to 90 nm, including subranges and fractions thereof.

[0038] In embodiments, the polydispersity of the lipid-encapsulated RNA nanoparticles does not exceed about 0.2.

[0039] In embodiments, the lipid portion of the lipid-encapsulated RNA nanoparticles further comprises one or more agents selected from the group consisting of helper lipids, cholesterol, and PEG-lipid conjugates.

[0040] In embodiments, the RNA is self-replicating RNA.

[0041] In embodiments, the method may further comprise lyophilizing the final diluted output solution.

[0042] The above-described embodiments may be combined in any combination to represent exemplary embodiments, which will be more fully understood hereinafter and further in the embodiments that follow.

[0043] Lipid-based formulations Therapies based on the intracellular delivery of nucleic acids to target cells face both extracellular and intracellular barriers. Naked nucleic acid materials cannot be easily administered systemically due to their toxicity, poor serum stability, rapid renal clearance, reduced uptake by target cells, phagocytic uptake, and ability to activate immune responses—all factors that preclude clinical development. When exogenous nucleic acid materials (e.g., mRNA) enter the human biological system, they are recognized as foreign pathogens by the reticuloendothelial system (RES) and are cleared from the blood circulation before they have a chance to encounter target cells, either inside or outside the vascular system. The half-life of naked nucleic acids in the bloodstream has been reported to be approximately several minutes (Kawabata K, Takakura Y, Hashida MPharm Res. 1995 Jun;12(6):825-30). Chemical modification and appropriate delivery methods can reduce uptake by RES and protect nucleic acids from degradation by ubiquitous nucleases, thereby increasing the stability and efficacy of nucleic acid-based therapeutics. In addition, RNA or DNA are anionic hydrophilic polymers that are not favorably taken up by cells, and they are also anionic on the surface. Therefore, the success of nucleic acid-based therapeutics depends largely on the development of vehicles or vectors that can efficiently and effectively deliver genetic material to target cells and achieve sufficient expression levels in vivo with minimal toxicity.

[0044] Furthermore, upon internalization into target cells, nucleic acid delivery vectors face challenges due to intracellular barriers, including endosomal entrapment, lysosomal degradation, nucleic acid removal from the vector, translocation across the nuclear membrane (for DNA), and release into the cytoplasm (for RNA). Thus, the success of nucleic acid-based therapeutics depends on the ability of the vector to deliver nucleic acids to target sites inside the cell and obtain sufficient levels of the desired activity, such as gene expression.

[0045] While some gene therapy approaches have successfully utilized viral delivery vectors (e.g., AAV), lipid-based formulations are increasingly recognized as one of the most promising delivery systems for RNA and other nucleic acid compounds due to their biocompatibility and ease of large-scale manufacturing. One of the most significant advances in lipid-based nucleic acid therapeutics occurred in August 2018, when patisiran (ALN-TTR02) became the first siRNA therapeutic approved by both the U.S. Food and Drug Administration (FDA) and the European Commission (EC). ALN-TTR02 is an siRNA formulation based on so-called stable nucleic acid lipid particle (SNALP) transfection technology. Despite the success of patisiran, delivery of nucleic acid therapeutics, including mRNA, using lipid formulations remains under development.

[0046] Some art-recognized lipid formulation delivery vehicles for nucleic acid therapeutics, according to various embodiments, include polymer-based carriers such as polyethyleneimine (PEI), lipidoid-containing formulations, lipid nanoparticles and liposomes, nanolysosomes, ceramide-containing nanolysosomes, multivesicular liposomes, proteolysosomes, both natural and synthetic delivery exosomes, natural, synthetic, and semi-synthetic lamellar bodies, nanoparticles, micelles, and emulsions.

[0047] These lipid formulations vary in structure and composition, and as can be expected in this rapidly evolving field, several different terms are used in the art to describe one kind of delivery vehicle.At the same time, the term lipid formulation is frequently conflated throughout scientific literature, and this inconsistent use causes confusion about the exact meaning of some terms of lipid formulation.Among several possible lipid formulations, liposome, cationic liposome and lipid nanoparticle are particularly described and defined herein for the purpose of this disclosure.

[0048] Liposomes Conventional liposomes are vesicles consisting of at least one bilayer and an internal aqueous compartment. The liposome bilayer is typically formed by amphiphilic molecules, such as synthetic or naturally occurring lipids, which contain spatially separated hydrophilic and hydrophobic domains (Lasic, Trends Biotechnol., 16:307-321, 1998). The liposome bilayer can also be formed by amphiphilic polymers and surfactants (e.g., polymersomes, niosomes, etc.). These generally exist as spherical vesicles and can range in size from 20 nm to several microns. Liposome formulations can be prepared as colloidal dispersions or lyophilized to reduce stability risks and improve shelf life for liposome-based drugs. Methods for preparing liposome compositions are known in the art and within the skill of those in the art.

[0049] Liposomes with only one bilayer are called unilamellar, while liposomes with two or more bilayers are called multilamellar. The most common types of liposomes are small unilamellar vesicles (SUVs), large unilamellar vesicles (LUVs), and multilamellar vesicles (MLVs). In contrast to liposomes, lysosomes, micelles, and reverse micelles are composed of a single layer of lipids. While liposomes are generally considered to have a single internal compartment, some formulations can be multivesicular liposomes (MVLs), which are composed of multiple discontinuous internal aqueous compartments separated by several lipid bilayers that do not share a common center.

[0050] Considering that liposomes are essentially analogs of biological membranes, liposomes have long been recognized as drug delivery vehicles due to their excellent biocompatibility and can be prepared from both natural and synthetic phospholipids (Int J Nanomedicine. 2014;9:1833-1843). When used as drug delivery vehicles, hydrophilic solutes dissolved in the liposome core cannot readily pass through the hydrophobic membrane of the bilayer, while hydrophobic compounds associate with the bilayer. Therefore, liposomes can be loaded with hydrophobic and / or hydrophilic molecules. When liposomes are used to deliver nucleic acids, such as RNA, the nucleic acid is contained within the liposome compartment in the aqueous phase.

[0051] Cationic Liposomes Liposomes can be composed of cationic, anionic, and / or neutral lipids. As an important subclass of liposomes, cationic liposomes are liposomes made entirely or partially from positively charged lipids, or more specifically, lipids containing both cationic and lipophilic moieties. In addition to the general characteristics outlined above for liposomes, the positively charged moieties of the cationic lipids used in cationic liposomes offer several advantages and some unique structural traits. For example, because the lipophilic moieties of cationic lipids are hydrophobic, they themselves separate from the aqueous interior of the liposome and associate with other nonpolar and hydrophobic moieties. Conversely, the cationic moieties associate with the aqueous medium and, more importantly, with polar molecules and species that can form complexes in the aqueous interior of the cationic liposome. For these reasons, cationic liposomes are increasingly being explored for use in gene transfer due to their ability to target negatively charged nucleic acids through electrostatic interactions, resulting in complexes that offer biocompatibility, low toxicity, and the potential for large-scale production necessary for in vivo clinical use. Cationic lipids suitable for use in cationic liposomes are listed herein below.

[0052] lipid nanoparticles In contrast to liposomes and cationic liposomes, lipid nanoparticles (LNPs) are structures containing a single monolayer or bilayer of lipids that encapsulate compounds in a solid phase. Thus, unlike liposomes, lipid nanoparticles do not have an internal aqueous or other liquid phase; instead, lipids from the bilayer or monolayer shell directly complex with the internal compound, encapsulating a solid core within. Lipid nanoparticles are typically spherical vesicles with a relatively uniform dispersion of shape and size. While the scientific literature varies in terms of what size constitutes a lipid particle as a nanoparticulate material, there is some agreement that lipid nanoparticles can have diameters ranging from 10 nm to 1000 nm. However, they are more commonly considered to be less than 120 nm or even 100 nm.

[0053] For lipid nanoparticle nucleic acid delivery systems, the lipid shell can be formulated to include ionizable cationic lipids that can complex with and associate with the negatively charged backbone of nucleic acid core.Ionizable cationic lipids with apparent pKa values ​​below about 7 have the advantage that the cationic lipids can be complexed with the negatively charged backbone of nucleic acid, and when positively charged, can be loaded into lipid nanoparticles at a pH value below the pKa of the ionizable lipid.Then, at physiological pH values, the lipid nanoparticles adopt a relatively neutral exterior, allowing the particles to significantly increase their circulation half-life after intravenous administration.In the context of nucleic acid delivery, lipid nanoparticles offer many advantages over other lipid-based nucleic acid delivery systems, including high nucleic acid encapsulation efficiency, strong transfection, improved tissue penetration and delivery of therapeutic agents, and low levels of cytotoxicity and immunogenicity.

[0054] Prior to the development of lipid nanoparticle delivery systems for nucleic acids, cationic lipids were extensively investigated as synthetic materials for delivering nucleic acid drugs. In these early efforts, nucleic acids were condensed with cationic lipids after mixing with each other at physiological pH to form lipid-nucleic acid complexes known as lipoplexes. However, lipoplexes proved unstable and characterized by a broad size distribution ranging from the submicron scale to several microns. Lipoplexes, such as Lipofectamine® reagent, have found reasonable utility for in vitro transfection. However, these first-generation lipoplexes have not proven useful in vivo. The large particle size and positive charge (imparted by cationic lipids) result in rapid plasma clearance, hemolysis, and other toxicities, as well as immune system activation.

[0055] Lipid mRNA formulation The mRNA disclosed herein, or a pharmaceutically acceptable salt thereof, can be incorporated into a lipid formulation (ie, a lipid-based delivery vehicle).

[0056] In the context of the present disclosure, lipid-based delivery vehicles typically serve to deliver the desired mRNA to target cells or tissues. The lipid-based delivery vehicle can be any suitable lipid-based delivery vehicle known in the art. In some embodiments, the lipid-based delivery vehicle is a liposome, a cationic liposome, or a lipid nanoparticle containing the mRNA of the present disclosure. In some embodiments, the lipid-based delivery vehicle comprises a nanoparticle or a bilayer of lipid molecules and the mRNA of the present disclosure. In some embodiments, the lipid bilayer preferably further comprises a neutral lipid or polymer. In some embodiments, the lipid formulation preferably comprises a liquid medium. In some embodiments, the formulation preferably further encapsulates a nucleic acid. In some embodiments, the lipid formulation preferably further comprises a nucleic acid and a neutral lipid or polymer. In some embodiments, the lipid formulation preferably encapsulates a nucleic acid.

[0057] The present disclosure provides lipid formulations comprising one or more therapeutic mRNA molecules encapsulated in the lipid formulation. In some embodiments, the lipid formulation comprises a liposome. In some embodiments, the lipid formulation comprises a cationic liposome. In some embodiments, the lipid formulation comprises a lipid nanoparticle.

[0058] In some embodiments, mRNA is completely encapsulated in the lipid portion of lipid formulation, so that the mRNA in lipid formulation is resistant to nuclease degradation in aqueous solution.In other embodiments, lipid formulations described herein are substantially non-toxic to mammals such as humans.

[0059] Lipid formulations of the present disclosure typically have a total lipid:RNA mass / mass ratio of about 1:1 to about 100:1, about 1:1 to about 50:1, about 2:1 to about 45:1, about 3:1 to about 40:1, about 5:1 to about 38:1, or about 6:1 to about 40:1, or about 7:1 to about 35:1, or about 8:1 to about 30:1; or about 10:1 to about 25:1, or about 8:1 to about 12:1; or about 13:1 to about 17:1; or about 18:1 to about 24:1; or about 20:1 to about 30:1. In some preferred embodiments, the total lipid:RNA mass / mass ratio is about 10:1 to about 25:1. The ratio can be any value or subvalue within the recited range, including the endpoints.

[0060] The lipid formulations of the present disclosure typically have a diameter of about 30 nm to about 150 nm, about 40 nm to about 150 nm, about 50 nm to about 150 nm, about 60 nm to about 130 nm, about 70 nm to about 110 nm, about 70 nm to about 100 nm, about 80 nm to about 100 nm, about 90 nm to about 100 nm, about 70 to about 90 nm, about 80 nm to about 90 nm, about 70 nm to about 80 nm, or about 30 nm, about 35 nm, about 40 nm, about 50 nm, about 60 nm, about 70 nm, about 80 nm, about 90 nm, about 70 nm, about 80 nm, about 80 nm, about 9 ... The lipid nanoparticles of the present disclosure have an average diameter of about 0 nm, about 45 nm, about 50 nm, about 55 nm, about 60 nm, about 65 nm, about 70 nm, about 75 nm, about 80 nm, about 85 nm, about 90 nm, about 95 nm, about 100 nm, about 105 nm, about 110 nm, about 115 nm, about 120 nm, about 125 nm, about 130 nm, about 135 nm, about 140 nm, about 145 nm, or about 150 nm, and are substantially non-toxic. The diameter can be any value or subvalue within the recited range, including the end points. In addition, when nucleic acids are present in the lipid nanoparticles of the present disclosure, they are resistant to degradation by nucleases in aqueous solution.

[0061] In a preferred embodiment, the lipid formulation comprises mRNA, a cationic lipid (e.g., one or more cationic lipids described herein or salts thereof), a phospholipid, and a conjugated lipid that inhibits particle aggregation (e.g., one or more PEG-lipid conjugates). The lipid formulation may also comprise cholesterol.

[0062] In nucleic acid-lipid formulation, mRNA can be completely encapsulated in the lipid portion of formulation, thereby protecting nucleic acid from nuclease degradation.In a preferred embodiment, the lipid formulation comprising mRNA is completely encapsulated in the lipid portion of lipid formulation, thereby protecting nucleic acid from nuclease degradation.In certain cases, the mRNA in lipid formulation is not substantially decomposed at least 20, 30, 45 or 60 minutes at 37 ℃ after particles are exposed to nuclease.In certain other cases, the mRNA in lipid formulation is not substantially decomposed at least 30, 45 or 60 minutes, or at least 2, 3, 4, 5, 6, 7, 8, 9, 10, 12, 14, 16, 18, 20, 22, 24, 26, 28, 30, 32, 34 or 36 hours at 37 ℃ after formulation is incubated in serum.In other embodiments, mRNA is complexed with the lipid portion of formulation.

[0063] In the context of nucleic acids, complete encapsulation can be determined by performing a membrane-impermeable fluorescent dye exclusion assay, which uses a dye that exhibits enhanced fluorescence when associated with nucleic acids. Encapsulation is determined by adding the dye to the lipid formulation, measuring the resulting fluorescence, and comparing it to the fluorescence observed upon the addition of a small amount of nonionic detergent. Detergent-mediated disruption of the lipid layer releases the encapsulated nucleic acid, allowing it to interact with the membrane-impermeable dye. Nucleic acid encapsulation can be calculated as E = (I0 - I) / I0, where I and I0 refer to the fluorescence intensity before and after the addition of detergent.

[0064] In other embodiments, the present disclosure provides nucleic acid-lipid compositions comprising a plurality of nucleic acid-liposomes, nucleic acid-cationic liposomes, or nucleic acid-lipid nanoparticles. In some embodiments, the nucleic acid-lipid compositions comprise a plurality of mRNA-liposomes. In some embodiments, the nucleic acid-lipid compositions comprise a plurality of mRNA-cationic liposomes. In some embodiments, the nucleic acid-lipid compositions comprise a plurality of mRNA-lipid nanoparticles.

[0065] In some embodiments, the lipid formulation comprises about 30% to about 100%, about 40% to about 100%, about 50% to about 100%, about 60% to about 100%, about 70% to about 100%, about 80% to about 100%, about 90% to about 100%, about 30% to about 95%, about 40% to about 95%, about 50% to about 95%, about 60% to about 95%, about 70% to about 95%, about 80% to about 95%, about 85% to about 95%, about 90% to about 95%, about 30% to about 90%, about 40% to about 90%, about 50% to about 90%, about 60% to about 95%, about 70% to about 95%, about 80% to about 95%, about 85% to about 95%, about 90% to about 95%, about 30% to about 90%, about 40% to about 90%, about 50% to about 90%, about 60% to about 95%, about 9 ... The lipid portion of the formulation contains mRNA fully encapsulated such that about 90%, about 70% to about 90%, about 80% to about 90%, or at least about 30%, about 35%, about 40%, about 45%, about 50%, about 55%, about 60%, about 65%, about 70%, about 75%, about 80%, about 85%, about 90%, about 91%, about 92%, about 93%, about 94%, about 95%, about 96%, about 97%, about 98%, or about 99% (or any fraction or range therein) of the mRNA is encapsulated therein. The amount can be any value or fraction within the recited range, including the endpoints.

[0066] Depending on the intended use of the lipid formulation, the proportions of the components may vary, and the delivery efficiency of a particular formulation can be measured using assays known in the art.

[0067] According to some embodiments, the expressible polynucleotide and mRNA constructs described herein are lipid-formulated. The lipid formulation is preferably selected from, but not limited to, liposomes, cationic liposomes, and lipid nanoparticles. In one preferred embodiment, the lipid formulation is: (a) an mRNA of the present disclosure; (b) cationic lipids, (c) aggregation-reducing agents (polyethylene glycol (PEG) lipids or PEG-modified lipids); (d) optionally a non-cationic lipid (such as a neutral lipid), and (e) Cationic liposomes or lipid nanoparticles (LNPs) optionally containing a sterol.

[0068] In some embodiments, the cationic lipid is an ionizable cationic lipid. In one embodiment, the lipid nanoparticle formulation consists of (i) at least one cationic lipid; (ii) a helper lipid; (iii) a sterol (e.g., cholesterol); and (iv) a PEG lipid in a molar ratio of about 20% to about 40% ionizable cationic lipid: about 25% to about 45% helper lipid: about 25% to about 45% sterol; and about 0.5% to 5% PEG lipid. Exemplary cationic lipids (including ionizable cationic lipids), helper lipids (e.g., neutral lipids), sterols, and ligand-containing lipids (e.g., PEG lipids) are described herein below.

[0069] The selection of specific lipids and their relative composition ratios depends on several factors, including the desired therapeutic effect, the intended in vivo delivery target, and the planned administration regimen and frequency. Generally, lipids that support high potency (i.e., therapeutic effect such as knockdown activity or translation efficiency) and biodegradability that allows rapid tissue clearance are most preferred. However, biodegradability may not be as important for formulations intended for only one or two administrations within a subject. Furthermore, the lipid composition may require careful modification so that the lipid formulation maintains its morphology during in vivo administration and transport to the intended target, while subsequently releasing the active agent upon uptake into target cells. Therefore, several formulations usually need to be evaluated to find the most promising lipid combinations, including the most promising lipid-to-active ingredient molar ratio and the most promising total lipid-to-active ingredient ratio.

[0070] Suitable lipid components and methods for producing lipid nanoparticles are well known in the art and are described, for example, in PCT / US2020 / 023442, US8,058,069, US8,822,668, US9,738,593, US9,139,554, PCT / US2014 / 066242, PCT / US2015 / 030218, PCT / 2017 / 015886, and PCT / US2017 / 067756, the contents of which are incorporated by reference in their entireties.

[0071] cationic lipids The lipid formulation preferably contains a cationic lipid suitable for forming cationic liposomes or lipid nanoparticles. Cationic lipids have been widely studied for nucleic acid delivery because they can bind to negatively charged membranes and induce uptake. Generally, cationic lipids are amphiphilic substances containing a positive hydrophilic head group, two (or more) lipophilic tails or steroid moieties, and a linker between these two domains. Preferably, the cationic lipid carries a net positive charge at approximately physiological pH. Traditionally, cationic liposomes have been the most commonly used non-viral delivery system for oligonucleotides, such as plasmid DNA, antisense oligos, and siRNA / small hairpin RNA-shRNA. Cationic lipids, such as DOTAP (1,2-dioleoyl-3-trimethylammonium-propane) and DOTMA (N-[1-(2,3-dioleoyloxy)propyl]-N,N,N-trimethyl-ammonium methylsulfate), can form complexes or lipoplexes with negatively charged nucleic acids through electrostatic interactions, providing high in vitro transfection efficiency.

[0072] In the lipid formulations of the present disclosure, the cationic lipid can be, for example, N,N-dioleyl-N,N-dimethylammonium chloride (DODAC), N,N-distearyl-N,N-dimethylammonium bromide (DDAB), 1,2-dioleoyltrimethylammonium propane chloride (DOTAP) (also known as N-(2,3-dioleoyloxy)propyl)-N,N,N-trimethylammonium chloride and 1,2-dioleyloxy-3-trimethylaminopropane chloride salt), N-(1-(2,3-dioleoyloxy)propyl)-N,N,N-trimethylammonium chloride, and 1,2-dioleyloxy-3-trimethylaminopropane chloride salt. (Dimethyl)-N,N,N-trimethylammonium chloride (DOTMA), N,N-dimethyl-2,3-dioleyloxy)propylamine (DODMA), 1,2-dilinoleyloxy-N,N-dimethylaminopropane (DLinDMA), 1,2-dilinolenyloxy-N,N-dimethylaminopropane (DLenDMA), 1,2-di-y-linolenyloxy-N,N-dimethylaminopropane (γ-DLenDMA), 1,2-dilinoleylcarbamoyloxy-3-dimethylaminopropane (DLin-C-DAP), 1,2-dilinoleyl 1,2-Dilinoleyloxy-3-(dimethylamino)acetoxypropane (DLin-DAC), 1,2-Dilinoleyloxy-3-morpholinopropane (DLin-MA), 1,2-Dilinoleoyl-3-dimethylaminopropane (DLinDAP), 1,2-Dilinoleylthio-3-dimethylaminopropane (DLin-S-DMA), 1-Linoleoyl-2-linoleyloxy-3-dimethylaminopropane (DLin-2-DMAP), 1,2-Dilinoleyloxy-3-trimethylaminopropane chloride salt (DLin-TMA.Cl), 1,2 -Dilinoleoyl-3-trimethylaminopropane chloride salt (DLin-TAP.Cl), 1,2-dilinoleyloxy-3-(N-methylpiperazino)propane (DLin-MPZ), or 3-(N,N-dilinoleylamino)-1,2-propanediol (DLinAP), 3-(N,N-dioleylamino)-1,2-propanediol (DOAP), 1,2-dilinoleyloxo-3-(2-N,N-dimethylamino)ethoxypropane (DLin-EG-DMA), 2,2-dilinoleyl-4-dimethylaminomethyl-[1,3]-dioxolane (DLin-K-DMA) or their analogs, (3aR,5s,6aS)-N,N-dimethyl-2,2-di((9Z,12Z)-octadeca-9,12-dienyl)tetrahydro-3aH-cyclopenta[d][1,3]dioxol-5-amine, (6Z,9Z,28Z,31Z)-heptatriaconta-6,9,28,31-tetraene-19 -yl 4-(dimethylamino)butanoate (MC3), 1,1'-(2-(4-(2-((2-(bis(2-hydroxydodecyl)amino)ethyl)(2-hydroxydodecyl)amino)ethyl)piperazin-1-yl)ethylazanediyl)didodecan-2-ol (C12-200), 2,2-dilinoleyl-4-(2-dimethylaminoethyl)-[1,3]-diol The compound may be 2,2-dilinoleyl-4-dimethylaminomethyl-[1,3]-dioxolane (DLin-K-DMA), (6Z,9Z,28Z,31Z)-heptatriaconta-6,9,28,31-tetraen-19-yl 4-(dimethylamino)butanoate (DLin-M-C3-DMA), 3-((6Z,9Z,28Z,31Z)-heptatriaconta-6,9,28,31-tetraen-19-yloxy)-N,N-dimethylpropan-1-amine (MC3 ether), 4-((6Z,9Z,28Z,31Z)-heptatriaconta-6,9,28,31-tetraen-19-yloxy)-N,N-dimethylbutan-1-amine (MC4 ether), or any combination thereof. Other cationic lipids include N,N-distearyl-N,N-dimethylammonium bromide (DDAB), 3P-(N-(N',N'-dimethylaminoethane)-carbamoyl)cholesterol (DC-Choi), N-(1-(2,3-dioleyloxy)propyl)-N-2-(sperminecarboxamido)ethyl)-N,N-dimethylammonium trifluoroacetate (DOSPA), dioctadecylamidoglycylcarboxyspermine (DOGS), 1,2-dioleoyl-sn-3-phosphoethanolamine (DOPE), 1,2-dioleoyl-3-dimethylammonium propane (DODAP), N-(1,2-dimyristyloxyprop-3-yl)-N,N-dimethylammonium trifluoroacetate (DOSPA), and N-(1-(2,3-dioleyloxy)propyl)-N-2-(sperminecarboxamido)ethyl)-N,N-dimethylammonium trifluoroacetate (DOSPA).Examples of suitable cationic lipids include, but are not limited to, N-dimethyl-N-hydroxyethylammonium bromide (DMRIE) and 2,2-dilinoleyl-4-dimethylaminoethyl-[1,3]-dioxolane (XTC). Additionally, commercially available preparations of cationic lipids can be used, such as, for example, Lipofectin (including DOTMA and DOPE, available from GIBCO / BRL) and Lipofectamine (including DOSPA and DOPE, available from GIBCO / BRL).

[0073] Other suitable cationic lipids are disclosed in International Publication Nos. WO09 / 086558, WO09 / 127060, WO10 / 048536, WO10 / 054406, WO10 / 088537, WO10 / 129709, and WO2011 / 153493; U.S. Patent Publication Nos. 2011 / 0256175, 2012 / 0128760, and 2012 / 0027803; U.S. Patent No. 8,158,601; and Love et al., PNAS, 107(5), 1864-69, 2010, the contents of which are incorporated herein by reference.

[0074] Other suitable cationic lipids include those with alternative fatty acid groups and other dialkylamino groups, such as those with different alkyl substituents (e.g., N-ethyl-N-methylamino- and N-propyl-N-ethylamino-). These lipids are part of a subcategory of cationic lipids called amino lipids. In some embodiments of the lipid formulations described herein, the cationic lipid is an amino lipid. In general, amino lipids with less saturated acyl chains are easier to size, especially when the complexes must be sized to about 0.3 microns or less for filter sterilization. Carbon chain lengths of C 14 ~C 22 Other scaffolds can be used to separate the amino group and the fatty acid or fatty alkyl portion of the amino lipid.

[0075] In some embodiments, the lipid formulation comprises a cationic lipid of Formula I according to Patent Application No. PCT / EP2017 / 064066, the disclosure of which is also incorporated herein by reference in this context.

[0076] In some embodiments, the amino lipids or cationic lipids of the present disclosure are ionizable and have at least one protonatable or deprotonatable group. As a result, the lipids have a positive charge at a pH below physiological pH (e.g., pH 7.4) and are neutral at a second pH (preferably above physiological pH). Of course, it is understood that the addition or removal of protons as a function of pH is an equilibrium process, and reference to charged or neutral lipids refers to the nature of the predominant species and does not require that all lipids exist in a charged or neutral form. Lipids having two or more protonatable or deprotonatable groups or that are zwitterionic are not excluded from use in the present disclosure. In certain embodiments, the protonatable lipids have a pKa of the protonatable group in the range of about 4 to about 11. In some embodiments, the ionizable cationic lipids have a pKa of about 5 to about 7. In some embodiments, the pKa of the ionizable cationic lipids is about 6 to about 7.

[0077] In some embodiments, the lipid formulation comprises an ionizable cationic lipid of Formula I [ka] or a pharmaceutically acceptable salt or solvate thereof, wherein R 5 and R 6 are each independently a straight-chain or branched C1-C 31 Alkyl, C2-C 31 Alkenyl, or C2-C 31 L is selected from the group consisting of alkynyl, and cholesteryl; 5 and L 6 are each independently a straight chain C1-C 20 Alkyl and C2-C20 alkenyl; X 5 is -C(O)O-, which results in -C(O)OR 6 is formed, or -OC(O)-, whereby -OC(O)-R 6 is formed; X 6 is -C(O)O-, which gives -C(O)OR 5 is formed, or -OC(O)-, whereby -OC(O)-R 5 is formed; X 7 is S or O; L 7 is absent or lower alkyl; R 4 is a straight-chain or branched C1-C6 alkyl; R 7 and R 8 are each independently selected from the group consisting of hydrogen and straight-chain or branched C1-C6 alkyl.

[0078] In some embodiments, X 7 is S.

[0079] In some embodiments, X 5 is -C(O)O-, which gives -C(O)OR 6 is formed, and X 6 is -C(O)O-, which gives -C(O)OR 5 is formed.

[0080] In some embodiments, R 7 and R 8 are each independently selected from the group consisting of methyl, ethyl, and isopropyl.

[0081] In some embodiments, L 5 and L 6 are each independently C1-C 10 In some embodiments, L 5 is C1-C3 alkyl, and L 6 is C1-C5 alkyl. In some embodiments, L 6is C1-C2 alkyl. In some embodiments, L 5 and L 6 Each is a straight chain C7 alkyl. In some embodiments, L 5 and L 6 are each a straight chain C9 alkyl.

[0082] In some embodiments, R 5 and R 6 are each independently alkenyl. In some embodiments, R 6 is alkenyl. In some embodiments, R 6 is a C2-C9 alkenyl. In some embodiments, the alkenyl contains a single double bond. In some embodiments, R 5 and R 6 Each is alkyl. In some embodiments, R 5 is a branched alkyl. In some embodiments, R 5 and R 6 are each independently selected from the group consisting of C alkyl, C alkenyl, and C alkynyl. 5 and R 6 are each independently, C 11 Alkyl, C 11 Alkenyl, and C 11 In some embodiments, R is selected from the group consisting of alkynyl. 5 and R 6 are each independently selected from the group consisting of C alkyl, C alkenyl, and C alkynyl. 5 is -CH((CH2) p CH3)2 or -CH((CH2) p CH3)((CH2) p-1 CH3), where p is 4 to 8. In some embodiments, p is 5 and L 5 is C1-C3 alkyl. In some embodiments, p is 6 and L 5 In some embodiments, p is 7. In some embodiments, p is 8 and L 5is C1-C3 alkyl. In some embodiments, R 5 is -CH((CH2) p CH3)((CH2) p-1 CH3) [wherein p is 7 or 8].

[0083] In some embodiments, R 4 is ethylene or propylene. In some embodiments, R 4 is n-propylene or isobutylene.

[0084] In some embodiments, L 7 is absent, R 4 is ethylene and X 7 is S and R 7 and R 8 Each is methyl. In some embodiments, L 7 is absent, R 4 is n-propylene, and X 7 is S and R 7 and R 8 Each is methyl. In some embodiments, L 7 is absent, R 4 is ethylene and X 7 is S and R 7 and R 8 are each ethyl.

[0085] In some embodiments, X 7 is S and X 5 is -C(O)O-, which gives -C(O)OR 6 is formed, and X 6 is -C(O)O-, which gives -C(O)OR 5 is formed, and L 5 and L 6 are each independently a straight chain C3-C7 alkyl; L 7 is absent, R 5 is -CH((CH2) p CH3)2 and R 6 is C7-C 12In some further embodiments, p is 6 and R 6 is a C9 alkenyl.

[0086] In some embodiments, the lipid formulation comprises: [ka] [ka] [ka] [ka] [ka] [ka] [ka] [ka] The ionizable cationic lipid comprises an ionizable cationic lipid selected from the group consisting of:

[0087] In some embodiments, any one or more of the lipids listed herein may be explicitly excluded.

[0088] Helper lipids and sterols The mRNA-lipid formulations of the present disclosure can include helper lipids, which can also be referred to as neutral lipids, neutral helper lipids, non-cationic lipids, non-cationic helper lipids, anionic lipids, anionic helper lipids, or zwitterionic lipids. Lipid formulations, particularly cationic liposomes and lipid nanoparticles, have been found to exhibit increased cellular uptake when helper lipids are present in the formulation (Curr. Drug Metab. 2014;15(9):882-92). For example, several studies have shown that neutral and zwitterionic lipids, such as 1,2-dioleoyl-sn-glycero-3-phosphatidylcholine (DOPC), dioleoyl-phosphatidylethanolamine (DOPE), and 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), which are more membrane-fusogenic (i.e., facilitate fusion) than cationic lipids, can affect the polymorphic phenotype of lipid-nucleic acid complexes, promoting the transition from a lamellar to a hexagonal phase, thereby inducing fusion and membrane disruption (Nanomedicine (Lond). 2014 Jan;9(1):105-20). In addition, the use of helper lipids can help reduce any potential adverse effects, such as toxicity and immunogenicity, associated with the use of many effective cationic lipids.

[0089] Non-limiting examples of non-cationic lipids suitable for the lipid formulations of the present disclosure include lecithin, phosphatidylethanolamine, lysolecithin, lysophosphatidylethanolamine, phosphatidylserine, phosphatidylinositol, sphingomyelin, egg sphingomyelin (ESM), cephalin, cardiolipin, phosphatidic acid, cerebrosides, dicetyl phosphate, distearoylphosphatidylcholine (DSPC), dioleoylphosphatidylcholine (DOPC), dipalmitoylphosphatidylcholine (DPPC), dioleoylphosphatidylglycerol (DOPG), dipalmitoylphosphatidylglycerol (DPPG), dioleoylphosphatidylethanolamine (DOPE), palmitoyloleoyl-phosphatidylcholine (POPC), palmitoyloleoyl-phosphatidylcholine (DPPG), dioleoylphosphatidylethanolamine (DOPE), palmitoyloleoyl-phosphatidylcholine (POPC), palmitoyloleoyl-phosphatidylcholine (DPPG), palmitoyloleoyl- ...DPPG), palmitoyloleoyl-phosphatidylethanolamine (DOPE), palmitoyloleoyl-phosphatidylcholine (POP Examples of suitable phospholipids include phospholipids such as palmitoyloleoylphosphatidylethanolamine (POPE), palmitoyloleoylphosphatidylglycerol (POPG), dioleoylphosphatidylethanolamine 4-(N-maleimidomethyl)-cyclohexane-1-carboxylate (DOPE-mal), dipalmitoyl-phosphatidylethanolamine (DPPE), dimyristoyl-phosphatidylethanolamine (DMPE), distearoyl-phosphatidylethanolamine (DSPE), monomethyl-phosphatidylethanolamine, dimethyl-phosphatidylethanolamine, dielaidoyl-phosphatidylethanolamine (DEPE), stearoyloleoyl-phosphatidylethanolamine (SOPE), lysophosphatidylcholine, dilinoleoylphosphatidylcholine, and mixtures thereof. Other diacylphosphatidylcholine and diacylphosphatidylethanolamine phospholipids can also be used. The acyl group in these lipids is preferably C 10 -C 24 It is an acyl group derived from a fatty acid having a carbon chain, for example, lauroyl, myristoyl, palmitoyl, stearoyl, or oleyl.

[0090] Further examples of non-cationic lipids include sterols such as cholesterol and its derivatives. One study concluded that, as a helper lipid, cholesterol increases the charge spacing of the lipid layer that interacts with nucleic acids, allowing the charge distribution to more closely match that of the nucleic acid. (JRSoc.Interface.2012 Mar 7;9(68):548-561). Non-limiting examples of cholesterol derivatives include polar analogs such as 5α-cholestanol, 5α-coprostanol, cholesteryl-(2'-hydroxy)-ethyl ether, cholesteryl-(4'-hydroxy)-butyl ether, and 6-ketocholestanol; non-polar analogs such as 5α-cholestan, cholestenone, 5α-cholestanone, and cholesteryl decanoate; and mixtures thereof. In a preferred embodiment, the cholesterol derivative is a polar analog such as cholesteryl-(4'-hydroxy)-butyl ether.

[0091] In some embodiments, the helper lipid present in lipid formulation comprises or consists of a mixture of one or more phospholipids and cholesterol or its derivatives.In some embodiments, the helper lipid present in lipid formulation comprises or consists of a mixture of one or more phospholipids and cholesterol or its derivatives.In still other embodiments, the helper lipid present in lipid formulation comprises or consists of cholesterol or its derivatives, for example, phospholipid-free lipid formulation.

[0092] Other examples of helper lipids include phosphorus-free containing lipids such as, for example, stearylamine, dodecylamine, hexadecylamine, acetyl palmitate, glycerol ricinoleate, hexadecyl stearate, isopropyl myristate, amphoteric acrylic polymers, triethanolamine lauryl sulfate, alkyl-aryl sulfate polyethyloxylated fatty acid amides, dioctadecyldimethylammonium bromide, ceramides, and sphingomyelin.

[0093] In some embodiments, the helper lipid comprises about 20 mol% to about 50 mol%, about 22 mol% to about 48 mol%, about 24 mol% to about 46 mol%, about 25 mol% to about 44 mol%, about 26 mol% to about 42 mol%, about 27 mol% to about 41 mol%, about 28 mol% to about 40 mol%, or about 29 mol%, about 30 mol%, about 31 mol%, about 32 mol%, about 33 mol%, about 34 mol%, about 35 mol%, about 36 mol%, about 37 mol%, about 38 mol%, or about 39 mol% (or any fraction or range therein) of the total lipid present in the lipid formulation.

[0094] In some embodiments, the helper lipid in the formulation comprises two or more types of helper lipid, and the total amount of helper lipids comprises about 20 mol% to about 50 mol%, about 22 mol% to about 48 mol%, about 24 mol% to about 46 mol%, about 25 mol% to about 44 mol%, about 26 mol% to about 42 mol%, about 27 mol% to about 41 mol%, about 28 mol% to about 40 mol%, or about 29 mol%, about 30 mol%, about 31 mol%, about 32 mol%, about 33 mol%, about 34 mol%, about 35 mol%, about 36 mol%, about 37 mol%, about 38 mol%, or about 39 mol% (or any fraction or range therein) of the total lipids present in the lipid formulation. In some embodiments, the helper lipid is a combination of DSPC and DOTAP. In some embodiments, the helper lipid is a combination of DSPC and DOTMA.

[0095] Cholesterol or cholesterol derivatives in a lipid formulation can comprise up to about 40 mol%, about 45 mol%, about 50 mol%, about 55 mol%, or about 60 mol% of the total lipids present in the lipid formulation. In some embodiments, cholesterol or cholesterol derivatives comprise about 15 mol% to about 45 mol%, about 20 mol% to about 40 mol%, about 30 mol% to about 40 mol%, or about 35 mol%, about 36 mol%, about 37 mol, about 38 mol, about 39 mol, or about 40 mol of the total lipids present in the lipid formulation.

[0096] The percentage of helper lipid present in the lipid formulation is a target amount, and the actual amount of helper lipid present in the formulation may vary, for example, by ±5 mol %.

[0097] Lipid formulations containing cationic lipid compounds or ionizable cationic lipid compounds may comprise, on a molar basis, about 20-40% cationic lipid compound, about 25-40% cholesterol, about 25-50% helper lipid, and about 0.5-5% polyethylene glycol (PEG) lipid, relative to the total lipid present in the formulation. In some embodiments, the composition comprises about 22-30% cationic lipid compound, about 30-40% cholesterol, about 30-40% helper lipid, and about 0.5-3% PEG lipid, relative to the total lipid present in the formulation.

[0098] lipid complex The lipid formulations described herein can further comprise lipid complexes. Complex lipids are useful for preventing particle aggregation. Suitable complex lipids include, but are not limited to, PEG-lipid complexes, cationic polymer-lipid complexes, and mixtures thereof. Furthermore, lipid delivery vehicles can be used for specific targeting by attaching ligands (e.g., antibodies, peptides, and carbohydrates) to their surface or to the end of the attached PEG chain (Front Pharmacol.2015 Dec 1;6:286).

[0099] In a preferred embodiment, the lipid conjugate is a PEG-lipid. The inclusion of polyethylene glycol (PEG) in lipid formulations as a coating or surface ligand, a technique called PEGylation, helps protect nanoparticles from the immune system and evade RES uptake ( Nanomedicine (London). 2011 Jun;6(4):715-28 ). PEGylation has been widely used to stabilize lipid formulations and their payloads through physical, chemical, and biological mechanisms. Detergent-like PEG lipids (e.g., PEG-DSPE) can enter the lipid formulation and form a hydration layer and a steric barrier on the surface. Based on the degree of PEGylation, the surface layer can generally be divided into two types: a brush-like layer and a mushroom-like layer. Regarding PEG-DSPE-stabilized formulations, PEG adopts a mushroom conformation at low PEGylation levels (usually less than 5 mol%) and transitions to a brush conformation as the PEG-DSPE content increases beyond a certain level ( J. Nanomaterials. 2011;2011:12 ). Increasing PEGylation has been shown to result in a significant increase in the circulating half-life of lipid formulations (Annu. Rev. Biomed. Eng. 2011 Aug 15;13():507-30; J. Control Release. 2010 Aug 3;145(3):178-81).

[0100] Suitable examples of PEG-lipids include, but are not limited to, PEG conjugated to dialkyloxypropyl (PEG-DAA), PEG conjugated to diacylglycerol (PEG-DAG), PEG conjugated to a phospholipid such as phosphatidylethanolamine (PEG-PE), PEG conjugated to ceramide, PEG conjugated to cholesterol, or derivatives thereof, and mixtures thereof.

[0101] PEG is a linear water-soluble polymer of ethylene PEG repeating units with two terminal hydroxyl groups. PEGs are classified according to their molecular weight and include: monomethoxypolyethyleneglycol (MePEG-OH), monomethoxypolyethyleneglycol-succinate (MePEG-S), monomethoxypolyethyleneglycol-succinimidylsuccinate (MePEG-S-NHS), monomethoxypolyethyleneglycol-amine (MePEG-NH2), monomethoxypolyethyleneglycol-tresylate (MePEG-TRES), monomethoxypolyethyleneglycol-imidazolyl-carbonyl (MePEG-IM), as well as compounds containing terminal hydroxyl groups instead of terminal methoxy groups (e.g., HO-PEG-S, HO-PEG-S-NHS, HO-PEG-NH2).

[0102] The PEG moiety of the PEG-lipid conjugates described herein may comprise an average molecular weight ranging from about 550 daltons to about 10,000 daltons. In certain instances, the PEG moiety has an average molecular weight of about 750 daltons to about 5,000 daltons (e.g., about 1,000 daltons to about 5,000 daltons, about 1,500 daltons to about 3,000 daltons, about 750 daltons to about 3,000 daltons, about 750 daltons to about 2,000 daltons). In preferred embodiments, the PEG moiety has an average molecular weight of about 2,000 daltons or about 750 daltons. The average molecular weight may be any value or subvalue within the recited range, including the endpoints.

[0103] In certain cases, PEG monomer can be optionally substituted with alkyl, alkoxy, acyl or aryl group.PEG can be directly conjugated to lipid, or can be linked to lipid via linker moiety.For example, any linker moiety suitable for linking PEG to lipid can be used, including non-ester-containing linker moiety and ester-containing linker moiety.In a preferred embodiment, the linker moiety is a non-ester-containing linker moiety. Suitable non-ester containing linker moieties include, but are not limited to, amide (-C(O)NH-), amino (-NR-), carbonyl (-C(O)-), carbamate (-NHC(O)O-), urea (-NHC(O)NH-), disulfide (-SS-), ether (-O-), succinyl (-(O)CCH2CH2C(O)-), succinamidyl (-NHC(O)CH2CH2C(O)NH-), ether, and combinations thereof (such as linkers containing both carbamate and amide linker moieties). In a preferred embodiment, a carbamate linker is used to attach PEG to the lipid.

[0104] In other embodiments, an ester-containing linker moiety is used to attach PEG to the lipid. Suitable ester-containing linker moieties include, for example, carbonate (-OC(O)O-), succinoyl, phosphate ester (-O-(O)POH-O-), and combinations thereof.

[0105] Phosphatidylethanolamines with various acyl chain groups of varying chain length and saturation can be conjugated to PEG to form lipid conjugates. Such phosphatidylethanolamines are commercially available or can be isolated or synthesized using conventional techniques known to those skilled in the art. 10 ~C 20Phosphatidylethanolamines containing saturated or unsaturated fatty acids with carbon chain lengths in the range of 0 to 100 are preferred. Phosphatidylethanolamines with mono- or di-unsaturated fatty acids, as well as mixtures of saturated and unsaturated fatty acids, can also be used. Suitable phosphatidylethanolamines include, but are not limited to, dimyristoyl-phosphatidylethanolamine (DMPE), dipalmitoyl-phosphatidylethanolamine (DPPE), dioleoyl-phosphatidylethanolamine (DOPE), and distearoyl-phosphatidylethanolamine (DSPE).

[0106] In some embodiments, the PEG-DAA conjugate is PEG-didecyloxypropyl (C 10 ) conjugate, PEG-dilauryloxypropyl (C 12 ) conjugate, PEG-dimyristyloxypropyl (C 14 ) conjugate, PEG-dipalmityloxypropyl (C 16 ) conjugate, or PEG-distearyloxypropyl (C 18 ) conjugates. In these embodiments, the PEG preferably has an average molecular weight of about 750 to about 2,000 daltons. In certain embodiments, the terminal hydroxyl group of the PEG is substituted with a methyl group.

[0107] In addition to the above, other hydrophilic polymers can be used in place of PEG. Examples of suitable polymers that can be used in place of PEG include, but are not limited to, polyvinylpyrrolidone, polymethyloxazoline, polyethyloxazoline, polyhydroxypropylmethacrylamide, polymethacrylamide and polydimethylacrylamide, polylactic acid, polyglycolic acid, and derivatized celluloses such as hydroxymethylcellulose or hydroxyethylcellulose.

[0108] In some embodiments, the lipid conjugates (e.g., PEG-lipids) comprise about 0.1 mol% to about 2 mol%, about 0.5 mol% to about 2 mol%, about 1 mol% to about 2 mol%, about 0.6 mol% to about 1.9 mol%, about 0.7 mol% to about 1.8 mol%, about 0.8 mol% to about 1.7 mol%, about 0.9 mol% to about 1.6 mol%, about 0.9 mol% to about 1.8 mol%, about 1 mol% to about 1.8 mol%, about 1 mol% to about 1.7 mol%, about 1.2 mol% to about 1.8 mol%, about 1.2 mol% to about 1.7 mol%, about 1.3 mol% to about 1.6 mol%, or about 1.4 mol% to about 1.6 mol% (or any fraction or range therein) of the total lipid present in the lipid formulation. In other embodiments, the lipid conjugates (e.g., PEG-lipids) comprise about 0.5%, 0.6%, 0.7%, 0.8%, 0.9%, 1.0%, 1.2%, 1.3%, 1.4%, 1.5%, 1.6%, 1.7%, 1.8%, 1.9%, 2.0%, 2.5%, 3.0%, 3.5%, 4.0%, 4.5%, or 5% (or any fraction or range therein) of the total lipids present in the lipid formulation. The amount can be any value or subvalue within the recited range, including the endpoints.

[0109] In some preferred embodiments, the PEG lipid is PEG550-PE. In some preferred embodiments, the PEG lipid is PEG750-PE. In some preferred embodiments, the PEG lipid is PEG2000-DMG.

[0110] The percentage of lipid-conjugates (e.g., PEG-lipids) present in the lipid formulations of the present disclosure is a target amount, and the actual amount of lipid-conjugates present in the formulation may vary, for example, by ±0.5 mol%. Those skilled in the art will understand that the concentration of lipid-conjugates may vary depending on the lipid-conjugates used and the rate at which the lipid formulation becomes fusogenic.

[0111] Mechanism of cellular uptake of lipid formulations Lipid formulations for intracellular delivery of nucleic acids, particularly liposomes, cationic liposomes, and lipid nanoparticles, are designed for cellular uptake by penetration of target cells, utilizing the endocytic mechanism of the target cells, and a certain amount of lipid delivery vehicle is delivered to the cytosol of the target cells. (Nucleic Acid Therapeutics, 28(3):146-157, 2018). Specifically, in the case of the mRNA lipid formulations targeted to hepatocytes described herein, the mRNA lipid formulation enters hepatocytes by receptor-mediated endocytosis. Prior to endocytosis, functional ligands, such as PEG lipids, are shed from the surface of the lipid delivery vehicle, thereby causing internalization into the target cells. During endocytosis, several portions of the cell's plasma membrane surround the vector, engulfing it in vesicles, which are then removed from the plasma membrane, enter the cytosol, and ultimately pass through the endolysosomal pathway. For the delivery vehicle that contains ionizable cationic lipid, the acidity increases as the endosome advances, resulting in the vehicle having a strong positive charge on the surface.Then, the interaction between delivery vehicle and endosomal membrane causes membrane function event, which leads to the cytosolic delivery of payload.For mRNA payload, the internal translation process of cell itself then translates mRNA into coded protein.The coded protein can then undergo further post-translational processing, including transport to targeted organelle or location within the cell.

[0112] By controlling the composition and concentration of the lipid complexes, the rate at which the lipid complexes are exchanged from the lipid formulation and, consequently, the rate at which the lipid formulation becomes fusogenic can be controlled. In addition, other variables, including, for example, pH, temperature, or ionic strength, can be used to vary and / or control the rate at which the lipid formulation becomes fusogenic. Other methods by which the rate at which the lipid formulation becomes fusogenic will be apparent to those skilled in the art upon reading this disclosure. Additionally, by controlling the composition and concentration of the lipid complexes, the liposome or lipid particle size can be controlled.

[0113] Manufacture of lipid formulations There are many different methods for preparing lipid formulations containing nucleic acids. (Curr. Drug Metabol. 2014, 15, 882-892; Chem. Phys. Lipids 2014, 177, 8-18; Int. J. Pharm. Stud. Res. 2012, 3, 14-20) The following techniques are briefly described here: thin film hydration, double emulsion, reverse phase evaporation, microfluidic preparation, double asymmetric centrifugation, ethanol injection, detergent dialysis, spontaneous vesicle formation by ethanol dilution, and encapsulation in preformed liposomes.

[0114] Thin Film Hydration In the thin film hydration (TFH) or Bangham method, lipids are dissolved in an organic solvent and then evaporated using a rotary evaporator to form a thin lipid layer. After layer hydration with an aqueous buffer solution containing the compound to be loaded, multilamellar vesicles (MLVs) are formed, which can be reduced in size to produce small or large unilamellar vesicles (LUVs and SUVs) by extrusion through a membrane or by sonication of the starting MLVs.

[0115] Double emulsion Lipid formulations can also be formed by double emulsion techniques, which involve dissolving lipids in a water / organic solvent mixture. An organic solution containing water droplets is mixed with an excess of aqueous medium to form a water-in-oil-in-water (W / O / W) double emulsion. After vigorous mechanical shaking, some of the water droplets collapse, resulting in large unilamellar vesicles (LUVs).

[0116] Reverse Phase Evaporation The reverse phase evaporation (REV) method also allows for the realization of LUVs loaded with nucleic acids. In this technique, phospholipids are dissolved in an organic solvent and an aqueous buffer to form a two-phase system. The resulting suspension is then briefly sonicated until the mixture becomes a clear, one-phase dispersion. After evaporating the organic solvent under reduced pressure, the lipid formulation is obtained. This method has been used to encapsulate various large and small hydrophilic molecules, including nucleic acids.

[0117] Preparation of microfluidics Unlike other bulk technologies, microfluidic methods offer the possibility of controlling the lipid hydration process. These methods can be classified into continuous-flow microfluidic and droplet-based microfluidic types according to the flow manipulation method. In the microfluidic hydrodynamic focusing (MHF) method, which operates in continuous-flow mode, lipids are dissolved in isopropyl alcohol and hydrodynamically focused at the microchannel intersection junction of two aqueous buffer streams. Vesicle size can be controlled by controlling the flow rate and, therefore, the lipid solution / buffer dilution process. A microfluidic device with three inlets and one outlet can be used to prepare oligonucleotide (ON) lipid formulations.

[0118] Double asymmetric centrifugation Double asymmetric centrifugation (DAC) differs from more conventional centrifugation by utilizing an additional rotation about its own longitudinal axis. Efficient homogenization is achieved by creating two overlapping movements. The sample is pushed outward, as in a conventional centrifuge, and then pushed toward the center of the vial by the additional rotation. Mixing the lipid and NaCl solution yields a viscous vesicular phospholipid gel (VPC), which is then diluted to obtain the lipid formulation dispersion. The size of the lipid formulation can be controlled by optimizing the DAC speed, lipid concentration, and homogenization time.

[0119] Ethanol injection The ethanol injection (EI) method can be used for nucleic acid encapsulation. This method involves the rapid injection of an ethanolic solution, dissolving lipids into an aqueous medium containing the nucleic acid to be encapsulated using a needle. Vesicles form spontaneously when the phospholipids are dispersed throughout the medium.

[0120] Detergent dialysis Nucleic acids can be encapsulated using detergent dialysis. Briefly, lipids and plasmids are dissolved in a detergent solution of appropriate ionic strength, and after the detergent is removed by dialysis, a stable lipid formulation is formed. Unencapsulated nucleic acids are then removed by ion exchange chromatography, and empty vesicles are removed by sucrose density gradient centrifugation. This technique is highly sensitive to the content of cationic lipids and the salt concentration of the dialysis buffer, and the method is also difficult to scale up.

[0121] Spontaneous vesicle formation upon ethanol dilution Stable lipid formulations can also be generated by spontaneous vesicle formation via ethanol dilution, in which stepwise or dropwise ethanol dilution results in the instantaneous formation of nucleic acid-loaded vesicles by the controlled addition of lipids dissolved in ethanol to a rapidly mixing aqueous buffer containing nucleic acid.

[0122] Encapsulation into preformed liposomes Nucleic acid entrapment can also be achieved by starting with preformed liposomes in two different ways: (1) simply mixing cationic liposomes with nucleic acid to obtain electrostatic complexes called "lipoplexes" (these can be used successfully to transfect cell cultures, but are characterized by low encapsulation efficiency and poor performance in vivo); and (2) liposome destabilization, in which absolute ethanol is slowly added to a suspension of cationic vesicles up to a concentration of 40%, followed by dropwise addition of nucleic acid to achieve loaded vesicles (however, the two main steps that characterize the encapsulation process are too sensitive and result in reduced particle size).

[0123] Formation of lipid-encapsulated RNA nanoparticles FIG. 1 shows an exemplary flow chart of a general method for producing lipid-encapsulated RNA nanoparticles as described herein.

[0124] The geometry of the mixed layer consists of a first tube for transporting an aqueous solution having an inner diameter as described herein and a second tube for transporting an ethanol (organic) solution, consisting of an ID as described herein, where, when using the mixinib module as described herein, the second (organic) tube intersects with the first (aqueous) tube at a perpendicular angle or near a perpendicular angle.

[0125] The method described herein provides, for example, an aqueous RNA solution comprising therapeutic large RNA, which is prepared under Good Manufacturing Practice (GMP) and solubilized in an aqueous solution containing buffer, for example, citrate.The method of the present invention also provides an organic solution comprising one or more lipids, for example, clinical-grade lipids synthesized under GMP, which is prepared by solubilizing lipids in a water-miscible organic solvent.In the method described herein, the water-miscible organic solvent is preferably a lower alkanol, for example, ethanol.Preferably, both solutions are sterile filtered, and their concentrations are adjusted.

[0126] The organic lipid solution is mixed with an aqueous solution containing nucleic acid to form lipid-encapsulated RNA nanoparticles having a lamellar morphology, for example, comprising a lipid bilayer. In one embodiment, the nucleic acid is encapsulated in the lipid-encapsulated RNA nanoparticles that form a lamellar structure.

[0127] The methods described herein involve vertically introducing a lipid solution into an aqueous solution in a mixing environment, preferably in a mixing module, and diluting the lipid solution with the aqueous solution to 10%-75% v / v ethanol, 12%-70% v / v ethanol, 14%-65% v / v ethanol, 16%-60% v / v ethanol, 18%-50% v / v ethanol, 20%-45% v / v ethanol, or 22%-30% v / v ethanol, resulting in the formation of lipid-encapsulated RNA nanoparticles in a turbulent flow.

[0128] After formation of the lipid-encapsulated RNA nanoparticles, the mixture is successively diluted with buffer to about 1 to about 10% v / v ethanol, or to 7.5%, 10%, or 15%, preferably less than 12.5% ​​ethanol, which further stabilizes the lipid-encapsulated RNA nanoparticles and increases nucleic acid encapsulation.

[0129] The lipid-encapsulated RNA nanoparticles are concentrated by tangential flow filtration, preferably by a hollow fiber filter. The concentrated lipid-encapsulated RNA nanoparticles are subjected to an ultrafiltration process to remove alkanol and exchange buffer. The nucleic acid concentration is adjusted by dilution. The resulting formulation is sterile filtered and filled into vials. This process will now be discussed in more detail below, using the steps illustrated in Figure 1.

[0130] Lipid solubilization and RNA dissolution In one embodiment, the lipid-encapsulated RNA nanoparticles produced by the methods described herein are in the form of multimolecular assemblies of RNA and lipids, where the RNA is at least partially encapsulated by ion pairing with cationic lipids.

[0131] In certain embodiments, the lipid nanoparticles described herein comprise four lipid components: a helper lipid; cholesterol; a PEG lipid; and a cationic lipid. Preferably, the helper lipid is DSPC, the PEG lipid is PEG-DMG, and the cationic lipid is an ionizable cationic lipid. In certain embodiments, the organic solvent concentration at which the lipid is solubilized is about 45% v / v to about 90% v / v. In certain preferred embodiments, the organic solvent is a lower alkanol. Suitable lower alkanols include, for example, methanol, ethanol, propanol, butanol, pentanol, isomers thereof, and combinations thereof. The solvent is preferably ethanol having a concentration of about 50% v / v to 90% v / v. The lipid may occupy a volume of about 1 mL / g to about 5 mL / g, or otherwise as described in the Examples below.

[0132] The lipids are solubilized at a suitable temperature, for example, using an overhead stirrer. In certain preferred embodiments, the RNA is contained in an aqueous solution (e.g., a buffer) and diluted to a final concentration.

[0133] Formation process of lipid-encapsulated RNA nanoparticles After preparation of the organic and aqueous solutions, they can be mixed together using the apparatus described in detail below. Briefly, the apparatus consists of a first tube for transporting the aqueous RNA solution and a second tube for transporting the organic lipid solution, with the second tube intersecting the first tube perpendicularly within a mixing module. The two solutions are pumped through the corresponding tubes by separate HPLC pumps and mixed in the vertical region of the first tube within the mixing module. The aqueous RNA solution is pumped at a rate 0.2 to 1 times faster than the organic lipid solution. Mixing the two solutions in the mixing region results in the formation of lipid-encapsulated RNA nanoparticles.

[0134] The pump speed and size of the first tube in the region of the mixing module result in a turbulent mixing process. In fluid mechanics, turbulence is fluid movement characterized by chaotic changes in pressure and flow velocity. This contrasts with laminar flow, which occurs when fluids flow in parallel layers, where there are no obstructions between these layers. Turbulent flow is always highly irregular, and the rapid availability of energy in turbulent flow tends to accelerate the homogenization (mixing) of a fluid mixture. The characteristic responsible for enhanced mixing and increased rates of mass, momentum, and energy transfer in a flow is called the "diffusion coefficient." Other characteristics of turbulent flow include "swirl," in which turbulence has a powerful three-dimensional vortex generation mechanism known as vortex stretching, and "dissipation," in which turbulence rapidly dissipates as kinetic energy is converted into internal energy by viscous shear stresses. Turbulent mixing is determined by the small-scale (compared to the parent flow) random movement of regions within a fluid, bringing them into closer or more distant relationship and allowing them to be more finely divided and mixed. The process described herein for mixing lipid and aqueous solutions results in encapsulation of RNA into lipid nanoparticles, which are formed simultaneously with the formation of lipid nanoparticles, with an encapsulation efficiency of over 95%.

[0135] The continuous process described herein is fully scalable. In one embodiment, lipid-encapsulated RNA nanoparticles with a median diameter of less than about 90 nm are formed without mechanical energy processes such as membrane extrusion, sonication, or microfluidization.

[0136] Lipid-encapsulated RNA nanoparticles The lipid-encapsulated RNA nanoparticles disclosed herein comprise a nanoparticle or bilayer of lipid molecules. In addition to cationic lipids (e.g., ionizable cationic lipids), the lipid-encapsulated RNA nanoparticles comprise neutral lipids or polymers.

[0137] In some embodiments, the RNA is fully encapsulated in the lipid portion of the lipid nanoparticle, such that the RNA within the lipid-encapsulated RNA nanoparticles is resistant to nuclease degradation in aqueous solution. In other embodiments, the lipid-encapsulated RNA nanoparticles described herein are substantially non-toxic to mammals, such as humans. Lipid-encapsulated RNA nanoparticles typically have a median diameter of 30 nm to 150 nm, 40 nm to 150 nm, 50 nm to 150 nm, 60 nm to 130 nm, 70 nm to 110 nm, or 70 nm to 90 nm. Lipid-encapsulated RNA nanoparticles described herein also typically have a lipid:RNA ratio (mass / mass) of 1:1 to 100:1, 1:1 to 50:1, 5:1 to 45:1, 10:1 to 40:1, 12:1 to 38:1, 15:1 to 45:1, 25:1 to 40:1, or 30:1 to 40:1. In some embodiments, the composition has a total lipid:RNA weight ratio of about 50:1 to 10:1. In some embodiments, the composition has a total lipid:RNA weight ratio of about 40:1 to 20:1. In some embodiments, the composition has a total lipid:RNA weight ratio of about 45:1 to 30:1. In some embodiments, the composition has a total lipid:RNA weight ratio of about 38:1 to 30:1.

[0138] In a preferred embodiment, the lipid particles comprise RNA, a cationic lipid (e.g., one or more cationic lipids described herein or salts thereof), a phospholipid, and a conjugated lipid (e.g., one or more PEG-lipid conjugates) that inhibit particle aggregation. The lipid-encapsulated RNA nanoparticles can also contain cholesterol. The lipid-encapsulated RNA nanoparticles can comprise at least 1, 2, 3, 4, 5, 6, 7, 8, 9, 10, or more different RNAs expressing one or more polypeptides.

[0139] In lipid-encapsulated RNA nanoparticles, RNA can be completely encapsulated in the lipid portion of the particle, thereby protecting RNA from nuclease degradation.In a preferred embodiment, lipid-encapsulated RNA nanoparticles comprise RNA that is completely encapsulated in the lipid portion of the particle, thereby protecting RNA from nuclease degradation.In certain cases, the RNA in lipid particles is not substantially degraded after exposing the particles to nuclease at 37 ℃ for at least 20, 30, 45 or 60 minutes.In certain other cases, the RNA in lipid particles is not substantially degraded after incubating the particles in serum at 37 ℃ for at least 30, 45 or 60 minutes, or at least 2, 3, 4, 5, 6, 7, 8, 9, 10, 12, 14, 16, 18, 20, 22, 24, 26, 28, 30, 32, 34 or 36 hours.In other embodiments, RNA is complexed with cationic lipids in lipid-encapsulated RNA nanoparticles. One advantage of the formulations of the present disclosure is that the lipid-encapsulated RNA nanoparticles are substantially non-toxic to mammals, such as humans.

[0140] Lipid particles are 30%-100%, 40%-100%, 50%-100%, 60%-100%, 70%-100%, 80%-100%, 90%-100%, 30%-95%, 40%-95%, 50%-95%, 60%-95%, 70%-95%, 80%-95%, 85%-95%, 90%-95%, 30%-90%, 40%-90%, 50%-90%, 60%-90%, 70%-90%, The particles comprise RNA completely encapsulated within the lipid portion such that 80%-90%, or at least 30%, 35%, 40%, 45%, 50%, 55%, 60%, 65%, 70%, 75%, 80%, 85%, 90%, 91%, 92%, 93%, 94%, 95%, 96%, 97%, 98%, or 99% (or any fraction or range therein) have RNA encapsulated therein.

[0141] Depending on the intended use of the lipid-encapsulated RNA nanoparticles, the proportions of the components may vary, and the delivery efficiency of a particular formulation can be measured using assays known in the art.

[0142] Dilution of lipid-encapsulated RNA nanoparticles The degree of RNA encapsulation can be improved by further diluting the lipid-encapsulated RNA nanoparticle suspension after mixing the organic lipid solution with the aqueous RNA solution and before removing the free RNA. This can be achieved by diluting the suspension with one or more buffers, for example, through one or more Y-connectors flowing into the output line. The buffers flowing into the one or more Y-connectors do not need to be the same.

[0143] The diluted lipid-encapsulated RNA nanoparticles can then optionally be collected in a container maintained at 15-20° C. and allowed to incubate for a few minutes to 2 hours before further dilution or concentration steps.

[0144] Sample concentration Diluted lipid-encapsulated RNA nanoparticles can be concentrated, for example, by tangential flow filtration (TFF) using hollow fiber membranes (mPES Kros membranes, Spectrum Laboratories, Inc., Rancho Dominguez, California), optionally by peristaltic pumps (based on the principle of magnetic levitation) or four-piston diaphragm pumps or centrifugal pumps.Methods for such concentration techniques are known in the art and will be readily apparent to those skilled in the art.

[0145] Removal of free RNA and buffer exchange Diafiltration against 7-10 volumes of 10 mM Tris, 50 mM NaCl, 9% sucrose (pH 7.5) can be followed by concentration to remove organic solvent and unbound RNA. Preferably, the diafiltration buffer is added through a heat exchanger to maintain the product temperature at 15-20°C. The formulation can be further concentrated to target a total formulated RNA concentration of greater than 3 mg / mL.

[0146] Sterile filtration and filling The RNA concentration in the lipid-encapsulated RNA nanoparticle formulation is then measured by IPRP-HPLC (Ion Pair Reverse Phase-High Performance Liquid Chromatography) and optionally adjusted to approximately 2 mg / mL (1.85-2.3 mg / mL) by dilution with a glycerol-containing buffer (described below) to a final 5% glycerol concentration in the formulation. The diafiltered lipid-encapsulated RNA nanoparticles are sterile filtered through a 0.2 μm sterilizing-grade filter (PES). The filtered formulation is then aseptically filled into glass bottles, stoppered and capped, and placed at -20°C or -70°C ± 5°C.

[0147] Device The description herein provides an apparatus for carrying out the above-described process. Figure 2 shows an example of an exemplary schematic diagram of an apparatus according to one embodiment of the description herein.

[0148] An aqueous solution containing RNA is transported through tubing by an HPLC pump. An organic solution containing lipids is transported through separate tubing by an HPLC pump. The organic solution is pumped into the aqueous solution at a 90-degree angle in a mixing module. The organic solution containing lipids is introduced into the aqueous solution in a flow perpendicular to the aqueous flow. This introduction at the correct angle occurs in a mixing module such as that shown in Figure 3, resulting in turbulent mixing under carefully controlled conditions to form lipid nanoparticle encapsulations of RNA in an acceptable manner in terms of particle size, dispersion, and encapsulation efficiency. The tubing containing the mixed lipid-RNA then transports the lipid-encapsulated RNA nanoparticles to a second mixing zone, for example, via polypropylene tubing that meets the dilution buffer at a 45-degree angle in the dilution zone. The diluted lipid-encapsulated RNA nanoparticles are collected in a stainless steel-coated container maintained at 15-20°C. The particles are further processed by tangential flow filtration, for example, using a septum or centrifugal pump.

[0149] In one embodiment, the mixing region is a mixing module in which the organic lipid solution is delivered to the flow of the aqueous RNA solution, preferably at an angle of about 90°. The first stainless steel tube transporting the aqueous RNA solution has a hole in the wall midway between its two ends. The second tube is fixed vertically by a filling through the hole in the wall of the first tube, allowing the liquid to be transported from the second tube into the interior of the first tube (see Figure 3). In a preferred embodiment, the flow rate of the aqueous RNA solution reduces shear forces and allows the integrity of large RNA to be maintained, thereby preparing lipid-encapsulated RNA nanoparticles with well-defined shapes and reproducible sizes. Vesicles with well-defined shapes and reproducible sizes can also be prepared by changing the flow rate of the fluid lines, for example, to ensure sufficient mixing in some cases.

[0150] FIG. 3 illustrates a mixing module and associated flow dynamics according to one embodiment.

[0151] The description herein provides a device with hollow fiber membranes (mPES Kros membranes, Spectrum Laboratories, Inc., Rancho Dominguez, California) and a four-piston diaphragm pump or centrifugal pump.

[0152] definition The term "anionic lipid" refers to a lipid that is negatively charged at physiological pH, including, but not limited to, phosphatidylglycerol, cardiolipin, diacylphosphatidylserine, diacylphosphatidic acid, N-dodecanoylphosphatidylethanolamine, N-succinylphosphatidylethanolamine, N-glutarylphosphatidylethanolamine, lysylphosphatidylglycerol, palmitoyloleoylphosphatidylglycerol (POPG), and other anionic modifying groups attached to neutral lipids.

[0153] The term "cationic lipid" refers to amphipathic lipids and their salts that have a positive hydrophilic head group; one, two, three, or more hydrophobic fatty acid or fatty alkyl chains; and a connector between these two domains. Ionizable or protonizable cationic lipids are usually characterized by their pK a It is protonated (i.e., positively charged) at pH below pK a The cationic lipids of the present disclosure may also be referred to as titratable cationic lipids. The cationic lipids may be "amino lipids" having a protonatable tertiary amine (e.g., pH-titrateable) head group. Some exemplary amino lipids include C 18 The cationic lipids may include alkyl chains, each independently having 0 to 3 (e.g., 0, 1, 2, or 3) double bonds, and an ether, ester, or ketal bond between the head group and the alkyl chain. Examples of cationic lipids include, but are not limited to, DSDMA, DODMA, DLinDMA, DLenDMA, γ-DLenDMA, DLin-K-DMA, DLin-K-C2-DMA (also known as DLin-C2K-DMA, XTC2, and C2K), DLin-K-C3-DMA, DLin-K-C4-DMA, DLen-C2K-DMA, y-DLen-C2K-DMA, DLin-M-C2-DMA (also known as MC2), DLin-M-C3-DMA (also known as MC3), and (DLin-MP-DMA) (also known as 1-Bl 1).

[0154] The term "complementary nucleotide base" refers to a pair of nucleotide bases that form hydrogen bonds with each other. Adenine (A) pairs with thymine (T) and, in RNA, with uracil (U), and guanine (G) pairs with cytosine (C). Complementary segments or strands of nucleic acid hybridize (i.e., bind by hydrogen bonds) with each other. "Complementary" means that a nucleic acid can form hydrogen bond(s) with another nucleic acid sequence by either conventional Watson-Crick or other non-conventional bonding modes.

[0155] The term "fully encapsulated" means that the nucleic acid (for example, mRNA) in nucleic acid-lipid particles is not significantly degraded after being exposed to serum or nuclease assays, which significantly degrade free RNA.When fully encapsulated, in a process that usually degrades 100% of free nucleic acid, preferably less than 25% of the nucleic acid in the particles is degraded, more preferably less than 10%, and most preferably less than 5% of the nucleic acid in the particles is degraded."Completely encapsulated" also means that the nucleic acid-lipid particles are not rapidly degraded into their component parts when administered in vivo.

[0156] The term "nucleic acid" refers to deoxyribonucleotides or ribonucleotides and polymers thereof in single- or double-stranded form. The term encompasses synthetic, natural, and non-naturally occurring nucleic acids containing known nucleotide analogs or modified backbone residues or linkages, which have similar binding properties as the reference nucleic acid, and which are metabolized in a manner similar to the reference nucleotide. Examples of such analogs include, but are not limited to, phosphorothioates, phosphoramidates, methyl phosphonates, chiral-methyl phosphonates, 2-O-methyl ribonucleotides, and peptide-nucleic acids (PNAs).

[0157] The term "delivery" refers to the act or manner of delivering a compound, substance, element, moiety, cargo, or payload.

[0158] The term "delivery agent" refers to any substance that facilitates, at least in part, the in vivo delivery of a polynucleotide to a targeted cell.

[0159] The term "recombinant" refers to a molecule that has been engineered to have traits or properties, whether structural or chemical, that differ from the starting, wild-type, or naturally occurring molecule.

[0160] "Expression" of a nucleic acid sequence refers to one or more of the following events: (1) production of an RNA template from a DNA sequence (e.g., by transcription), (2) processing of the RNA transcript (e.g., by splicing, editing, 5' capping, and / or 3' end processing), (3) translation of the RNA into a polypeptide or protein, and (4) post-translational modification of the polypeptide or protein.

[0161] The term "hydrophobic lipid" refers to a compound having a non-polar group, including but not limited to saturated and unsaturated aliphatic hydrocarbon groups, which may be substituted with one or more aromatic, alicyclic, or heterocyclic group(s). Suitable examples include, but are not limited to, diacylglycerol, dialkylglycerol, N-N-dialkylamino, 1,2-diacyloxy-3-aminopropane, and 1,2-dialkyl-3-aminopropane.

[0162] The term "lipid" refers to organic compounds containing esters of fatty acids, characterized by being insoluble in water but soluble in many organic solvents. Lipids are generally divided into at least three classes: (1) "simple lipids," which include fats, oils, and waxes; (2) "complex lipids," which include phospholipids and glycolipids; and (3) "derived lipids," such as steroids.

[0163] The term "lipid delivery vehicle" refers to a lipid formulation that can be used to deliver a therapeutic nucleic acid (e.g., mRNA) to a desired target site (e.g., cell, tissue, organ, etc.). The lipid delivery vehicle can be a nucleic acid-lipid particle, which can be formed from a cationic lipid, a non-cationic lipid (e.g., phospholipid), a conjugated lipid that prevents particle aggregation (e.g., PEG lipid), and optionally cholesterol. Typically, the therapeutic nucleic acid (e.g., mRNA) can be encapsulated in the lipid portion of the particle to protect it from enzymatic degradation.

[0164] The term "lipid-encapsulated" refers to lipid particles that provide a therapeutic nucleic acid, e.g., mRNA, that is fully encapsulated, partially encapsulated, or both. In a preferred embodiment, the nucleic acid (e.g., mRNA) is fully encapsulated in the lipid particle.

[0165] The term "lipid complex" refers to a complex lipid that inhibits lipid particle aggregation. Such lipid complexes include, but are not limited to, PEG conjugated to dialkyloxypropyl (e.g., PEG-DAA complex), PEG conjugated to diacylglycerol (e.g., PEG-DAG complex), PEG conjugated to cholesterol, PEG conjugated to phosphatidylethanolamine, and PEG-lipid complexes such as ceramide, cationic PEG lipid, polyoxazoline (POZ)-lipid complex, PEG conjugated to polyamide oligomer, and mixtures thereof. PEG or POZ can be directly conjugated to lipid or linked to lipid via a linker moiety. Any linker moiety suitable for linking PEG or POZ to lipid can be used, including, for example, non-ester-containing linker moieties and ester-containing linker moieties. In certain preferred embodiments, non-ester-containing linker moieties such as amide or carbamate are used.

[0166] The terms "amphipathic lipid" or "amphiphilic lipid" refer to a lipid material in which the hydrophobic portion faces the hydrophobic phase, while the hydrophilic portion faces the aqueous phase. The hydrophilic character comes from the presence of polar or charged groups, such as carbohydrate, phosphate, carboxyl, sulfato, amino, sulfhydryl, nitro, hydroxyl, and other equivalent groups. Hydrophobicity can be imparted by the inclusion of nonpolar groups, including, but not limited to, long-chain saturated and unsaturated aliphatic hydrocarbon groups, and such groups substituted with one or more aromatic, alicyclic, or heterocyclic group(s). Examples of amphipathic compounds include, but are not limited to, phospholipids, aminolipids, and sphingolipids.

[0167] The term "messenger RNA" (mRNA) means any polynucleotide that encodes a protein or polypeptide of interest and can be translated to produce the encoded protein or polypeptide of interest in vitro, in vivo, in situ, or ex vivo.

[0168] "Modified" refers to a change in the state or structure of a molecule of the present disclosure. Molecules can be modified in many ways, including chemical, structural, and functional ways. In one embodiment, an mRNA molecule of the present disclosure is modified by the introduction of non-natural nucleosides and / or nucleotides, e.g., as they relate to the natural ribonucleotides A, U, G, and C. Non-standard nucleotides, such as cap structures, are not considered "modified," even though they may differ in chemical structure from the A, C, G, and U ribonucleotides.

[0169] The term "nucleotide" refers to natural (standard) and modified bases known in the art. Such bases are typically located at the 1' position of the nucleotide sugar moiety. Nucleotides typically comprise a base, a sugar, and a phosphate group. Nucleotides can be unmodified or modified at the sugar, phosphate, and / or base moieties (nucleotide analogs, modified nucleotides, non-natural nucleotides, non-standard nucleotides, and others are also referred to; see, e.g., Usman and McSwiggen, supra; Eckstein, et al., International PCT Publication No. WO 92 / 07065; Usman, et al., International PCT Publication No. WO 93 / 15187; Uhlman & Peyman, supra, all of which are incorporated herein by reference). Some examples of modified nucleobases known in the art exist, as summarized in Limbach, et al., Nucleic Acids Res. 22:2183, 1994. Some non-limiting examples of base modifications that can be incorporated into nucleic acid molecules include inosine, purine, pyridin-4-one, pyridin-2-one, phenyl, pseudouracil, 2,4,6-trimethoxybenzene, 3-methyluracil, dihydrouridine, naphthyl, aminophenyl, 5-alkylcytidines (e.g., 5-methylcytidine), 5-alkyluridines (e.g., ribothymidine, 5-halouridine (e.g., 5-bromouridine), or 6-azapyrimidines or 6-alkylpyrimidines (e.g., 6-methyluridine), propyne, and others (Burgin, et al., Biochemistry 35:14090, 1996; Uhlman & Peyman, supra). In this embodiment, "modified base" refers to a nucleotide base other than adenine, guanine, cytosine, thymine, and uracil at the 1' position, or their equivalents.

[0170] The term "open reading frame" or "ORF" refers to a nucleic acid sequence (DNA or RNA) capable of encoding a polypeptide of interest. An ORF often begins with an initiation codon, ATG, and ends with a nonsense codon or a stop codon or signal.

[0171] The term "RNA" refers to a molecule containing at least one ribonucleotide residue. "Ribonucleotide" refers to a nucleotide having a hydroxyl group at the 2' position of a β-D-ribo-furanose moiety. This term includes double-stranded RNA, single-stranded RNA, isolated RNA such as partially purified RNA, essentially pure RNA, synthetic RNA, recombinantly produced RNA, as well as altered RNA that differs from naturally occurring RNA by the addition, deletion, substitution, and / or alteration of one or more nucleotides. Such alterations can include the addition of non-nucleotide material, such as to the end(s) of an interfering RNA, or internally, for example, at one or more nucleotides of the RNA. Nucleotides in the RNA molecules of the present disclosure can also include non-standard nucleotides, such as non-naturally occurring nucleotides or chemically synthesized nucleotides or deoxynucleotides. These altered RNAs can be referred to as analogs or analogs of naturally occurring RNA. As used herein, the terms "ribonucleic acid" and "RNA" refer to molecules containing at least one ribonucleotide residue, including siRNA, antisense RNA, single-stranded RNA, microRNA, mRNA, non-coding RNA, and polyvalent RNA.

[0172] The term "targeted cell" refers to any one or more cells of interest. The cells may be found in vitro, in vivo, in situ, or in a tissue or organ of an organism. The organism may be an animal, preferably a mammal, more preferably a human, and most preferably a patient.

[0173] The term "therapeutic agent" means any agent that has a therapeutic, diagnostic, and / or prophylactic effect and / or induces a desired biological and / or pharmacological effect when administered to a subject.

[0174] The term "monomer" refers to a single unit, e.g., a single nucleic acid, that can combine with another molecule, the same or different, to form an oligomer. In some embodiments, the monomer can be an unlocked nucleic acid, i.e., a UNA monomer.

[0175] The term "neutral lipid" refers to lipid species that exist in either an uncharged or neutral zwitterionic form at a selected pH. At physiological pH, such lipids include, for example, diacylphosphatidylcholine, diacylphosphatidylethanolamine, ceramide, sphingomyelin, cephalin, cholesterol, cerebrosides, and diacylglycerol.

[0176] The term "non-cationic lipid" means an amphipathic lipid or a neutral lipid or an anionic lipid, as described herein.

[0177] The term "oligomer" can be used interchangeably with "polynucleotide" and refers to a molecule containing at least two types of monomers, including oligonucleotides such as DNA and RNA. In the case of oligomers containing RNA monomers and / or unlocked nucleic acid (UNA) monomers, the oligomers of the present disclosure can contain sequences in addition to the coding sequence (CDS). These additional sequences can be untranslated sequences, i.e., sequences that are not converted into protein by the host cell. These untranslated sequences can include a 5' cap, a 5' untranslated region (5' UTR), a 3' untranslated region (3' UTR), and a tail region, e.g., a polyA tail region. As described in more detail herein, any of these untranslated sequences can contain one or more UNA monomers—these UNA monomers cannot be translated by the host cell machinery. In the context of the present disclosure, "mRNA sequence," "mRNA sequence," "translatable polynucleotide," or "translatable compound" refers to a sequence that includes a region that can be converted into a protein or a fragment thereof, e.g., a coding region of an RNA.

[0178] The term "translatable" can be used interchangeably with the term "expressible" and refers to the ability of a polynucleotide, or portion thereof, to be converted into a polypeptide by a host cell. As understood in the art, translation is the process by which ribosomes in the cytoplasm of a cell make a polypeptide. In translation, messenger RNA (mRNA) is decoded by tRNA in the ribosomal complex to make a specific amino acid chain, or polypeptide. Furthermore, the term "translatable," when used herein in reference to an oligomer, means that at least a portion of the oligomer (e.g., the coding region (also known as coding sequence or CDS) of the oligomer sequence) can be converted into a protein or fragment thereof.

[0179] The term "translation efficiency" refers to a measure of the production of a protein or polypeptide by translation of an mRNA sequence in vitro or in vivo. The present disclosure provides a range of mRNA sequence molecules that can contain one or more UNA monomers and multiple nucleic acid monomers, and the mRNA sequences can be expressed to provide a polypeptide or protein.

[0180] Therapeutically Effective Outcome: As used herein, the term "therapeutically effective outcome" means an outcome sufficient to treat, ameliorate the symptoms of, diagnose, prevent, and / or delay the onset of an infection, disease, disorder, and / or condition in a subject suffering from or susceptible to the infection, disease, disorder, and / or condition. [Example]

[0181] Additional embodiments of the present disclosure are described in further detail in the following examples, which are not intended to limit the scope of the claims in any way.

[0182] Example 1: Preparation of large RNA-encapsulating lipid nanoparticles Dissolving RNA and lipid excipients This example outlines some general conditions used to prepare LNP-encapsulated large RNA. The lipid excipients (ionizable cationic lipid / cationic lipid:phosphate lipid:cholesterol:PEG lipid) were weighed and dissolved in 200-proof alcohol (at molar ratios of 50:X:48.5 to X:1.5, with X = 7, 10, or 13) at 40°C until dissolution was complete; dissolution time was not to exceed 4 hours. After visible dissolution, the solution was allowed to equilibrate to room temperature, and then filtered through a 0.2 μm polyethersulfone (PES) filter into a jacketed glass or stainless steel vessel. The nominal lipid concentration at this stage was 5–125 mg / mL.

[0183] The RNA was diluted in 5 mM citrate (pH 4.0) buffer containing 0-300 mM NaCl. The solution was then filtered through a 0.2 μm PES filter. The concentration of large RNA at this stage is approximately 0.096-0.765 mg / mL.

[0184] Nanoparticle formation by T-shaped stainless steel mixing module Large RNA-encapsulated lipid nanoparticles were formed by mixing an ethanolic lipid solution with an aqueous RNA solution at a controlled rate in a T-shaped stainless steel mixing module ("T-module"). Mixing involves flowing the ethanolic and aqueous solutions through the mixing module, which consists of a second tube connected perpendicularly to the first tube. An output solution containing a mixture of the two solutions flows in the direction of the original RNA flow.

[0185] The total lipid to mRNA weight ratio was set to approximately 35.88:1, although this weight ratio may vary depending on the exact size of the large RNA used and the desired lipid composition. Those skilled in the art will understand that the process described herein can be applied to lipid compositions containing any suitable combination of lipids in any suitable molar and weight ratio to RNA. A high-pressure piston pump (Knauer) was used to control the addition rate for each solution, with the lipid and mRNA solutions added at flow rates of 30-75 and 90-225 mL / min, respectively. The two streams were combined in a stainless steel mixing module at a total flow rate of 120-300 mL / min. Peek tubing was used for the high-pressure piston pump, with an ID of 0.03-0.08 inches for the RNA stream and 0.01-0.03 inches for the lipid stream.

[0186] Nanoparticle formation using a multi-inlet vortex mixer Large RNA-encapsulated lipid nanoparticles were formed by mixing an ethanolic solution of lipid with an aqueous solution of RNA at a controlled rate in a multi-inlet vortex mixer (MIVM, Holland).

[0187] The weight ratio of total lipid to mRNA was 35.88:1. An HPLC pump was used to control the addition rate of each solution, with the lipid and mRNA solutions added to each stream at a flow rate of 20-50 mL / min. The four streams were combined in a stainless steel mixing module at a total flow rate of 80-200 mL / min. Peek tubing was used for the high-pressure piston pumps, with an ID of 0.02-0.8 inches for the RNA stream and 0.01-0.03 inches for the lipid stream.

[0188] Stabilization of nanoparticles by stepwise dilution The nanoparticles thus formed are stabilized by successive in-line dilutions with buffer: first with 45 mM phosphate (pH 6.5) buffer delivered at a flow rate of 80-600 mL / min, followed by 50 mM HEPES or Tris, 50 mM NaCl, 9% (w / v) sucrose (pH 8.0) delivered at a flow rate of 240-2700 mL / min.

[0189] Concentration and buffer exchange The diluted nanoparticle formulation obtained as described above is concentrated and diafiltered against a 50 mM HEPES / 20 mM Tris, 50 mM NaCl, 9% (w / v) sucrose (pH 8.0) buffer solution by tangential flow filtration using a modified PES hollow fiber membrane with a 100 kDa MWCO. This process step ensures ethanol removal and buffer exchange. The temperature of the formulation during concentration and diafiltration is maintained between 16 and 25°C. Once ethanol removal is confirmed by Alco-Screen Alcohol Test Strips, the concentrated solution is filtered through a 0.2 μm PES filter into a glass bottle to remove potential large particulates and microbial contaminants. A sample of this filtered bulk product is collected for in-process RNA concentration analysis. The bulk product is stored at 2 to 8°C until the concentration is adjusted.

[0190] Concentration, filling, and freezing Adjust the concentration of RNA in the formulation to a target concentration of 0.2 mg / mL by adding 50 mM HEPES / 20 mM Tris, 50 mM NaCl, 9% (w / v) sucrose (pH 8.0) buffer containing glycerol to a final concentration of 6.3% (w / v) of glycerol in the buffer.

[0191] After concentration adjustment, the adjusted bulk product is filtered through a 0.2 μm PES sterilizing grade filter into a sterile collection container.

[0192] The product is aseptically filled into 2 mL Type I borosilicate glass vials with a fill volume of 1 mL (0.2 mL overfill), stoppered, and capped. All vials are frozen to ≦-55°C using a freezer dryer at a controlled rate of 0.5°C / min, or directly to -70°C. Vials are stored in a freezer maintained at -70°C ± 10°C.

[0193] Concentration adjustment, LYO excipient addition, filling, and lyophilization The RNA concentration of the formulation is adjusted to a target concentration of 0.1-0.2 mg / mL by adding 50 mM HEPES / 20 mM Tris, 50 mM NaCl, and 9% (w / v) sucrose (pH 8.0) buffer containing the appropriate LYO excipients. The formulation can then be stored at 2-8°C, -20°C, or -70±10°C prior to the lyophilization process or directly lyophilized.

[0194] Dynamic Light Scattering The mean particle size (z) and polydispersity index (PDI) of the lipid nanoparticle formulations used in the examples were measured by dynamic light scattering on a Malvern Zetasizer Nano ZS (United Kingdom).

[0195] RiboGreen assay The encapsulation efficiency of the lipid nanoparticle formulations was characterized using a RiboGreen fluorometric assay. RiboGreen is a proprietary fluorescent dye (Molecular Probes / Invitrogen, now a division of Life Technologies, part of Thermo Fisher Scientific of Eugene, Oregon, United States) used in the detection and quantification of nucleic acids, including both RNA and DNA. In its free form, RiboGreen exhibits nearly no fluorescence and has a negligible absorbance signature. Upon binding to nucleic acids, the dye fluoresces at an intensity several orders of magnitude greater than the unbound form. The fluorescence can then be detected by a sensor (fluorometer), allowing the nucleic acids to be quantified.

[0196] Acceptable LNP physicochemical properties Further experiments were performed, as described in the Examples below, to evaluate the impact of various reagents, process parameters, and equipment configurations on the quality of LNP-encapsulated large RNA formulations. The quality of these formulations was assessed by evaluating the acceptable particle size (Z or Z average), polydispersity index (PDI), and encapsulation efficiency (% Encapsulated) of the formulations. Various tested compositions were screened for meeting threshold properties, including acceptable particle size (less than 150 nm, with less than 120 nm being most preferred), PDI (<0.2), and high encapsulation efficiency (>85%).

[0197] Example 2: LNPs containing large RNAs - Initial studies In initial runs, LNPs were produced using a Precision Nano Assembler (A benchtop formulation system, Precision Nanosystems, Inc., Vancouver, BC, Canada). The composition contained cationic lipid:DSPC:cholesterol:PEG lipid in a molar ratio of 50:7:41.5:1.5. The mRNA was diluted in 5 mM citrate buffer (pH 4.0). The total lipid:RNA weight ratio was 35.2, and the combined flow rate of both streams was 12 mL / min with an EtOH:water ratio of 1:3. Dilution, purification, and concentration steps were as described in Example 1, with the first dilution ratio being 1:2 and the second dilution ratio being 1:3. Initial results comparing typical small and large nucleotide sequences are shown below in Table 1. [Table 1]

[0198] As can be seen in Table 1, using the same process, larger RNAs were encapsulated resulting in larger particle sizes, higher polydispersity index (PDI), and lower encapsulation efficiency (% encap) percentages.

[0199] Example 3: Effect of adding salt In this example, the use of NaCl in the citrate buffer was observed to improve the quality of large RNA-encapsulated lipid nanoparticles. The composition, formulation module, and mRNA were the same as those described in Example 2. The results are summarized in Table 2 below.

[0200] In the table below, "[Lipid IP] mM" refers to the in-process concentration of the two lipids after mixing the two streams without any dilution. [Table 2]

[0201] With these process improvements, a citrate buffer at pH 4.0 containing 10 mM NaCl was selected for further development because the pH 4.0 buffer also maintained good mRNA purity and integrity.

[0202] The effect of adding salt to the citrate buffer was observed across formulation compositions and different large self-replicating RNAs. It was also discovered that this buffer composition was easy to transfer to mid-scale formulation systems, including those using a multi-inlet vortex mixer. However, the buffer composition was not compatible with large-scale formulation systems, such as those using a T-type mixing module. These further results are summarized in Tables 3-5.

[0203] For formulation using a multi-inlet vortex mixer in this example, four streams were combined in a stainless steel mixing module as described in Example 1 at a total flow rate of 120 mL / min, with a lipid:RNA flow rate ratio of 1:3. Peek tubing was used for the high-pressure piston pumps with an ID of 0.03 inches for the RNA stream and 0.01 inches for the lipid stream. Dilution ratios were 1:2 and 1:3, and purification and concentration steps were as described in Example 1.

[0204] For formulations using the scalable T-module in this example, the two streams were combined in a stainless steel mixing module as described in Example 1 at a total flow rate of 300 mL / min, with a lipid:RNA flow rate ratio of 1:3. Peek tubing was used for the high-pressure piston pumps with an ID of 0.03 inches for the RNA stream and 0.01 inches for the lipid stream. Purification and concentration steps were performed as described in Example 1. [Table 3] [Table 4] [Table 5]

[0205] Example 4: Effect of Addition of Salt and pH of First Dilution Buffer To improve the process, salt addition and the pH of the first dilution buffer (45 mM phosphate buffer) were evaluated. The formulation composition and process were as described in Example 2 using a Precision Nano Assembler.

[0206] As shown in Table 6, adding NaCl to the phosphate buffer did not further improve the quality of the LNPs, resulting in larger particle sizes, higher polydispersity index (PDI), and lower encapsulation efficiency (% encap) percentages. [Table 6]

[0207] Further evaluations were performed, including varying the pH of the phosphate buffer. For this formulation, a pH of 6.5 was selected based on the pKa of the ionizable cationic lipid (approximately 6.4). As shown in Table 7, this change dramatically improved the encapsulation efficiency of the lipid nanoparticles. A molar ratio of ionizable cationic lipid:DSPC:CHOL:PEG-DMG2000 of 50:13:35.5:1.5 was used in this formulation. [Table 7]

[0208] Example 5: Effect of flow rate in MIVM and T-module systems The molar ratio of ionizable cationic lipid:DSPC:CHOL:PEG-DMG2000 was 50:10:38.5:1.5, and the weight ratio of total lipid:RNA was approximately 35.78:1. Luciferase self-replicating RNA (construct number pARM2807, 9693nt) was used in this example. The formulation process for the MIVM system and the T-module system was as described in Example 3, respectively.

[0209] Surprisingly, lower flow rates worked better for large RNA-encapsulated formulations in both the MIVM system and the formulation module system. The lower the flow rate, the smaller the particle size (Z-ave) and PDI, and the higher the % encap (as shown in Tables 8 and 9). To obtain both acceptable production rates and acceptable LNP quality, 100 mL / min for the MIVM system and 160 mL / min for the T-module system were selected for further development. [Table 8] [Table 9]

[0210] The findings in this example were opposite to those observed with the small RNA-encapsulated lipid nanoparticle formulations shown in Table 1. Faster flow rates resulted in smaller particle sizes and lower PDIs for the small RNA-encapsulated lipid nanoparticles. For the process with results for small RNA-encapsulated LNPs in Table 10, siRNAs with a length of 23 nt were used. [Table 10]

[0211] Example 6: Production of large RNA-encapsulated lipid nanoparticles in a scalable MIVM and formulation module system All of the improvements mentioned in Examples 2-5 were applied across a scalable formulation system (MIVM and T module) to develop LNP-encapsulated large RNAs (luciferase self-replicating RNA, pARM2807, 9693 nt; and another self-replicating RNA, 11,665 nt), which are applicable to different compositions (Table 11). Unless otherwise specified, the formulation conditions and processes in this example were as described in previous examples. [Table 11]

[0212] To meet the production requirements, the feasibility of a fully in-line setup for continuous production was tested. Longer tubing was used between the first and second dilutions to increase the hold time between both dilutions. This was based on the teachings of previous examples. As shown in Table 12, the quality of the large RNA-encapsulated LNPs was not affected by the fully in-line setup. [Table 12]

[0213] Example 7: Scale-up of the manufacturing process for large RNA-encapsulated lipid nanoparticles using MIVM and T-module formulation systems The formulation process described in Example 6 was shown to work well for the production of large RNA-encapsulated lipid nanoparticles, but batch volumes and production rates indicated a need for further improvement. In embodiments of this example, multiple approaches were taken to achieve this goal.

[0214] All formulation compositions, conditions, and processes in this example were the same as in Example 6 unless otherwise specified.

[0215] In-process concentration increase by adding NaCl in citrate buffer In this embodiment, the in-process concentration was increased for scale-up, and the relationship between the in-process concentration and the NaCl concentration in the citrate buffer was discovered (shown in Tables 13 and 14). This finding spans different manufacturing modules. Increasing the in-process concentration formulation required a higher NaCl concentration in the citrate buffer, which resulted in smaller particle size, PDI, and higher % encap. However, once a threshold was reached, no further increase in the high NaCl concentration was necessary. An appropriate NaCl concentration in the citrate buffer was necessary for the production of large RNA-encapsulated lipid nanoparticles, which was compatible with all in-line setups, considering the possibility of continuous large-scale manufacturing. [Table 13] [Table 14]

[0216] Decreasing the dilution ratio for scale-up To further reduce the batch volume, a lower dilution ratio was evaluated. As described in Example 1, the manufacturing process for large RNA-encapsulated lipid nanoparticles required a two-step dilution. In all previous examples, a 1:2 dilution with 45 mM phosphate buffer and a 1:3 dilution with 50 mM HEPES buffer (pH 8.0) containing 50 mM NaCl and 9% sucrose were performed. In this embodiment, lower dilution ratios were tested within the range that maintained appropriate ethanol concentrations and pH in the diluted formulation. An 8-hour hold time before the purification process (TFF) was also tested to ensure formulation stability during the large-scale manufacturing purification process (Table 15).

[0217] As shown in Table 15, the physicochemical properties of the formulation were not affected by the reduced dilution ratio of this process. After 8 hours of holding, the physicochemical properties of the formulation were also maintained. In this example, the purity and integrity of the mRNA were also measured by a fragment analyzer, ensuring that the ability of the mRNA to be tested was maintained during manufacturing. The purity and integrity of the mRNA are reported relative to the purity and integrity of the mRNA before encapsulation.

[0218] At 12 mM lipid [IP] (1:1.5 phosphate dilution followed by 1:2.5 HEPES buffer dilution) containing 50 mM NaCl in 5 mM citrate buffer (pH 4.0), the LNPs were of good quality, with stable physicochemical properties and mRNA purity for at least 8 hours before starting the purification process (TFF), ensuring stability of the LNPs during the manufacturing process. [Table 15]

[0219] Example 8: Tubing configuration and backpressure for the production of large RNA-encapsulated lipid nanoparticles For the production of smaller RNA-encapsulated lipid nanoparticles, smaller tubing (0.01, 0.02 inch ID) was often used to provide back pressure for the high-pressure piston pump used in the module. A clamp on the tubing connecting the Peek tubing to the mixing module was often used to avoid oscillations in the RNA flow. This example uses the manufacturing process developed in Example 7, but shows flaws when using 0.02 inch ID Peek tubing for the RNA flow and a clamp on the tubing between the Peek tubing and the T-module. The effects of tubing configuration and back pressure were then evaluated (Table 16). In this example, all formulation compositions, conditions, and processes were the same as in Example 7 unless otherwise noted. [Table 16]

[0220] As shown in Table 16, when using 0.02" peak tubing for RNA flow and 0.01" tubing for lipid flow during production, particle size and PDI were surprisingly large, but %encap was surprisingly small. To understand which line made the difference, the investigational setup: RNA 0.03 (50 cm); lipid: 0.01 (30 cm) was used to produce quality LNPs. Comparing these three setups, the results clearly showed that the tubing for the RNA flow had a significant impact on LNP quality. However, it was unclear whether the tubing ID, high pressure, or both were responsible.

[0221] Therefore, a series of tests with different RNA flow tubing lengths and sizes was performed (Table 17). The results showed that even at extremely short lengths, when using 0.02 inch tubing, LNP quality was affected. [Table 17]

[0222] Further investigations were performed and the results are shown in Table 18. For formulation, removing the clamps was preferable. However, when using 0.03" ID tubing for both the line and the long tubing in the RNA line (90 cm) to avoid vibration, the results were unfavorable. When using medium length tubing (48 cm), maintaining back pressure with clamps in the RNA line through the mixing module while mixing in combination with smaller tubing (0.01" ID) for the lipid flow was better for formulation. A pressure of approximately 70 psi was safe for formulation. [Table 18]

[0223] Example 9: Possible mechanism for the findings of Examples 1 to 8 Taking all of the above findings into consideration, this example discusses a possible mechanism. Because these findings only apply to large RNAs (approximately 6,000-13,000 nt), the quality of large RNA-encapsulated lipid nanoparticles was affected by the RNA concentration in the citrate buffer, the NaCl concentration in the citrate buffer, the tubing size of the RNA + citrate stream, and the flow rate of the RNA stream. The hypothesis is that large RNAs (approximately 6,000-13,000 nt) are more sensitive to shear stress and shear rate.

[0224] This finding can be explained using the following equation:

[0225] Shear rate:

number

[0226] Volume flow rate Q; inner pipe radius r.

[0227] Shear stress: With respect to a Newtonian fluid wall,

number

[0228] As will be appreciated by those skilled in the art, additional fluid dynamic viscosities and calculations can be evaluated.

[0229] Example 10: Further improvement and scale-up of the manufacturing process for large RNA-encapsulated lipid nanoparticles Unless otherwise stated, in this example, all formulation conditions and processes were the same as in Example 7, and in this example, the peak tubing size for the RNA flow had an ID of 0.04 inches.

[0230] Based on the teachings of Examples 1-9, further improvements were realized by design. In this realization, the batch volume was dramatically reduced, the production rate was greatly improved, and the quality of the large RNA-encapsulated lipid nanoparticles was improved. When the total flow rate was sufficiently fast (300 mL / min), a clamp could be applied to the RNA flow alone to avoid vibration. A pressure of approximately 80 psi for the RNA flow was shown to be a safe condition, but exceeding 100 psi adversely affected the quality of the lipid nanoparticles.

[0231] Tables 19 and 20 demonstrate significant improvements in the manufacturing process for large RNA-encapsulated lipid nanoparticles, designed based on the teachings of Examples 1-9. By combining increased flow rate, NaCl concentration in the citrate buffer, and tubing ID for the RNA flow, the batch volume was dramatically reduced while adequately maintaining LNP quality. Using a total flow rate of 300 mL / min, no clamps were required for the RNA flow. [Table 19] [Table 20]

[0232] Example 11: Further improvement and scale-up of the manufacturing process for large RNA-encapsulated lipid nanoparticles in alternative buffer systems The preparation of lyophilized RNA-encapsulated LNPs is important to provide stability to the drug product in a particular environment. Part of the lyophilization process involves preparing a suspension of LNPs in an appropriate matrix. U.S. Application No. 17 / 402,077 describes a method for lyophilizing lipid nanoparticle-encapsulated RNA, which is incorporated herein by reference. To develop a lyophilized large RNA-encapsulated lipid nanoparticle drug product, a manufacturing process for large RNA-encapsulated lipid nanoparticles in a Tris buffer system was developed based on the teachings of Examples 1-10. The formulation conditions and process setup were the same as in Example 10, with the entire in-line setup having a hold time of 15-25 seconds. The only difference was that the second dilution buffer was 50 mM Tris (pH 8.0) containing 50 mM NaCl and 9% sucrose, and the diafiltration buffer was 20 mM Tris (pH 8.0) containing 50 mM NaCl and 9% sucrose (Table 21).

[0233] The teachings of Examples 1-10 were all applicable to formulation in a Tris buffer system. [Table 21]

[0234] Example 12: Addition of EDTA during dilution and final validation of the manufacturing process of large RNA-encapsulated lipid nanoparticles In this example, all formulation conditions and process setup were the same as in Example 11, except where otherwise specified. The effect of adding EDTA during the second dilution and removing it during diafiltration was tested in this example, and a final highly scaled-up manufacturing process was confirmed with a 20 second hold time between the first and second dilution, dilution ratios of 1:15 and 1:2, and a lipid IP concentration of 28 mM, with and without EDTA addition, to produce high-quality large RNA-encapsulated lipid nanoparticles (Table 22). [Table 22]

[0235] Example 13: Effect of manufacturing process improvements on large RNA-encapsulated lipid nanoparticles on in vivo efficacy In this embodiment, Figure 4 shows that not only were the physicochemical properties of the large RNA-encapsulated lipid nanoparticles well maintained and improved using the scaled-up process described herein, but also the in vivo efficacy was maintained and improved. Table 23 shows that the manufacturing differences between these two processes, all formulation conditions, and other process setups other than those described in Table 23, were the same as in Example 7.

[0236] In this embodiment, 50 μL of large RNA-encapsulated lipid nanoparticles containing RNA encoding the COVID spike protein were injected intramuscularly into both legs of Balb / c mice at the doses shown in Figure 4. There were five mice per group. After 40 days, serum was collected and tested using a Luminex assay—a bead-based multiplexed immunoassay system in a microplate format—to confirm the expression levels of COVID spike protein antibodies. [Table 23]

[0237] Further Considerations The foregoing description is provided to enable one skilled in the art to implement the various configurations described herein. There may be many other ways of implementing the subject technology. The various functions and elements described herein may be separated differently than shown without departing from the scope of the subject technology. Various modifications to these configurations will be readily apparent to those skilled in the art, and the general definitions defined herein may be applicable to other configurations. Thus, many changes and modifications may be made to the subject technology by one skilled in the art without departing from the scope of the subject technology.

[0238] While the detailed description contains many specific examples, these should not be construed as limiting the scope of the subject technology, but merely as illustrating different examples and aspects of the subject technology. It should be understood that the scope of the subject technology includes other embodiments not discussed in detail above. Various other modifications, changes, and variations can be made in the arrangement, operation, and details of the methods and apparatus of the subject technology disclosed herein without departing from the scope of the present disclosure. In addition, a device or method need not solve every problem that can be solved (or have every advantage that can be realized) by different embodiments of the present disclosure to be within the scope of the present disclosure. As used herein, "can" and its derivatives should be understood to mean "possibly" or "optionally," as opposed to categorical possibility.

Claims

1. a) flowing an aqueous solution containing RNA into a first tube having an inner diameter (ID) of about 0.01 inches to about 0.08 inches, wherein the pH of said aqueous solution ranges from about 3.0 to about 4.5, with an optional NaCl concentration of up to about 300 mM; the RNA comprises about 6,000 to about 13,000 nucleotides; b) flowing an ethanol solution containing lipids through a second tube having an ID of about 0.01 inches to about 0.04 inches at a flow rate of about 0.2 to about 1 times the flow rate of the aqueous solution through the first tube, wherein the lipids comprise cationic lipids; c) mixing the ethanol solution with the aqueous solution, said mixing producing an output solution flowing through said first tube containing a turbulent flow of said RNA and said lipids at about 10% to 75% ethanol (v / v); the lipid-encapsulated RNA nanoparticles have a bilayer structure; A method for producing the lipid-encapsulated RNA nanoparticles, comprising:

2. a) flowing an aqueous solution containing RNA into a first tube having a first inner diameter (ID), the RNA comprises about 6,000 to about 13,000 nucleotides; b) flowing an ethanol solution containing lipids through a second tube having a second inner diameter (ID) at a flow rate of about 0.2 to about 1 times the flow rate of the aqueous solution through the first tube, wherein the lipids comprise cationic lipids; c) mixing the ethanol solution with the aqueous solution, the first ID and the second ID and the flow rates through the first tube and the second tube are selected to produce shear forces low enough to protect the integrity of the RNA; said mixing producing an output solution flowing through said first tube containing a turbulent flow of said RNA and said lipids at about 10% to 75% ethanol (v / v); the lipid-encapsulated RNA nanoparticles have a bilayer structure; A method for producing the lipid-encapsulated RNA nanoparticles, comprising:

3. 3. The method of claim 1, wherein the mixing comprises flowing the ethanol solution and the aqueous solution through a mixing module comprising the second tube perpendicularly connected to the first tube.

4. 3. The method of claim 1 or 2, wherein the mixing comprises passing the ethanol solution and the aqueous solution through a multi-inlet vortex mixer.

5. 5. The method of claim 1, wherein the concentration of RNA in the aqueous solution ranges from about 85 micrograms / mL to about 2100 micrograms / mL.

6. 6. The method of claim 1, wherein the concentration of lipid in the ethanol solution ranges from about 5.0 mg / mL to about 125 mg / mL.

7. 7. The method of any one of claims 1 to 6, wherein the aqueous solution is pumped through the first tube by a first pump and the ethanol solution is pumped through the second tube by a second pump at a back pressure of about 200 psi or less.

8. The method of claim 1 , wherein the first tube has an ID ranging from about 0.02 inches to about 0.03 inches and the second tube has an ID ranging from about 0.01 inches to about 0.02 inches.

9. The method of claim 1 , wherein the first tube has an ID of about 0.02 inches and the second tube has an ID of about 0.01 inches.

10. The method of claim 1 , wherein the first tube has an ID of about 0.03 inches and the second tube has an ID of about 0.01 inches.

11. The method of claim 2, wherein the first tube has an ID ranging from about 0.01 inches to about 0.08 inches and the second tube has an ID ranging from about 0.01 inches to about 0.04 inches.

12. The method of claim 2, wherein the first tube has an ID ranging from about 0.02 inches to about 0.03 inches and the second tube has an ID ranging from about 0.01 inches to about 0.02 inches.

13. 13. The method of any one of claims 1 to 12, wherein the aqueous solution is pumped at a flow rate ranging from about 40 mL / min to about 375 mL / min.

14. 14. The method of any one of claims 1 to 13, wherein the ethanol solution is pumped at a flow rate ranging from about 10 mL / min to about 75 mL / min.

15. 15. The method of any one of claims 1 to 14, wherein the output solution has a total flow rate in the range of about 120 mL / min to about 300 mL / min.

16. 16. The method of any one of claims 1 to 15, wherein the aqueous solution, the ethanol solution, and the output solution are maintained at a temperature in the range of about 10°C to about 25°C.

17. 17. The method of any one of claims 1 to 16, further comprising pumping a first dilution buffer and introducing the dilution buffer into the output solution, thereby mixing the dilution buffer with the output solution to create a first diluted output solution.

18. 18. The method of claim 17, further comprising pumping a second dilution buffer into the first diluted output solution to form a final diluted output solution, wherein there is a delay between pumping the first dilution buffer and pumping the second dilution buffer.

19. 20. The method of claim 18, wherein the delay is from about 0.1 to about 30 seconds, and the delay is created by a length of tubing.

20. the first dilution buffer solution comprising: a) a buffer having a pH of about 5.5 to about 7.0; b) optionally, a sodium chloride concentration of up to about 100 mM; and The method according to any one of claims 17 to 19, comprising:

21. the second dilution buffer comprises: a) a buffer having a pH of about 7.4 to 8.0; b) optionally, a sodium chloride concentration of up to about 100 mM; and The method according to any one of claims 18 to 20, comprising:

22. 22. The method of any one of claims 18 to 21, wherein the second buffer comprises up to 15% w / v sucrose.

23. 23. The method of any one of claims 18 to 22, wherein the second buffer comprises up to about 0.5% w / v of an antioxidant.

24. 24. The method of any one of claims 18 to 23, wherein the second buffer comprises up to 20 mM of a chelating agent.

25. 25. The method of any one of claims 17 to 24, wherein the first diluted output solution comprises about 1.0% to about 10.0% ethanol.

26. 26. The method of any one of claims 17 to 25, wherein the first dilution buffer is pumped at a flow rate of about 80 mL / min to about 900 mL / min.

27. 27. The method of any one of claims 18 to 26, wherein the second dilution buffer is pumped at a flow rate of about 240 mL / min to about 5400 mL / min.

28. The cationic lipid has the structure of Formula I: 【Chemistry 1】 or a pharmaceutically acceptable salt or solvate thereof. [In the formula, R ５ and R ６ are each independently a straight-chain or branched C １ -C ３１ Alkyl, C ２ -C ３１ alkenyl, or C ２ -C ３１ selected from the group consisting of alkynyl, and cholesteryl; L ５ and L ６ are each independently a linear C １ -C ２０ Alkyl and C ２ -C ２０ alkenyl, X ５ is —C(O)O— or —OC(O)—, X ６ is —C(O)O— or —OC(O)—, X ７ is S or O, L ７ is absent or lower alkyl, R ４ is a straight or branched C １ -C ６ alkyl, and R ７ and R ８ are each independently hydrogen and a straight-chain or branched C １ -C ６ and alkyl.

29. 29. The method of any one of claims 1 to 28, wherein the lipid-encapsulated RNA nanoparticles have an average particle size ranging from about 50 nm to about 120 nm.

30. 30. The method of any one of claims 1 to 29, wherein the lipid-encapsulated RNA nanoparticles have an average particle size ranging from about 70 nm to about 90 nm.

31. 31. The method of any one of claims 1 to 30, wherein the polydispersity of the lipid-encapsulated RNA nanoparticles does not exceed about 0.

2.

32. 32. The method of any one of claims 1 to 31, wherein the lipid portion of the lipid-encapsulated RNA nanoparticles further comprises one or more agents selected from the group consisting of helper lipids, cholesterol, and PEG-lipid conjugates.

33. 33. The method of any one of claims 1 to 32, wherein the RNA is self-replicating RNA.

34. 34. The method of any one of claims 1 to 33, further comprising lyophilizing the final diluted output solution.

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