Patent Application
20240384269
The invention relates generally to therapeutic compositions that include nucleic acid nanoparticles for delivery of cargo and methods of using the same.
Nucleic acids (NAs) nanotechnology offers an exciting opportunity to assemble structures with precise control of their properties at the nanoscale. By harnessing the base-pairing interactions, the size, shape and placement of cargo molecules of the self-assembled construct can be fully controlled. Interestingly, the resulting structures have different properties compared to their linear components. For example, they can resist enzymatic degradation or can be retained in tissues longer. Some structures have even been shown to enter cells without the need for transfection agents. Therefore, NA constructs have gained interest for various biological applications, including imaging, sensing, and delivery. They offer a particularly interesting platform to improve NA therapeutics delivery. Indeed, while NA therapeutics, made of linear oligonucleotides, have a huge potential to address unmet pharmaceutical needs, they still suffer from challenges that need to be urgently addressed: their delivery is restricted to the liver, they have poor cellular uptake and are retained in the endosome.
By integrating NAs therapeutics within the programmable assemblies, the PK/PD properties are expected to be altered, resulting in new distribution and cellular uptake profiles.
Indeed, the shape, size, charge, and hydrophobicity are known parameters that affect tissue distribution. While the shape and size can be tuned by changing structural design, the other parameters can be changed by integrating chemical modifications. Oligonucleotides are synthesized with precise sequence control and offer a platform to introduce selected chemical modifications at the desired stoichiometry. For example, 2′F-modifications are introduced in RNA strands to increase nuclease resistance. One of the main goals of affecting PK/PD properties is to achieve precise tissue and cell-targeting-a particularly urgent need in the context of cancer treatments, where current therapies are efficient, but cause too many off-target side effects.
In this invention, we describe how to introduce chemical modifications within the composition in an efficient manner, to systematically screen the resulting effects of both the position and the nature of the modification. Ultimately, this will result in favorable PK/PD properties and improve therapeutics delivery. This invention is particularly exciting in the context of tumor delivery of NA therapeutics, but not limited to, and can be applied to the delivery of all types of therapeutics, as the composition can be conjugated with small molecules, peptides, antibodies and any relevant therapies.
Use of phosphoramidite chemistry to alter the physicochemical characteristics of nucleic acid nanoparticles Generation of RNA strands and functionalization post-synthetically is ideal for a research and development setting, however there may be issues with scale-up, particularly for some bioconjugation reactions. To overcome these potential pitfalls, it would be extremely useful to develop a new type of phosphoramidite to which any modification could be attached. These should be subdivided into two distinct categories: universal internal modifications and polymeric tails that can drastically alter both the pharmacokinetics (PK) and pharmacodynamics (PD) of the nucleic acid nanoparticles (for example
The universal internal modification will be inosine modified with an appropriate reactive click handle. Inosine is considered to be a “universal base”, as it can form hydrogen bonds with all four canonical bases (G. Butora, D. M. Kenski, A. J. Cooper, W. Fu, N. Qi, J. J. Li, W. M. Flanagan, I. W. Davies, Nucleoside Optimization for RNAi: A High-Throughput Platform, J. Am. Chem. Soc. 133 (2011) 16766-16769). From a synthetic point of view, gram-scale synthesis of this universal base would be simpler than modifying all four bases (A, G, C, U). Alternatively, to inosine, any of the following moieties might be used as a universal base: 2′-deoxynebularine. 3-nitropyrrole 2′-deoxynucleoside, 5′-nitroindole 2′-deoxynucleoside (D. Loakes, D. M. Brown, 5-Nitroindole As an Universal Base Analogue, Nucleic Acids Res. 22 (1994) 4039-4043), 6H, 8H-3,4-dihydro-pyrimido[4,5-c][1,2] oxazin-7-one (P) and 2-amino-9-(2-deoxy-β-ribofuranosyl)-6-methoxyaminopurine.
The aforementioned modifications could be used for coupling in solution. However, a solid phase synthesis approach is also possible. Solid phase oligonucleotide synthesis allows one to introduce a wide variety of different chemically modified moieties into the sequence utilizing phosphoramidite building blocks. The presence of a 4,4′-dimethoxytrityl (DMT) group at the modification of interest allows further elongation of a growing oligonucleotide resulting in formation of an oligonucleotide that can contain different types of customized moieties, such as spacers, linkers or molecules that alter the overall physicochemical properties (K. Bartosik, K. Debiec, A. Czarnecka, E. Sochacka, G. Leszczynska, Synthesis of nucleobase-modified RNA oligonucleotides by post-synthetic approach, Molecules. 25 (2020)).
One very interesting exploitation of such modifications in the nanoparticle field is an approach to generate a desired conjugate straight during the solid phase oligonucleotide synthesis (T. Yamamoto, C. Terada, K. Kashiwada, A. Yamayoshi, M. Harada-Shiba, S. Obika, Synthesis of Monovalent N-Acetylgalactosamine Phosphoramidite for Liver-Targeting Oligonucleotides, Curr. Protoc. Nucleic Acid Chem. 78 (2019) e99); (I. Cedillo, D. Chreng, E. Engle, L. Chen, A. K. McPherson, A. A. Rodriguez, Synthesis of 5-GalNAc-conjugated oligonucleotides: A comparison of solid and solution-phase conjugation strategies, Molecules. 22 (2017) 1-12). In contrast to post-synthetic conjugation in solution, this approach allows one to potentially reduce the number of impurities, time and resources and can significantly improve the overall yield of a conjugation.
Herein, we present a successful incorporation of a disulfide linkage into the growing oligonucleotide to increase an overall yield of siRNA-oligonucleotide conjugation when compared to standard approach in solution.
As an extension to the solid phase approach, a potentially straightforward way of altering PK/PD without having too much of an effect on assembly would be to develop a simplified “modifier”-type molecule that can be extended to form oligophosphates. Synthesis of these types of molecules has been explored in the literature (D. de Rochambeau, Y. Sun, M. Barlog, H. S. Bazzi, H. F. Sleiman, Modular Strategy To Expand the Chemical Diversity of DNA and Sequence-Controlled Polymers, J. Org. Chem. 83 (2018) 9774-9786) and is known in the art (US 2019/0060324), however, using them for specific modulation of PK/PD properties has not been widely explored. The flexibility and control of solid phase phosphoramidite synthesis will also allow for a high degree of fine-tuning and will be possible at a high-throughput level.
In addition to these novel PK/PD altering phosphoramidites, the proposed invention will also utilise bioconjugation strategies using known and novel compounds, which are described herein (M. L. W. J. Smeenk, J. Agramunt, K. M. Bonger, Recent developments in bioorthogonal chemistry and the orthogonality within, Curr. Opin. Chem. Biol. 60 (2021) 79-88); ([1] B. L. Oliveira, Z. Guo, G. J. L. Bernardes, Inverse electron demand Diels-Alder reactions in chemical biology, Chem. Soc. Rev. 46 (2017) 4895-4950). Functionalization can be performed according to proposed strategies either on pre-assembled constructs or directly to the core strands followed by assembly. The reactive moiety can be introduced more than once to the 5′ end of the core strand or as various combinations of reactive moieties following the principles of orthogonal labelling.
Alterations to Nucleic Acid Nanoparticles to Alter their Physicochemical Properties
Accessing the right organ is one of the most important steps for any delivery agent. The organs can be targeted by modulating the charge, size, and the shape of the agent, along with the protein corona. Peptides are naturally occurring biopolymers that possess different amino acid side chains with varying desirable properties for charge, size, and nature. Depending on the chosen amino acids, peptides can be classified as positively charged, negatively charged or neutral and hydrophilic or hydrophobic peptides. Such a diverse set of peptides can contribute to alter PK/PD properties of cargo (see for example
L-RNA is the left-turning and mirror image version of natural RNA, as opposed to the naturally occurring right-turning version called D-RNA (
Many different chemical strategies are currently used to successfully deliver a molecule of interest into a biological target. As a result of bioconjugation strategies (see for example Table 2 and Table 3), upon which a new covalent bond is formed, many targeting units, i.e., peptides, can be attached to a therapeutic molecule in order to increase internalization into tumor cells. The nature of a chemical bond allows scientists to predict not only the design of the most efficient pathway for successful delivery but also to control the mechanism of release of the therapeutic moiety. (E. I. Vrettos, T. Karampelas, N. Sayyad, A. Kougioumtzi, N. Syed, T. Crook, C. Murphy, C. Tamvakopoulos, A. G. Tzakos, Development of programmable gemcitabine-GnRH pro-drugs bearing linker controllable “click” oxime bond tethers and preclinical evaluation against prostate cancer, Eur. J. Med. Chem. 211 (2021) 113018); (M. Dirin, E. Urban, B. Lachmann, C. R. Noe, J. Winkler, Concise postsynthetic preparation of oligonucleotide-oligopeptide conjugates through facile disulfide bond formation, Future Med. Chem. 7 (2015) 1657-1673).
Increasing the complexity of the delivery system by adding more units, especially if more than one chemical approach has been utilized, increases the complexity of the synthesis, and can have a counterproductive effect on biological activity. To overcome any potential issues, the applied synthetic strategy should be designed and executed in a way that the newly formed molecule is compatible with all subsequent modifications and therefore the previously introduced bonds remain intact throughout the whole manufacturing process in addition to surviving metabolic pathways further downstream.
The main goal of that application is to improve the loading capacity, targeting and controlled release of each component as a result of attaching a therapeutic together with targeting moiety into the drug delivery system. This can be achieved by implementing effective design changes to the nucleic acid nanoparticle.
The efficiency of a drug delivery vehicle can be estimated with how much drug reaches the target. Minimum dosage to reach therapeutic activity is the target. Increasing the number of therapeutic cargoes loaded onto the drug delivery systems, e.g., prepare multivalent systems, is one way to achieve this. Multivalency aims at delivering multiple drugs at once for synergistic effects and/or higher therapeutic index. Multivalent strategies have been shown to increase efficacy, e.g., divalent siRNAs where effects can be seen up to 6 months post-treatment (J. F. Alterman, B. M. D. C. Godinho, M. R. Hassler, C. M. Ferguson, D. Echeverria, E. Sapp, R. A. Haraszti, A. H. Coles, F. Conroy, R. Miller, L. Roux, P. Yan, E. G., Knox, A. A. Turanov, R. M. King, G. Gernoux, C. Mueller, H. L. Gray-Edwards, R. P. Moser, N. C. Bishop, S. M. Jaber, M. J. Gounis, M. Sena-Esteves, A. A. Pai, M. DiFiglia, N. Aronin, A. Khvorova, A divalent siRNA chemical scaffold for potent and sustained modulation of gene expression throughout the central nervous system, Nat. Biotechnol. 37 (2019) 884-894). Yet, common synthetic approaches span from two extremes: either two to three molecules can be attached, or hundreds (G. Yamankurt, R. J. Stawicki, D. M. Posadas, J. Q. Nguyen, R. W. Carthew, C. A. Mirkin, The effector mechanism of siRNA spherical nucleic acids, Proc. Natl. Acad. Sci. U.S.A 117 (2020) 1312-1320). Multivalent systems carrying therapeutic moieties with perfect control over their identity and numbers are still lacking (for example, the ability to attach three different drugs in different ratios). Currently attachment of functional RNAs to nucleic acid-based delivery vehicles is achieved by strand hybridization or using various bioconjugation chemistries such as CuAAC, IEDDA, SPAAC, etc. (Q. Hu, S. Wang, L. Wang, H. Gu, C. Fan, DNA Nanostructure-Based Systems for Intelligent Delivery of Therapeutic Oligonucleotides, Adv. Healthc. Mater. 7 (2018) 1-19); (H. Zhang, G. S. Demirer, H. Zhang, T. Ye, N. S. Goh, A. J. Aditham, F. J. Cunningham, C. Fan, M. P. Landry, DNA nanostructures coordinate gene silencing in mature plants, Proc. Natl. Acad. Sci. U.S.A 116 (2019) 7543-7548); (H. Xue, F. Ding, J. Zhang, Y. Guo, X. Gao, J. Feng, X. Zhu, C. Zhang, DNA tetrahedron-based nanogels for siRNA delivery and gene silencing, Chem. Commun. 55 (2019) 4222-4225); (K. Astakhova, R. Ray, M. Taskova, J. Uhd, A. Carstens, K. Morris, “Clicking” Gene Therapeutics: A Successful Union of Chemistry and Biomedicine for New Solutions, Mol. Pharm. 15 (2018) 2892-2899).
The proposed approach will simplify how to increase loading capacity, while allowing to retain full control over the stoichiometry. Additionally, the approach makes processes more environmentally friendly and sustainable by minimizing the amounts of material required for delivery vehicles, cargo to be loaded (such as functional RNAs) as well as reagents needed.
The invention shows a novel use of incorporation of branching phosphoramidites in adding more functional NAs such as siRNA, mRNA, miRNA, shRNA, InRNA, antisense oligonucleotides, aptamers etc. to the composition. Branching units will be introduced in one or more of the nucleic acid nanoparticle component strands (for example
Scaffolded origami offers a programmable nanoscale platform for the controlled self-folding of nucleic acids into arbitrary geometric shapes with precisely defined properties (
In origami nanostructures, a scaffold nucleic acid molecule of up to several thousand bases in length is packed into bundles of double-helical structure. The “glue” that holds these bundles together is Watson-Crick base pairing between complementary sequence segments. Typically, though not necessarily required, hundreds of staple strands are designed to hybridize to two or more segments of the scaffold, thereby creating crosslinks between neighboring helix bundles (
Staple strands can be manufactured by standard solid-phase oligonucleotide synthesis, which allows on-column incorporation of backbone and nucleotide modifications such as 2′Fluoro (2′F), 2′O-Methyl (2′OMe) and phosphorothioate linkages. Biological methods for staple strand production based on bacteriophages (F. Praetorius, B. Kick, K. L. Behler, M. N. Honemann, D. Weuster-Botz, H. Dietz, Biotechnological mass production of DNA origami, Nature. 552 (2017) 84-87) or rolling circle amplification (C. Ducani, C. Kaul, M. Moche, W. M. Shih, B. Hogberg, Enzymatic production of “monoclonal stoichiometric” single-stranded DNA oligonucleotides, Nat. Methods. 10 (2013) 647-652); (T. L. Schmidt, B. J. Beliveau, Y. O. Uca, M. Theilmann, F. Da Cruz, C.-T. Wu, W. M. Shih, Scalable amplification of strand subsets from chip-synthesized oligonucleotide libraries, Nat. Commun. 6 (2015) 8634) have also been published but are primarily used for the synthesis of unmodified strands.
By introducing modifications in staple strands, the molecular characteristics of origami nanostructures can be tuned. For example, but without limitation, a targeting group can be attached to target specific cells. Similarly, a cell penetrating peptide or lipid can be attached to help overcome certain physical barriers like the endosome. Staple strands modified with 2′F, 2′OMe and/or phosphorothioate linkages may be used to control immunomodulation and serum stability.
Independent of the staple strands, the folding of the nucleic acid into a more compressed structure could make it less immunogenic and less prone to nuclease degradation (in comparison to single stranded mRNA), improving the safety and half-life of the nucleic acid drug. Furthermore, the size and shape of the origami could be designed to direct NA therapeutics to specific organs. For instance, particles with sizes of more than 100 nm are likely to accumulate in the spleen and liver. Hence, the specific folding of the origami gives control over the biodistribution and potential therapeutic targets.
Methods for the Decoration of NA Origamis with Functional Groups
In compositions of the invention, cargo molecules (see for example Table 5) may be attached to nucleic acid origami structures, functional elements, or both via linkers. The attachments may be covalent or non-covalent. The attachments may be reversible. Particularly useful are reversible attachments that bind the cargo molecule to the nanoparticle or functional element while the composition is being transported to a target and then release the cargo molecule from the nanoparticle or functional element when the cargo molecule has been delivered to the target. Examples of reversible linkers that may be used in compositions of the invention include acetals, acid-labile linkages, amino esters, azide-alkyne bonds, biotin-streptavidin linkages, disulfide bonds, dithiopyridyls, enzymatically cleavable linkages, hydrazones, imines, maleic anhydrides, maleimides, nucleotide base pairs, ribozyme linkages, Schiff-base linked imidazoles, thioethers, and triethylene glycol (see for example Table 4).
Efficient delivery of nucleic acid therapeutics to target cells remains one of the greatest challenges in the field. Naked administration can lead to uptake by macrophages, dendritic cells, and lung epithelial cells (M. Y. T. Chow, Y. Qiu, J. K. W. Lam, Inhaled RNA Therapy: From Promise to Reality. Trends Pharmacol Sci. 41(2020) 715-729). To achieve cellular internalization in other cell types, however, delivery vehicles are required. LNPs are currently the leading choice and the most clinically advanced vehicles. A typical LNP consists of (i) an amino lipid that aids NA encapsulation, cellular uptake, endosomal escape, and improves tolerability, (ii) a phospholipid that stabilizes the bilayer and contributes to endosomal escape, (iii) cholesterol or a sphingolipid for enhanced stability; and (iv) polyethylene glycol (PEG) to reduce nonspecific binding to proteins and increase bioavailability.
(G) RNA and/or DNA drug, e.g., mRNA. (H) Complementary binding of staple to nucleic acid drug. Modifications to staple free end is designed to alter physiochemical properties. (I) Staple complementary binding from nucleic acid drug to another nucleic acid cargo. (J) Multiple cargoes can be linked onto the origami construct.
In certain embodiments, compositions of the invention include nanoparticles. As used herein, “nanoparticle” refers to particles having dimensions that are measured on the nanometer scale. For example, a nanoparticle may have a diameter, length, width, or depth of from 1 to 1000 nm.
RNA nanoparticles are formed from the ordered arrangement of individual RNA molecules having defined secondary structures. RNA molecules form a variety of structural motifs, such as pseudoknots, kissing hairpins, and hairpin loops, that affect both the geometry of the molecule and its ability to form stable interactions with other RNA molecules via base pairing. Typically, individual RNA molecules have double-stranded regions that result from intramolecular base pairing and single-stranded regions that can for base pairs with other RNA molecules or can otherwise bind to other types of molecules.
Various RNA nanostructures having ordered two-dimensional or three-dimensional structures are known, including, for example and without limitation, nanoarrays, nanocages, nanocubes, nanoprisms, nanorings, nanoscaffolds, and nanotubes. Nanorings may be symmetrical structures that include 3, 4, 5, 6, 7, 8, or more RNA molecules arrayed around an axis. Thus, nanorings may be trimers, tetramers, pentamers, hexamers, heptamers, oxamers, or higher-numbered polymers. Nanorings may be circular, triangular, square, pentagonal, hexagonal, heptagonal, octagonal, or otherwise polygonal in shape. Other types of RNA nanoparticles, such as sheets, cages, dendrimers and clusters, are also possible and within the scope of the invention. “Nanoscaffold” refers generally to a nanostructure to which other molecules can be attached. RNA nanoparticles of various structural arrangements are described in, for example, WO 2005/003,293; WO 2007/016,507; WO 2008/039,254; WO 2010/148,085; WO 2012/170,372; WO 2015/042,101; WO 2015/196,146; WO 2016/168,784; and WO 2017/197,009, the contents of each of which are incorporated herein by reference. Nucleic acid nanoparticles may contain naturally occurring nucleotides, or they may contain chemically modified nucleotides (for example
In certain embodiments, compositions of the invention include phosphoramidites that provide stimuli-responsive characteristics to the nucleic acid nanoparticle. The first aspect of the invention relates to a compound of formula (I):
Examples of suitable groups for R′ are provided below:
R″ may be synthesized from the group consisting of, but not limited to, ADIBO-PEG4, N-[(1R,8S,9s)-bicyclo[6.1.0]non-4-yn-9-ylmethyloxycarbonyl]-1,8-diamino-3,6-dioxaoctane, (1R,8S,9s)-bicyclo[6.1.0]non-4-yn-9-ylmethanol, bromoacetamido-dPEG®4-amido-DBCO, bromoacetamido-dPEG®12-amido-DBCO, bromoacetamido-dPEG®24-amido-DBCO, dibenzocyclooctyne-acid, dibenzocyclooctyne-N-hydroxysuccinimidyl ester, dibenzocyclooctyne-PEG4-acid, dibenzocyclooctyne-PEG4-alcohol, dibenzocyclooctyne-PEG4-N-hydroxysuccinimidyl ester, (4-(1,2,4,5-tetrazin-3-yl)phenyl)methanamine hydrochloride, (E)-cyclooct-4-enol, (E)-cyclooct-4-enyl 2,5-dioxo-1-pyrrolidinyl carbonate, 2,5-Dioxo-1-pyrrolidinyl 5-[4-(1,2,4,5-tetrazin-3-yl)benzylamino]-5-oxopentanoate, 5-[4-(1,2,4,5-tetrazin-3-yl)benzylamino]-5-oxopentanoic acid, 5-norbornene-2-acetic acid succinimidyl ester, 5-norbornene-2-endo-acetic acid, methyltetrazine-NHS ester, methyltetrazine-PEG4-NHS ester, TCO PEG4 succinimidyl ester, TCO-amine, tetrazine-PEG5-NHS ester, alkyne-PEG5-acid, (R)-3-amino-5-hexynoic acid hydrochloride, (S)-3-amino-5-hexynoic acid hydrochloride, (S)-3-(boc-amino)-5-hexynoic acid, N-boc-4-pentyne-1-amine, boc-propargyl-Gly-OH, 3-ethynylaniline, 4-ethynylaniline, propargylamine hydrochloride, propargyl chloroformate, propargyl-N-hydroxysuccinimidyl ester, propargyl-PEG2-acid, 3-(4-azidophenyl)propionic acid, 3-azido-1-propanamine, 3-azido-1-propanol, 4-carboxybenzenesulfonazide, O-(2-aminoethyl)-O′-(2-azidoethyl)heptaethylene glycol, O-(2-aminoethyl)-O′-(2-azidoethyl)nonaethylene glycol, O-(2-aminoethyl)-O′-(2-azidoethyl)pentaethylene glycol, azido-dPEG®4(n)acid (where n could be 4, 8, 12, 24), azido-dPEG® (n)-amine (where n could be 7, 11, 23, 35), azido-dPEG®4(n) NHS ester (where n could be 4, 8, 12, 24), azido-dPEG® (n)-TFP ester (where n could be 4, 8, 12, 24, 36), 2-[2-(2-azidoethoxy)ethoxy]ethanol, O-(2-azidoethyl)-O-[2-(diglycolyl-amino)ethyl]heptaethylene glycol, O-(2-azidoethyl)heptaethylene glycol, O-(2-azidoethyl)-O′-methyl-triethylene glycol, O-(2-azidoethyl)-O′-methyl-undecaethylene glycol, 17-azido-3,6,9,12,15-pentaoxaheptadecan-1-amine, 14-azido-3,6,9,12-tetraoxatetradecanoic acid, 11-azido-3,6,9-trioxaundecan-1-amine, bromoacetamido-dPEG® (n)azide (where n could be 3, 11, 23).
Other examples of phosphoramidites included are provided below.
In addition to direct attachment of singular cargo molecules at each attachment point on a nanoparticle, compositions of the present invention may also be used to modify oligonucleotides so that they can be linked to cargo molecules, which are then linked to other cargo molecules (for example
These linked cargo molecules, also referred to as ‘Combinatorial chains’, could include, but are not limited to, molecules that promote a function and/or biological effect inside or outside a cell (e.g., IRES, ribosomal recruitment, cytokine stimulation), molecules that promote entry into a cell (e.g., peptides, endosomal escape compounds), molecules that bind to target cells (e.g., aptamers, antibodies, ligands), cytotoxic compounds (e.g., cytotoxic nucleosides), molecules that express a gene product inside a cell (e.g., mRNA), chemotherapeutic compounds (e.g., alkylating agents, antimetabolites, topoisomerase inhibitors), molecules that silence or alter a gene inside a cell (e.g., siRNA, miRNA, antisense therapy, lncRNA), CRISPR molecules (e.g., gRNA, Cas9 protein, Cas9 mRNA), small molecule therapies (e.g., protein-tyrosine kinase inhibitors, proteasome inhibitors), proteins, peptides, and diagnostic agents.
The labile nature of the linkages (see for example Table 4) will allow for these chains to be broken in response to certain stimuli, thereby releasing the payload only when desired. Furthermore, the high reactivity imparted by the reactive handles will allow for comparatively easy assembly of these complex constructs (for example
In certain embodiments, compositions of the invention include the building blocks that are used to create oligonucleotides and their modifications. DNA and RNA relies on a molecular self-assembly process that is driven by supramolecular interactions between four units that are placed in a defined order. Extending the structural diversity of these recognition units, and even replacing them with alternate functionalities, allows for precise control of the oligo structure at both strand and assembly level (M. Vybornyi, Y. Vyborna, R. Haner, DNA-inspired oligomers: From oligophosphates to functional materials, Chem. Soc. Rev. 48 (2019) 4347-4360); (A. Al Ouahabi, L. Charles, J.-F. Lutz, Synthesis of Non-Natural Sequence-Encoded Polymers Using Phosphoramidite Chemistry, J. Am. Chem. Soc. 137 (2015) 5629-5635). The contents of which are incorporated herein by reference. Chemical alternatives to nucleic acids are indispensable in generating materials that are amenable to a high degree of fine tuning with regards to their physicochemical characteristics.
Nanoparticles may contain any monomeric building blocks that are introduced via phosphoramidite chemistry for direct alteration of its physicochemical properties (for example
In addition to monomeric building blocks that affect the physical properties of the nanoparticle, the nanoparticle may contain an oligomeric component that has monomeric building blocks functionalized with reactive handles (for example
The aforementioned modifications are ideal for alteration of the physicochemical properties of the nucleic acid nanoparticle and would be ideal to add as a tail extending from the central core. In some cases, however, it may be beneficial to incorporate the modifications within the central core itself. These modifications can be incorporated pre- or post-synthetically and would usually require a functionalized nucleotide so that the appropriate conjugation chemistries can occur. In our prior invention (SIX-003/00US 34514/9), we outlined the reactive handles needed for several click chemistry strategies at both strand and nucleotide level. These modifications were on the canonical bases adenine (A), cytosine (C), guanine (G), and thymine (T)/uridine (U). From a manufacturing perspective, the synthesis of multiple modifications on multiple bases would be costly and time consuming. Therefore, it would be beneficial to choose a “universal” base onto which these modifications are attached. Universal bases are used to reduce the degeneracy of the four canonical bases. The most widely used are 2′-deoxyinosine and 2′-deoxynebularine. 3-Nitropyrrole 2′-deoxynucleoside and 5′-nitroindole 2′-deoxynucleoside are also used to some extent. Unfortunately, all these bases have a destabilizing effect on DNA/RNA duplexes, so the search for a true universal base is still widely underway. 6H, 8H-3,4-dihydro-pyrimido[4,5-c][1,2] oxazin-7-one (P) and 2-amino-9-(2-deoxy-β-ribofuranosyl)-6-methoxyaminopurine (dK) show considerable promise as degenerate bases and can effectively H-bond to A or G and C or T, respectively ((P. K. T. Lin, D. M. Brown, Synthesis and duplex stability of oligonucleotides containing cytosine-thymine analogues, Nucleic Acids Res. 17 (1989) 10373-10383); (D. M. Brown, P. K. Thoo Lin, Synthesis and duplex stability of oligonucleotides containing adenine-guanine analogues, Carbohydr. Res. 216 (1992) 129-139)).
The composition may include variants of any given universal base. The base will be attached to a chemically or enzymatically modified nucleotide. Universal nucleotides may be modified on the sugar, at the 2′ position, on the phosphate, or on the universal base (for example
mRNA Origami
Nucleic acid (NA) therapies aim to cure genetic or acquired diseases caused by aberrant gene expression. Broadly, the therapeutic approaches developed to date can be classified into three main categories. The first category, gene therapy, involves the introduction of corrective genetic material to restore the expression of a missing or defective gene. The second strategy, RNA-based therapy, involves delivery of nucleic acids that reduce the expression levels of defective or overexpressed messenger RNAs (mRNAs), or alternatively provide functional mRNA molecules to increase the expression levels of underexpressed or missing proteins. The third approach, gene editing, allows scientists to correct mutations in endogenous DNA or mRNA sequences.
Compared with DNA gene therapies, the use of RNA therapeutics is considered markedly safer. Not only is there no risk of stable genome integration, but RNA also possesses a short half-life in vivo and is readily degraded by ribonucleases (RNases), ensuring its activity is non-permanent. Moreover, due to the predominantly cytoplasmic localization of RNA, transport across the nuclear membrane is not required, which facilitates delivery. The FDA approval of six antisense oligonucleotides (ASOs), four small interfering RNA (siRNA) therapeutics and the recent success of two mRNA vaccines against COVID-19 demonstrates the therapeutic potential of RNA drugs and, in particular, has put mRNA drugs into spotlight.
Messenger RNA is a temporary copy of genetic information that is copied from DNA and translated into a protein. Mature mRNA is a single-stranded polynucleotide with an average length of 2,000-2,500 bases (T. Ota, et al., Complete sequencing and characterization of 21,243 full-length human cDNAs, Nat Genet. 36 (2004), 40-45). It is characterized by a 5′ 7-methylguanosine cap (m7G), which protects the mRNA from degradation and promotes translation initiation, 5′ and 3′ untranslated regions (UTRs) that flank the protein-coding open-reading frame (ORF), and a 3′ polyA-tail that regulates mRNA stability. Synthetic mRNA can be produced by in vitro transcription (IVT). To prevent immunostimulation, modified nucleobases are introduced during IVT.
Despite the advantages of nucleic acid drugs and continuous progress in the field, mRNA delivery in vivo remains a major challenge. The intrinsically negative charge of mRNA prevents its translocation across negatively charged cell membranes. Moreover, mRNA needs to be protected from enzymatic degradation by ubiquitously expressed RNases. Thirdly, due to mRNA's large size of approximately 105-106 Da, encapsulation in delivery vehicles is more difficult to achieve than for smaller payloads (K. A. Hajj, K. A. Whitehead, Tools for translation: non-viral materials for therapeutic mRNA delivery. Nat. Rev. Mater. 2 (2017) 17056) (C. Zeng, C. Zhang, P. G. Walker, Y. Dong, Formulation and Delivery Technologies for mRNA Vaccines, in: Current Topics in Microbiology and Immunology, Springer, Berlin, Heidelberg, 2020).
Various delivery strategies have been developed to overcome these bottlenecks. (1) The most clinically advanced systems are lipid nanoformulations such as liposomes and lipid nanoparticles that encapsulate the mRNA in a hydrophilic interior surrounded by a protective outer layer of lipids. Although efficacious and successful in the clinics, these delivery vehicles are often associated with toxicity and immunogenicity in vivo, which can be mitigated by using ionizable over cationic lipids. Furthermore, they often provide only limited control over particle size, may suffer from high batch-to-batch variability, and display low encapsulation efficiencies. (2) Viral lentiviruses, adeno-associated viruses and virus-like replicon particles have also been employed as nucleic acid carriers. Whilst allowing efficient cytoplasmic delivery of mRNAs, their application is limited by unwanted immune responses and issues with large-scale production. (3) Cationic polymer shuttles deliver nucleic acids into the cytosol via electrostatic interactions. However, their use is associated with toxicities related to high molecular weight, highly branched formulations, and aggregation. (4) Other delivery strategies based on transcript-activated matrices, exosomes, peptides and nanoemulsions have been reviewed (K. A. Hajj, K. A. Whitehead, Tools for translation: non-viral materials for therapeutic mRNA delivery, Nat. Rev. Mater. 2 (2017) 17056); (C. Zeng, C. Zhang, P. G. Walker, Y. Dong, Formulation and Delivery Technologies for mRNA Vaccines, in: Current Topics in Microbiology and Immunology, Springer, Berlin, Heidelberg, 2020); (T. C. Roberts, R. Langer, M. J. A. Wood, Advances in oligonucleotide drug delivery, Nat Rev Drug Discov 19 (2020), 673-694); (S. Uchida, F. Perche, C. Pichon, and H. Cabral, Nanomedicine-Based Approaches for mRNA Delivery, Molecular Pharmaceutics 17 (2020), 3654-3684).
Overall, there remains an unmet need in the art for improved nucleic acid compositions suitable for general clinical use. Current research is largely devoted to fine tuning the composition of delivery vehicles and enhancing the tolerability thereof. The present invention, in contrast, describes the use of NA nanotechnology to precisely tweak the characteristics of the NA drug molecule itself. By exploiting the programmability of NA base-pairing, compact structures of controlled shape, size and complexity can be formed. Compaction occurs through a process of molecular self-folding termed origami, in which a single-stranded DNA or RNA molecule (scaffold, e.g., an mRNA) hybridizes to one or more DNA or RNA molecules, for example hundreds of short complementary 20-60-mer staple strands. (P. Rothemund, Folding DNA to create nano-scale shapes and patterns, Nature. 440 (2006) 297-302); (S. Douglas, H. Dietz, T. Liedl, et al. Self-assembly of DNA into nanoscale three-dimensional shapes. Nature 459 (2009) 414-418); (N. Seeman, “Nanomaterials based on NA”, An. Ref Biochem. 79 (2010) 65-87); (X. Qi, X. Liu, L. Matiski, R. R. Del Villar, T. Yip, F. Zhang, S. Sokalingam, S. Jiang, L. Liu, H. Yan, and Y. Chang, ACS Nano 14 (2020), 4727-4740).
The present invention provides compositions and methods that can be used to reversibly compact nucleic acids into defined origami shapes and sizes with tunable pharmacokinetic and pharmacodynamic (PK/PD) properties. The compactness of the origami structure may protect the therapeutic moiety against chemical, biochemical or mechanical stresses and increase its resistance against nucleases. Size, rigidity, and shape of the construct can be varied to modulate packaging and achieve a favorable PK/PD profile. In addition, each of the origami building blocks can be selectively modified to tune the PK/PD properties of the origami structure, such as serum stability, biodistribution and cellular uptake. Upon internalization by living cells, the construct may dissociate and release the decompacted therapeutic molecule(s) (
Other objects, features, and advantages of the present invention will be apparent to one of skill in the art from the following detailed description and figures.
The nucleic acid nanoparticles used in this invention are interchangeably referred to as Mergo.
RNA Strands Covered in this Invention
Oligonucleotides were synthesized on 1-10 μmol scale using a K&A synthesizer (H-16). All protocols were modified depending on the sequence requirements. Phosphoramidites and CPGs with standard protecting groups were purchased from ChemGenes and Glen Research. Adenosine phosphoramidites containing amino acids, amino acid analogues, PEGs and hydrocarbon chains were synthesized in-house. The detritylation step was carried out with 3% TCA in DCM, followed by coupling with 0.1M phosphoramidite solutions and 0.25M BMT in MeCN. Capping was performed using THF/lutidine/acetic anhydride (80/10/10) as capping A and 16% N-methylimidazole in THE as capping B, respectively. The oxidation step was accomplished with 0.02 M iodine solution in THF/Pyr/water (90.6/0.4/9).
All synthesized oligonucleotides were cleaved and deprotected using aq. methylamine/ammonium hydroxide solution (1:1) for 3 h at RT for a solid support with a first base attached or for 1 h at 65° C. for a universal CPG. The removal of tert-butyl silyl protecting groups was performed by incubating an intermediate product in DMSO Et3N·3HF for 3 h at 65° C. Crude oligonucleotides were subsequently precipitated from ethanolic solution containing sodium acetate. After 2 h at −70° C. the precipitate was harvested by 25 min centrifugation at 4° C. (14,000 rpm). The supernatant was separated, and the remaining pellet was washed repeatedly with 70% EtOH. After a final wash, the crude sample was dried under vacuum in a speedvac and redissolved in water for purification.
Crude RNA strands were purified either by IEX-HPLC or by IP-RP HPLC.
IEX was carried out with a preparative DNAPac PA200 (ThermoFisher), 22×250 mm column, or PL-SAX (Agilent) 22×150 mm 1000 Å column at 75° C. with a flow rate of 15 mL/min and UV detection at 260 nm. Elution was performed with a linear gradient selected based on crude impurity profile, determined by analytical testing using either a DNAPac PA200RS UPLC column or PL-SAX analytical column. Buffer A: 25 mM Tris HCl, pH 8.0, 20% acetonitrile, 10 mM sodium perchlorate; buffer B: 25 mM Tris HCl, pH 8.0, 20% acetonitrile, 600 mM sodium perchlorate, OR, Buffer A: 25 mM Tris HCl, pH 8.0, 20% acetonitrile, 25 mM sodium chloride; buffer B: 25 mM Tris HCl, pH 8.0, 20% acetonitrile, 1.5M sodium chloride.
RP-HPLC was carried out with a BEH C18 300 Å (Waters) 19×150 mm at 60° C., with a flow rate of 25 mL/min and UV detection at 260 nm. Buffer A: TEAA (0.1 M, pH=7); buffer B: MeCN, OR, Buffer A: HAA (0.1M, pH7); buffer B: MeCN.
Fractions containing RNA were assessed for purity by analytical PAGE, IEX and RP-HPLC, then pooled and subject to final QC on PAGE, IEX and RP-HPLC, acetonitrile removed in vacuo. The purified oligos were then desalted with Gel-Pak desalting columns (Glen). The solution was lyophilized, and the RNA dissolved in nuclease-free water for concentration determination by UV absorbance and quality assessment via denaturing PAGE.
5′ amino—10% DEA solution in MeCN was applied onto the oligonucleotide while still on CPG. After 5 min treatment the column was rinsed with MeCN and processed further.
5′ Cy3—MMTr group at 5′-end of Cy3 containing sequences was removed during RPC MMT-ON purification.
5′ cholesterol modification—10% DEA solution in MeCN was applied onto the oligonucleotide while still on CPG. After 5 min treatment the column was rinsed with MeCN, and the protecting group (DMT) was removed while still on solid support prior to cleavage and deprotection steps.
The key scaffold in this work was assembled according to a standard protocol. Equimolar amounts of the 5 different strands, C-1.0, C-2.0, C-3.0, C-4.0 and C-5.0 (and sub-variants in Table 6) were combined in PBS+MgCl2 (2 mM) buffer, with a final concentration of 10 μM. The 5 strands were annealed to each other at 95° C. for 5 min then slowly cooled down to 15° C. The scaffold was then analyzed by native polyacrylamide gel electrophoresis (PAGE) and dynamic light scattering (DLS) (vide infra).
For PAGE, the assembled scaffold was electrophoresed on native PAGE (6%) in 1×TBMg (890 mM Tris Borate+20 mM Mg(OAc)2, pH=8.3) at a constant voltage of 100 V. Gel bands were visualized using GelRed™. 10 pmol of structures was loaded. 2 μL of glycerin (70% in H2O) was added to samples before loading.
For DLS, the assembled scaffold was analyzed using a Malvern Zetasizer Nano S ZEN 1600 Nano Particle Size Analysis—20 μL of samples were used, and intensity was recorded. Average of three trials was calculated. All measurements were carried out at 25° C. Samples were centrifuged at 12000 rpm for 5 minutes before analysis to remove dust and debris.
The key scaffold used in this invention has been further refined to reduce the overall RNA content by 19%, which allows for more cost-effective manufacturing. The optimized characteristics are given in
Solid phase oligonucleotide synthesis of core strand conjugated to siRNA using C-6 disulfide modifier.
Synthesis: The sequence containing disulfide linkage was synthesized using the following reagents: 3% TCA in DCM, 0.25M Hyacinth BMT solution, CAP A (THF/lutidine/acetic anhydride), CAP B (16% N-methylimidazole), 0.02M Iodine/Py/water.
Phosphoramidites: 2′-tBDSilyl Adenosine (n-bz) CED phosphoramidite, 2′-tBDSilyl Cytidine (n-acetyl) CED phosphoramidite, 2′-tBDSilyl Guanosine (n-ibu) CED phosphoramidite, 2′-tBDSilyl Uridine CED phosphoramidite, 2′-Fluoro-2′-deoxyCytidine (n-ac) CED phosphoramidite, 2′-Fluoro-2′-deoxy Uridine CED phosphoramidite, Thymidine CED phosphoramidite
5′-thiol modifier C6: formula (VIII):
Deprotection: AMA, rt, 3 h. TEA×3HF, 65° C.
Quality control of a raw material is provided in
Purification method: The sequence was purified using IEX chromatography, using DNAPac_PA100 22×250 mm at 75° C.
Isolation yield: 25%.
To install the appropriate reactive groups to enable conjugation chemistry, 5′ amino modified RNA strands were treated with heterobifunctional NHS-linkers containing the same.
The amino-modified oligonucleotide was prepared as a stock solution or dry aliquot. The heterobifunctional NHS-ester (NHS-SM) was dissolved at a concentration of 100 mM in anhydrous DMSO.
Amino-modified oligonucleotide was diluted to a final concentration of 100-200 μM, followed by the addition of DMSO (50% total volume), bicarbonate buffer (0.5 M, pH=8.4, 20% total volume) and NHS-SM (5-20 eq). The reaction mixture was agitated at 30° C. for 1-3 h and was then purified by RP-HPLC. With higher volumes, EtOH precipitation and resuspension in H2O is recommended.
The amino-modified oligonucleotide was prepared as a stock solution or dry aliquot. The heterobifunctional NHS-ester (NHS-SM) was dissolved at a concentration of 100 mM in anhydrous DMF.
Amino-modified oligonucleotide was diluted to a final concentration of 200-500 μM, followed by the addition of DMF (35% total volume), sodium chloride/bicarbonate buffer (100 mM NaCl, 0.05 M, pH=8.4, 30% total volume) and NHS-Tetrazine (5-20 eq). The reaction mixture was agitated at 30° C. for 1 h and was then purified by EtOH precipitation and resuspension in H2O.
Norbornene modified core strand C-4.4 (5 nmol, 1.0 eq, 1400 μM final concentration) was mixed with siRNA functionalized via tetrazine-NHS (5-1.5, 15 nmol, 1.6 eq) in PBS buffer. The reaction mixture was agitated at RT for 12 h, followed by purification with IEX chromatography, using DNAPac_PA100 22×250 mm column at 75° C., at a flow rate of 25 mL/min. 40% to 60% B in 30 min (A: 0.1 M NaCl pH 7, B: 1.0M NaCl), fractions containing product were concentrated and desalted, resulting in 44% isolated yield.
1H NMR spectra were recorded at 400 MHz. 13C NMR spectra were recorded at 100 MHz. Chemical shifts (δ) are quoted in units of parts per million (ppm) downfield from tetramethylsilane and are referenced to a residual solvent peak. (CDCl3 (δH: 7.26, δC: 77.0)). Coupling constants (J) are quoted in units of Hertz (Hz). The following abbreviations are used within 1H NMR analysis: s=singlet, d=doublet, t=triplet, q=quartet, pent=pentet, m=multiplet, dd=doublet of doublets, dt=doublet of triplets. Spectra recorded at 400 (1H NMR) and 100 (13C NMR) were carried out by the Imperial College London Department of Chemistry NMR Service.
Low- and high-resolution mass spectrometry (EI, CI, FAB) were recorded at Imperial College London. Measurements carried out by the Imperial College Department of Chemistry Mass Spectrometry Service used a Micromass Platform II and Micromass AutoSpec-Q spectrometer.
Flash column chromatography was carried out on BDH silica gel 60, particle size 0.040-0.063 mm. Thin layer chromatography (TLC) was performed on pre-coated aluminum backed or glass backed plates (Merck Kieselgel 60 F254), and visualized with ultraviolet light (254 nm) or potassium permanganate (KMnO4), vanillin or phosphomolybdic acid (PMA) stains.
Synthesized according to a procedure outlined by Varenikov and co-workers (A. Varenikov, M. Gandelman, Organotitanium Nucleophiles in Asymmetric Cross-Coupling Reaction: Stereoconvergent Synthesis of Chiral α-CF 3 Thioethers, J. Am. Chem. Soc. 141 (2019) 10994-10999). Colorless oil obtained (18.2 g, 92%). 1H NMR (400 MHz, Chloroform-d) δ 3.65 (t, J=6.6 Hz, 4H), 2.70 (dd, J=7.9, 6.8 Hz, 4H), 1.81-1.77 (m, 2H), 1.76-1.65 (m, 4H), 1.59 (dq, J=7.9, 6.6 Hz, 4H), 1.51-1.32 (m, 8H).
Synthesized according to a modified procedure found in the art (US2011/263526). A solution of DCC (1.15 g in 5 mL anhydrous DCM, 5.61 mmol) was added dropwise to a stirred solution of 5-norbornene-2-carboxylic acid (500 mg, 3.62 mmol), 6,6′-disulfanediylbis(hexan-1-ol) (1.93 g, 7.25 mmol) and DMAP (89 mg, 0.72 mmol) in anhydrous DCM (20 mL) over 5 min at 0° C. The reaction mixture was then stirred at 0° C. for 3 h. Upon completion (TLC: 25% EtOAc/pentane), the reaction mixture was filtered. The filtrate was then washed with water (3×20 mL) and brine (3×20 mL). The organic layer was then dried (MgSO4) and concentrated in vacuo. The crude residue was then purified by column chromatography (20 to 30% EtOAc/pentane), affording the title compound as a colorless oil (469 mg, 34%). 1H NMR (400 MHz, Chloroform-d) δ 6.22 (dd, J=5.7, 3.1 Hz, 1H), 5.94 (dd, J=5.7, 2.9 Hz, 1H), 4.04 (td, J=6.6, 4.2 Hz, 2H), 3.23 (dq, J=3.4, 1.8 Hz, 1H), 2.95 (ddd, J=12.6, 4.7, 3.0 Hz, 2H), 2.71 (td, J=7.3, 2.2 Hz, 5H), 1.93 (ddd, J=12.6, 9.3, 3.7 Hz, 1H), 1.72 (dt, J=7.2, 4.0 Hz, 4H), 1.51-1.37 (m, 14H), 1.36-1.26 (m, 1H); HRMS ES+ (m/z): [M]+ calc'd for C20H34O3: 386.6090; found: 386.6097.
6-((6-Hydroxyhexyl)disulfaneyl)hexyl bicyclo[2.2.1]hept-5-ene-2-carboxylate (496 mg, 0.87 mmol) and N,N-diisopropylethylamine (451 mg, 609 μL, 3.49 mmol) were dissolved in anhydrous DCM (15 mL) and stirred over activated molecular sieves for 1 h at 0° C. 2-Cyanoethyl N,N-diisopropylchlorophosphoramidite (413 mg, 1.74 mmol) was added and the reaction mixture was stirred for 30 min at 0° C., and was then slowly warmed to RT over 1.5 h. Upon completion (TLC: 25% EtOAc/pentane), the reaction mixture was washed with sat. NaHCO3 (3×20 mL). The organic layer was then dried (MgSO4) and concentrated in vacuo, and the crude product was purified by column chromatography (10% EtOAc/pentane+1% Et3N), affording the title compound was a colorless oil (341 mg, 67%). 1H NMR (400 MHz, Chloroform-d) δ 6.21 (dd, J=5.7, 3.1 Hz, 1H), 5.94 (dd, J=5.7, 2.9 Hz, 1H), 4.04 (td, J=6.6, 4.0 Hz, 2H), 3.92-3.78 (m, 2H), 3.75-3.54 (m, 4H), 3.23 (dd, J=4.1, 2.3 Hz, 1H), 3.03-2.88 (m, 2H), 2.73-2.67 (m, 6H), 2.07 (s, 1H), 1.92 (ddd, J=11.8, 9.3, 3.7 Hz, 1H), 1.70 (d, J=7.2 Hz, 3H), 1.66-1.61 (m, 4H), 1.45-1.39 (m, 8H), 1.30 (t, J=4.4 Hz, 1H), 1.21 (dd, J=6.8, 4.1 Hz, 14H); 31P NMR (162 MHz, Chloroform-d) δ 147.26; HRMS ES+ (m/z): [M]+ calc'd for C29H51O4PS2: 586.3028; found: 586.8304.
This compound may be incorporated into any designs shown in
N-(9-((2R,3R,4R,5R)-5-((bis(4-methoxyphenyl)(phenyl)methoxy)methyl)-4-hydroxy-3-(prop-2-yn-1-yloxy)tetrahydrofuran-2-yl)-9H-purin-6-yl)benzamide (2′O propargyl A) (1 equiv.) and the azide (R—N3) (1.2 equiv.) were dissolved in THE (0.1 M reaction concentration). Copper(II) sulfate (0.085 equiv.) and sodium ascorbate (0.1 equiv.) were added under N2 and the mixture was stirred for 16 h. Upon completion, a 5% solution of EDTA was added and the reaction mixture was extracted with ethyl acetate. The organic layer was dried (MgSO4) and concentrated in vacuo. The crude residue was then purified by column chromatography to afford the title compound.
PK/PD modulating nucleosides, whereby R imparts the biological activity and affects the biodistribution, are given in
Following a procedure outlined in the art (B. Ross, Q. Song, Process of Purifying Phosphoramidites, U.S. Pat. No. 7,030,230 B2, 2006.), the nucleoside (1 equiv.) was dissolved in anhydrous DMF. Activated molecular sieves were added and the suspension was stirred for 30 min, followed by the addition of 3-((bis(diisopropylamino)phosphaneyl)oxy)propanenitrile (1.5 equiv.) and 5-(ethylthio)-1H-tetrazole (0.82 equiv.). When dissolved, 1-methyl-1H-imidazole (8.3 equiv.) was added, and the reaction mixture was stirred at RT for 5 h.
The following work-up was specifically for 150 mL of DMF as the reaction solvent. Care should be taken to adjust the volumes as appropriate.
When complete, triethylamine (15 mL) was added, and the reaction mixture was diluted with DMF (50 mL). Water (25 mL) was added, and the mixture was extracted with hexane (3×150 mL). The aqueous layer was separated and was diluted with water (75 mL) and was then extracted with toluene (3×225 mL). The upper organic layer was then separated and was washed with DMF:water (60:40 v/v, 3×225 mL) and water (3×150 mL). The upper layer was dried (MgSO4), filtered and concentrated in vacuo, affording the phosphoramidite, which was often pure enough for direct use.
In a mixture of H2O (48 mL) and DCM (81 mL), NaN3 (19.9 g, 300 mmol) was dissolved. This solution was cooled on ice. Triflic anhydride (16.9 g, 10.1 mL 60 mmol) was slowly added. This reaction mixture was stirred vigorously for 2 h at room temperature. The water-layer was extracted with DCM (2×50 mL). These combined DCM-layers were washed with saturated aq. Na2CO3-solution. Histamine (5.52 g, 184 mmol), K2CO3 (16.5 g, 120 mmol) and CuSO4·5H2O (47.8 mg, 0.3 mmol) were dissolved in H2O (96 mL) and MeOH (192 mL). If the amine salt is used instead of the free amine, 1 additional equivalent of K2CO3 was added per acidic proton. The freshly made triflic azide in DCM was added. The reaction was stirred overnight at room temperature. The mixture was extracted with DCM (3×200 mL) and water (50 mL). The DCM-layers were combined, dried over Na2SO4, filtered, and concentrated in vacuo to give crude azide. A slightly yellow oil was obtained (4.12 g, 87%). The product was pure enough for further synthetic manipulations. An analytical sample was obtained by column chromatography (5% MeOH/DCM) in the dark. This afforded the product as a colorless oil. 1H NMR (400 MHz, Chloroform-d) δ 8.39 (br s, 1H), 7.66 (br s, 1H), 6.95 (br s, 1H), 3.58 (d, J=6.7 Hz, 2H), 2.90 (t, J=6.7 Hz, 2H). LRMS (ESI+) m/z (%): 138.1 [M]+ (100), 139.1 [M+H]+ (10).
Alternatively, following a protocol described in the art (E. D. Goddard-Borger, R. V. Stick, An Efficient, Inexpensive, and Shelf-Stable Diazotransfer Reagent: Imidazole-1-sulfonyl Azide Hydrochloride, Org. Lett. 13 (2011) 2514-2514), imidazole-1-sulfonyl azide hydrochloride (377 mg, 1.8 mmol) was added to a stirred suspension of histamine (170 mg, 1.5 mmol), K2CO3 (414 mg, 3 mmol) and CuSO4·5H2O (3.75 mg, 15 μmol) in MeOH (7.5 mL). Upon completion of the reaction, water (5 mL) was added, and the reaction mixture was extracted with DCM (3×5 mL). The organic layer was then dried (MgSO4) and concentrated, affording the crude title compound as a brown oil (360 mg, 95%), which was used without further purification. 1H NMR (400 MHz, Chloroform-d) δ 7.64 (s, 1H), 6.92 (s, 1H), 3.61 (t, J=6.7 Hz, 2H), 2.97-2.88 (m, 2H); LRMS ES+ (m/z): [M]+ calc'd for C5H7N5; 137.1 found: 138.1 [M+H]+.
NaH (60% in mineral oil) (41 mg, 1 mmol) was added to THE (4 mL). A solution of 4-(2-azidoethyl)-1H-imidazole (93 mg, 0.68 mmol) in 2 mL THE was added and the resultant suspension was stirred for 3 h at RT. A solution of chloromethyl pivalate (150 mg, 1 mmol) in THE (2 mL) was then added. After 1 h at RT, H2O (0.5 mL) was added, and the reaction mixture was concentrated in vacuo. The crude residue was redissolved in EtOAc, and the organic layer was washed with water (3×10 mL), brine (3×10 mL), dried (MgSO4) and concentrated in vacuo. The crude residue was purified by column chromatography (50% EtOAc in pentane) to afford the title compound as a colorless oil (150 mg, 88%). 1H NMR (400 MHz, Chloroform-d) δ 7.63 (d, J=1.4 Hz, 1H), 6.93 (s, 1H), 5.80 (s, 2H), 3.57 (t, J=6.8 Hz, 2H), 2.85 (td, J=6.8, 0.8 Hz, 2H), 1.18 (s, 9H); 13C NMR (101 MHz, Chloroform-d) δ 177.7, 139.6, 138.0, 116.8, 67.6, 50.6, 38.7, 28.1, 26.8. LRMS ES+ (m/z): [M]+ calc'd for C11H17N5O2; 251.3 found: 252.3 [M+H]+
Following general procedure A, the title compound was afforded as a colorless foam (4.53 g, 94%) after column chromatography (5% MeOH/DCM). 1H NMR (400 MHz, Chloroform-d) δ 8.71 (s, 1H), 8.19 (s, 1H), 8.07-7.98 (m, 2H), 7.62-7.47 (m, 4H), 7.43-7.38 (m, 2H), 7.33-7.16 (m, 9H), 6.84-6.77 (m, 4H), 6.21 (d, J=5.0 Hz, 1H), 5.67 (d, J=1.9 Hz, 2H), 4.88-4.79 (m, 2H), 4.73 (d, J=12.9 Hz, 1H), 4.59 (d, J=5.5 Hz, 2H), 4.45 (s, 1H), 4.27 (d, J=3.9 Hz, 1H), 4.03 (s, 1H), 3.76 (s, 6H), 3.48 (dd, J=10.6, 3.3 Hz, 1H), 3.38 (dd, J=10.6, 4.3 Hz, 1H), 3.07 (s, 2H), 2.44 (s, 1H), 1.12 (s, 9H); 13C NMR (101 MHz, Chloroform-d) δ 177.6, 164.7, 158.5, 152.7, 151.6, 149.6, 144.5, 143.6, 141.9, 138.3, 135.7, 135.6, 133.7, 132.8, 130.1, 130.1, 128.8, 128.2, 127.9, 126.9, 123.5, 122.9, 117.1, 113.2, 87.0, 86.6, 84.5, 81.9, 69.8, 67.6, 64.3, 63.2, 55.3, 55.2, 53.5, 49.8, 38.7, 29.0, 26.8.
Following general procedure B, the title compound was afforded as a colorless foam (77%). Further purification was not required. 1H NMR (400 MHz, CDCl3) δ 9.23 (d, J=5.9 Hz, 1H), 8.69 (d, J=6.1 Hz, 1H), 8.13 (d, J=8.5 Hz, 1H), 8.08-7.92 (m, 2H), 7.63-7.57 (m, 1H), 7.55-7.47 (m, 3H), 7.44-7.36 (m, 2H), 7.33-7.14 (m, 7H), 6.86-6.77 (m, 4H), 6.75 (dd, J=4.2, 1.2 Hz, 1H), 6.17 (dd, J=6.7, 5.8 Hz, 1H), 5.68 (d, J=3.1 Hz, 2H), 4.91 (dt, J=11.2, 5.4 Hz, 1H), 4.82 (dd, J=12.6, 7.3 Hz, 1H), 4.69 (ddd, J=20.8, 11.8, 3.7 Hz, 2H), 4.54 (td, J=7.3, 2.1 Hz, 2H), 4.39 (dd, J=24.6, 3.7 Hz, 1H), 3.97-3.81 (m, 1H), 3.77 (dd, J=3.1, 0.7 Hz, 6H), 3.73-3.47 (m, 2H), 3.33 (ddd, J=10.6, 4.2, 1.8 Hz, 1H), 3.10-3.02 (m, 2H), 2.94 (s, 1H), 2.87 (d, J=0.6 Hz, 1H), 2.66-2.60 (m, 1H), 2.37 (dd, J=12.6, 6.2 Hz, 1H), 1.26 (dd, J=6.8, 5.6 Hz, 1H), 1.17 (d, J=6.8 Hz, 8H), 1.12 (d, J=1.5 Hz, 9H), 1.04 (d, J=6.8 Hz, 4H); 13C NMR (101 MHz, CDCl3) δ 177.6, 164.7, 158.5, 152.7, 151.7, 149.5, 144.4, 144.1, 143.9, 142.1, 138.4, 138.2, 135.7, 135.6, 135.5, 133.8, 132.7, 130.1, 130.1, 128.8, 128.3, 128.2, 127.9, 126.9, 123.6, 122.8, 122.6, 117.9, 117.4, 116.9, 113.2, 87.0, 86.9, 86.7, 86.6, 84.4, 84.2, 80.1, 79.6, 71.2, 70.7, 70.6, 67.6, 64.2, 63.1, 62.8, 59.0, 58.9, 58.1, 57.9, 55.2, 49.6, 43.4, 43.3, 43.1, 38.7, 36.5, 29.1, 29.1, 26.8, 24.8, 24.7, 24.6, 20.4, 20.4, 20.2; 31P NMR (162 MHz, CDCl3) δ 150.7, 150.4.
Following a procedure outlined in the art (H. Schotte, Verfahren zur Darstellung von Diacylisothioharnstoff-S-alkylaethern, DE1925C036959D 19250717, 1928.), (ethylsulfanyl)methanimidamide hydrobromide (21.3 g, 115 mmol) was dissolved in dry pyridine (110 mL) and benzoyl chloride (32.3 g, 230 mmol) was added at 0° C. Upon completion, the reaction mixture was poured into water. The formed crystals were filtered off and washed with EtOH and Et2O, affording the title compound (29.7 g, 82%). 1H NMR (400 MHz, DMSO) δ 13.58 (s, 1H), 8.25-8.11 (m, 2H), 7.94-7.84 (m, 2H), 7.75-7.48 (m, 6H), 3.37 (s, 1H), 3.19 (q, J=7.3 Hz, 2H), 1.37 (t, J=7.3 Hz, 3H); 13C NMR (101 MHz, DMSO) δ 175.1, 167.9, 164.5, 135.6, 133.4, 132.8, 131.5, 129.5, 128.9, 128.3, 127.6, 25.1, 13.4. LRMS ES+ (m/z): [M]+ calc'd for C17H16N2O2S; 312.4 found: 313.1 [M+H]+
To a stirred solution of ethyl (E)-N,N-dibenzoylcarbamimidothioate (8.43 g, 27 mmol) in MeCN (50 mL), 4-azidobutan-1-amine (3.49 g, 30 mmol) was added. The reaction mixture was stirred at ambient temperature for 2 h and concentrated in vacuo. The crude mixture was purified by column chromatography DCM/Pentane (20-100%). Affording the title compound (9.01 g, 82%) 1H NMR (400 MHz, DMSO) δ 14.38 (s, 1H), 9.54 (t, J=5.9 Hz, 1H), 8.26-8.18 (m, 2H), 8.02-7.94 (m, 2H), 7.80-7.72 (m, 1H), 7.72-7.63 (m, 2H), 7.62-7.53 (m, 1H), 7.49 (ddt, J=8.2, 6.7, 1.2 Hz, 2H), 3.67 (q, J=6.6 Hz, 2H), 3.42 (t, J=6.7 Hz, 2H), 1.82-1.71 (m, 2H), 1.66 (dddd, J=13.7, 8.2, 6.6, 1.7 Hz, 2H); 13C NMR (101 MHz, DMSO) δ 177.6, 167.5, 156.8, 137.9, 134.2, 132.5, 132.4, 129.9, 129.6, 128.6, 127.9, 50.8, 40.8, 26.4, 26.2. LRMS ES+ (m/z): [M]+ calc'd for C19H20N6O2; 364.4 found: 365.1 [M+H]+.
Following general procedure A, the title compound was afforded as a colorless foam (40%) after purification by column chromatography (1 to 10% MeOH/DCM). 1H NMR (400 MHz, DMSO) δ 14.35 (s, 1H), 11.24 (s, 1H), 9.51 (t, J=5.9 Hz, 1H), 8.66 (s, 1H), 8.54 (s, 1H), 8.21-8.14 (m, 2H), 8.07-8.02 (m, 2H), 8.01 (s, 1H), 7.98-7.94 (m, 2H), 7.76-7.70 (m, 1H), 7.69-7.61 (m, 3H), 7.57-7.50 (m, 3H), 7.48 (m, 2H), 7.38-7.33 (m, 2H), 7.28-7.16 (m, 7H), 6.87-6.78 (m, 4H), 6.19 (d, J=4.8 Hz, 1H), 5.76 (s, 1H), 5.42 (d, J=6.1 Hz, 1H), 4.83 (t, J=5.0 Hz, 1H), 4.76 (d, J=12.3 Hz, 1H), 4.67 (d, J=12.3 Hz, 1H), 4.54 (q, J=5.3 Hz, 1H), 4.36 (t, J=7.0 Hz, 2H), 4.14 (q, J=4.6 Hz, 1H), 3.71 (d, J=1.1 Hz, 6H), 3.64 (q, J=6.6 Hz, 2H), 3.36 (s, 8H), 3.25 (d, J=4.8 Hz, 2H), 1.88 (p, J=7.1 Hz, 2H), 1.69-1.58 (m, 2H); 13C NMR (101 MHz, DMSO) δ 176.8, 166.6, 165.2, 157.6, 155.9, 151.5, 151.3, 150.1, 144.4, 143.2, 142.7, 137.0, 135.2, 135.0, 133.3, 132.9, 132.0, 131.6, 131.5, 129.3, 129.0, 128.7, 128.1, 128.0, 127.8, 127.4, 127.3, 127.1, 126.2, 125.4, 123.5, 112.7, 85.9, 83.4, 79.2, 68.7, 63.1, 62.8, 54.6, 54.5, 48.6, 26.7, 25.3. LRMS ES+ (m/z): [M]+ calc'd for C60H57N11O9; 1076.2 found: 774.2 [M+2H-DMTr]+.
Following general procedure B, the title compound was afforded as a colorless foam (90%) following purification by column chromatography (60% EtOAc/pentane). 1H NMR (400 MHz, CDCl3) δ 14.50-14.46 (m, 1H), 9.51 (s, 1H), 9.03 (s, 1H), 8.70 (d, J=5.6 Hz, 1H), 8.27-8.20 (m, 2H), 8.15 (d, J=8.3 Hz, 1H), 8.06-7.96 (m, 4H), 7.67-7.58 (m, 1H), 7.62-7.37 (m, 11H), 7.37-7.22 (m, 6H), 7.26-7.15 (m, 1H), 6.85-6.75 (m, 4H), 6.19 (t, J=5.4 Hz, 1H), 4.95 (dt, J=13.3, 5.3 Hz, 1H), 4.86 (dd, J=12.5, 6.8 Hz, 1H), 4.79-4.62 (m, 2H), 4.47-4.35 (m, 1H), 4.38-4.28 (m, 2H), 3.97-3.79 (m, 1H), 3.77 (dd, J=2.5, 0.6 Hz, 6H), 3.73-3.63 (m, 2H), 3.66-3.48 (m, 3H), 3.35 (ddd, J=10.6, 4.3, 1.8 Hz, 1H), 2.62 (t, J=6.3 Hz, 1H), 2.38 (t, J=6.4 Hz, 1H), 2.04 (s, 3H), 2.04-1.94 (m, 2H), 1.92 (s, 1H), 1.71 (q, J=7.4, 6.9 Hz, 2H), 1.20-1.12 (m, 9H), 1.05 (d, J=6.8 Hz, 3H); 13C NMR (101 MHz, CDCl3) δ 178.6, 168.4, 164.6, 158.6, 157.2, 152.6, 151.8, 149.5, 144.4, 142.1, 142.0, 137.6, 135.7, 133.6, 132.8, 132.1, 131.9, 130.1, 129.4, 129.2, 128.9, 128.3, 128.2, 128.1, 127.9, 127.9, 127.8, 127.0, 122.7, 122.4, 118.0, 113.2, 87.1, 86.6, 84.3, 80.1, 63.0, 62.8, 60.4, 59.0, 58.8, 55.3, 55.2, 49.8, 43.4, 43.3, 43.1, 40.4, 27.6, 26.3, 24.8, 24.7, 24.6, 21.1, 20.4, 20.2, 14.2; 31P NMR (162 MHz, CDCl3) δ 150.64, 150.34.
Na-(((9H-fluoren-9-yl)methoxy)carbonyl)-Nt-trityl-L-histidine (185 mg, 299 μmol) was activated with HATU (125 mg, 329 μmol), 1H-benzo[d][1,2,3]triazol-1-ol hydrate (50.4 mg, 329 μmol) and DIPEA (116 mg, 898 μmol) in DMF (2 mL) for 20 min. A solution of 17-azido-3,6,9,12,15-pentaoxaheptadecan-1-amine (110 mg, 359 μmol) was then added in DMF (1 mL). The reaction mixture was stirred for 3 h. When complete (TLC: 10% MeOH/DCM), the reaction mixture was extracted with Et2O (3×10 mL) and DCM (10 mL). The combined organic mixtures were then concentrated and the crude residue was purified by flash chromatography with a gradient of 50% EtOAc/pentane to 10% MeOH/DCM, affording the title compound as a colorless oil (203 mg, 75%). LRMS ES+ (m/z): [M]+ calc'd for C52H57N7O8: 908.1; found: 930.5 ([M+Na]+).
(9H-fluoren-9-yl)methyl(S)-(1-azido-19-oxo-21-(1-trityl-1H-imidazol-4-yl)-3,6,9,12,15-pentaoxa-18-azahenicosan-20-yl)carbamate (203 mg, 262 μmol) was dissolved in a 1:1 mixture of DCM/diethylamine (2 mL) and was stirred at RT for 90 min. Upon completion (TLC: 10% MeOH/DCM), the reaction mixture was concentrated in vacuo and was resuspended in DCM, followed by further concentration (×2). The crude residue was then purified by column chromatography (5 to 10% MeOH/DCM) to afford the title compound as a colorless foam (101 mg, 66%). LRMS ES+ (m/z): [M]+ calc'd for C37H47N7O6: 685.8; found: 686.4.
(S)-2-amino-N-(17-azido-3,6,9,12,15-pentaoxaheptadecyl)-3-(1-trityl-1H-imidazol-4-yl)propanamide (101 mg, 147 μmol) was dissolved in a 3:1 mixture of 4 M HCl in dioxane/MeOH and the reaction mixture was stirred at 60° C. for 2 h. Upon full conversion (TLC: 10% MeOH/DCM), the solvent was removed under a stream of nitrogen and the crude residue was triturated in Et2O, affording the title compound as a colorless foam (69 mg, 98%). 1H NMR (400 MHz, CDCl3) δ 13.89 (s, 1H), 13.49 (s, 1H), 8.96 (s, 1H), 8.61 (s, 1H), 8.33 (s, 2H), 7.65 (s, 1H), 4.70 (s, 1H), 3.81-3.63 (m, 18H), 3.43 (s, 2H), 3.35-3.08 (m, 4H); LRMS ES+ (m/z): [M]+ calc'd for C18H33N7O6: 443.7; found: 444.7.
To a stirred solution of pyrazole-1-carboxamidine hydrochloride (374 mg, 2.55 mmol) and DIPEA (356 mg, 479 μL, 2.75 mmol) in DCM (2 mL), 17-azido-3,6,9,12,15-pentaoxaheptadecan-1-amine (766 mg, 2.50 mmol) was added. The reaction mixture was stirred at RT for 26 h and, upon completion, was concentrated in vacuo. The crude residue was then purified by column chromatography (5 to 10% 7N NH3 in MeOH/DCM to 30% to overcome issues with dragging on the column). The title compound was isolated as a brown oil. 1H NMR (400 MHz, DMSO) δ 7.85-6.92 (m, 2H), 3.64-3.46 (m, 20H), 3.40 (dd, J=5.6, 4.3 Hz, 2H), 3.29 (q, J=5.3 Hz, 2H); 13C NMR (101 MHz, DMSO) δ 157.8, 70.3, 70.2, 70.2, 70.2, 69.7, 69.1, 50.5, 41.3. LRMS ES+ (m/z): [M]+ calc'd for C13H28N6O5: 349.4; found: 350.4.
Click reactions were performed in 1:1 mixtures 2 M TEAA:DMSO using standard protocols with Cu2SO4 and TTIPTA (E. Paredes, S. R. Das, Click chemistry for rapid labeling and ligation of RNA, ChemflioChem. 12 (2011) 125-131).
The following solutions were prepared: Deprotection solution: 20% piperidine in DMF; Activator solution: 0.25 M HATU in DMF; Basic solution: 2,6-lutidine (2.05 mL)+DIPEA (1.96 mL) in DMF (5.54 mL) Capping solution: Ac2O (0.92 mL)+2,6-lutidine (1.3 mL) in DMF (18 mL); Amino acid solution: 0.2 M in DMF.
Pre-loaded amino-based resin (as described above) (50 mg) was swelled in DMF (3 mL) at rt for 30 min. The DMF was then drained, and the resin was immersed in 20% piperidine in DMF (this step was repeated). The resin was then washed with DMF (3×3 mL), DCM (3×3 mL) and again with DMF (3×3 mL). In a separate vessel, the desired amino acid solution (1.29 mL), HATU (452 μL, 4.5 equiv.) and base solution (110 μL) were mixed and then added to the resin. The resultant suspension was then agitated at rt for 30 min, the syringe was flushed, and the coupling step was repeated. Coupling success was monitored with the Kaiser test. Following successful coupling, the resin was washed with DMF (3×3 mL), DCM (3×3 mL) and DMF (3×3 mL). The resin was then immersed in capping solution (vide supra) for 5 min. The syringe was flushed, and the resin was washed with DMF (3×3 mL), DCM (3×3 mL) and DMF (3×3 mL). The process was then repeated (from the deprotection step) until the desired sequence was obtained.
Cleavage from the resin was achieved by submerging it in a mixture of TFA/phenol/water/TIPS (88/5/5/2) and agitating for 3 h, followed by dropwise precipitation into ice cold diethyl ether. The resultant precipitate was then dissolved in acetic acid and lyophilized, affording the desired peptide as the acetate salt.
Purification of RNA-peptide conjugates was carried out by IEX preparative HPLC using a PL-SAX (Agilent) 22×150 mm 1000 Å column at 75° C. with a flow rate of 15 mL/min and UV detection at 260 nm. Elution was performed with a linear gradient selected based on impurity profile, determined by analytical testing using either a DNAPac PA200RS UPLC column or PL-SAX analytical column. Buffer A: 25 mM Tris HCl, pH 8.0, 20% acetonitrile, 10 mM sodium perchlorate; buffer B: 25 mM Tris HCl, pH 8.0, 20% acetonitrile, 600 mM sodium perchlorate.
Although dispersing the modifications throughout a nucleic acid nanoparticle structure has its advantages, some modifications might not be suitable for internal positioning. In such instances, 5′ modification of the oligonucleotide can help mitigate any potential issues. 5′ modification has been shown extensively in the literature. The following examples are incorporated herein by reference and include small molecules (E. Paredes, M. Evans, S. R. Das, RNA labeling, conjugation and ligation, Methods. 54 (2011) 251-259), peptides (K. Klabenkova, A. Fokina, D. Stetsenko, Chemistry of peptide-oligonucleotide conjugates: A review, Molecules. 26 (2021) 1-36), polymers (F. Xiao, Z. Wei, M. Wang, A. Hoff, Y. Bao, L. Tian, Oligonucleotide-Polymer Conjugates: From Molecular Basics to Practical Application, Springer International Publishing, 2020) and lipids (X. Li, K. Feng, L. Li, L. Yang, X. Pan, H. S. Yazd, C. Cui, J. Li, L. Moroz, Y. Sun, B. Wang, X. Li, T. Huang, W. Tan, Lipid-oligonucleotide conjugates for bioapplications, Natl. Sci. Rev. 7 (2020) 1933-1953).
The structure-function relationship of oligonucleotides has inspired the development of alternative functional materials that utilize the phosphodiester backbone (N. Appukutti, C. J. Serpell, High definition polyphosphoesters: Between nucleic acids and plastics, Polym. Chem. 9 (2018) 2210-2226). Rather than forming chains of nucleotides, functional monomers are linked together with phosphodiesters via phosphoramidite chemistry. Although there are size limitations to these polymers, it is possible to get completely monodisperse, sequence-defined materials in good yields. A powerful way of introducing modifications at the oligonucleotide level is through the introduction of oligophosphate polymers at the 5′ end (
Incorporation of the modification post-synthetically may come with some challenges and full conversion may be difficult to achieve. To avoid these potential issues, the PK/PD modulating modification may also be incorporated directly onto the functional monomer. The central core may be a tertiary amine or serinol-based. Proposed synthetic routes are given in
The oligophosphate may be conjugated to the 5′ end of an oligonucleotide via a click handle that is incorporated within the oligophosphate chain. This might be at either terminal position or any given internal position.
RNA therapeutics may be conjugated with highly modified oligophosphate strands, as outlined in
Additionally, direct modification of the 2′ position of nucleosides may also be carried out to form a more naturally derived modifying polymer. For example, highly modified XNA strand C-1.4 was coupled to S-1.3, whereby the long strand (C-1.4) was modified with 2′OMe throughout and the PPIB sense strand (S-1.3) was modified with 2′OMe and PTO. The purification of these species is non-trivial and required extensive IEX method development. Heavily modified conjugated strands, particularly strands that have a high loading of 2′OMe, tend to co-elute with the starting material. (
The nucleic acid nanoparticles described in this invention may incorporate xeno nucleic acids (XNAs) in the backbone of the component oligonucleotides. XNAs are chemically modified nucleic acid analogues, whereby the sugar component is either modified or replaced. 2′F and 2′OMe modified nucleosides are classed as XNAs and these are incorporated into many aspects of the current invention. Additional XNA modifications that may be incorporated into the nucleic acid nanoparticles include, but are not limited to, FNA, FANA, 2′,4′-diFANA, 2′OMe, MOE, 2S-MOP, LNA, AmNa, R-5′-Me-LNA, S-5′Me-LNA, methylene cLNA, N-MeO-amino BNA, 2′4′-BNANC, N-Me-aminooxy BNA, 2′4′-BNAcoc, 2′4′-BNAcocPh, tricyclo DNA, HNA, FHNA, S-cEt, s-cMOE, CeNA, F-CeNA, Me-SRNA, MOE-SRNA, TNA, UMA, WNA, GuNA.
Nucleic acid nanoparticles were imaged by Atomic Force Microscopy using a Bruker Dimension FastScan XR using Bruker FastScan D cantilevers. To immobilize the Mergo, 6 μL of sample at 10 μM were added to a freshly cleaved mica disk with 30 μl of NiCl2 buffer. After 30 min of incubation, excess sample was removed by performing three washes with NiCl2 buffer. Imaging was performed in 60 μL of NiCl2 buffer. The AFM images are shown in
Stoichiometric amounts of the different strands (5 to 7) were combined in the assembly buffer (PBS+MgCl2 (2 mM)), with a final construct concentration of 10 μM. The strands were annealed to each other at 95° C. for 5 min then slowly cooled down to 4° C. (2.5° C./min), using a PCR thermocycler. The scaffold was then analyzed by native polyacrylamide gel electrophoresis (PAGE). For PAGE, the assembled scaffold was electrophoresed on native PAGE (6%) in 1×TBMg (890 mM Tris Borate+20 mM Mg(OAc)2, pH=8.3) at a constant voltage of 100 V. Gel bands were visualized using Cy3, then stained with GelRed™. 10 pmol of structures was loaded. 2 μL of glycerin (70% in H2O) was added to samples before loading. The resultant native PAGE is shown in
Particle size (hydrodynamic diameter, d) and surface charge (zeta potential) were analyzed on a Zetasizer Ultra instrument (Malvern, UK). We used the diffusion barrier technique to measure 20 μL of sample, in a DTS1070 (Malvern, UK) measurement cuvette. The measurement was performed at 37° C. The analyzed material was set to ‘Proteins’, which reliably measures the electrophoretic mobility of proteins or other fragile samples. Attenuation and voltage selection was set to automatic and equilibration time to 90 sec. The monomodal analysis mode was used for data analysis. The results of this analysis are shown in
Experiments were carried out using a Cary3500 UV-vis spectrophotometer, in quartz cuvettes (rectangular, 10 mm, 70 μL). Concentration of Mergos was 0.5 μM in assembly buffer. 70 μL of silicon oil was pipetted on the top of Mergo solution to prevent evaporation. Absorbance was measured at 260 nm and 375 nm and detected in increments of 1° C. from 15° C. to 95° C. Heating and cooling was done at a rate of 2.5° C./min. Melting temperatures are calculated by taking the temperatures corresponding to the derivative maxima of the curves obtained. These results are shown in
EMSA. To each 1 μL aliquot of 10 μM assembled Mergo (prepared as described previously), was added the desired number of equivalents of human serum or cerebrospinal fluid diluted in 1×PBS. Solutions were incubated at 37° C. for 30 minutes before adding 2 μL of 70% (v/v) glycerine solution to aid PAGE loading. Samples were analyzed by 6% native PAGE, ran for 1 h at 100 V at room temperature (
Gel analysis. PAGE gels were imaged under the Cy3 channel (532 nm) to see the fluorophore-labelled products and after that, under the GelRed channel (staining with GelRed® 1× solution, for approximately 10 minutes). Shifts to a lower mobility reveals binding to serum proteins and formation of a so-called ‘protein corona’.
Band quantification. The intensity of individual bands was quantified using Image Lab software. Boxes were manually drawn around each band of interest and intensity was extracted for each one. The intensity of the bands was normalized to intensity of the band at t=0 h. Data was then analyzed using GraphPad Prism software. EC50 shift equation was used to determine EC50 (Y=Bottom+(Top−Bottom)/(1+(EC/X){circumflex over ( )}HillSlope), where X is the concentration, Y is the band intensity).
Table 12 as provided in
Snake Venom Phosphodiesterase stability assays (
RNase III stability experiments (
Gel analysis. PAGE were imaged under Cy3 channel (532 nm) to see the fluorophore-labelled products and after that, under the GelRed channel (staining with GelRed 1× solution, for approximately 10 minutes).
Band quantification. The intensity of individual bands was quantified using Image Lab and/or ImageJ software. Boxes were manually drawn around each band of interest and intensity was extracted for each one. In stability experiments, background intensity was considered. The intensity of the bands was normalized to intensity of the band at t=0 h. Data was analyzed using GraphPad Prism software. First-order decay kinetics were assumed to calculate half-life.
Transfections of human A549 lung carcinoma cells were performed either as forward transfections with Lipofectamine 2000 (11668027, Invitrogen, Thermo Fisher Scientific) (where indicated) in 24-well plates or as reverse transfections in 96-well plates using Lipofectamine RNAiMAX (13778150, Invitrogen, Thermo Fisher Scientific) as transfection reagent. The siGENOME RISC-Free Control (D-001220-01-05, Dharmacon) was used as a non-targeting transfection control (NTC).
For forward transfections, A549 cells in logarithmic growth phase were plated at 5,000 cells/well in a 24-well plate on the day prior to transfection. One day later, the cell culture medium was aspirated and replaced by 200 μL of fresh DMEM/F12 medium (11330032, Gibco, Thermo Fisher Scientific) supplemented with 10% (v/v) heat-inactivated fetal bovine serum (F9665, Sigma Aldrich, Merck) and 1% (v/v) Penicillin-Streptomycin solution (15140122, Gibco, 15140122). Mergo constructs or free siRNA were diluted in Opti-MEM™ I Reduced Serum Medium (31985070, Gibco, Thermo Fisher Scientific) to 6× the final concentration. Likewise, Lipofectamine 2000 reagent was diluted 1:100 in Opti-MEM and incubated for 5 min at room temperature. Equal volumes of RNA dilution and lipofectamine dilutions were then combined and, after an incubation period of 20 minutes at room temperature, 100 μL of oligomer-Lipofectamine 2000 complexes were added to each well containing cells and medium. The cells were incubated at 37° C. in a CO2 incubator for 48 hours, washed with cold PBS and the plate frozen at −80° C. Total RNA was extracted using RNeasy Plus Mini kits (74136, Qiagen), reverse transcribed with random primers using Superscript III reverse transcriptase (18080093, Invitrogen, Thermo Fisher Scientific) and the cDNA was then subjected to real-time PCR on a Quantstudio 5 thermal cycler (Applied Biosystems, Thermo Fisher Scientific) using PowerUp SYBR Green Master Mix (A25742, Applied Biosystems, Thermo Fisher Scientific).
For reverse transfections, 5 μL of RNA diluted in OptiMEM to 20× the final concentration was combined with 0.2 μL of Lipofectamine RNAiMAX and 14.8 μL Opti-MEM in each well of a 96-well plate. After a 15-minute incubation at room temperature, 80 μL of A549 cells in DMEM/F12 supplemented with 10% FBS without antibiotics were added to reach a final cell density of 4,000 cells per well. Approximately 48 hours later, the cells were either subjected to RNA extraction as described above or processed using the FastLane Cell SYBR® Green Kit (216213, Qiagen) or the Luna® Cell Ready One-Step RT-qPCR Kit (E3030S, New England Biolabs) as indicated, according to manufacturer's instructions. One-step RT-qPCR was performed on a Quantstudio 5 thermal cycler or a qTOWER3 84 instrument (Analytik Jena). Primer sequences were hPPIB forward (5′-GTTTGGCAAAGTTCTAGAGG-3′), hPPIB reverse (5′-ACATCCTTCAGGGGTTTATC-3′), hRPLP0 reverse (5′-CTTCGCTGGCTCCCACTT-3′) and hRPLP0 forward (5′-CCATTGAAATCCTGAGTGATGTG-3′).
All transfections and qPCR runs were performed in technical triplicate in two independent experiments unless otherwise stated. The data is shown in
Using Lipofectamine RNAiMAX (13778150, Thermo Fisher Scientific) as a transfection reagent, human A549 lung carcinoma cells were reverse transfected with Mergos loaded with one to two mono- or di-siRNAs at a cell density of 4,000 cells per well in a 96-well plate according to manufacturer's instructions. Mergos were transfected at equal concentration (0.1 nM) independent of the number of siRNA molecules attached. For comparison, unloaded mono- or di-siRNAs were transfected at concentrations equivalent to 1× (0.1 nM), 2× (0.2 nM) and 4× (0.4 nM) the base concentration. siGENOME RISC-Free Control (D-001220-01-05, Dharmacon) was used as a non-targeting transfection control (NTC). After 48 hours, the cells were washed with cold PBS and the plate frozen at −80° C. Total RNA was extracted using RNeasy Plus Mini kits (74136, Qiagen), reverse transcribed with random primers using Superscript III reverse transcriptase (18080093, Invitrogen, Thermo Fisher Scientific) and the cDNA was then subjected to real-time PCR on a Quantstudio 5 thermal cycler (Applied Biosystems, Thermo Fisher Scientific) using PowerUp SYBR Green Master Mix (A25742, Applied Biosystems, Thermo Fisher Scientific). For statistical analysis, 1-way repeated measures ANOVA was performed with Tukey's post-hoc test.
The results of this experiment are shown in
The GFP-GAL9 assay was also utilized to determine endosomal escape (
Stable Hela cells expressing GFP-GAL9 were generated by lentiviral transduction. Cells were seeded at 20×103 cells/well (96-well) and incubated with lentivirus packaged with the GFP-GAL9-355 vector as per manufacturer's instructions. Cells were incubated for 48 h before the addition of 1 g/ml puromycin to select for stably integrated cells. The generated Hela cells stably expressing GFP-GAL9 were seeded at 7.5×103 cells/well (96-well) and incubated with 200 nM of each Mergo (SQ) or 75 μM chloroquine. At the end of each experiment (0, 1, 4, 8 and 24 h) cells were washed with phosphate buffered saline (PBS) and fixed with 4% paraformaldehyde (PFA) for 10 min at room temperature (RT). After fixation, cells were washed with PBS, permeabilized with 0.1% Triton-X for 10 min, washed with PBS and incubated with 2% BSA in PBS for 30 min. Nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI), washed with PBS, mounted on microscope slides and imaged using a confocal microscope. Images were processed on Columbus software. Data were plotted on GraphPad Prism 9.3.1.
All experimental studies involving animals were approved by the UK Home Office. Female BALB/c mice (weight: approximately 20 g, 8-12 weeks old) received two tail vein injections of 200 μL Mergo at 10 ml/kg (day 0 and day 3). Each injection corresponds to 2 nmol of Mergo at a concentration of 10 μM. Mergos carry a Cy3 fluorescent dye. Vehicle injections were used as negative control and state of the art Lipid Nanoparticles (LNP) were used as comparison. Animals were group housed as appropriate in the animal facility and maintained under a 12 h light/dark cycle with free access to food and water, where temperature and humidity were controlled according to Home Office regulations.
On day 7, the animals were sacrificed by cervical dislocation and the liver, kidney, spleen, pancreas, lung and heart were excised and imaged for 5 sec, on an Azure biosystems c600. The imager was set at the Cy3 filter with a 127 nm resolution. The fluorescent intensity of the organs was analyzed on ImageJ Fiji. As the Cy3 intensity depends on the chemical modifications of the Mergo, the data were corrected to the corresponding Cy3 intensity. Organ autofluorescence was assessed by the control—vehicle injected animals. Mergo fluorescence was subsequently normalized to the control vehicle.
Organs stored in RNAlater (Sigma, R0901) were pierced using 2 mm punches to assess the gene silencing by mRNA quantification. The pierced tissues were lysed using QuantiGene Sample Processing Kit, Tissues (Invitrogen, QS0106) according to the manufacturer's instructions using a Tissue Lyzer II (Qiagen). mRNA was detected according to the Quantigene 2.0 protocol using the following probe sets: mouse HPRT (SB-15463), mouse PPIB (SB-10002). All data were plotted on GraphPad Prism 9.3.1. The data from these experiments is shown in
Blood, typically 100 μl, was collected into sodium heparin, from mice via the tail vein, at 2 h post injection and plasma prepared by centrifugation (14,000 rpm, 4° C., 5 min.). Cytokine levels, using manufactured bespoke kits, were determined by MAGPIX Luminex system. Clinical chemistry of plasma ALT/AST was completed using a Beckman Coulter instrument. Mice were weighed prior to treatment and on day 7 post initiation of the treatment. The data from these experiments are shown in
The core nanostructure will comprise L-RNA as shown in
To study the proof of concept of the RNA origami, a commercially available mRNA encoding for eGFP was bought from Trilink to be used as scaffold mRNA. As the single-stranded staple strands offer a lot of possibilities in terms of composition and complementarity regions, the decision was made to study their fundamental properties. Antisense oligos were designed (Sequences are given in Table 13) complementary to the open reading frame of the eGFP mRNA. The same sequence as unmodified DNA, PS-DNA, 2′OMe-DNA and unmodified RNA (
600 fmol of mRNA were assembled with 3 pmol of antisense oligos in 1 μM PBS. DNA oligo, thiophosphorylated (ps) DNA oligo, RNA oligo and 2′OMe-DNA oligo was compared.
The mRNA was incubated with the antisense species for 5 min at 80° C., followed by a 30 min temperature gradient 80 to 25° C. The assembly was then transfected in A549 cells and fluorescence was measured 22 h after transfection on Tecan Infinite 200 Pro. The next step was to assess how the introduction of folding on the scaffold mRNA through RNA and/or 2′OMe-DNA single-stranded staples influences the translation efficiency. Translation should be more inhibited when the folding is induced on regulatory elements present in the 5′ and/or 3′ untranslated regions and less inhibited when folding is occurring in the open reading frame. Based on
To test this, 600 fmol of mRNA were assembled with 3 pmol of 2′OMe-DNA staple strands in 1 μM PBS. The mRNA was incubated with the antisense species for 5 min at 80° C., followed by a 30 min temperature gradient 80 to 25° C. The assembly was then transfected in A549 cells and fluorescence was measured 22 h after transfection on Tecan Infinite 200 Pro.
Modulation of translation efficiency based on the hybridized region of the mRNA scaffold both on the open reading frame and 3′UTR is observed (
A semi-assembled version of the 10HB_rectangle (variant P with staples R19-25) was tested for RNase H resistance. For this purpose, 2′OMe mixmers with a gap space of DNA nucleotides between the 2′OMe modifications (Table 14) were acquired. 2′OMe modified and non-modified DNA staple strands (20 pmol each strand) were added to separate tubes eGFP mRNA (800 fmol). The mixtures were assembled via a 1 h gradient from 75 to 20° C. After assembly the volumes of the 2′OMe modified and the non-modified assay were quartered. To the aliquots 0, 1.25, 2.5, and 5 U of RNAse H were added together with RNase H buffer. The mixtures were incubated at 25° C. for 30 min and cleavage was analyzed via native 2% agarose gel (
To fold an origami applying a mRNA as a scaffold and short DNAs as staple strands, an 8-9-8-8 crossover strand layout for the design of the 2D rectangular origami 10HB_rectangle was used. This is used to compensate for the different helix geometry that comes with the change from DNA-DNA duplex (B-helix, 10.5 bp/turn) to RNA-DNA (A-helix, 11.0 bp/turn) (see
The chosen sequence for the scaffold was an mRNA that codes for an enhanced green fluorescence protein (eGFP) with the 5′-UTR of HIV envelope glycoprotein and 3′-UTR of hemoglobin alpha-2 (see Table 15). The mRNA was purchased from TriLink Biotechnologies. The sequence for the DNA staples that were used to fold the different structures out of the eGFP mRNA are shown in Table 15. The individual DNA staple strands were purchased from IDT. For effective translation, the mRNA was 5′-capped (cap1) and poly(A)-tailed. The designs were successfully tested in oxDNA on their molecular dynamic parameters (see
Assembly of mRNA Origami Constructs
600 fmol mRNA was assembled with 12 pmol DNA mix (either rectangle, tube or block mixture) in 10 mM Tris/HCl (pH 7.0) and 120 mM NaCl. The mixture was incubated for 5 min at 75° C., slowly (2° C./min) to 65° C. and very slowly (1° C./min) cooled down to 20° C. The samples were purified via 50 MWCO spin filter (4 wash steps, 10 min, 12,000 rcf) and the assembly was verified via band-shift assay on a 2% agarose gel (
AFM Imaging of mRNA Origami (
A freshly cleaved mica surface was preincubated with 10 mM NiOAc solution for 20 sec and washed three times with TE-buffer. The origami samples were highly diluted in TE-buffer. An aliquot of this diluted sample was put on the mica surface and incubated for 10 min prior to the imaging.
The chosen sequence for the scaffold is an mRNA that codes for an enhanced green fluorescence protein (eGFP) with the 5′-UTR of HIV envelope glycoprotein and 3′-UTR of hemoglobin alpha-2 (see Table 15). The mRNA was purchased from TriLink Biotechnologies. The sequence for the DNA staples that were used to fold the different structures out of the eGFP mRNA are shown in Table 16. The individual DNA staple strands were purchased from IDT. For effective translation, the mRNA was 5′-capped (cap1) and poly(A)-tailed.
For the ‘Handle_basic’ design (
Given that circular RNA has been shown to be more stable than linear mRNA, the delivery of circular mRNA scaffolds (mRNA with its ends covalently linked) with increasing number of staples (Circular_2staples, Circular_4staples) will also be tested (
The branched siRNA was designed to increase the therapeutic loading capacity of the nucleic acid constructs and allow for the synthesis of more potent therapeutics in a more sustainable way (i.e., by reducing waste). This methodology will also enhance Mergo versatility and speed of development, i.e., ability to readily adapt to delivery of multiple, different cargo types, creating an intelligent delivery system that goes beyond the limitations of current standards. The designs utilised in this invention are outlined in
The branching unit was incorporated into the oligonucleotide sequence using solid phase oligonucleotide synthesis. The branching unit allows attachment of more than one therapeutic moiety at a given location.
Alternatively, a double siRNA approach is also used which includes connections of two siRNA units with a linker poly thymidine (poly-T) in the form of a combinatorial chain (
Stoichiometric amounts of the different strands (5 to 7) were combined in the assembly buffer (PBS+MgCl2 (2 mM)), with a final construct concentration of 10 μM. The strands were annealed to each other at 95° C. for 5 min then slowly cooled down to 4° C. (2.5° C./min), using a PCR thermocycler. The scaffold was then analyzed by native polyacrylamide gel electrophoresis (PAGE). For PAGE, the assembled scaffold was electrophoresed on native PAGE (6%) in 1× TBMg (890 mM Tris Borate+20 mM Mg(OAc)2, pH=8.3) at a constant voltage of 100 V. Gel bands were visualized using Cy3, then stained with GelRed™. 10 pmol of structures was loaded. 2 μL of glycerin (70% in H2O) was added to samples before loading (
References and citations to other documents, such as patents, patent applications, patent publications, journals, books, papers, web contents, have been made throughout this disclosure. All such documents are hereby incorporated herein by reference in their entirety for all purposes.
Various modifications of the invention and many further embodiments thereof, in addition to those shown and described herein, will become apparent to those skilled in the art from the full contents of this document, including references to the scientific and patent literature cited herein. The subject matter herein contains important information, exemplification, and guidance that can be adapted to the practice of this invention in its various embodiments and equivalents thereof.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/IB2022/000218 | 4/15/2022 | WO |
| Number | Date | Country | |
|---|---|---|---|
| 63175185 | Apr 2021 | US |
