
==== Front
Nucleic Acids Res
Nucleic Acids Res
nar
Nucleic Acids Research
0305-1048
1362-4962
Oxford University Press

39149897
10.1093/nar/gkae681
gkae681
AcademicSubjects/SCI00010
Chemical Biology and Nucleic Acid Chemistry
Rational design of base, sugar and backbone modifications improves ADAR-mediated RNA editing
Lu Genliang Wave Life Sciences, Cambridge, MA, USA

Shivalila Chikdu Wave Life Sciences, Cambridge, MA, USA

Monian Prashant Wave Life Sciences, Cambridge, MA, USA

Yu Hui Wave Life Sciences, Cambridge, MA, USA

Harding Ian Wave Life Sciences, Cambridge, MA, USA

Briem Stearne Wave Life Sciences, Cambridge, MA, USA

Byrne Michael Wave Life Sciences, Cambridge, MA, USA

Faraone Alyse Wave Life Sciences, Cambridge, MA, USA

Friend Stephen Wave Life Sciences, Cambridge, MA, USA

Huth Olivia Wave Life Sciences, Cambridge, MA, USA

Iwamoto Naoki Wave Life Sciences, Cambridge, MA, USA

Kawamoto Tomomi Wave Life Sciences, Cambridge, MA, USA

Kumarasamy Jayakanthan Wave Life Sciences, Cambridge, MA, USA

Lamattina Anthony Wave Life Sciences, Cambridge, MA, USA

Longo Kenneth Wave Life Sciences, Cambridge, MA, USA

McCarthy Leah Wave Life Sciences, Cambridge, MA, USA

McGlynn Andrew Wave Life Sciences, Cambridge, MA, USA

Molski Allison Wave Life Sciences, Cambridge, MA, USA

Pan Qianli Wave Life Sciences, Cambridge, MA, USA

Pu Tom Wave Life Sciences, Cambridge, MA, USA

Purcell-Estabrook Erin Wave Life Sciences, Cambridge, MA, USA

Rossi Jeff Wave Life Sciences, Cambridge, MA, USA

Standley Stephany Wave Life Sciences, Cambridge, MA, USA

Thomas Carina Wave Life Sciences, Cambridge, MA, USA

Walen Alexandra Wave Life Sciences, Cambridge, MA, USA

Yang Hailin Wave Life Sciences, Cambridge, MA, USA

Kandasamy Pachamuthu Wave Life Sciences, Cambridge, MA, USA

https://orcid.org/0000-0002-4690-7543
Vargeese Chandra Wave Life Sciences, Cambridge, MA, USA

To whom correspondence should be addressed. Tel: +1 6179492900; Fax: +1 6179492901; Email cvargeese@wavelifesci.com
Correspondence may also be addressed to Pachamuthu Kandasamy. Email: pkandasamy@wavelifesci.com
The first three authors should be regarded as Joint First Authors.

23 9 2024
16 8 2024
16 8 2024
52 17 1006810084
25 7 2024
20 7 2024
19 4 2024
© The Author(s) 2024. Published by Oxford University Press on behalf of Nucleic Acids Research.
2024
https://creativecommons.org/licenses/by-nc/4.0/ This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial License (https://creativecommons.org/licenses/by-nc/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited. For commercial re-use, please contact journals.permissions@oup.com

Abstract

AIMers are short, chemically modified oligonucleotides that induce A-to-I RNA editing through interaction with endogenous adenosine deaminases acting on RNA (ADAR) enzymes. Here, we describe the development of new AIMer designs with base, sugar and backbone modifications that improve RNA editing efficiency over our previous design. AIMers incorporating a novel pattern of backbone and 2′ sugar modifications support enhanced editing efficiency across multiple sequences. Further efficiency gains were achieved through incorporation of an N-3-uridine (N3U), in place of cytidine (C), in the ‘orphan base’ position opposite the edit site. Molecular modeling suggests that N3U might enhance ADAR catalytic activity by stabilizing the AIMer-ADAR interaction and potentially reducing the energy required to flip the target base into the active site. Supporting this hypothesis, AIMers containing N3U consistently enhanced RNA editing over those containing C across multiple target sequences and multiple nearest neighbor sequence combinations. AIMers combining N3U and the novel pattern of 2′ sugar chemistry and backbone modifications improved RNA editing both in vitro and in vivo. We provide detailed N3U synthesis methods and, for the first time, explore the impact of N3U and its analogs on ADAR-mediated RNA editing efficiency and targetable sequence space.

Graphical Abstract

Graphical Abstract

Wave Life Sciences
==== Body
pmcIntroduction

The class of nucleic acid medicines known as RNA medicines has risen in prominence considerably over the last decade. RNA medicines are chemically modified RNA-based molecules that are used to alter the properties or function of specific genes, transcripts, or proteins. One of the more promising emerging RNA medicine modalities is RNA editing, in which synthetic oligonucleotides direct enzymes to alter the base sequences of target mRNAs (1,2). The most advanced RNA editing approaches engage adenosine deaminase acting on RNA (ADAR) enzymes (3).

ADAR enzymes convert adenosine (A) to inosine (I) in double stranded RNAs (3). The translation machinery reads inosine (I) as guanosine (G); therefore, correction of pathogenic G-to-A point mutations is an attractive therapeutic application. Using synthetic oligonucleotides to direct site-specific ADAR-mediated point mutation correction has several potential advantages, including reversibility, cell cycle independence, avoidance of double-stranded DNA breaks, and circumvention of the permanent off-target effects seen with DNA editing (4,5).

A major challenge for the therapeutic development of RNA editing is that ADAR enzymes have innate bias toward target adenosines within specific sequence contexts, limiting the targetable sequence space (6). More specifically, ADAR enzymes exhibit ‘preferences’ for the 5′- and 3′-nucleotide context of the target A, leading to extremely low editing efficiency for some sequences. Both ADAR1 and ADAR2 prefer an A with a 5′-nearest neighbor that is uridine (U > A > C > G) (3); ADAR1 and ADAR2 have similar but not identical 3′-nearest neighbor preferences (G > C ≈ A > U and G > C > U ≈ A, respectively) (7). The 5′- and 3′-nearest neighbor sequence preferences derive from their impact on flipping of the target A into the active site for editing, rather than through direct sequence detection, suggesting that the efficiency of base flipping is a driver of editing efficiency overall (3,8).

ADAR activity is also affected by the base opposite to its target A, which is known as the ‘orphan base’ position (8). ADAR enzymes prefer A-C mismatches in the edit site. This preference arises from interactions between the orphan base and a conserved glutamic acid residue (E488 in ADAR2, E1008 in ADAR1) responsible for flipping the target A into the active site for deamination. Glutamate can hydrogen (H) bond with C, but clashes with G or A (9). While glutamate can form a similar H bonding pattern with U in the orphan position, it comes at an energetic cost to base flipping due to U–A base pairing (9). The importance of the glutamate-orphan base interaction for catalytic activity is supported by work showing glutamine substitution of the residue, E488Q or E1008Q, results in hyperactive ADAR2 and ADAR1, respectively. Importantly, the hyperactivity of ADAR2 E488Q is attributed to enhanced H bonding between Q488 and the orphan base C (9). In sum, the efficiency of base flipping and H bonding between the orphan base and ADAR residue E488/E1008 are crucial factors contributing to editing efficiency.

Our previous work demonstrated that the efficiency of ADAR-mediated RNA editing could be improved through the incorporation of phosphoryl guanidine (PN) backbone modifications in stereopure, short, fully chemically modified oligonucleotides called AIMers (10). The goal of the current study is to further improve the efficiency and potency of AIMer-mediated RNA editing by exploring new chemistry patterns of sugar, backbone, and base. We describe a new AIMer chemistry design, which features a pattern of more distributed sugar and backbone modifications (AIMer-Distributed, AIMer-D) compared to our previously reported segmented sugar and backbone modification design (AIMer-Segmented, AIMer-S). We also investigated, for the first time, the impact of incorporating N-3-uridine (N3U, also referred to as isouridine) in oligonucleotides that elicit A-to-I editing via ADAR enzymes. Based on structural modeling, we hypothesized that incorporating N3U into the orphan position in AIMers might deliver RNA editing efficiency gains by mimicking the RNA-enzyme interactions present in the hyperactive ADAR1/ADAR2 mutants.

Materials and methods

Structural modeling

The crystal structure of the Human ADAR2 deaminase domain in complex with an RNA duplex (PDB code: 7KFN) was retrieved from the Protein Data Bank (4,11). The Protein Preparation Wizard in Schrödinger Maestro (version 12.9.123, installed in a Valence VWS-1690441-SMD workstation with CentOS 7) was used to prepare the downloaded structure. Next, the chemical modifications on each nucleobase were incorporated opposite to the edited base (placed optimally to minimize steric clashes with neighboring side chains) using the structure editing option in Maestro. Finally, the duplex-protein complex model with all the modifications was energy-minimized using the OPLS4 force field as implemented in Maestro Protein Preparation Wizard using a root mean square deviation (RMSD) Convergence criteria of 0.3 Å on heavy atoms. All drawings were generated using the Maestro software.

Calculation of pKa values

The Jaguar program was used to optimize the gas phase geometries using DFT-D3 and 6–31G level of theory. The optimized structures were used to calculate the pKa using the jaguar pKa module in Schrödinger Maestro (version 12.9.123, installed in a Valence VWS-1690441-SMD workstation with CentOS 7). Each nucleobase with a methyl group at the N1 position for C, U and T and N3 position for N3U and its analogs was used for the analysis.

Compound synthesis

General conditions

Flash column chromatography was performed using a Teledyne ISCO Combi System with pre-packed RediSep Teledyne ISCO silica gel cartridges. In the case of L-PSM amidite, the cartridge was washed with 2.5% triethylamine in acetonitrile (MeCN) and 2.5% triethylamine in ethyl acetate (EtOAc) followed by loading solvent containing 2.5% triethylamine to neutralize the silica prior to loading the crude samples for purification. All moisture sensitive reactions were carried out under anhydrous conditions using dry glassware, anhydrous solvents, and under argon atmosphere. ESI-MS spectra were recorded on a Waters SQD2 system. HRMS (ESI) spectra were recorded on a Waters Xevo G2-XS QTof Quadrupole Mass Spectrometer. 1H, 19F and 31P NMR spectra were recorded at 400 or 600, 376 or 565, and 162 or 243 MHz, respectively. Spectra are reported as ppm and are referenced to the solvent resonances in deuterated chloroform (CDCl3), or deuterated dimethyl sulfoxide (DMSO-d6).

For synthesis of l-phenylsulfonylmethyl (PSM) 2′-deoxy 5-substituted N-3-uridine 7b–d, 2′-O-tert-butyldimethylsilyl (TBS) N-3-uridine 18a, 3′-OTBS N-3-uridine 18b, 2′-MOE N-3-uridine 20, arabino N-3-uridine 26, (S/R) glycol nucleic acid (GNA) N-3-uridine 29a and 29b, 2′-fluoroarabino N-3-uridine 33, 2′-deoxy-5-(prop-1-yn-1yl) N-3-uridine 38 and 2′-deoxyl 6-methyl N-3-uridine 42 amidites see Supplementary Materials and Methods.

Synthesis of L-PSM 2′-deoxy N-3-uridine amidite: 7a

3-((2R,4S,5R)-4-Hydroxy-5-(hydroxymethyl)tetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione: 3a

NaH (6.62 g, 165.45 mmol), 60% in mineral oil was added slowly to a solution of compound 2a (17.0 g, 110.3 mmol) in acetonitrile (MeCN) (1700 mL) under argon at 0°C and stirred at 0°C for 30 minutes. After 1-chloro-3,5-di(4-chlorbenzoyl)-2-deoxy-d-ribose 1a (66.0 g, 156.30 mmol) was added to the reaction mixture, the reaction mixture was stirred for 30 min at 0°C and at 65°C for 3 h. The reaction mixture was cooled to room temperature and filtered. The filtrate was concentrated and purified by flash chromatography over silica gel column chromatography to give a mixture of alpha and beta isomers, which was triturated with EtOAc (66 ml) and MeCN (3.3 ml) at 60°C (4 times) to yield beta isomer 3a (13.5 g, 24%) as a white solid. Proton nuclear magnetic resonance (1H NMR) (400 MHz, DMSO-d6) δ 11.17 (br s, 1H), 8.02–7.88 (m, 4H), 7.61–7.56 (m, 2H), 7.55–7.50 (m, 2H), 7.46 (d, J = 7.7 Hz, 1H), 6.72 (dd, J = 4.3, 9.0 Hz, 1H), 5.77 (td, J = 5.8, 8.0 Hz, 1H), 5.60 (d, J = 7.5 Hz, 1H), 4.66–4.57 (m, 1H), 4.52–4.46 (m, 1H), 4.44–4.36 (m, 1H), 3.03 (tt, J = 4.2, 8.9 Hz, 1H), 2.48–2.40 (m, 1H); electrospray ionization mass spectrometry:MS (ESI), 527.0 [M + Na]+.

3-((2R,4S,5R)-4-Hydroxy-5-(hydroxymethyl)tetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione: 4a

Compound was described previously (12). Sodium methoxide (NaOMe) (10.82 g, 200.4 mmol) was added slowly to a solution of compound 3a (40.5 g, 80.15 mmol) in 580 ml of methanol (MeOH) at 15°C and was stirred at room temperature for 1 h. The reaction mixture was adjusted to neutral by addition of ammonium chloride, filtered, washed with MeOH. This solution was concentrated and purified by flash chromatography over silica gel eluted with dichloromethane (DCM)/MeOH to yield compound 4a (17.0 g, 93%) as a white solid. 1H NMR (400 MHz, DMSO-d6) δ 11.07 (s, 1H), 7.42 (d, J = 7.6 Hz, 1H), 6.53 (dd, J = 6.4, 8.0 Hz, 1H), 5.54 (d, J = 7.6 Hz, 1H), 5.06 (d, J = 4.1 Hz, 1H), 4.54 (t, J = 5.4 Hz, 1H), 4.29 (dd, J = 3.9, 6.9 Hz, 1H), 3.69–3.63 (m, 1H), 3.57 (td, J = 4.0, 11.4 Hz, 1H), 3.48–3.40 (m, 1H), 2.74–2.62 (m, 1H), 1.91 (ddd, J = 4.5, 8.2, 12.8 Hz, 1H); MS (ESI), 227.1 [M–H]−.

3-((2R,4S,5R)-5-((Bis(4-methoxyphenyl)(phenyl)methoxy)methyl)-4-hydroxytetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione: 5a

To a stirred solution of compound 4a (17.0 g, 74.50 mmol) in pyridine (250 ml) was added 4,4-dimethoxytrityl chloride (DMTrCl) (25.24 g, 74.50 mmol) and then stirred at room temperature overnight. The reaction mixture was diluted with water, extracted with EtOAc, dried over magnesium sulfate (Mg2SO4), filtered and concentrated under reduced pressure to give a residue, which was purified by flash chromatography over silica gel (petroleum ether/ethyl acetate = 10:1, 0:1, then DCM: MeOH = 20:1, 5% TEA) to yield compound 5a (31.0 g, 76%) as a white solid. 1H NMR (400MHz, DMSO-d6) δ 11.04 (br s, 1H), 7.48–7.35 (m, 3H), 7.30–7.15 (m, 7H), 6.92–6.77 (m, 4H), 6.59 (dd, J = 4.3, 8.9 Hz, 1H), 5.55 (d, J = 7.6 Hz, 1H), 5.06 (d, J = 5.4 Hz, 1H), 4.31–4.19 (m, 1H), 3.86–3.77 (m, 1H), 3.73 (d, J = 2.1 Hz, 6H), 3.21 (dd, J = 8.0, 9.6 Hz, 1H), 3.06 (dd, J = 3.4, 9.8 Hz, 1H), 2.59 (ddd, J = 4.3, 8.3, 12.9 Hz, 1H), 2.06–2.00 (m, 1H); 13C NMR (151 MHz, DMSO-d6) δ 162.97, 157.97, 157.94, 150.67, 145.15, 141.19, 135.91, 135.78, 129.74, 129.72, 127.78, 127.65, 126.51, 113.04, 112.99, 100.17, 85.52, 85.23, 80.39, 71.37, 64.77, 54.99, 54.97, 37.22; HRMS (ESI/Q-TOF) m/z: [M + Na]+ Calculated for C30H30N2O7Na 553.1951; Found 553.1943.

3-((2R,4S,5R)-5-((Bis(4-methoxyphenyl)(phenyl)methoxy)methyl)-4-(((1S,3S,3aS)-3-((phenylsulfonyl)methyl)tetrahydro-1H,3H-pyrrolo[1,2-c][1,3,2]oxazaphosphol-1-yl)oxy)tetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione: 7a

Triethylamine (19.57 ml, 140.42 mmol) and (3S,3aS)-1-chloro-3-((phenylsulfonyl)methyl)tetrahydro-1H,3H-pyrrolo[1,2-c][1,3,2]oxazaphosphole 6 [113.59 ml of 0.89 M in tetrahydrofuran (THF), 101.1 mmol] was added to a solution of dry compound 5a (29.8 g, 56.17 mmol) in THF (200 ml). The reaction mixture was stirred at room temperature for 2.5 h. Thin layer chromatography (TLC) and liquid chromatography–mass spectrometry (LC–MS) showed that the reaction was complete. The reaction was quenched with water (794 μl) and dried over anhydrous MgSO4. The mixture was filtered through celite, and the filtrate was concentrated to yield the crude product which was purified by column chromatography over silica gel eluting with 50–100% EtOAc in hexanes (each mobile phase contained 1% triethylamine) to yield 7a ((40.4 g, 88%) as a white foam. 1H NMR (600 MHz, CDCl3) δ 9.28 (bs, 1H), 7.87 (dd, J = 8.3, 1.4 Hz, 2H), 7.63–7.57 (m, 1H), 7.53–7.44 (m, 4H), 7.37–7.33 (m, 4H), 7.23 (t, J = 7.8 Hz, 2H), 7.19–7.13 (m, 1H), 6.81–6.75 (m, 4H), 6.70 (dd, J = 8.4, 5.1 Hz, 1H), 6.64 (d, J = 7.7 Hz, 1H), 5.53 (d, J = 7.7 Hz, 1H), 4.96 (q, J = 6.1 Hz, 1H), 4.87 (dq, J = 12.8, 5.6 Hz, 1H), 3.93 (td, J = 5.9, 3.9 Hz, 1H), 3.752 (s, 3H), 3.749 (s, 3H), 3.62 (dq, J = 11.7, 6.0 Hz, 1H), 3.44–3.25 (m, 5H), 2.94 (qd, J = 10.0, 4.0 Hz, 1H), 2.85 (ddd, J = 13.2, 8.0, 5.2 Hz, 1H), 2.23 (ddd, J = 13.6, 8.6, 5.5 Hz, 1H), 1.85–1.79 (m, 1H), 1.76–1.69 (m, 1H), 1.65–1.59 (m, 1H), 1.12–1.02 (m, 1H); 13C NMR (151 MHz, CDCl3) δ 162.90, 158.50, 152.37, 145.19, 139.38, 139.03, 136.41, 136.35, 134.11, 130.34, 129.39, 128.45, 128.33, 127.81, 126.78, 113.12, 113.10, 102.30, 86.07, 84.90 (d, J = 2.7 Hz, 1C), 81.46, 74.92 (d, J = 10.4 Hz, 1C), 73.51 (d, J = 5.5 Hz, 1C), 66.40 (d, J = 3.3 Hz, 1C), 63.76, 58.41 (d, J = 3.8 Hz, 1C), 55.32, 46.35 (d, J = 36.0 Hz, 1C), 36.45 (d, J = 3.8 Hz, 1C), 27.53, 26.11 (d, J = 3.8 Hz, 1C); 31P NMR (243 MHz, CDCl3) δ 149.14; HRMS (ESI/Q-TOF) m/z: [M + Na]+ Calculated for C42H44N3O10PSNa 836.2383; Found 836.2408.

Synthesis of L-PSM 2′-OMe N-3-uridine amidite: 14

(2R,3R,4R,5R)-2-((Benzoyloxy)methyl)-5-(2,6-dioxo-3,6-dihydropyrimidin-1(2H)-yl)tetrahydrofuran-3,4-diyl dibenzoate: 8

NaH (7.92 g) was added to a solution of compound 2a (22.88 g, 148.47 mmol) in MeCN (2000 ml) at 0°C and stirred at 0°C for 30 minutes under nitrogen. Compound 1b (52 g, 98.98 mmol) was added slowly for 30 min at room temperature, stirred at 40°C for 1 h and at 60°C for 2 h. TLC indicated compound 1b was consumed completely. After filtration, the solvent was removed under reduced pressure to yield the crude compound, which was purified by flash chromatography over silica gel (petroleum ether: ethyl acetate = 1:0 to 0:1) to yield compound 8 (15.0 g, 27% yield) as a yellow solid. 1H NMR (400 MHz, DMSO-d6) δ 11.40 (s, 1H), 7.98 (d, J = 7.6 Hz, 2H), 7.91 (d, J = 7.6 Hz, 2H), 7.83 (d, J = 7.6 Hz, 2H), 7.69–7.57 (m, 3H), 7.54 (d, J = 7.5 Hz, 1H), 7.50–7.42 (m, 4H), 7.42–7.34 (m, 2H), 6.51 (s, 1H), 6.12–6.05 (m, 2H), 5.68 (d, J = 7.6 Hz, 1H), 4.72–4.62 (m, 2H), 4.57–4.48 (m, 1H); MS (ESI), 579.3 [M + Na]+.

3-((2R,3R,4S,5R)-3,4-Dihydroxy-5-(hydroxymethyl)tetrahydrofuran-2-yl)-1-(4-methoxybenzyl)pyrimidine-2,4(1H,3H)-dione: 9

Potassium carbonate (K2CO3) (7.45 g, 53.91 mmol) and 4-methoxybenzyl chloride (6.19 g, 39.53 mmol) were added to a solution of compound 8 (20 g, 35.94 mmol) in dimethyl formamide (DMF, 300 ml). The mixture was stirred at 25°C for 12 h. The reaction mixture was diluted with water and extracted with ethyl acetate (EtOAc). The combined organic layers were dried over sodium sulfate (Na2SO4), filtered and concentrated under reduced pressure to give a residue, which was purified by flash chromatography over silica gel (petroleum ether: ethyl acetate = 15:1 to 0:1) to yield (2R,3R,4R,5R)-2-((benzoyloxy)methyl)-5-(3-(4-methoxybenzyl)-2,6-dioxo-3,6-dihydropyrimidin-1(2H)-yl)tetrahydrofuran-3,4-diyl dibenzoate (23 g, 95% yield) as a yellow oil; MS (ESI), 700.3 [M + Na]+. To a solution of (2R,3R,4R,5R)-2-((benzoyloxy)methyl)-5-(3-(4-methoxybenzyl)-2,6-dioxo-3,6-dihydropyrimidin-1(2H)-yl)tetrahydrofuran-3,4-diyl dibenzoate (23.0 g, 34.0 mmol) in MeOH (300 ml) was added NaOMe (5.51 g, 101.97 mmol). The mixture was stirred at 25°C for 3 h. The ammonium chloride (NH4Cl, 5.6 g) was added to the reaction mixture and stirred at room temperature for 10 min, and then concentration of the reaction mixture yielded the crude residue which was purified by flash chromatography over silica gel (petroleum ether: ethyl acetate = 5:1 to 0:1 to ethyl acetate: MeOH = 3:1) to yield compound 9 (12.0 g, 92% over 2 steps) as a yellow oil. 1H NMR (400 MHz, DMSO-d6) δ 7.84 (d, J = 7.9 Hz, 1H), 7.28 (d, J = 8.6 Hz, 2H), 6.98–6.87 (m, 2H), 6.07 (d, J = 3.4 Hz, 1H), 5.70 (d, J = 7.9 Hz, 1H), 5.05 (br s, 1H), 4.87–4.77 (m, 2H), 4.58 (s, 1H), 4.43 (dd, J = 3.5, 5.8 Hz, 1H), 4.09 (t, J = 6.3 Hz, 1H), 3.73 (s, 3H), 3.70–3.52 (m, 2H), 3.45–3.29 (m, 2H); MS (ESI), 365.2 [M + H]+.

1-(4-Methoxybenzyl)-3-((6aR,8R,9R,9aR)-2,2,4,4-tetraisopropyl-9-methoxytetrahydro-6H-furo[3,2-f][1,3,5,2,4]trioxadisilocin-8-yl)pyrimidine-2,4(1H,3H)-dione: 10

Chloro-[chloro(diisopropyl)silyl]oxy-diisopropyl-silane (13.51 g, 42.82 mmol) was added slowly to a solution of compound 9 (12.0 g, 32.94 mmol) in pyridine (200 ml) at 0°C for 1 h under nitrogen. The mixture was stirred at 25°C for 11 h. The reaction mixture was filtered and concentrated under reduced pressure to give a residue, which was purified by flash chromatography over silica gel (petroleum ether: ethyl acetate = 20:1 to 0:1) to yield 3-((6aR,8R,9R,9aS)-9-hydroxy-2,2,4,4-tetraisopropyltetrahydro-6H-furo[3,2-f][1,3,5,2,4]trioxadisilocin-8-yl)-1-(4-methoxybenzyl)pyrimidine-2,4(1H,3H)-dione (15.0 g, 75%) as a yellow oil. Sodium hydride (NaH, 988.61 mg, 24.72 mmol, 60% in mineral oil) and iodomethane (MeI, 7.02 g, 49.44 mmol) were added to a solution of 3-((6aR,8R,9R,9aS)-9-hydroxy-2,2,4,4-tetraisopropyltetrahydro-6H-furo[3,2-f][1,3,5,2,4]trioxadisilocin-8-yl)-1-(4-methoxybenzyl)pyrimidine-2,4(1H,3H)-dione (5.0 g, 8.24 mmol) in DMF (200 ml) at 0°C under nitrogen (N2). The mixture was stirred at 15°C for 10 h. The reaction mixture was quenched with NH4Cl, diluted with water, and extracted with EtOAc. The combined organic layers were dried over Na2SO4 , filtered and concentrated under reduced pressure to give a residue, which was purified by flash chromatography over silica gel (petroleum ether: ethyl acetate = 10:1 to 0:1) to yield compound 10 (4.3 g, 84%) as a yellow oil. 1H NMR (400 MHz, CDCl3) δ 7.23 (d, J = 8.6 Hz, 2H), 7.07 (d, J = 8.0 Hz, 1H), 6.91–6.87 (m, 2H), 6.26 (d, J = 1.6 Hz, 1H), 5.64 (d, J = 8.0 Hz, 1H), 5.06 (dd, J = 5.9, 9.1 Hz, 1H), 4.90 (d, J = 14.6 Hz, 1H), 4.69 (d, J = 14.5 Hz, 1H), 4.22 (dd, J = 1.5, 5.9 Hz, 1H), 4.07 (d, J = 2.8 Hz, 1H), 4.01–3.96 (m, 1H), 3.86–3.80 (m, 4H), 3.51 (s, 3H), 1.17–1.01 (m, 28H); MS (ESI), 621.4 [M + H]+.

3-((2R,3R,4R,5R)-4-Hydroxy-5-(hydroxymethyl)-3-methoxytetrahydrofuran-2-yl)-1-(4-methoxybenzyl)pyrimidine-2,4(1H,3H)-dione: 11

Tetra-n-butylammonium fluoride (TBAF, 1.0 M in THF, 61.20 ml) was added to a solution of compound 10 (19.0 g, 30.60 mmol) in THF (200 ml) at 0°C and the mixture was stirred at 0°C for 4 h. The reaction mixture was filtered and concentrated under reduced pressure to give a residue, which was purified by flash chromatography over silica gel (petroleum ether: ethyl acetate = 1:1 to 0:1 to ethyl acetate: MeOH = 1:1) to yield compound 11 (10.9 g, 94% yield) as a white solid. 1H NMR (400 MHz, DMSO-d6) δ 7.84 (d, J = 7.9 Hz, 1H), 7.29 (d, J = 8.6 Hz, 2H), 6.93 (d, J = 8.6 Hz, 2H), 6.12 (d, J = 3.3 Hz, 1H), 5.71 (d, J = 7.9 Hz, 1H), 4.88 (d, J = 7.1 Hz, 1H), 4.83 (d, J = 7.1 Hz, 2H), 4.58 (t, J = 5.8 Hz, 1H), 4.27–4.14 (m, 2H), 3.74 (s, 3H), 3.66 (dt, J = 3.3, 6.6 Hz, 1H), 3.60 (ddd, J = 3.4, 5.3, 11.7 Hz, 1H), 3.39 (td, J = 6.3, 12.0 Hz, 1H), 3.30 (s, 3H); MS (ESI), 401.2 [M + Na]+.

3-((2R,3R,4R,5R)-4-Hydroxy-5-(hydroxymethyl)-3-methoxytetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione: 12

Compound was described previously (13). Ceric ammonium nitrate (CAN) (39.48 g, 72.02 mmol) was added to a solution of compound 11 (10.9 g, 28.81 mmol) in MeCN (400 ml) followed by water (100 ml). The mixture was stirred at 60°C for 3 h. The reaction mixture was filtered and concentrated under reduced pressure to give a residue, which was purification by preparative high performance liquid chromatography (prep-HPLC,column: Welch Xtimate C18 250*70mm#10 μm; mobile phase A,10 mM NH4HCO3 in water and mobile phase B, acetonitrile; B%: 0–10%, 20 min) to yield compound 12 (3.6 g, 49%) as a white solid. 1H NMR (400 MHz, DMSO-d6) δ 11.17 (brs, 1H), 7.46 (d, J = 7.6 Hz, 1H), 6.10 (d, J = 3.1 Hz, 1H), 5.58 (d, J = 7.6 Hz, 1H), 4.86 (d, J = 6.9 Hz, 1H), 4.58 (t, J = 5.8 Hz, 1H), 4.24–4.16 (m, 2H), 3.66 (dt, J = 3.3, 6.3 Hz, 1H), 3.60 (ddd, J = 3.4, 5.3, 11.8 Hz, 1H), 3.44–3.36 (m, 1H), 3.31 (s, 3H); MS (ESI), 281.1 [M + Na]+.

3-((2R,3R,4R,5R)-5-((Bis(4-methoxyphenyl)(phenyl)methoxy)methyl)-4-hydroxy-3-methoxytetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione: 13

4,4′-Dimethoxytrityl chloride (DMT-Cl, 4.59 g, 13.55 mmol) was added to a solution of compound 12 (3.5 g, 13.55 mmol) in pyridine (50 ml). The mixture was stirred at 15°C for 6 h. The reaction mixture was filtered and concentrated under reduced pressure to give a residue, which was purified by flash chromatography over silica gel (petroleum ether:ethyl acetate = 10:1 to 0:1 to ethyl acetate:MeOH = 5:1), 5% triethanolamine (TEA) to yield compound 13 (6.3 g, 83%) as a white solid. 1H NMR (400 MHz, CDCl3) δ 9.36 (brs, 1H), 7.47 (d, J = 7.8 Hz, 2H), 7.35 (d, J = 8.6 Hz, 4H), 7.26–7.22 (m, 2H), 7.20–7.15 (m, 1H), 6.85–6.74 (m, 5H), 6.36 (d, J = 1.8 Hz, 1H), 5.62 (d, J = 7.8 Hz, 1H), 4.54–4.45 (m, 1H), 4.21 (dd, J = 1.9, 6.6 Hz, 1H), 3.93 (dt, J = 3.1, 7.1 Hz, 1H), 3.77 (s, 6H), 3.47 (s, 3H), 3.43 (dd, J = 3.0, 10.1 Hz, 1H), 3.31 (dd, J = 6.6, 10.1 Hz, 1H), 2.70 (d, J = 9.1 Hz, 1H);13C NMR (151 MHz, CDCl3) δ 162.51, 158.53, 152.27, 145.06, 139.26, 136.25, 136.23, 130.28, 130.26, 128.36, 127.87, 126.83, 113.19, 113.16, 102.28, 86.27, 85.89, 83.39, 81.51, 70.68, 64.51, 58.42, 55.31; HRMS (ESI/Q-TOF) m/z: [M + Na]+ Calculated for C31H32N2O8Na 583.2057; Found 583.2056.

3-((2R,3R,4R,5R)-5-((Bis(4-methoxyphenyl)(phenyl)methoxy)methyl)-3-methoxy-4-(((1S,3S,3aS)-3-((phenylsulfonyl)methyl)tetrahydro-1H,3H-pyrrolo[1,2-c][1,3,2]oxazaphosphol-1-yl)oxy)tetrahydrofuran-2-yl)pyrimidine-2,4(1H,3H)-dione: 14

To a solution of compound 13 (3.00 g, 5.35 mmol) in THF (18 ml) was added triethylamine (1.72 ml, 12.3 mmol) and (3S,3aS)-1-chloro-3-((phenylsulfonyl)methyl)tetrahydro-1H,3H-pyrrolo[1,2-c][1,3,2]oxazaphosphole 6 (0.90 M in THF, 9.51 ml, 8.56 mmol). The reaction mixture was stirred at room temperature for 2.5 h. TLC and LC–MS showed the reaction was complete. The reaction was quenched with water (48 μl) and anhydrous MgSO4 was added. The mixture was filtered through celite, and the filtrate was concentrated to yield the crude product which was purified by normal phase flash chromatography over silica gel eluting with 30–100% EtOAc in hexanes (each mobile phase contained 2.5% triethylamine) to yield 14 (3.02 g, 67%) as a white foam. 1H NMR (600 MHz, CDCl3) δ 8.99 (s, 1H), 7.88 (d, J = 7.8 Hz, 2H), 7.59 (t, J = 7.5 Hz, 1H), 7.51–7.44 (m, 4H), 7.34 (d, J = 8.6 Hz, 4H), 7.23 (t, J = 7.6 Hz, 2H), 7.16 (t, J = 7.3 Hz, 1H), 6.82–6.74 (m, 5H), 6.35 (d, J = 3.0 Hz, 1H), 5.57 (d, J = 7.7 Hz, 1H), 5.08 (q, J = 6.1 Hz, 1H), 4.87 (dt, J = 9.5, 6.8 Hz, 1H), 4.35 (dd, J = 6.2, 3.1 Hz, 1H), 4.09 (dq, J = 6.8, 3.3 Hz, 1H), 3.75 (s, 6H), 3.66 (dq, J = 11.6, 6.0 Hz, 1H), 3.43 (s, 3H), 3.41–3.31 (m, 4H), 3.19 (dd, J = 10.4, 6.0 Hz, 1H), 2.88 (qd, J = 9.7, 4.1 Hz, 1H), 1.80 (dtt, J = 12.1, 8.0, 3.5 Hz, 1H), 1.75–1.66 (m, 1H), 1.61–1.54 (m, 1H), 1.10–1.03 (m, 1H); 13C NMR (151 MHz, CDCl3) δ 162.79, 158.50, 152.18, 145.15, 139.65, 139.42, 136.40, 136.27, 134.05, 130.33, 129.31, 128.44, 128.35, 127.83, 126.78, 113.15, 113.13, 102.17, 86.21, 81.98 (d, J = 2.7 Hz, 1C), 80.91, 75.27 (d, J = 10.4 Hz, 1C), 71.17 (d, J = 3.3 Hz, 1C), 66.30 (d, J = 3.3 Hz, 1C), 63.81, 58.54 (d, J = 3.8 Hz, 1C), 58.38, 55.31, 46.16, 46.09 (d, J = 35.4 Hz, 1C), 27.53, 26.17 (d, J = 3.8 Hz, 1C); 31P NMR (243 MHz, CDCl3) δ 148.59; HRMS (ESI/Q-TOF) m/z: [M + Na]+ Calculated for C43H46N3O11PSNa 866.2489; Found 866.2508.

Oligonucleotide synthesis

Chemically modified stereorandom and stereopure oligonucleotides having chimeric PS/PO/PN backbones were prepared using methods similar to those described in recent publications (10,14–16). No changes were required for the N-3-uridine modification. Base deprotection of 8-oxo-adenosine required extended time in ammonium hydroxide solution to reach completion. Incorporation of triantennary GalNAc (Supplementary Figure S1) was achieved using previously reported methods (10). Oligonucleotides were purified by anion exchange chromatography with a 20 mM sodium hydroxide in water:acetonitrile (4:1) and 2.5 M sodium chloride, 20 mM sodium hydroxide in water:acetonitrile (4:1) mobile phase system. Desired fractions were pooled and desalted against water for injection via a G-25 Sephadex column. Mass spectrometry analysis was performed on a Novatia Oligo HTCS or Waters SQ Detector 2 system. Oligonucleotide concentration was determined by measuring absorbance at 260 nm on a Spectramax plate reader. Stereoselectivity is determined at the dimer synthesis stage, as described previously (14,15,17). The sequences, chemistries, and predicted and measured mass of all oligonucleotides can be found in Supplementary Tables S1–S4.

Animal work

Animal experiments were performed at Biomedical Research Models, Inc. dba Biomere (Worcester, MA) in compliance with an approved protocol under Biomere's Institutional Animal Care and Use Committee. Mice were on a 12-h light-dark cycle. Food (lab diet 5001) and water were available ad libitum. Housing rooms were maintained at 20–26°C and relative humidity was 30–70%. Male C57BL6-ADAR1-p110 mice (8 weeks old) were subcutaneously administered 10 mg/kg of each AIMer on days 0, 2 and 4. In-life assessments included body weight measurements and cage side observations by technician and veterinary staff. No clinical observations were noted and no moribund animals (body weight loss greater than 20%) were noted during study. Animals were euthanized on day 7 per testing facility standard operating procedure (briefly, animals will be euthanized by CO2 asphyxiation followed by exsanguination). Liver tissue was collected at day 7.

Humanized ADAR1-p110 mice and ADAR1-p110/NSG-PiZ mice

ADAR1-p110 mice were generated by Biocytogen (Waltham, MA). Human ADAR1-p110 Transcript-variant-4 cDNA was cloned into a Rosa26 targeting vector. Mice stably expressing ADAR1-p110 from the Rosa26 endogenous promoter were generated through zygotic injection of the Rosa26-ADAR1-P110 targeting vector, Rosa26-CRISPR-gRNA, and Cas9 mRNA in a C57BL/6 background strain.

Hemizygous C57BL/6-ADAR1-p110 mice were crossed with NSG-PiZ mice (Jackson Laboratory, Cat. No. 028 842), which express mutant human SERPINA1 (Glu342Lys mutation), to generate mice hemizygous for both the PiZ and ADAR1 alleles.

Primary hepatocyte extraction

C57BL/6-ADAR1-p110 mice or C57BL/6-ADAR1-p110/NSG-PiZ mice were used to obtain primary mouse hepatocytes expressing human ADAR-p110 or both human ADAR-p110 and SERPINA1-PiZ allele. Primary hepatocyte extraction was performed by Biomere (Worcester, MA). Gymnotic delivery of AIMers to hepatocytes and measurement of RNA editing were performed as described below. Biological duplicates of each condition were performed.

Tissue processing for oligonucleotide quantification

Animal tissues were dissected and flash-frozen in pre-weighed Eppendorf tubes. Final tissue weight was measured before initiation of each experiment. For lysis, lysis buffer (10 mM Tris, 100 mM NaCl, 5 mM EDTA, 0.5% nonidet P-40 supplemented with 2 mg/ml proteinase K) were added to the thawed tissues and homogenized at 4°C using Precellys (Bertin Instruments) until no tissue pieces could be observed. Roughly 30–50 μl of tissue lysates were saved in 96-well plates for oligonucleotide quantification by hybridization enzyme-linked immunosorbent assay (ELISA). Two biological replicates were performed for each condition.

Oligonucleotide quantification by hybridization ELISA

The following probes were utilized to selectively quantify the oligonucleotides by hybridization ELISA. Capture probe: /5AmMC12/ GTGCCGCAGTGGATC; Detection probe: T + T + T + AT + ATT + CTT + A + AT + C/3BioTEG/ (Integrated DNA Technologies, Coralville, IA). Maleic anhydride-activated 96-well plates (Pierce 15 110) were coated with 50 μl of capture probe at 500 nM in 2.5% NaHCO3 (Gibco, 25080-094) for 2 h at 37°C. The plate was then washed 3 times with phosphate buffered saline supplemented with 0.1% Tween-20 (PBST), blocked with 5% fat free milk-PBST at 37°C for 1 h. Payload oligonucleotide was serially diluted into matrix. This standard and the original samples were diluted with lysis buffer so that the oligonucleotide amount was less than 200 ng/ml in all samples. The diluted samples (20 μl) were mixed with 180 μl of 333 nM detection probe diluted in PBST and then denatured (65°C for 10 min, 95°C for 15 min, then a 4°C hold). Next, 50 μl of the denatured samples were added to blocked ELISA plates in duplicate and incubated overnight at 4°C. After 3 washes with PBST, 50 μl of streptavidin-AP (Southern Biotech, 7100-04) diluted 1:2000 in PBST was added per well and incubated at room temperature for 1 h. Following extensive washes with PBST, 100 μl of AttoPhos (Promega S1000) was added, incubated at room temperature in the dark for 10 min and read on a plate reader (Molecular Device, M5; excitation 435 nm, emission 555 nm). The amount of oligonucleotide in the samples was calculated according to the standard curve by 4-parameter regression analysis.

Endogenous RNA-editing assays

To evaluate RNA editing of endogenous transcripts under gymnotic or N-acetylgalactosamine (GalNAc)-mediated uptake, AIMers were added to the media at the desired concentration at the time primary mouse hepatocytes isolated from C57BL/6-ADAR1-p110 mice or C57BL/6-ADAR1-p110/NSG-PiZ mice were seeded (30 000-hepatocytes per well; collagen-coated 96-well plate). RNA was collected 48 h later.

RNA was harvested from cells using Promega SV 96 Total RNA Isolation System according to the manufacturer's protocol. RNA was eluted in a final volume of 50 μl. Eluted RNA was incubated at 95°C for 5 min and then immediately placed on ice. 9 μl of RNA was then used to generate cDNA using a High-Capacity cDNA Reverse Transcription Kit (Life Technologies, Cat. No. 4 388 950) in a 20 μl reaction according to manufacturer's protocol. 2 μl of cDNA was used to amplify the transcript of interest using Phusion High-Fidelity DNA Polymerase (Thermo Fisher, F530) and transcript-appropriate primers. Primers for PCR are shown in Supplementary Table S5. PCR products were purified using ExoSAP-IT Express (ThermoFisher, 75001.1) and analyzed by Sanger sequencing (Genewiz, Cambridge MA). Percentage of editing was calculated from Sanger sequencing traces using a custom workflow, modified from EditR (18). Briefly, the area under the curve for each position across each base (A, T, G or C) was directly extracted from the sequencing file using sangerseqR (19). The percentage of editing was calculated using the following equation: 100 × (area A/sum area A + T + C + G).

Statistical analyses

All statistical analyses for this manuscript were run in the R computing environment (v 3.6.0) (20) and the KNIME (Konstanz Information Miner) platform (v 4.3.2) (21). When the sample size was ≥3, assumptions of equal variance were tested using Levene's tests across all experimental factors and assumptions of normality were tested with Shapiro-Wilk tests of model residuals and per condition (R package rstatix, v 0.7.0.999; https://rpkgs.datanovia.com/rstatix/). The normality assumption was found to be violated if residuals and all conditions featured significant Shapiro–Wilk tests (P < 0.05). For experiments with a sample size of 2, unequal variance was assumed. For multi-factor comparisons of non-longitudinal data, type III 2-way or three-way analysis of variance (ANOVA) were initially run to investigate all possible interactions between factors (R package car, v 3.0–7) (22). If non-significant interactions were identified, type II ANOVAs featuring only main effects were run. In the event of unequal variance (Levene's test P< 0.05) or a sample size of 2 per group, Welch's 1-way ANOVA was run in place of typical one-way ANOVA, while two-way and three-way ANOVA were White-adjusted to allow for unequal variance. To compare groups in each experiment, Tukey HSD post hoc tests were run using the stats R package (20) for one-factor experiments where equal variance assumptions were found to hold, while two-tailed pairwise and reference-specific post hoc tests were extracted from the corresponding regression model via the multcomp R package for multi-factor experiments (v 1.4–13) (23). If significant interactions were determined via ANOVA, groups were compared per stratum, while treatment main effects were compared after adjusting for covariates in the event of non-significant interactions. Non-Tukey HSD post-hoc P values were Bonferroni-corrected for multiple hypotheses. For experiments featuring unequal variance, robust HC3 covariance estimation was implemented in post-hoc comparisons to allow heteroscedasticity (R package sandwich, v 2.5-1) (24–26). To compare data collected in two experiments, a Three-way ANOVA was run using construct, dose, and experiment as factors. To quantify the proportion of variance explained by each main effect and interaction on RNA editing level in the nearest neighbor experiment, ETA2 (η2) values were extracted from a full-interaction ANOVA model investigating the effects of concentration, orphan base, AIMer design, 3′-nearest neighbor, and 5′-nearest neighbor. Four-parameter log-logistic functions were utilized for fitting dose response curves and calculating and statistically comparing relative EC50, absolute EC50, and upper plateau (maximum) percent editing estimates through the R package drc (v 3.0–1) (27) with statistical significance determined by non-overlapping 95% confidence intervals. The delta method and the t-distribution were utilized to calculate asymptotics-based 95% confidence intervals.

Results

N3U in the orphan position increases predicted H bonding with ADAR

ADAR1 and ADAR2 possess a conserved, functionally equivalent residue responsible for A base-flipping (E1008 in ADAR1, E488 in ADAR2) (29) and substitution of either E1008 or E488 with glutamine leads to higher activity than wild-type ADAR1 or ADAR2, respectively (30). Enhanced H bonding between ADAR2 E488Q and the orphan site C is thought to drive hyperactivity in the E488Q ADAR2 mutant by promoting base flipping of the target A (8). In crystal structure studies, ADAR2 Q488 was observed to recognize the orphan base C by donating H bonds to the cytosine N3 and O2 atoms (9). Therefore, we hypothesized that we might induce similar interactions between the homologous residue in wild type ADAR2 E488/ADAR1 E1008 and the orphan base by incorporating a modified base in the orphan site that mimics this pattern of H bonding. If the modified base in the orphan site also maintained low base pairing with the target A, it could promote base flipping and thereby improve editing efficiency (4). We hypothesized that an N3-substituted uridine might satisfy these conditions, as the structure possesses N3H and O2 atoms, which could form H bonds with ADAR2 E488 / ADAR1 E1008, while forming only a single H bond with the target base A. To test this hypothesis, we used modeling based on the available ADAR2 crystal structures to examine whether a simple N3-substituted uridine, N3U, could support enhanced H bonding with wild type ADAR2 E488.

We generated models of the ADAR2 active site (PDB ID: 7KFN) (4) with orphan site nucleotides containing 2′-deoxyribose (2′-H) sugar and either C, U, N3U, or T bases to understand H bond interactions between the modified base at the orphan position and E488 (Figure 1A). Using the maestro pKa module, pKa values of each nucleobase with N1-Me for C, U and T and N3-Me for N3U were calculated and the reported experimental pKa values with N1-sugar pKa values for C, U and T are shown in parentheses. Compared to C, N3U shortens the length of one H bond contact with E488 and lowers the pKa of N–H at the N1 position. The closer contact might be explained by the different sugar pucker conformations observed for the two nucleosides (Supplementary Figure S2). The N3U nucleoside forms a sugar pucker that is between C2′-endo and C3′-endo conformation, whereas the C nucleoside forms a C3′-endo conformation, resulting in the N3U nucleoside having a larger dihedral angle of N-C1′–C2′–C3′ that may position N3U closer to E488. Overall, this suggests a stronger interaction between N3U and E488, which should facilitate more efficient base-flipping compared to C. Importantly, while U and T are also anticipated to have lower pKa's and stronger H bonding to E488 compared to C (Figure 1A), they form two H bonds with the edit site A, thus imposing an energetic cost to base flipping (Figure 1B). By contrast, N3U is predicted to form only one H bond with A (Figure 1B), supporting the hypothesis that N3U might promote more efficient base flipping than either C, U or T in the orphan base position. Of note, our modeling illustrates the N3U base in the anti conformation; if N3U must rotate from syn to anti before fitting into the active site, this would cause an energetic penalty to flipping out the target A and would be anticipated to negatively impact the benefit N3U confers on catalytic activity. However, ADAR1 and ADAR2 are double-stranded RNA binding enzymes (28), and N3U must adopt an anti conformation to form Watson–Crick base pairing with the target RNA. Since this must occur before the ADAR enzyme recognizes the duplex, we anticipate that N3U base rotation does not impact target A base flipping activity.

Figure 1. Structure-guided AIMer orphan site design. (A) Models generated from PDB structures of ADAR2 (PDB ID: 7KFN) with 2′-deoxy (2′-H) C, 2′-H U, 2′-H N3U, and 2′-H T demonstrate H bond interactions between the base at the orphan position and E488 in ADAR2 (purple). (B) Model of predicted base-pairing between the edit site A base and different orphan bases.

ADAR1 is widely expressed across tissues, thus the potential impact of N3U on ADAR1-mediated RNA editing is also of great therapeutic interest. Unfortunately, the lack of published high-resolution structures of ADAR1 prevents modeling of N3U in the ADAR1 catalytic site. However, ADAR1 possesses a conserved residue (E1008) that corresponds to ADAR2 E488 (29) and substitution of glutamate with glutamine (E1008Q) similarly leads to higher activity than wild-type ADAR1 (30). Thus, we hypothesize that N3U in the orphan position would lead to enhanced catalytic activity for both ADAR1 and ADAR2.

Collectively, our modeling suggests that N3U in the orphan base position would stabilize the base flipped conformation, thus increasing ADAR catalytic efficiency.

Synthesis of 2′-deoxy N-3-uridine, 5a, and L-PSM 2′-deoxy N-3-uridine amidite, 7a

To assess the impact of introducing N3U in the AIMer orphan site position, the corresponding N3U phosphoramidite monomer was needed. We developed a new synthetic method for 2′-deoxy N3U 5a and L-PSM 2′-deoxy N3U amidite 7a (Scheme 1). In brief, the reaction of 1-chloro-3,5-di(4-chlorbenzoyl)-2-deoxy-d-ribose 1a with 1-acetylpyrimidine-2,4(1H,3H)-dione 2a and NaH provided the mixture of alpha and beta isomers. The anomeric mixture was purified by column chromatography, followed by trituration with EtOAc and MeCN afforded the pure beta isomer 3a. Deprotection of 4-cholorbenzoyl group of the pure beta isomer 3a using NaOMe afforded 2′-deoxy N3U 4a in 93% yield and the structure was confirmed by X-ray crystallography (Figure 2, Supplementary Table S6, Supplementary Dataset S1). Protection of the 5′-OH of 2′-deoxy N3U 4a with DMTrCl gave 5′-DMTr protected 2′-deoxy N3U 5a. The stereopure amidite L-PSM 2′-deoxy N3U 7a was synthesized from the reaction of 5a with L-PSM-Cl 6 in good yield.

Scheme 1. Synthesis of L-PSM 2′-deoxy N3U amidite, 7a. Conditions:a, 2a, NaH, MeCN, 24%; b, NaOMe, MeOH, 93%; c, DMTrCl, pyridine, 76%; d, 6, TEA, THF, 88%.

Figure 2. X-ray crystallographic structure of 2′-deoxy N3U as a dimer complexed with ethanol. (A) absolute configuration and (B) structure model derived from X-ray crystallography.

Synthesis of 2′-OMe-N-3-uridine, 13, and L-PSM 2′-OMe-N-3-uridine amidite, 14

A new synthetic method for 2′-OMe N-3-uridine 13 and L-PSM 2′-OMe N-3-uridine 14 was developed (Scheme 2). The reaction of (2R,3R,4R,5S)-2-((benzoyloxy)methyl)-5-bromotetrahydrofuran-3,4-diyl dibenzoate 1b with 2a and NaH afforded a mixture of alpha and beta isomers, which was purified by flash chromatography over silica gel to afford pure beta isomer 8. Protection of amino group of N-3-uridine 8 using p-methoxybenzyl chloride (PMBCl) and deprotection of benzoyl group using NaOMe afforded compound 9 in good yield over two steps. Protection of 3′, 5′-dihydroxyl group of N-3-uridine 9 using 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane (TIPDSiCl2) and methylation of 2′-hydroxyl group using MeI gave compound 11 in good yield. Deprotection of silyl group using TBAF and PMB group using ceric ammonium nitrate (CAN) followed by protection of 5′-OH using DMTrCl afforded 2′-OMe N3U 13. L-PSM 2′-OMe N3U 14 amidites were synthesized via the reaction of 13 with L-PSMCl 6 in good yield.

Scheme 2. Synthesis of L-PSM 2′-OMe N3U amidite 14. Conditions: a, 2a, NaH, MeCN, 27%; b, PMBCl, K2CO3, DMF; c, NaOMe, MeOH, 92% over 2 steps; d, TIPDSiCl2, pyridine; e, NaH, MeI, DMF, 84% over 2 steps; f, TBAF, THF, 94%; g, CAN, MeCN and water, 49%; h, DMTrCl, pyridine, 83%; i, 6, TEA, THF, 67%.

N3U and the AIMer-D chemistry pattern improve editing efficiency across sequences

An acknowledged challenge of the ADAR field is the need for approaches that counter the natural nearest neighbor sequence biases of ADAR enzymes and enable efficient editing of challenging sequences (7,9,31,32). We hypothesized that if N3U in the orphan base position enhanced the catalytic efficiency of ADAR, it should enhance editing independent of nearest neighbor sequences. To test this hypothesis, we treated hepatocytes from mice expressing ADAR1-p110 with AIMers targeting mouse UDP-glucose pyrophosphorylase 2 (Ugp2) as an exemplary transcript. In the orphan position, N3U was compared to C for various nearest neighbor combinations. To explore the generality of the impact of N3U, the edit region combinations were applied to two sugar chemistry patterns: either our previously described segmented AIMer sugar pattern, referred to as AIMer-S (10), or another screening format reported herein possessing distributed sugar modifications, called AIMer-D (Figure 3A, Supplementary Table S1).

Figure 3. N3U and AIMer-D chemistry enhance editing efficiency in vitro. (A) Schematic of backbone configurations and 2′-sugar modifications for the AIMer-S and AIMer-D designs, including the orphan site (black arrow) and edit region (black dashed box), which includes the orphan site and 5′ and 3′ nearest neighbors. (B) Percentage of Ugp2 RNA editing shown with respect to AIMer (0.3 and 3 μM doses) with variable orphan base (C or N3U) and chemistry format (AIMer-S or AIMer-D) in primary mouse hepatocytes. Light blue bars, orphan base C; Dark blue bars, orphan base N3U. Data are presented as mean of n= 3, error bars represent SEM. (C) Violin plot illustrating the major effect of the orphan position base (C versus N3U) on editing at 0.3 and 3 μM doses. Lines connect complexes with identical 5′ and 3′ nearest neighbors, chemistry format, and dose. Horizontal lines in each violin plot represent 25%, 50% and 75% quartiles according to density estimates. Data are presented as mean of n= 3. (D) Violin plot illustrating the major effect of the chemistry format (AIMer-S vs. AIMer-D) on editing at 0.3 and 3 μM doses. Lines connect complexes with identical 5′- and 3′-nearest neighbors, orphan base, and dose. Horizontal lines in each violin plot represent 25%, 50% and 75% quartiles according to density estimates. Data are presented as mean of n= 3. For AIMer sequences and chemistries and statistical analyses see Supplementary Table S1, Supplementary Dataset S2.

A key feature of the AIMer-D format is that it possesses a greater number of 2′-F sugar modifications, compared to AIMer-S, which are distributed throughout the AIMer molecule. We introduced this design based on work in other oligonucleotide modalities, which have shown that incorporation of 2′-F ribose modifications favor a C3′-endo sugar conformation, which is more RNA-like than 2′-OMe, and provides more metabolic stability to the oligonucleotide compared to natural RNA (33).

The impact of both AIMer orphan site base modification (N3U vs C) and sugar modification pattern (AIMer-S versus AIMer-D) were examined for combinations of different 5′- and 3′-nearest neighbors to the target A (Figure 3B). AIMers combining both the AIMer-D chemistry pattern and N3U supported highly efficient editing for some nearest neighbor combinations (up to 89.2% editing; 5′-C, 3′-G, 3 μM dose) and, in some cases, striking improvements in editing efficiency compared to AIMers combining the AIMer-S chemistry pattern and C (Mean editing 5′-G, 3′-G, 3 μM dose: AIMer-S, C 4.0%; AIMer-D, N3U 87.9%). The orphan site base (N3U versus C) was a main effect that explained the highest proportion of variance (28.1%), with N3U conferring a higher mean percent RNA editing than C at both AIMer concentrations for all sequences tested, although the magnitude of increase varied (Figure 3C). The chemistry mask (AIMer-S versus AIMer-D) was the second most impactful main effect, explaining 24.9% of the variance. Similar to the general effect of N3U, AIMer-D conferred a higher mean percent RNA editing compared to the AIMer-S for most sequences tested (Figure 3D; detailed statistical analysis of sources of variance is provided in Supplementary Dataset S2). Together, the data show that the AIMer-D sugar chemistry mask and orphan site N3U broadly enhance editing efficiency across nearest neighbor sequences compared to AIMer-S and C, respectively, and that their effects appear to be largely additive.

N3U enhances editing efficiency in vitro and in vivo

We next evaluated the impact of N3U on RNA editing with N-acetylgalactosamine (GalNAc)-conjugated AIMers, and whether N3U is compatible with 2′-O-methyl (2′-OMe) sugar modification in the orphan site position (Supplementary Figure S1, Supplementary Table S2, Figure 4A). Conjugating GalNAc to oligonucleotides increases their uptake in hepatocytes and has been shown to improve oligonucleotide therapeutic potency in humans (34). It was previously shown that AIMers with 2′-H, C in the orphan site are taken up in hepatocytes and direct higher potency RNA editing compared to AIMers with 2′-OMe, C in the orphan site (10). To understand the compatibility of N3U with 2′-OMe, we initially compared editing in primary hepatocytes from ADAR1-p110 mice using AIMers directing editing of endogenous mouse Ugp2 mRNA. The AIMers varied only in their orphan site nucleotide, possessing either N3U, 2′-OMe (UGP2-471); N3U, 2′-H (UGP2-470); C, 2′-OMe (UGP2-467); or C, 2′-H (UGP2-469). Consistent with our findings using unconjugated AIMers, we observed a shift in editing activity with the N3U, 2′-H orphan site AIMer compared to the C, 2′-H orphan site AIMer (Figure 4B, UGP2-470 absolute EC50 = 0.014 μM; UGP2-469 plateaued with <50% of Ugp2 edited). Importantly, 2′-OMe did not significantly impact editing when N3U was the orphan base (UGP-470, absolute EC50= 0.014 μM [95% CI: 0.006–0.021] vs. UGP-471, absolute EC50 = 0.010 μM [95% CI: 0.008–0.012]), whereas 2′-OMe reduced maximum editing when combined with C (Figure 4B, UGP2-467 maximum 15.6% [95% CI: 11.0–20.2] editing vs. UGP2-469 maximum 28.3% [95% CI: 27.3–29.2] editing). For associated statistical analyses, see Supplementary Dataset S2. This suggests that, in vitro, AIMers with N3U as the orphan base support more efficient editing than those with C, and N3U and 2′-OMe are compatible in the orphan site.

Figure 4. N3U is compatible with 2′-OMe modification in the orphan site and enhances editing in vivo. (A) Diagrams of GalNAc-conjugated AIMers illustrating chemical modifications for each Orphan site is indicated by a black box. (B) Dose-response curve (thick lines) showing percentage of endogenous Ugp2 RNA editing with respect to GalNAc-AIMer concentration in primary mouse hepatocytes isolated from hemizygous ADAR1-p110 mice. Data shown are the mean with SEM, n = 3 for each condition; 95% confidence intervals indicated with dashed lines. (C) Percent editing of Ugp2 mRNA in vivo in 8-week-old male ADAR1-p110 hemizygous knock-in mice dosed with PBS (black) or GalNAc-AIMer (10 mg/kg) subcutaneously at days 0, 2 and 4. Liver tissue was harvested on day 7 after first dose. Data are shown as mean with SEM, n = 5. Stats: One-way ANOVA followed by Tukey HSD post hoc tests. * P < 0.01; **** P < 0.0001; ns, not significant. (D) Concentration of AIMer from liver of mice in panel C at day 7. Data shown as mean with SEM. Stats: One-way ANOVA followed by Tukey HSD post hoc tests vs. ACTB-816. ns, not significant. All pairwise comparisons were computed but only a subset is shown; see Supplementary Dataset S2 for complete statistical analysis. For AIMer sequence and chemistry information, see Supplementary Table S2.

Next, we wanted to investigate the impact of N3U on RNA editing in mouse hepatocytes in vivo (Figure 4C). Consistent with in vitro data, AIMers with either orphan N3U, 2′-H (UGP2-470) or N3U, 2′-OMe (UGP2-471) gave robust editing compared to AIMers with orphan C, 2′-H (UGP2-469), leading to >70% maximal editing in vivo (Mean editing: UGP2-469 14.6%; UGP2-470 56.1%; UGP2-471 70.8%). C, 2′-H at the orphan position (UGP2-469) conferred a modest but statistically significant increase in mean Ugp2 mRNA editing compared to the non-targeting control (mean editing: UGP2-469 14.6%; ACTB-816 4.3%, P < 0.05), which was not observed with the 2′-OMe sugar modification (mean editing UGP2-467 8.8%, P= ns compared to ACTB-816). Importantly, while AIMers with N3U supported higher maximal editing, they did not result in detectable bystander editing (Supplementary Figure S3). To understand if differences in tissue accumulation contributed to observed editing efficiency, liver tissue samples from AIMer-treated mice were analyzed for AIMer accumulation (Figure 4D). All AIMers showed similar concentrations in liver tissue, and tissue concentrations were generally consistent compared to other studies with a similar dosing schedule (10). These results confirmed the superiority of N3U over C in the orphan position in vitro and in vivo, and the application of N3U enables the use of 2′-OMe in the orphan position.

N3U enhances editing of a disease-relevant target, SERPINA1

The scope of potential therapeutic targets for ADAR-based RNA editing is limited by the nearest neighbor sequence bias of ADAR enzymes. One example is the c. 1096G > A (p.Glu366Lys) variant in the SERPINA1 gene, commonly referred to as the Z allele, which causes α1-antitrypsin deficiency (35). The 1096G > A variant could potentially be reverted (to the M allele) through A-to-I editing; however, the less-preferred nearest neighbor context of 1096A (5′-C, 3′-A) hinders editing efficiency. To confirm that incorporation of N3U could improve editing for a disease-relevant target, we designed AIMers to edit the SERPINA1 mRNA encoded by the Z allele, which harbors an E366K missense mutation that causes α1-antitrypsin deficiency in humans, as a benchmark (Supplementary Table S3). The sequence used for these AIMers is based on a SERPINA1-targeting AIMers that we previously described, which supported RNA editing in primary mouse hepatocytes (10). We evaluated these AIMers in primary hepatocytes from a transgenic mouse model expressing both the human SERPINA1 Z allele and ADAR1-p110. We initially sought to understand the impact of N3U, separately from the benefit of the AIMer-D pattern, by comparing the impact of different base modifications in AIMers with the AIMer-S pattern. SERPINA1-763, an AIMer with the AIMer-S sugar chemistry pattern, N3U, 2′-H, significantly increased SERPINA1 editing compared to SERPINA1-505, with AIMer-S, C, 2′-H (Figure 5A, mean editing at 3 μM dose: SERPINA1-763 24.0%; SERPINA1-505 11.3%, P < 0.01 when adjusting for both doses). By comparison, SERPINA1-563, incorporating the base analog 8-oxo-adenosine, 2′-H in the AIMer orphan site, did not significantly enhance SERPINA1 editing compared to AIMers with orphan site C, 2′-H (mean editing at 3 μM dose: SERPINA1-563 12.2%; SERPINA1-505, 11.3%; P = ns when adjusting for both doses), consistent with our previously published results using a different AIMer design (10).

Figure 5. AIMers with orphan site N3U modification support editing of the SERPINA1 transcript. (A) Percentage of SERPINA1 Z mRNA editing in primary mouse hepatocytes (hemizygous for SERPINA1 Z and ADAR1-p110), treated gymnotically with AIMer-S format AIMers (0.3 or 3 μM) possessing the indicated base in the orphan site (C, 8-oxo-adenosine, or N3U). Orphan site indicated with black dashed box. Data are presented as mean with SEM; n = 2. Stats: White-adjusted Two-way ANOVA with post-hoc comparison of construct main effects to SERPINA1-505 allowing heteroscedasticity. ** P < 0.01, ns, not significant. (B) Percentage of SERPINA1 RNA editing shown with respect to AIMer (0.03, 0.3 or 3 μM) with the indicated sugar and backbone modifications outside of the edit region. Data are presented as mean with SEM; n ≥ 4. Stats: White-adjusted three-way ANOVA with post-hoc comparison to SERPINA1-822 of construct effect per dose (controlling for experiment) while allowing heteroscedasticity. * P < 0.05, **** P < 0.0001, ns not significant. (C) Percentage of SERPINA1 RNA editing shown with respect to AIMer (0.03 or 0.3 μM) with the indicated sugar and backbone modifications in the edit region (black dashed box). Data are presented as mean with SEM; n = 2. Stats: White-adjusted two-way ANOVA followed by post hoc test comparisons of construct main effects per dose to SERPINA1-1769 allowing heteroscedasticity. ** P < 0.01, *** P < 0.001, **** P < 0.0001, ns, not significant. (D) Percentage SERPINA1 RNA editing shown with respect to AIMer (0.03 or 0.3 μM) with the indicated base and sugar modification combinations at the orphan site (black dashed box). Data are presented as mean with SEM; n = 2. Stats: White-adjusted Two-way ANOVA followed by post hoc test comparisons to SERPINA1-1113 of construct main effects allowing heteroscedasticity. * P < 0.05, ** P < 0.01, ns not significant. Dose: △, 0.03 μM; ○, 0.3 μM; □, 3 μM. All pairwise comparisons were computed but only a subset is shown; see Supplementary Dataset S2 for complete statistical analysis. For AIMer sequence and chemistry information, see Supplementary Table S3.

Next, we employed structure activity relationship (SAR) studies to further improve SERPINA1 editing efficiency of AIMers with the AIMer-D chemistry design and N3U in the orphan position. In other oligonucleotide modalities, judiciously placed 2′-OMe sugar modifications, as well as PN and phosphodiester (PO) backbone linkages, can improve oligonucleotide stability, potency, and/or durability in a modality-dependent fashion (14–16). Moreover, in our prior RNA editing work, we found that AIMer PN linkages can have a position-dependent impact on RNA editing efficiency in a luciferase reporter system (10). Therefore, we investigated whether integrating 2′-OMe, PN, and PO outside of the edit region improved editing efficiency in SERPINA1-targeting AIMers with N3U, 2′-H in the orphan site (Supplementary Table S3, Figure 5B).

Compared to the baseline AIMer-D design (SERPINA1-822), AIMers with an additional modest number of PO and PN linkages and 2′-OMe sugar modifications outside of the edit region (SERPINA1-979, SERPINA1-989) supported significantly improved SERPINA1 editing efficiency (mean editing for 3 μM dose: SERPINA1-822 40.1%; SERPINA1-979 55.0%, P < 0.05; SERPINA1-989 61.9%, P < 0.0001).

Building on the AIMer that supported the most efficient editing (SERPINA1-989), we next investigated the impact of incorporating different backbone linkage chemistries within the edit region and the compatibility of N3U with 2′-OMe in the orphan site (Supplementary Table S3, Figure 5C, D). In a structural study of human ADAR2 bound to a double stranded RNA substrate, it was reported that the majority of the contacts between ADAR2 and the complementary strand were mediated by the phosphodiester backbone in the edit region, therefore we hypothesized that backbone modifications in the edit region might impact RNA editing efficiency (9). Compared to SERPINA1-1769 (edit region PO linkages), SERPINA1-1783 (Sp PS), SERPINA1-989 (Sp PS and Sp PN) and SERPINA-1061 (Sp PS and Rp PN) improved editing efficiency (mean editing at 0.3 μM dose: SERPINA1-1769 18.0%; SERPINA1-1783 46.7%, P < 0.01; SERPINA1-989 50.5%, P < 0.0001; SERPINA1-1061 47.0%, P < 0.0001) (Figure 5C). In the same experiment, the orphan base N3U (SERPINA1-989) supported improved editing efficiency over C (SERPINA1-1113) even when N3U was combined with 2′-OMe sugar modification in the orphan site (SERPINA1-1642) (Figure 5D, mean editing at 0.3 μM dose: SERPINA1-1113 38.6%; SERPINA1-989 50.5%; SERPINA1-1642 48.6%). These data are consistent with results seen in the baseline AIMer-D design, showing that N3U is compatible with 2′-OMe in the orphan site in multiple AIMer chemistry contexts.

These results show that AIMers with N3U in the orphan base exhibit improved editing efficiency over those with C across multiple editing targets, and that additional SAR analyses may further enhance RNA editing efficiency.

N3U supports chemical flexibility in the AIMer orphan site

We also explored the chemical flexibility of the orphan position in N3U-based AIMers. Given that N3U supported comparable editing with either 2′-H or 2′-OMe in the orphan site, we investigated whether N3U might support similar levels of SERPINA1 editing when combined with other common 2′ sugar modifications in AIMers with identical base sequences (Figure 6A, Supplementary Table S4). Data presented in Figure 6A come from the same experiment presented in Figures 5C, D.

Figure 6. AIMers with orphan site N3U modification support RNA editing for diverse edit region sequences and chemistries. (A) Percentage SERPINA1 mRNA editing shown with respect to AIMer (0.03 or 0.3 μM) with the indicated sugar modification in the orphan site (black dashed box) in primary mouse hepatocytes (hemizygous for SERPINA1 Z and ADAR1-p110). Chemical structures for the evaluated sugar modifications are shown. AIMer sequence: 5′-CCCAGCAGCUUCAGUCCCUUUCT[N3U][I]UCGAU-3′. Data presented as mean with SEM; n = 2. Stats: White-adjusted Two-way ANOVAs followed by pairwise post hoc comparisons of main effects of constructs to SERPINA1-989 allowing heteroscedasticity. ** P < 0.01, ns not significant. (B) Percentage of SERPINA1 mRNA editing shown with respect to AIMer (0.03 or 0.3 μM) with the indicated sugar modifications at the orphan site (black dashed box). AIMer sequence: 5′-CCCAGCAGCUUCAGUCCCUUUCT[N3U][I]UCGAU-3′. Data presented as mean with SEM; n = 2. Stats: White-adjusted Two-way ANOVAs followed by comparison of main effects of constructs per dose to SERPINA1-989 allowing heteroscedasticity. * P < 0.05; ** P < 0.01; *** P < 0.001; **** P < 0.0001; ns, not significant. (C) Percentage of SERPINA1 mRNA editing shown with respect to AIMer (0.03 or 0.3 μM) with the indicated N3U analogs in the orphan base position (black dashed box). Data presented as mean with SEM; n= 2. Stats: White-adjusted Two-way ANOVAs followed by pairwise post hoc comparisons of main effects of constructs to SERPINA1-989 allowing heteroscedasticity. (D) Percentage of SERPINA1 RNA editing shown with respect to AIMer (0.03 or 0.3 μM) with the indicated bases in the orphan site and 3′-nearest neighbor position (black dashed box). Data presented as mean with SEM; n = 2. Stats: White-adjusted two-way ANOVAs followed by pairwise post hoc test comparisons of main effects of constructs allowing heteroscedasticity. * P < 0.05; *** P < 0.001; ns, not significant. All pairwise comparisons were computed but only a subset is shown; see Supplementary Dataset S2 for complete statistical analysis. Dose: △, 0.03 μM; ○, 0.3 μM. For AIMer sequence and chemistry information, see Supplementary Table S4. 2′-FANA: 2′- fluoro-arabino; 2′-MOE: 2′-methoxyethyl; S/R-GNA: S/R glycol nucleic acid; N3T: N-3-thymidine.

AIMers with arabinose (SERPINA1-1346) and 2′- fluoro-arabino (2′-FANA) (SERPINA1-1749) preserved editing activity compared to 2′-H (SERPINA1-989), with only the bulky 2′-O-methoxyethyl (2′-MOE) (SERPINA1-1750) leading to slightly reduced editing (Mean editing at 0.3 μM dose: SERPINA1-989 50.5%, P = ns; SERPINA1-1346 48.9%, P = ns; SERPINA1-1749 42.7%, P = ns; SERPINA1-1750 35.8%, P < 0.01 when adjusting for dose).

Since AIMers with N3U and common 2′ sugar modifications were generally efficient editors, we wanted to explore whether AIMers with N3U and additional sugar modifications in the orphan site could also support editing (Figure 6B). Of the sugar modifications tested (ribose, 3′-hydroxyl, (S)-glycol nucleic acid, (R)-glycol nucleic acid), all resulted in >50% mean editing at the highest dose tested (mean editing at 0.3 μM dose: SERPINA1-989 77.4%; SERPINA1-1637 76.7%; SERPINA1-1638 53.1%; SERPINA1-1649 70.2%; SERPINA1-1650 53.86%). Overall, these data suggest that N3U generally supports editing in the context of diverse sugar modifications, with limited impact on editing efficiency.

In the same experiment, we investigated the impact of various substitutions at the 5-position of U to see whether N3U analogs with altered electron density (as shown by the calculated pKa values, Supplementary Table S7), such as N-3-thymidine (N3T), 5-ethyl N3U, 5-fluoro-N3U, 5-propynyl N3U, might support similar or improved editing efficiency (Supplementary Table S4, Figure 6C). AIMers containing the N3U analogs in the orphan base supported SERPINA1 editing comparably to those with N3U (mean editing at 0.3 μM: SERPINA1-989 77.4%; SERPINA1-1640 75.3%; SERPINA1-1643 76.0%; SERPINA-1644 79.2%, and SERPINA1-1647 76.2%, P = ns). This suggests that N3U may represent a family of modified bases that can support highly efficient RNA editing by AIMers, lending greater chemical flexibility to AIMer design.

Finally, we wanted to understand how N3U at the orphan base would influence the impact of incorporating a less-preferred sequence context. Previously, for a SERPINA1 AIMer, incorporating I in the position directly 3′ to the orphan site supported higher editing efficiency compared to AIMers with G immediately 3′ to the orphan position (10). We wanted to know how N3U would impact the inclusion of a less-preferred base (G) immediately 3′ to the orphan base position (Supplementary Table S4, Figure 6D). Data presented in Figure 6D come from the same experiment presented in Figures 5C,D and 6A. Consistent with our previous results (10), AIMers with a G base directly 3′ to the orphan position (SERPINA1-1760) directed reduced editing compared to those with I (SERPINA1-1113), when C was the orphan base (Mean editing at 0.3 μM: SERPINA1-1760 28.9%; SERPINA1-1113 38.6%; P <0.05). By contrast, when N3U was in the orphan position, AIMers with either 3′ G (SERPINA1-1655) or 3′ I (SERPINA1-989) supported similar editing (Mean editing at 0.3 μM: SERPINA1-1655 51.7%; SERPINA1-989 50.47%; P = ns). Thus, N3U in the orphan position increased editing activity in the context of a G immediately 3′ to the orphan base. These results might be at least partially explained by the observation that duplexes of edited mRNA and the 5′-[N3U]G-3′ AIMer (SERPINA1-1655) have a reduced melting temperature (Tm, 70.57°C) compared to duplexes with 5′-CG-3′ (SERPINA1-1760, 72.77°C, Supplementary Table S8). Taken together, these data suggest that using N3U or its analogs in the orphan base position may add chemical flexibility to the range of sugar and base modifications that can be incorporated in and around the orphan site.

Discussion

ADAR-mediated RNA editing is an emerging therapeutic modality. The purpose of this study was to improve AIMer-based RNA editing efficiency, building on our previous work (10), which demonstrated that control over backbone modifications could support RNA editing with AIMers. To our knowledge, this is the first time that the impact of N3U has been explored in RNA editing. To enable these investigations, we needed to develop methods to synthesize and incorporate N3U in AIMers. We developed novel, scalable synthetic methods to manufacture N3U and its analogs, including amidites for incorporation into synthetic oligonucleotides of any kind. With our methods, N3U is simpler to incorporate into oligonucleotides than 6-amino-5-nitro3-(1′-β-d-2′-deoxyribofuranosyl)-2(1H)-pyridone (also referred to as 2′-deoxy Benner's Base Z, or 2′-deoxy Z), which has previously been used in the orphan position to improve ADAR-mediated RNA editing efficiency in vitro (4).

N3U represents a family of modified bases that support efficient RNA editing

We identify a class of modified bases (based on N3U) that enhance A-to-I RNA editing potency when deployed in the orphan base position. To our knowledge, the N3U base has not been explored in the context of oligonucleotides and we hypothesized that this modification could beneficially impact ADAR-based RNA editing. The N3U base was selected because it appears to create a H-bonding pattern that mimics the interaction between the typical orphan base C and the Q488 residue of the hyperactive E488Q ADAR2 mutant. Previous work suggested that the E488Q H-bonding pattern supported hyperactivity by promoting flipping of the target base into the enzyme's active site. Here, we demonstrate that the incorporation of N3U increased editing efficiency across many edit region sequences and for multiple target mRNAs. This is consistent with previous in vitro data showing that 2′-deoxy Z, another modified base that is proposed to mimic the E488Q H-bonding pattern, also supports more efficient editing (4). Our data support the hypothesis that N3U in the orphan position may increase the catalytic efficiency of ADAR-mediated A-to-I editing, perhaps by stabilizing the base-flipped conformation, which is a major overall driver of editing efficiency. Importantly, there was no evidence that N3U impacted AIMer uptake or tissue exposure in vivo in liver (Figure 4D), consistent with the hypothesis that N3U impacts editing efficiency by enhancing the catalytic activity of ADAR. Additionally, our observation that N3U reduces the melting temperature of mRNA-AIMer duplexes post-editing suggests the possibility that N3U may contribute to editing efficiency by increasing enzymatic turnover. Future studies will be needed to confirm and characterize the proposed benefit of N3U for base flipping efficiency.

One of the biggest challenges for advancing RNA editing as a therapeutic modality is the nearest neighbor bias of ADAR enzymes. ADAR enzyme bias for specific 5′- and 3′-nearest neighbor sequences is thought to reflect the influence of these sites on base flipping (8,9,36). Our data suggests that the N3U base modification can partially address this challenge. While differences in editing efficiencies were still observed for N3U-containing AIMers with the AIMer-D chemistry format, all nearest neighbor combinations saw improved editing with N3U, including difficult-to-edit combinations. In one case, mean editing increased from 2% with orphan C to >82% with orphan N3U using AIMers with otherwise identical sequences and chemistry formats. Enhanced editing efficiency across a broad range of nearest neighbor sequences can help to expand the overall target space for A-to-I editing via endogenous ADAR enzymes.

Interestingly, editing was not negatively impacted when uracil 5-substituted analogs (i.e. N3U analogs) with altered electron density, as shown by the calculated pKa values (Supplementary Table S7), were incorporated in the orphan position. Thus, N3U analogs may offer additional flexibility in AIMer design.

In addition to the gains in potency and sequence flexibility in the edit region, N3U supports chemical flexibility in the orphan base position. In therapeutic oligonucleotides various 2′ modifications have been used to provide nuclease stability to improve pharmacological properties. We demonstrate that N3U supports efficient editing when combined with a wide variety of 2′-sugar modifications, including 2′-OMe. In contrast, orphan site C may be less flexible in some contexts, as Ugp2-targeting AIMers with orphan site C, 2′-OMe, supported lower editing compared to AIMers with C, 2′-H, in mouse hepatocytes both in vitro and in vivo. In mice, these AIMers showed no differences in PK, suggesting that the orphan site sugar might influence editing through impacting target engagement or impacting base flipping efficiency. However, for the SERPINA1-targeting AIMers there was no significant difference in editing efficiency between AIMers with orphan site C, 2′-H and those with C, 2′-OMe, and substituting N3U, 2′-OMe has a smaller benefit than was seen for Ugp2-targeting AIMers. This suggests that the detrimental impact of orphan site 2′-OMe may depend not only on the orphan base (N3U or C), but also the sequence, sugar, and backbone context of the entire AIMer, which may all contribute to target engagement and base flipping efficiency.

Still, that N3U supports high-potency editing when combined with multiple sugar modifications suggests potential opportunities to optimize for other pharmacological properties without sacrificing potency, which is just one of several features that impact the development of oligonucleotides as therapeutics.

AIMer-D is a useful format for target-specific screening and optimization

The benefits of the AIMer-D design appear to derive, at least partially, from a separate mechanism than N3U, as efficiency gains observed with the two modifications (base and backbone) were largely additive. We speculate that the design might increase the stability and/or uptake of the AIMer, and/or enhance AIMer engagement with RNA as observed previously with similar constructs for exon skipping oligonucleotides (15). In particular, the 2′-F sugar modification is known to be more like RNA than 2′-OMe, and it also stabilizes oligonucleotides compared to natural RNA through nuclease resistance (33), so the inclusion of 2′-F throughout the full AIMer may facilitate interactions with the target mRNA while increasing AIMer stability. Teasing out the mechanism(s) by which the AIMer-D design enhances editing efficiency requires exploration in future studies.

While the AIMer-D design broadly enhances editing efficiency across many sequences, the specific pattern of sugar and backbone modifications that maximize editing efficiency for each target may depend on the oligonucleotide sequence. We show that the AIMer-D design supports efficient editing of both Ugp2 and the exemplary target SERPINA1, however for SERPINA1 AIMers we found that additional changes to the sugar and backbone modification pattern can further improve editing efficiency. The introduction of two additional PN linkages in the AIMer improved editing compared to AIMers with PN linkages confined to the termini. Similarly, the introduction of Sp PS in the edit region, alone or in combination with either Rp PN or Sp PN linkages, increased editing compared to PO linkages. Optimization of 2′ modifications outside of the edit region further enhanced editing efficiency. Given these observations, it is likely that sequence also impacts the optimal modification pattern. Thus, we propose that the AIMer-D design could serve as a consistent launch point that could be further optimized to generate AIMers tailored to a specific transcript to maximize editing efficiency while preserving other essential pharmacological properties.

Conclusion

In summary, we have described a scalable synthetic method to manufacture novel N3U and its analogs, including amidites for incorporation into synthetic oligonucleotides using standard solid phase synthesis. We demonstrate that N3U can improve RNA editing when positioned as the orphan base of an AIMer. The AIMer-D sugar and backbone modification format also provided benefits to editing efficiency, suggesting the AIMer-D format may be a useful screening format for additional optimization. Combining N3U with an AIMer-D design enabled maximal editing of over 85% in mouse hepatocytes in vitro and ∼70% in mouse liver in vivo. The impact of N3U on RNA editing was observed across multiple targets, with a variety of nearest neighbor sequences, indicating that it can overcome, at least in part, the natural sequence bias of ADAR enzymes. However, the magnitude of the benefit of N3U varies, and is influenced by the chemical and sequence context of the AIMer and target. Overall, the versatility of N3U and its analogs creates helpful flexibility for AIMer design, broadening the avenues for optimizing other important pharmacological properties while preserving potency. Thus, the application of N3U to RNA editing greatly advances the therapeutic potential of this emerging modality.

Supplementary Material

gkae681_Supplemental_Files

Acknowledgements

The authors thank Milinda Samaraweera for setting up the structure modeling analysis, Khoa Luu for providing thermal denaturing data (Tm), Brooke Koshel for providing HRMS data and the following colleagues for supplying oligonucleotides used in this study: Jason Dufresne, Frank Favaloro, Gina Lein, Amber Lindsey, and Richard Looby. JetPub Scientific Communications LLC, supported by Wave Life Sciences, and Nicole Neuman, employee of Wave Life Sciences, assisted the authors in the preparation of this manuscript.

Data availability

Source data for Figures 3B–D, 4B–D, 5A–D and 6A–D are provided with the paper.

Atomic coordinates and structure factors for the reported crystal structure has been deposited with the CCDC with the refcode 2355432.

Supplementary data

Supplementary Data are available at NAR Online.

Funding

Wave Life Sciences. Funding for open access charge: Wave Life Sciences.

Conflict of interest statement. All authors were employees of Wave Life Sciences during completion of this work. P.M., M.S., C.S., G.L., D.B., J.G., A.H., N.I., J.K., P.K., A.L., S.S., H.Y. and C.V. have patent applications (WO 2018/237 194, WO 2015107425, and WO 2020/191 252); G.L., P.M., C.S., I.H., C.V., P.K., N.I., S.S. and A.L. have a patent application (WO 2023/201 095); and G.L., H.Y. and P.K. have a patent application related to this work.

Notes

Present address: Tom Pu, Northeastern University, Boston, MA, USA.
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References

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