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

39162233
10.1093/nar/gkae711
gkae711
AcademicSubjects/SCI00010
Molecular and Structural Biology
Efficient circularization of protein-encoding RNAs via a novel cis-splicing system
https://orcid.org/0000-0003-0204-0236
Qi Shaojun Department of mRNA Sciences, Suzhou Abogen Biosciences Co., Ltd., Suzhou 215123, China

Wang Huiming Department of mRNA Sciences, Suzhou Abogen Biosciences Co., Ltd., Suzhou 215123, China

Liu Guopeng Department of mRNA Sciences, Suzhou Abogen Biosciences Co., Ltd., Suzhou 215123, China

Qin Qianshan Department of mRNA Sciences, Suzhou Abogen Biosciences Co., Ltd., Suzhou 215123, China

https://orcid.org/0000-0003-4532-647X
Gao Peng Department of mRNA Sciences, Suzhou Abogen Biosciences Co., Ltd., Suzhou 215123, China

Ying Bo Department of mRNA Sciences, Suzhou Abogen Biosciences Co., Ltd., Suzhou 215123, China

To whom correspondence should be addressed. Tel: +86 15716135948; Fax: +86 (0512)69150735; Email: bo.ying@abogenbio.com
Correspondence may also be addressed to Peng Gao. Tel: +86 17712624683; Email: peng.gao@abogenbio.com
23 9 2024
20 8 2024
20 8 2024
52 17 1040010415
16 8 2024
24 7 2024
12 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 reprints@oup.com for reprints and translation rights for reprints. All other permissions can be obtained through our RightsLink service via the Permissions link on the article page on our site-for further information please contact journals.permissions@oup.com

Abstract

Circular RNAs (circRNAs) have emerged as a promising alternative to linear mRNA, owing to their unique properties and potential therapeutic applications, driving the development of novel approaches for their production. This study introduces a cis-splicing system that efficiently produces circRNAs by incorporating a ribozyme core at one end of the precursor, thereby eliminating the need for additional spacer elements between the ribozyme and the gene of interest (GOI). In this cis-splicing system, sequences resembling homologous arms at both ends of the precursor are crucial for forming the P9.0 duplex, which in turn facilitates effective self-splicing and circularization. We demonstrate that the precise recognition of the second transesterification site depends more on the structural characteristics of P9.0 adjacent to the ωG position than on the nucleotide composition of the P9.0-ωG itself. Further optimization of structural elements, like P10 and P1-ex, significantly improves circularization efficiency. The circRNAs generated through the cis-splicing system exhibit prolonged protein expression and minimal activation of the innate immune response. This study provides a comprehensive exploration of circRNA generation via a novel strategy and offers valuable insights into the structural engineering of RNA, paving the way for future advancements in circRNA-based applications.

Graphical Abstract

Graphical Abstract

Suzhou Abogen Biosciences Co., Ltd
==== Body
pmcIntroduction

Messenger RNA (mRNA) has emerged as a promising tool in therapeutics, particularly highlighted by the success of mRNA vaccines in combating infectious diseases like COVID-19 (1–4). In vitro transcribed (IVT) mRNAs offer a versatile platform for rapid and customizable protein expression, showcasing their potential for various therapeutic applications. However, challenges such as mRNA stability, unwanted immunogenicity and limited expression persistence in vivo have spurred the exploration of alternative synthetic RNA molecules for therapeutic purposes (5–7).

Circular RNA (circRNA) represents a unique class of single-stranded RNA molecules characterized by a covalently closed loop structure. Naturally occurring circRNAs are generally formed through a non-canonical splicing process known as ‘back-splicing’ (8,9). Unlike linear mRNAs, circRNAs do not require a 5′-cap or 3′-poly(A) tail for stability, and their closed-loop structure confers resistance to exonuclease-mediated degradation, resulting in an extended half-life compared to linear mRNA counterparts (10–12).

While circRNAs are predominantly noncoding, recent studies have demonstrated the potential for some circRNAs to undergo translation into proteins (13,14). Synthetic circRNAs with cap-independent translation elements, such as internal ribosome entry sites (IRES)(15,16) or N6-methyladenosine (m6A) modifications (17), have been shown to facilitate protein translation (18,19), even with tissue or cell-type specificity (20). The delivery of circRNAs via lipid nanoparticles (LNPs) enables sustained in vivo expression, offering advantages over linear mRNAs (21). Moreover, circRNAs demonstrate reduced immunogenicity and prolonged translation duration, making them attractive candidates for therapeutic interventions (18,22).

Two primary methods for in vitro circRNA preparation are Enzyme ligation-mediated and ribozyme-mediated circularization (8,23). The former method, which utilizes enzymes such as T4 DNA ligase and T4 RNA ligases, often requires a complementary splint to bring RNA ends together for catalysis (24). However, this approach can be inefficient for large RNA molecules and may lead to the formation of undesired intermolecular end-joining byproducts, necessitating complex optimization strategies (23,24). Alternatively, ribozyme-mediated circularization, such as the group I intron, offers a promising avenue for circRNA synthesis (16). Group I introns catalyze self-splicing reactions independently of spliceosome assistance through a process known as cis-splicing (25,26) (Supplementary Figure S1A). The 5′ splice site, as marked by a conserved u-G pair, undergoes nucleophilic attack by the 3′-OH group of exogenous GTP bound at the G-binding site of the intron. Subsequently, a conformational change displaces the guanosine from the G-binding site by the 3′-terminal omega G (ωG) that signifies the 3′ splice site. The 3′-OH group of the terminal residue of the 5′ exon then participates in a reaction that mirrors the initial step. Ultimately, the 5′ and 3′ exons are joined, releasing the intron (25).

The structural elements of group I introns (25,27,28), such as duplexes P1 to P8, play a crucial role in forming the scaffold (P4-P6) and catalytic core (P3, P7ṇP8). Duplex P1, consisting of 4–6 bp from both intronic and 5′ exonic sequence (‘GNNNNN’ in the intron, also known as the internal guide sequence (IGS) and ‘nnnnnu’ in 5′ exon), presents a ‘u-G’ wobble base pair at the 5′ splice site (29) (30). The P1 extension region (P1ex) influences the speed of the 5′ splicing reaction and site recognition (31). Duplex P10 forms after the first transesterification reaction, involving base pairing between the 5′ intron and 3′ exon, which is crucial for the second transesterification site (32)(Supplementary Figure S1A). In some instances, enhancing P9.0 or P9.2 structures can improve the accuracy of the 3′ splice site (30). The linker elements connecting P9.0 and P9.2 are also essential for the ribozyme-catalyzed trans-splicing reaction (33).

The permuted intron and exon (PIE) method leverages modified group I introns to achieve efficient RNA circularization by facilitating transesterification reactions at defined splice sites (16). This approach eliminates the need for protein ligases, simplifying reaction conditions and purification methods (18). CircRNAs produced using the PIE method exhibit prolonged translation duration, expanding their potential applications in both in vitro and in vivo settings (18). Despite its advantages, the PIE system may introduce additional fragments that could trigger immune responses and require further optimization (34). The spatial arrangement of ribozyme fragments in the PIE system necessitates precise folding and spacer sequences to prevent interference from internal sequences (16,20), highlighting the need for streamlined circularization methods. Efforts towards developing simpler, faster, safer, and more accurate ribozyme-mediated circularization techniques are essential to advance the field of circRNA-based therapeutics and unlock their full potential in various biomedical applications.

This study presents a novel cis-splicing system designed for the efficient circularization of RNAs encoding foreign proteins. We achieved precise circularization reactions without the need for additional spacer elements by integrating the ribozyme moiety as a whole at one end of the precursor. Our efforts were concentrated on developing and identifying the sequence composition and structural features adjacent to the second transesterification site, which is crucial for achieving precise circularization. By delineating these essential elements, we have successfully elucidated the requirements for accurate circularization and highlighted the system's versatility in designing splicing sites. This work advances the development of synthetic circRNAs with less unwanted immune responses and longer translation duration, laying the groundwork for their potential therapeutic applications.

Materials and methods

Plasmid construction

The sequences of homologous arms, group I intron fragments, IRES, and open reading frame (ORF) elements were chemically synthesized by GenScript Biotech Corporation. These fragments can be assembled and cloned into a plasmid backbone containing the T7 promoter using the Gibson assembly method (NEB, NEBuilder HiFi DNA Assembly kit, Cat #E5520S). The sequence information of the synthetic fragments is provided in Supplementary Table S1. Additionally, all the precursor designs mentioned in this study were detailed in Supplementary Table S2, including the recognizers at the 5′ and 3′ ends and the sequence elements composing P1, P1ex and P10.

CircRNA synthesis

The linearized DNAs were utilized as templates for in vitro transcription (IVT) using the HiScribe T7 High Yield RNA Synthesis Kit (NEB, Cat #E2040S) in the presence of unmodified NTPs. The IVT reaction can achieve co-transcriptional circularization by adjusting the magnesium ion (Mg2+) concentration (in this study, IVT was conducted under conditions with a final Mg2+ concentration exceeding 26 mM, such as 36, 46, 56 mM). IVT was carried out at 37°C for 3 h, followed by treatment with DNase I (KACTUS, Cat # DNI-EE001-12) for 30 min at 37°C to eliminate DNA templates. The RNA was purified either by precipitation with 7.5 M LiCl or column purification using a Monarch RNA cleanup kit (NEB, Cat # T2040) to eliminate excess NTPs, salts in the IVT buffer, and any potential small RNA fragments generated during IVT.

For an optional secondary circularization step, the co-transcriptional circularization product was first heated to 70°C for 5 min, followed by immediate cooling on ice for 5 min. Subsequently, GTP (Glycogen, Cat# MR-1002) was added to the RNA to a final concentration of 2 mM, along with a magnesium-containing buffer (comprising 50 mM Tris–HCl, pH 7.5, Thermofisher, Cat# 15567027; 10 mM MgCl2, Thermofisher, Cat# AM9530G; and 1 mM DTT, Sigma, Cat# 43816). The reaction mixture was incubated at 55°C for 12 min, after which the RNA was purified using the method previously described.

The purified IVT samples underwent treatment with RNase R to enrich and confirm the production of circular RNA. A single reaction system (total volume of 50 μl) was prepared as follows: 5 μl of 10× RNase R reaction buffer (KACTUS, Cat # RNR-EE001) and 2 μl of RNase R were added to IVT RNAs totaling 20 μg (with volume adjusted to 50 μl using water). Following a 20-min incubation at 37°C, the products were purified using the Monarch RNA cleanup kit (NEB, Cat # T2040) and quantified using a NanoDrop.

Subsequently, 150 ng of RNA per sample was mixed at a 1:1 ratio by volume with 2× GLB II (Thermofisher, Cat # AM8546G) to a final volume of 20 μl/well. The mixture was heated to 75°C for at least 2 min, then cooled on ice for at least 3 min. The RNA was then separated on a precast 2% E-Gel EX Agarose Gel (Invitrogen, Cat #G401002) using an E-Gel Power Snap Electrophoresis System (Invitrogen, Cat # G8300) with the E-Gel EX 1% - 2% program, utilizing ssRNA ladder (NEB, Cat # N0362S) as a reference standard. Bands were visualized under blue light transillumination.

mRNA synthesis

The linearized plasmids were used as in vitro transcription templates. mRNA was synthesized using the HiScribe T7 High Yield RNA Synthesis Kit (NEB, Cat #E2040S) with the following modifications: CleanCap AG (TriLink, N-7113) was included at a final concentration of 4 mM, and N1-methyl pseudouridine (m1ψ) (Glycogene, Cat #MR-3002) completely replaced UTP. One microgram of linearized plasmid template was used per 20 μl of IVT reaction. Reactions were incubated at 37°C for 3 h. Subsequently, the DNA templates were degraded by adding 2 μl of DNaseI per IVT reaction and incubating for 30 min at 37°C. The resulting mRNA was then purified using column purification before further use.

Fragment analyzer

A fragment analyzer (FA) was applied to evaluate the products. Specifically, purified RNAs (circular or linear) were further analyzed in the RNA mode with capillary electrophoresis with Agilent 5200 or 5300 Bioanalyzer. Samples were diluted to an appropriate concentration and analyzed according to the manufacturer's instructions (Agilent, DNF-471 RNA Kit, 15 nt). Agilent ProSize Data Analysis Software was utilized to analyze the results. In brief, quantifying the circular RNA component in a sample is based on the migration velocity of different components (such as precursors and circular RNA) in a capillary and the corresponding signal intensity when it passes through a detector. This method calculates the percentage of the signal containing the circular RNA component among all detected signals (Supplementary Figure S1C). RNase R can digest linear RNA components, significantly enhancing the signal corresponding to circular RNA. It is important to note that FA cannot effectively distinguish between nicked RNA and circRNA of similar sizes. Therefore, the purity obtained from FA analysis should encompass circRNA and nicked RNA (both splicing products). For descriptive convenience, the results from FA can also be considered a measure of splicing efficiency.

Capillary quantitative analysis

Capillary quantitative analysis (SCIEX, PA800 Plus) was employed to detect circRNAs. CircRNA samples, dissolved in nuclease-free water, were diluted to a concentration of 10 ng/μl using Sample Loading Solution (SLS) (SCIEX, 608082). In the case of complex samples, additional denaturation steps were carried out (70°C for 3 min followed by a 2-min incubation on ice). Subsequently, 100 μl of the treated sample was utilized for further detection, which was conducted on the PA 800 Plus system (SCIEX, A66528) utilizing the RNA 9000 Purity & Integrity Kit (SCIEX, C48231). Following sample loading, components of the input were separated via capillary electrophoresis and detected using the LIF detector under the following conditions: 50 psi, 30 kV, 25°C, for 40 min, resulting in distinct peak signals at successive time points. The RNA 9000 Ladder (SCIEX, AM7150) served as a size reference for the sample bands, with the area percentage (%) and quality (bp) of each component determined through integrated quantification of the PA 800 Plus system.

Cell culture and transfection

HuH7 (SCSP-526, Cell Bank/Stem Cell Bank, Chinese Academy of Sciences) or HEK293 (CRL-1573, ATCC) and A549 (CCL-185, ATCC) cells were maintained in DMEM (Thermo Fisher, Cat # 10313021) and RPMI-1640 (Thermo Fisher, Cat # 11875093) supplemented with 10% fetal bovine serum (Thermo Fisher, Cat #10099141C), respectively. To detect RNA protein expression, 2×104 HuH7, HEK293 or A549 cells were seeded onto 96-well black plates with a clear flat bottom (Corning, Kennebunk, ME) 24 h before transfection. An equal total amount of each RNA sample (100 ng/well) was transfected into the cells using TransIT transfection reagent (Mirus, Cat #MIR 2225) following the manufacturer's instructions.

Protein expression analysis

Cells were observed and photographed for the reporter gene GFP 24 h post-transfection using a fluorescence microscope (OLYMPUS, CKX53).

For luminescence assays, 20 ul of supernatant was collected daily post-transfection for subsequent analysis, followed by complete medium replacement in the cell culture wells. To detect luminescence from Gaussia luciferase, 20 ul of tissue culture medium was transferred to a flat-bottomed white-walled plate (Corning). 30 ul of ONE-Glo™ EX Luciferase reagent, including stabilizer (Promega, Cat #E8120), was added to each sample, and luminescence was measured on a SpectraMax iD3 Microplate Reader (Molecular Devices). Human erythropoietin was detected by solid phase sandwich ELISA (ACROBiosystems, Cat #CEA-C027), essentially according to the manufacturer's instructions.

Reverse transcription, qPCR and PCR

Total RNA was isolated from transfected cells using the FlysisAmp Cells-to-CT 1-Step Probe Kit (Vazyme, Cat #CL131) according to the manufacturer's instructions, followed by TaqMan probe-based qPCR analysis. Gene-specific TaqMan primers were employed for amplification detection, including GAPDH (Thermo Fisher, Cat #Hs99999905_m1), IFN-β1 (Thermo Fisher, Cat #Hs01077958_s1), IL-6 (Thermo Fisher, Cat #Hs00714131_m1), and RIG-I (Thermo Fisher, Cat #Hs01061436_m1). In the case of SYBR Green-based qPCR, total RNA was extracted using the FlysisAmp Cells-to-CT 1-Step SYBR Green Kit (Vazyme, Cat #CL132) according to the manufacturer's instructions. qPCR reactions were performed on the Applied Biosystems 7500 instrument (Thermo Fisher, Cat #4351105). The primers used for SYBR Green-based qRT-PCR are listed in Supplementary Table S1. Threshold cycle values (Ct) were determined for each sample using the comparative Ct method, and gene expression levels were normalized to the housekeeping gene GAPDH or ACTB and relative to the untransfected or mock-transfected controls.

For RT-PCR, in vitro-synthesized circular RNA was reverse transcribed using the HiScript III 1st Strand cDNA Synthesis Kit (Vazyme, Cat #R312), and the resulting cDNA was subsequently used as a template for PCR amplification, with the crossing BSS primers listed in Supplementary Table S1

dsRNA ELISA

The sandwich enzyme-linked immunosorbent assay (sandwich ELISA) (Vazyme, Cat #DD3509-01) was employed to quantify the levels of dsRNA in mRNA or circRNA synthesized in vitro, following the manufacturer's instructions with modifications. In brief, the capture antibody F2 (mouse IgG2a monoclonal antibody) was immobilized on microtiter plates to form a solid-phase antibody. The dsRNA standards and test samples were added to the solid-phase antibody-coated microtiter plate, followed by a 2-h incubation at room temperature. After washing, the plates were further incubated with the detection antibody M5 (purified mouse IgM monoclonal antibody) for 1 h at room temperature. Subsequently, the plates were rewashed and incubated for 1 h with the horseradish peroxidase-conjugated secondary antibody to form the ‘capture antibody-antigen-enzyme-labeled detection antibody’ complex. Following washing, a colorimetric substrate was added for color development. The reaction was terminated, and the absorbance was measured using a SpectraMax iD3 Microplate Reader (Molecular Devices). The absorbance values were directly proportional to the amount of dsRNA in the samples.

IFN-β response in BJ fibroblasts

IFN-β response in BJ fibroblasts is used to qualitatively measure innate immune response following the protocol as previously described (35). BJ fibroblasts (ATCC, Cat #CRL-2522) were cultured in EMEM (ATCC, Cat #30-2003) growth media with L-glutamine (Thermo Fisher, Cat #25030081) supplemented with 10% heat-inactivated fetal bovine serum (Thermo Fisher, Cat #10100147C). Cells were seeded in 96-well flat-bottom cell culture plates (NEST, Cat #701001) at 20000 cells per well for 24 h before transfection. Cells were transfected with mRNA (250 ng per well) using TransIT transfection reagent (Mirus, Cat #MIR 2225). Cell culture supernatants were harvested 48 h after transfection for analyses. According to the manufacturer's protocol, IFN-β production in the supernatant was detected via standard sandwich ELISA (Beyotime, cat # PI572).

Statistics

Statistics were performed using GraphPad Prism. Comparisons of >2 groups were performed using one-way ANOVA adjusted for Dunnett's or Tukey's multiple comparisons. Information on statistical tests performed, exact values of n, and how significance was defined is available in the figure legends. For cell culture experiments, n is defined as cell cultures treated with separately prepared transfection solutions on different days.

Flow cytometry

Huh7 or HEK293 cells were harvested for GFP analysis (untransfected cells as negative control), and 5×104 total cell events were detected on NovoCyte (Agilent, NovoCyte 2060R). The GFP positive population gating was performed using the nadir between the positive and negative peaks, with consistent gates for all comparison experiments. Flow cytometry analysis was performed using NovoExpress 1.4.1 (Agilent, https://www.agilent.com.).

Next-generation sequencing analysis

To address the high error rates commonly observed at the beginnings and ends of raw reads, the first 10 bases at the head and the last base at the tail were trimmed using the Cutadapt tool (https://cutadapt.readthedocs.io/en/stable/). Following this initial step, the universal Illumina adaptors were removed using Trimmomatic (http://www.usadellab.org/cms/page=trimmomatic). Subsequently, the NGSQCToolkit (http://www.nipgr.res.in/ngsqctoolkit.html) was employed to filter out reads of low quality, ensuring that only high-quality reads proceeded to the subsequent analysis phase. In the subsequent step, reads that successfully passed the quality control checks were aligned to the reference sequence utilizing the BWA software (https://mybiosoftware.com/bwa-burrows-wheeler-aligner-short-long-reads.html). Finally, insertions and deletions (indels) were detected by the GATK Mutect2 tool (https://gatk.broadinstitute.org/hc/en-us/articles/360037593851-Mutect2).

Results

CircRNA preparation through cis-splicing reaction

Synthetic circRNAs are generated from linear pre-mRNA transcripts using enzyme ligation or ribozyme (like group I intron) mediated circularization techniques. The classical PIE circularization system divides a group I intron into two parts and positions them on both sides of the targeted RNA sequence. This arrangement requires that the two separated intron segments are able to fold properly without interference from adjacent sequences. Additional non-structural segments, also known as spacers, are incorporated between the target RNA sequence and intron fragments to ensure correct folding and circularization (Supplementary Figure S1B) (16).

To better preserve the structure of introns and minimize the introduction of exogenous spacer sequences, we designed a system in which the entire ribozyme core (including scaffold and catalytic domains) is positioned on one side of the RNA precursor for circularization. This design mirrors the self-splicing Group I intron's natural process that connects the exon regions on both sides of the intron through cis-splicing, thus named the cis-circularization system. We initially selected the Tetrahymena ribozyme (referred to as ribozyme T hereafter) for preliminary validation due to its well-studied structure (36) and splicing mechanism (32) (37): the u-G pairing in the P1 duplex defines the first transesterification site, while the structural domains P2-P8, including the scaffold and catalytic functions, play a crucial role. The P9.0 duplex and the ωG at the 3′ intron end define the second transesterification site. The complete sequence of P1 to P8 containing the intact first transesterification site is placed at the 3′ end of the precursor as the core of the ribozyme, while the 5′ half of P9.0 follows. The other half of the sequence at the 3′ ends P9.0 is placed at the 5′ end of the precursor RNA. We hypothesize that the P9.0 sequences at both ends of the precursor can facilitate the second transesterification by coming into proximity and forming a duplex in space. Ultimately, after two complete transesterification reactions, the ends of the GOI region are connected to form a closed circular RNA structure (Figure 1A).

Figure 1. The cis-splicing system can effectively mediate precursor RNA circularization. (A) Shows a schematic diagram of a cis-splicing circularization system based on ribozyme (group I intron). R1 can pair with R2 to form a P9.0 duplex, a P9.2 duplex mimic, and a double-stranded region through base pairing between Homology arm 1 and Homology arm 2. Additionally, derived from natural sequences, Spacer 1 and Spacer 2 serve as linker structures between the P9.2 and P9.0 duplexes. The secondary structure near the splicing reaction site is shown on the right, with the P10 duplex expected to form after the first step reaction. (the design is called Full-ωG in this context). (B–G):(B, E) The schematic diagram of the circularization elements is illustrated using ribozyme T as an example. In (B), the back splicing site is inside the ORF-GFP, while in (E), it is within the IRES. The design of all R1 and R2 is consistent with that of the precursor Full-ωG (A). (C) and (F) demonstrate the product purity with or without RNase R digestion, as detected by fragment analyzer (FA). Specifically, (C) corresponds to the precursor (B), and (F) corresponds to the precursor (E). (D) and (G) show the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. BSS, back-splicing site.

To ensure the formation of the P9.0 duplex and the accuracy of the second transesterification site, we initially incorporated recognizer elements R1 and R2 at both ends of the precursor, retaining portions of the P9.0 and P9.2 sequences connected with linker sequences (spacer 1 and spacer 2), as shown in Figure 1A (This original precursor design is referred to as ‘Full-ωG’ here, see in Supplementary Table S2). We integrated a half P10 in the linker adjacent to the P1 duplex, enabling it to base pair with the sequence from the 5′ end of the GOI to form the P10 duplex after the first transesterification reaction (Figure 1A). Furthermore, we substituted the half P1 ‘nnnnnu’ sequence with that from the 3′ end of the GOI, ensuring no additional sequences remain in the GOI post-splicing. The versatility of the ‘nnnnnu’ sequence allows it to be located at various positions within the GOI, either in ORF or IRES sequences (Figure 1B and E).

To assess the RNA circularization efficiency of this construct, we performed in vitro transcription (IVT) and circularization reactions using a DNA template containing the HRVB3 IRES and a gene encoding GFP as the GOI, along with the previously mentioned circularization elements. The ‘nnnnnu’ sequence was located inside the protein-coding sequence and positioned at the 3′ end of the gene of interest (GOI) as shown in Figure 1B (unless otherwise noted, ‘nnnnnu’ sequences are located within the coding sequence throughout this work). We adapted a strategy to allow the simultaneous circularization of the IVT transcribed precursor RNA by adjusting the magnesium ion concentration to specific levels (36, 46 and 56 mM). Subsequent enrichment of the circular RNA by RNase R resulted in a total purity of over 90% (detected by Fragment Analyzer; see method and Supplementary Figure S1C), demonstrating the system's effectiveness in facilitating RNA circularization (Figure 1C).

Previous studies reported that circular RNA preparation often results in the formation of nicked RNAs, which can be separated and detected from circular RNAs of similar molecular size by the E-Gel EX system instead of standard electrophoretic methods (e.g. Fragment Analyzer, agarose gel) (38). Similarly, in the products of this cis-circularization system, the presence of nicked RNA, precursor RNA, and spliced ribozyme fragments can be observed through E-Gel analysis, as shown in Figure 1D. The treatment of RNase R digestion can significantly reduce the linear impurities, resulting in purer circular RNA products (Figures 1D and Supplementary Figure S1D).

Next, we positioned the back splicing site (BSS) within the IRES sequence, as shown in Figure 1E. Precursor RNA molecules from the IVT reaction were capable of self-splice to form circular products, which were further enriched by RNase R treatment (Figures 1F, G, and Supplementary Figure S1D). This approach demonstrates versatility in design and efficacy in generating circRNAs. According to previous studies (16,18), purified precursor RNA molecules obtained through IVT typically require an additional self-splicing reaction to achieve efficient circularization. Although optimizing the magnesium ion concentration during IVT allows for co-transcriptional circularization (Figures 1C and F), we attempted to further improve the circularization efficiency by adding a second circularization step after the initial co-transcriptional circularization event. The results showed that after RNase R enrichment, the yield of circRNA from direct co-transcriptional circularization products was higher than that from performing an additional self-splicing reaction. This suggests that the co-transcriptional circularization method is more efficient and cost-effective (Supplementary Figure S1E).

Simplified design of recognizers R1 and R2 for efficient RNA circularization

While we have shown that placing the ribozyme core at one end of the precursor molecule (specifically at the 3′ end) facilitates RNA self-splicing circularization, we continued to include some P9.2 and spacer elements at R1 and R2 of the precursor. To simplify the design further, we initially removed the P9.2 duplex from R1 and R2 (referred to as ‘ΔP9.2-ωG’) (Figure 2A), with the back splicing site designed within the IRES, similar to Figure 1E. We hypothesized that the P9.2 duplex would function as a homologous arm in this context. Despite the absence of P9.2, circularization of the precursor RNA occurred (Figure 2B), and the resulting circularized RNA was capable of expressing GFP in cells (Figure 2C). However, the splicing efficiency was lower (35.2%) compared to the design that includes P9.2 (59.5% at 46 mM Mg2+; Figures 1E and 2B). We then modified the design by retaining P9.2 as part of a homologous arm to bring the two ends of the precursor closer but removed the spacers between P9.2 and P9.0 (referred to as ‘ΔSpacers-ωG’) (Figure 2D), with BSS designed within the ORF, similar to Figure 1C. E-Gel results revealed that precursor molecules lacking spacers still underwent self-splicing circularization, and treatment with RNase R effectively enriched the circRNA to a final purity of 85.0% (Figure 2E) and improved its cellular expression (Supplementary Figure S2A). These findings suggest that the spacers in R1 and R2 are not crucial for circularization. Furthermore, we eliminated all auxiliary components except the essential P9.0 and ωG sites needed for the second transesterification site recognition (referred to as ‘P9.0-ωG’) (Figure 2F). The modified precursor could still self-splice into a circular form, be enriched by RNase R treatment, and express the reporter gene in cells (Figures 2G and H). However, the splicing efficiency was lower than designs incorporating homologous arms (50.2% versus 54.0%), with final purity decreasing from 85.0% to 65.2% (Figure 2E and G).

Figure 2. The cis-splicing system can be simplified. (A) Shows the schematic diagram of the circularization element with ribozyme T as an example—the P9.2 is removed (referred to as ‘ΔP9.2-ωG’). (B) Shows the product purity with or without RNase R digestion detected by fragment analyzer (FA). (C) Shows the Huh7 cell expression (detected by Flow Cytometer for FITC-GFP) of products prepared under different Mg2+ concentrations after treatment with RNase R (data presented as means ± SEM; n = 3). (D) Shows the schematic diagram of the circularization element with ribozyme T as an example. The recognizer can also mediate circularization without Spacers (referred to as ‘ΔSpacers-ωG’). (E–H):(E) and (G) show the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. The product purity of FA analysis is presented in percentage form. BSS, back-splicing site. (F) The recognizer can mediate circularization without P9.2, homology arm elements, and spacers (the design is referred to as P9.0-ωG in this context). (H) the fluorescence image of GFP expression in Huh7 cells transfected with circularized samples (before and after RNase R treatment). Scale bar: 50 μm.

After confirming the cis-circularization system with ribozyme T, we extended our validation to other group I intron-derived ribozymes. The ribozyme from Pneumocystis carinii (referred to as ribozyme P hereafter) was chosen for its well-documented ability to catalyze trans-excision-splicing, a process that removes a targeted sequence from within an RNA transcript (30,39). We integrated the core structural domain of ribozyme P at one end of the precursor, retaining the double base pair of P9.0 (Watson-Crick or wobble pairing) in R1 and R2, and introduced homology arms on both sides to facilitate circularization (Supplementary Figure S2B). The precursor based on ribozyme P effectively underwent self-splicing circularization, with increased magnesium ion concentration enhancing circRNA formation. Specifically, the splicing efficiency increased from 2.4% to 21.4% as the Mg2+ concentration rose from 36 to 56 mM. (Supplementary Figures S2C and D). The resulting product could be enriched by RNase R treatment and was capable of expressing the reporter gene in cells (Supplementary Figure S2E). When we retained only P9.0 at both ends of the precursor without homology arms, the precursor could still undergo ribozyme P-mediated self-splicing and express reporter gene in cells after RNase R treatment, confirming the production of intact circRNAs (Supplementary Figures S2F–H).

These findings demonstrate that in the cis-circularization system, R1 and R2 require only a homology arm to bring the ends of the precursor sufficiently close for circularization.

The proximity of P9.0 and ωg affects the efficiency and accuracy of the second trans-esterification reaction

Our previous findings indicated that R1 and R2 help bring the two ends of the precursor closer, aiding the ribozyme in catalyzing the second trans-esterification reaction precisely, likely related to P9.0 duplex formation (40,41). We hypothesize that the establishment of a double-stranded region, facilitated by the 5′ and 3′ homology arm sequences near the ωG site, is crucial for improving circularization efficiency. In a precursor design named ‘Loop-ωG,’ we eliminated P9.0 and substituted it with a non-structured sequence (e.g. consisting of ‘AC’ bases), disrupting duplex formation at the ωG site (Figure 3A). E-Gel analysis revealed more abundant nicked/intermediated RNAs relative to circRNA after the splicing reaction of Loop-ωG precursor, although the RNA integrity of the unresolvable nicked RNA and circRNA mixture determined by FA was still 63.9%. (lane 1 in Figure 3B). The ribozyme T band in E-Gel suggests that the Loop-ωG precursor successfully underwent the step one reaction to cleave the ribozyme core. However, it may not efficiently progress from the first to the second transesterification reaction, leading to a notable reduction in circular RNA production (Figure 3B).

Figure 3. Impact of P9.0 structural design variants on circularization efficiency and RNA splicing. (A–F): (A, D) the schematic diagram of the circularization element using Tetrahymena as a model. (A) The molecular recognizer cannot facilitate efficient circularization without a paired structure (the design is referred to as Loop-ωG in this context). (D) Reintroducing paired structures (Arms) in R1 and R2 can reinstate circularization (the design is called Arm-Loop-ωG in this context). The BSS is within the open reading frame (ORF). (B) and (E) show the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. The product purity of FA analysis is presented in percentage form. (C) and (F) show the fluorescence image of GFP expression in Huh7 cells transfected with circularized samples (before and after RNase R treatment). Scale bar: 50 μm. (G) Schematically illustrates the process of conducting RNA-seq analysis directly on circularized samples following column purification. After a two-step transesterification reaction, the results depict the splicing region of the GOI, encompassing the BSS, detected mutation types (deletion, insertion), and the corresponding probabilities of their occurrences, presented in percentages. InDel refers to insertion-deletion events. (H) Illustrates the circularization element using Tetrahymena as a model. In this depiction, P9.0 is substituted with a homologous arm adjacent to the ωG residue (the design is called Arm-ωG in this context). (I) Shows the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. The product purity of FA analysis is presented in percentage form. (J) Displays the RNA-seq results showcasing the connecting region of the gene of interest (GOI) after splicing. This visualization includes the detected mutation types at the circularization site, such as insertion-deletion events (InDel).

The inefficiency of the reaction was confirmed by quantifying the circRNAs and nicked RNAs (or intermediates) in the samples using Capillary Quantitative Analysis (PA800 Plus, SCIEX) (Supplementary Figure S3A). Here, we used the precursor ΔSpacers-ωG as a reference, as its R1 and R2 contain homologous arms, P9.2 and P9.0 (as depicted in Figure 2D). The ΔSpacers-ωG produced circRNA, accounting for 36% of the total products, with only 30% being nicked RNAs. In contrast, the Loop-ωG precursor resulted in less than 4% circRNA production, with nicked RNAs representing up to 63.9% (Supplementary Figure S3A). In vitro cell transfection experiments showed that circRNAs produced from Loop-ωG precursor resulted in fewer GFP-positive cells compared to the P9.0-ωG precursor (Figures 3C and 2H), likely due to reduced circular RNA production (Figures 3B and 2G). These results indicate that the reaction at the ωG site might require the formation of a P9.0 duplex or proximity of the precursor 5′ and 3′ ends to proceed effectively. To test this hypothesis, we reintroduced homology arms into the Loop-ωG precursor to bring the ends closer together, resulting in the Arm-Loop-ωG precursor (Figure 3D). Unlike Full-ωG, the Arm-Loop-ωG construct exhibits a distinct Loop region that substitutes for the P9.0 duplex found in Full-ωG and is devoid of the Spacers that bridge the Arm and P9.0 elements (Figures 1A and 3D).This modification, facilitating the duplex formation through the close spatial alignment of precursor ends, improved circularization efficiency to 28.4%, significantly higher than the 3.9% observed with the Loop-ωG precursor (Supplementary Figure S3A). Subsequent E-Gel analysis and cell transfection experiments demonstrated that the circRNA from Arm-Loop-ωG could be enriched by RNase R treatment, leading to increased reporter gene expression in cells (Figure 3E and F and Supplementary Figure S3A).

To further verify the sequence integrity at the splicing sites after the two-step transesterification reactions of Loop-ωG and Arm-Loop-ωG precursors, we analyzed the circularized products using RNA-seq (Figure 3G). Notably, the Loop-ωG precursor exhibited a deletion of approximately 13 nucleotides located downstream of the BSS position, corresponding to the region downstream of the second transesterification ωG site, and occurred at a frequency of 2.4%. In contrast, the Arm-Loop-ωG precursor resulted in an additional 1 to 2 ‘G’ bases after the splicing site at a much less frequent frequency of 0.48%. Notably, no mutations at the circularization site were found in the ΔSpacers-ωG design, as verified by Sanger sequencing and RNA-seq analysis (Figures 2D, 3G, and Supplementary Figure S3B), aligning with the high splicing accuracy previously reported (16,22).

Upon confirming the importance of the ωG residue near P9.0 for precise splicing, we questioned whether the efficiency of the splicing reaction was related to the nucleotide composition of the P9.0 duplex (e.g. the natural P9.0 duplex (32)). To address this question, we substituted P9.0 with other homology arms, ensuring their proximity to ωG to facilitate the reaction (referred to as Arm-ωG precursor)(Figure 3H). This setup enables a two-step transesterification reaction (Figure 3I), with no insertion or deletion detected at the splicing site (Figure 3J), suggesting that the accuracy of the second transesterification reaction site depends more on the proximity of ωG to P9.0 than on the specific nucleotide composition of P9.0.

Diversity in splicing site design (first and second transesterification reactions)

We focused on R1 and R2 to ascertain that the P9.0 duplex adjacent to ωG is a critical determinant for the second transesterification reaction. To explore the diversity of splicing site design, we initially swapped the sequence of the natural P9.0 between R1 and R2 while maintaining their pairing relationship (referred to as Swapped P9.0-ωG) (Supplementary Figure S4A). The findings demonstrate that the precursor maintained a self-splicing efficiency of 58.8% after changing the P9.0 di-nucleotides (Supplementary Figure S4B). CircRNA products from this configuration could be selectively enriched through RNase R treatment (Supplementary Figure S4B) and were capable of expressing the reporter gene in cells (Supplementary Figure S4C), demonstrating the design flexibility of P9.0. Additionally, as ωG is typically the last nucleotide at the 3′ end of most group I introns and participates in the second transesterification reaction (42,43), we questioned whether other nucleotides could replace ωG in a cis-circularizing system. To test this, while maintaining the design of Arm-P9.0 in R1 and R2 (consolidate the homology arm and P9.2 into the term ‘Arm’ due to their similar function), we generated precursors with various ωN (N = C, A or U) substitutions, termed Arm-P9.0-ωN (Figure 4A). Replacing ωG with other nucleotides did not significantly affect the two-step transesterification reaction, maintaining an efficiency of 55% or higher (Figures 2E and 4B). Furthermore, RNA-seq analysis of products from ωA/C/U as a representative revealed no mutations or InDels at the circularization site (Supplementary Figure S4D). These results indicate that the specific nucleotide at the second transesterification reaction site can vary as long as adjacent homology arms are present to facilitate ribozyme self-splicing.

Figure 4. Exploring the flexibility of splicing sites in the cis-circularization system. (A) Shows the schematic diagram of the circularization element with Tetrahymena as an example. The ωG is substituted with ωN (A/C/U) (the design is referred to as Arm-ωG precursor in this context). (B) Shows the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. The product purity of FA analysis is presented in percentage form. (C) Shows the schematic diagram of the circularization elements with Tetrahymena as an example. The P1 duplex between GOI and IGS includes the c-A wobble base pair (the design is referred to as P1-cA precursor in this context)(BSS is set in IRES). (D) Show the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. The product purity of FA analysis is presented in percentage form.

Next, we examined the compatibility of wobble-base pairs other than ‘u-G’ at the first transesterification site within the P1 duplex in the cis-splicing system. Prior research has demonstrated the potential of various non-Watson–Crick base pairings in the P1 region within trans-splicing systems, with the ‘u-G’ wobble pairing showing the highest efficiency (39). To assess the efficiency of alternative wobble pairings in the cis-splicing system, we tested the ‘c-A’ pairing, which exhibited the second-highest reactivity in the trans system, following ‘u-G’ (39). Upon substituting ‘u-G’ with ‘c-A’ at the first transesterification site in the P1 duplex of the cis-splicing system (Figure 4C), the P1-cA precursor underwent a precise two-step transesterification reaction (Supplementary Figure S4D), resulting in the circularization with an efficiency of at least 54% under the conditions tested (Figure 4D). Furthermore, both precursors with the first transesterification site as ‘u-G’ or ‘c-A’ reached a total purity of at least 70% after RNase R treatment (Figures 1F and 4D). These results indicate that the first transesterification site within the P1 duplex of the cis-splicing system can accommodate various wobble-base pairs besides ‘u-G’, thereby broadening the sequence selectivity for the circularization site within the GOI region.

Enhancing the efficiency of group I intron circularization through P1-ex and P10 duplex

During the transition from the first to the second step of the group I intron cis-splicing, conformational changes primarily involve base pairing between partial sequences of the 3′ exon and 5′ intron, leading to the formation of the P10 duplex, essential for the second transesterification reaction (32,44)(Supplementary Figure S1A). Additionally, the P1 extension (P1-ex) can enhance the efficiency and accuracy of the first-step reaction, with its structural stability associated with P10 formation (31). To investigate the necessity of P10 and P1-ex in the cis-splicing system, we removed the P1-ex duplex and half-P10 from the IGS (the precursor is denoted as ΔP1exΔP10) (Figure 5A). This modification significantly increased the presence of incomplete reaction products, including precursors and nicked /intermediate products (Figure 5B). Nevertheless, circRNAs were still produced and were capable of expressing the reporter gene in cells (Figures 5B and C).

Figure 5. Regulatory roles of P1-ex and P10 in group I intron circularization. (A-F): (A) and (D) the schematic diagram of the circularization element using Tetrahymena as a model. (A) The P1-ex and P10 are removed (denoted as ΔP1exΔP10 in the context). (D) P1-ex-1 is removed, and P1-ex-2 is retained. (denoted as Truncated P1ex-P10 in the context). The removed sequence elements are highlighted in blue. The BSS is within the open reading frame (ORF). (B) and (E) Show the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. The product purity of FA analysis is presented in percentage form. (C, F) The Huh7 cell expression (detected by Flow Cytometer for FITC-GFP) of products prepared under 46 mM Mg2+ concentration after treatment with RNase R (data presented as means ± SEM; n = 3).

To further investigate the roles of P1-ex and P10, we retained only a portion of the P10 sequence within the IGS (P1-ex-2) (Figure 5D). This modification led to the formation of a shorter P1 extension and a truncated P10 duplex (referred to as Truncated P1ex-P10), resulting in the restoration of efficient circularization and reporter gene expression compared to the ‘ΔP1exΔP10’ precursor (Figures 5B, E and F). These findings indicate that the conformational change between the first and second steps of the transesterification reaction and the circularization efficiency can be adjusted by modulating by varying the structure (such as sequence and length) of the P1-ex and P10 duplexes.

Prolonged protein expression and minimal innate immune stimulation of CircRNA prepared by the cis-splicing system

CircRNA, due to its unique structural characteristics, is more stable in cells than linear mRNA. We engineered circRNA containing EVB-IRES and the gene encoding human erythropoietin (hEpo) and compared its long-term cellular expression with N1-methylpseudouridine (m1ψ)-base modified linear mRNA counterpart. CircRNAs can be produced on a small-scale (20 μl) to medium-scale (100 ml) with high efficiency (Figure 6A). In a comparative study of RNA cellular expression over 6 days, we observed a significant drop in mRNA expression level within 24 h post-transfection. In contrast, circRNA maintained stable expression for at least 6 days (Figure 6B).

Figure 6. Circular RNA vs. mRNA: protein expression and immune response. (A) and (C) Show the migration mode of the splicing products in the 2% E-Gel EX system. The lines indicate the product types of the corresponding bands, respectively. RNase R digests precursor RNAs but not circular RNAs, allowing circular RNAs to be enriched. The product purity of FA analysis is presented in percentage form. (B, D) hEpo expression stability over 6 days in HEK293 cells (B) and GLuc expression stability over seven days in A549 cells (D), transfected with m1ψ-mRNA or unmodified circRNA. The data are presented as means ± SDs, n = 3; statistical significance indicated by ****P< 0.0001, ***P< 0.001 as determined by two-way ANOVA followed by a Dunnett's test. (E) The expression levels of GLuc in A549 cells transfected with mRNA or circRNA for 7 days. AUC represents the area under the curve. The data are presented as means ± SDs, n = 3; statistical significance indicated by ****P< 0.0001 as determined by one-way ANOVA followed by a Dunnett's test. (F) dsRNA ELISA determined dsRNA content. The data are presented as means ± standard deviations; n = 2 technical replicates; statistical significance indicated by ****P< 0.0001 as determined by one-way ANOVA followed by a Tukey's test.

The PIE-based RNA circularization system is currently the most extensively investigated method for RNA circularization (8,16). CircRNAs produced from this system have been shown to elicit a minimal innate immune response and sustained protein expression profile compared to linear mRNAs (18). To compare the cis-circularization system with the PIE system, we generated circRNAs containing the same IRES-ORF (EVB-Gaussia luciferase) using both frames (Figure 6C). We then compared their protein expression over time, using purified m1ψ-modified mRNA as a benchmark. The A549 cell line, known for its innate immune-related RNA sensors (18,22), was chosen to facilitate a more accurate comparison of exogenous RNA expression under innate immune activation (18). In A549 cells, circRNAs produced by the PIE and Cis systems demonstrated longer expression durations and significantly higher overall expression levels over seven days compared to the m1ψ-modified mRNA (Figure 6D and E).

Efficient expression and repeated administration of synthetic RNAs for therapeutic purposes necessitate minimizing innate immune activation (35,45,46). To assess the activation of the innate immune system, we conducted dsRNA content analysis and IFN-β response assays in BJ fibroblasts using purified circRNAs and mRNA (35). The results revealed minimal residual dsRNA in the circRNA samples (Figure 6F), and the level of IFN-β response was comparable to that elicited by mRNA (Supplementary Figure S5A). Furthermore, we also validated in A549 cells that circRNAs prepared using the cis-splicing system do not strongly activate the expression of innate immune-related factors (Supplementary Figure S5B). These results suggest that circRNAs induce minimal innate immune activation, even in the absence of m1ψ-base modification, which is consistent with previous studies (18,22). Aside from ribozyme-mediated circularization, other methods, such as T4 RNA ligase-mediated circularization, do not induce strong innate immune activation (34,47,48). To compare the Cis-splicing system with other circularization systems, we took circPOLR2A as an example. We circularized it using three different methods: T4 RNA ligase 1 (Lig), PIE, and Cis, ensuring that the resulting RNA sequence was consistent with the previously designed circPOLR2A_J (EPIC) (Supplementary Figure S5C) (47). As expected, the purified circPOLR2A_J did not trigger strong innate immune responses independent of the circularization method (Supplementary Figure S5C). These results demonstrate that circular RNA (coding or noncoding) prepared using the Cis system can evade significant inflammatory factor activation without the need for modified nucleotide incorporation.

Discussion

Ligase and ribozyme-mediated ligations are common approaches for in vitro preparation of circular RNAs. Protein ligase, such as T4 RNA ligase 1, catalyzes the formation of a phosphodiester bond between the 5′-monophosphate and 3′-hydroxyl groups of an RNA molecule, resulting in a closed-loop RNA. However, for longer RNA sequences, a complementary splint (a DNA or RNA oligo) is often necessary to facilitate the proximity of both RNA ends during the ligation reaction. Moreover, optimizing the ligation reaction is necessary to ensure efficient ligation while minimizing non-specific byproduct formation. In contrast, ribozyme-mediated RNA circularization utilizes an RNA structure-dependent self-splicing reaction. This method does not require a ligase or splint, simplifying the reaction setup and facilitating scale-up (8,23,24). Furthermore, ribozyme-mediated systems like PIE (16,47) and Cis systems (Supplementary Figure S6B) are capable of circularizing longer RNA sequences, thereby enhancing their suitability for various applications.

Ribozyme-mediated RNA circularization can be effectively executed using the PIE method, which is based on the group I intron mechanism. CircRNAs synthesized by the PIE method have demonstrated low immunogenicity and longer translation duration, which broaden their potential applications (8,18,20,21). The PIE system splits the ribozyme into two segments and places them at the RNA precursor's 5′ and 3′ ends, requiring that ribozyme fragments are correctly folded and spatially aligned to form the complete catalytic domain. However, the internal sequence structure may interfere with the ribozyme structure at both ends, requiring additional spacer sequences to separate the internal and ribozyme fragments (20,21). Moreover, the two-step transesterification reaction depends on the interaction between partial exonic sequences (E1 and E2) in certain group I introns, such as those in Anabaena, posing potential concerns for innate immune activation and safety (34). Precursors designed using the PIE approach have relatively fixed back-splicing sites (e.g. E1 and E2 junctions), limiting optimization mainly to the homologous arms and spacer regions. This could be challenging for longer and more complex sequences (22).

In our study, the main ribozyme is integrated as a single unit at one end of the precursor, enabling a precise and efficient circularization process without needing additional spacers between the GOI and the ribozyme. While the ribozyme can be positioned in the 5′ region of the precursor (5′ cis) to form the P1 duplex, our design places it in the 3′ region (3′-cis), where it forms a structure similar to the P9.0 duplex. To assess the outcomes of these design strategies, we used the same GOI region with identical splicing sites within the IRES, ensuring the consistency of the sequence in the P1 duplex and P9.0-ωG to eliminate base composition effects (e.g. GC contents). These two setups have the same first and second transesterification sites, differing only in the position of the ribozyme within the precursor (Supplementary Figure S6C and Figure 1E). The results demonstrated that both 3′ cis and 5′ cis designs facilitate the completion of the two-step transesterification reaction with comparable efficiency (Supplementary Figure S6C). However, RNase R did not effectively remove nicked RNA in both systems (Supplementary Figure S6C). AS a 3′ to 5′ exoribonuclease, the efficiency of RNase R is mainly related to the 3′ end structure of RNA (49). For nicked RNA digestion, we observed variations in RNase R efficiency depending on the location of the BSS (such as in the IRES or ORF) (Figures 1D, G, and Supplementary Figure S6C), which may be associated with the sequence structure at the splicing site. Future studies should explore and compare the mechanisms of nicked RNA formation in various circularization systems. Based on this understanding, efforts can be made in sequence design and reaction processes to minimize nicked RNA generation. Additionally, optimizing digestion conditions, such as adjusting ion components to unfold or reduce RNA secondary structures, could enhance RNase R’s digest efficiency (50,51).

Previous studies have shown that removing P9.0 or P10 individually does not affect the second transesterification reaction, but their simultaneous removal does inhibit it (52), indicating the importance of P9.0–P10 region in this process. Our study has revealed that the accuracy of the second transesterification reaction site is more closely related to the proximity of ωG and P9.0 rather than to the nucleotide composition of P9.0. One possible reason is that the introduction of unpaired structures between ωG and P9.0 could disrupt the formation of the P9.0 duplex (stem pairing between P7-P9) (32,36), thereby weakening the interaction between ωG and the guanosine binding site in the P7 duplex, ultimately diminishing the accuracy of the second transesterification reaction site (52). These results suggest that the accuracy of the splicing site relies more on structure integrity than the sequence (53).

Furthermore, it has been observed that substituting ωG with other nucleotides inhibits the second transesterification reaction's efficiency without affecting the splicing site's accuracy (52). Previous reports also suggest that replacing the 3′ end ωG with ωA in a group I intron from Anabaena does not affect the accuracy or efficiency of the second transesterification reaction, though it may reverse the first transesterification reaction, possibly due to the affinity between the G-binding site and ωN (54). In our cis-splicing system, substituting ωN does not affect splicing accuracy (illustrated by ωA/C/U sequencing data) and does not significantly reduce circularization efficiency. This may be attributed to the homologous arms at both ends of the precursor, which aid in ribozyme's conformational changes and stability after the first transesterification, allowing recognition of the second transesterification site determined by P10 and P9.0 (42,43).

For the first transesterification reaction, studies on ribozyme T or P intron self-splicing and trans-excision-splicing (TES) reactions have shown that alternative wobble pairings can replace the classical ‘u-G’ pairing. However, this substitution is often accompanied by a significant decrease in splicing efficiency (39,53) and the formation of cryptic products (39). We explored substituting ‘c-A’ for ‘u-G’ in the P1 duplex and found that the precursor molecule could still efficiently undergo self-splicing and circularization, albeit with a slightly decreased efficiency compared to u-G (Figure 1F and 4D). This result indicates that the cis-splicing system is highly flexible when designing the first transesterification site, offering more possibilities for choosing back-splicing sites within the GOI region.

Besides the crucial roles of P1 and P9.0 in splice site recognition, P10 also impacts the second transesterification reaction in the ribozyme P, with its length variation affecting reaction efficiency (55). In contrast, P10 appears dispensable in ribozyme T (22). In our cis-splicing system, the precursor can complete the two-step circularization reaction in the absence of P10, but reintroduction of the P10 duplex significantly improves efficiency. These studies also demonstrate that while the presence of P10 is not obligatory, it plays a role in improving splicing reaction efficiency. The P1-ex structure contributes to the accuracy of the first-step reaction, but overly stable P1-ex duplexes may impede the progress of the second-step reaction. Conversely, a less stable P1-ex duplex might facilitate P10 formation but could compromise splicing accuracy (31). Therefore, the cis-splicing system can tailor the combination of P10 and P1-ex with varying structural strengths based on the characteristics of different GOI sequences to achieve optimal designs that balance both high efficiency and precision for the synthesis of circRNA therapeutics.

One of the key advantages of in vitro-prepared circRNAs as therapeutic agents is their ability to evade innate immune recognition, thereby avoiding the activation of cytokines that could pose severe safety concerns (8). This immune evasion property, however, may depend significantly on the circularization methods (34) and the purification processes employed (18,48). Both PIE and Cis systems have been observed to generate linear RNA byproducts, which can robustly activate the innate immune system. This activation is likely due to the presence of 5′-triphosphate groups or double-stranded RNA structures that are recognized by immune sensors such as RIG-I (18). To address this issue, purification methods and treatments with enzymes such as alkaline phosphatase can be employed. These steps effectively reduced contaminating linear RNA species that contain 5′-triphosphate groups, resulting in high-purity circRNA that is less likely to trigger strong innate immune responses (18). This underscores the immunologically silent feature of in vitro-synthesized circular RNA, which holds promise for developing new therapeutic modalities.

Supplementary Material

gkae711_Supplemental_Files

Acknowledgements

Author contributions: S.J.Q., P.G. and B.Y. conceived of the approach. S.J.Q., P.G. and H.M.W. designed the research. S.J.Q., H.M.W. performed experiments and analyzed data. G.P.L., Q.S.Q. conducted computational and statistical analysis. The manuscript was drafted by S.J.Q. and revised by P.G., with input and approval from all authors added.

Data availability

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Contacts, peng.gao@abogenbio.com (P.G.) or bo.ying@abogenbio.com (B.Y.). All unique/stable reagents generated in this study are available from the Contacts with a completed Material Transfer Agreement. The data supporting the findings of this study are available within the article and its supplementary materials. The raw data for RNA-seq have been deposited in NCBI’s Gene Expression Omnibus and are accessible through GEO Series accession number GSE263525.

Supplementary data

Supplementary Data are available at NAR Online.

Funding

Suzhou Abogen Biosciences Co., Ltd. Funding for open access charge: Suzhou Abogen Biosciences Co., Ltd.

Conflict of interest statement. S.J.Q., P.G. and B.Y. are co-inventors on pending patent applications related to the cis circularization system. P.G., S.J.Q., H.M.W., G.P.L. and Q.S.Q. are employees of Suzhou Abogen Biosciences. B.Y. is the CEO of Suzhou Abogen Biosciences.
==== Refs
References

1. Pardi N. , HoganM.J., PorterF.W., WeissmanD. mRNA vaccines - a new era in vaccinology. Nat. Rev. Drug Discov. 2018; 17 :261–279.29326426
2. Barbier A.J. , JiangA.Y., ZhangP., WoosterR., AndersonD.G. The clinical progress of mRNA vaccines and immunotherapies. Nat. Biotechnol. 2022; 40 :840–854.35534554
3. Zhang N.N. , LiX.F., DengY.Q., ZhaoH., HuangY.J., YangG., HuangW.J., GaoP., ZhouC., ZhangR.R.et al . A thermostable mRNA vaccine against COVID-19. Cell. 2020; 182 :1271–1283.32795413
4. Zhao H. , ZhangN.N., HannawiS., ZhuA.R., XiongX.C., HuangY.J., YuD.D., ChenC.J., DaiJ., AbuqutaA.et al . Neutralization of Omicron XBB.1 by booster vaccination with BA.4/5 monovalent mRNA vaccine. Cell Discov. 2024; 10 :7.38191591
5. Leppek K. , ByeonG.W., KladwangW., Wayment-SteeleH.K., KerrC.H., XuA.F., KimD.S., TopkarV.V., ChoeC., RothschildD.et al . Combinatorial optimization of mRNA structure, stability, and translation for RNA-based therapeutics. Nat. Commun. 2022; 13 :1536.35318324
6. Qureischi M. , MohrJ., Arellano-VieraE., KnudsenS.E., VohidovF., Garitano-TrojaolaA. mRNA-based therapies: preclinical and clinical applications. Int. Rev. Cell Mol. Biol. 2022; 372 :1–54.36064262
7. Rohner E. , YangR., FooK.S., GoedelA., ChienK.R. Unlocking the promise of mRNA therapeutics. Nat. Biotechnol. 2022; 40 :1586–1600.36329321
8. Niu D. , WuY., LianJ. Circular RNA vaccine in disease prevention and treatment. Signal Transduct. Target Ther. 2023; 8 :341.37691066
9. Qu L. , YiZ., ShenY., LinL., ChenF., XuY., WuZ., TangH., ZhangX., TianF.et al . Circular RNA vaccines against SARS-CoV-2 and emerging variants. Cell. 2022; 185 :1728–1744.35460644
10. Kristensen L.S. , AndersenM.S., StagstedL.V.W., EbbesenK.K., HansenT.B., KjemsJ. The biogenesis, biology and characterization of circular RNAs. Nat. Rev. Genet. 2019; 20 :675–691.31395983
11. Enuka Y. , LauriolaM., FeldmanM.E., Sas-ChenA., UlitskyI., YardenY. Circular RNAs are long-lived and display only minimal early alterations in response to a growth factor. Nucleic Acids Res. 2016; 44 :1370–1383.26657629
12. Liu C.X. , ChenL.L. Circular RNAs: characterization, cellular roles, and applications. Cell. 2022; 185 :2016–2034.35584701
13. Chen C.K. , ChengR., DemeterJ., ChenJ., Weingarten-GabbayS., JiangL., SnyderM.P., WeissmanJ.S., SegalE., JacksonP.K.et al . Structured elements drive extensive circular RNA translation. Mol. Cell. 2021; 81 :4300–4318.34437836
14. Liu C.X. , ChenL.L. Expanded regulation of circular RNA translation. Mol. Cell. 2021; 81 :4111–4113.34686312
15. Fan X. , YangY., ChenC., WangZ. Pervasive translation of circular RNAs driven by short IRES-like elements. Nat. Commun. 2022; 13 :3751.35768398
16. Wesselhoeft R.A. , KowalskiP.S., AndersonD.G. Engineering circular RNA for potent and stable translation in eukaryotic cells. Nat. Commun. 2018; 9 :2629.29980667
17. Yang Y. , FanX., MaoM., SongX., WuP., ZhangY., JinY., YangY., ChenL.L., WangY.et al . Extensive translation of circular RNAs driven by N(6)-methyladenosine. Cell Res. 2017; 27 :626–641.28281539
18. Wesselhoeft R.A. , KowalskiP.S., Parker-HaleF.C., HuangY., BisariaN., AndersonD.G. RNA circularization diminishes immunogenicity and can extend translation duration In vivo. Mol. Cell. 2019; 74 :508–520.30902547
19. Shen L. , YangJ., ZuoC., XuJ., MaL., HeQ., ZhouX., DingX., WeiL., JiangS.et al . Circular mRNA-based TCR-T offers a safe and effective therapeutic strategy for treatment of cytomegalovirus infection. Mol. Ther. 2024; 32 :168–184.37974400
20. Feng Z. , ZhangX., ZhouJ., LiQ., ChuL., DiG., XuZ., ChenQ., WangM., JiangX.et al . An in vitro-transcribed circular RNA targets the mitochondrial inner membrane cardiolipin to ablate EIF4G2(+)/PTBP1(+) pan-adenocarcinoma. Nat Cancer. 2024; 5 :30–46.37845485
21. Chen R. , WangS.K., BelkJ.A., AmayaL., LiZ., CardenasA., AbeB.T., ChenC.K., WenderP.A., ChangH.Y. Engineering circular RNA for enhanced protein production. Nat. Biotechnol. 2023; 41 :262–272.35851375
22. Lee K.H. , KimS., SongJ., HanS.R., KimJ.H., LeeS.W. Efficient circular RNA engineering by end-to-end self-targeting and splicing reaction using Tetrahymena group I intron ribozyme. Mol. Ther. Nucleic Acids. 2023; 33 :587–598.37637208
23. Chen X. , LuY. Circular RNA: biosynthesis in vitro. Front. Bioeng. Biotechnol. 2021; 9 :787881.34917603
24. Petkovic S. , MullerS. RNA circularization strategies in vivo and in vitro. Nucleic Acids Res. 2015; 43 :2454–2465.25662225
25. Vicens Q. , CechT.R. Atomic level architecture of group I introns revealed. Trends Biochem. Sci. 2006; 31 :41–51.16356725
26. Ikawa Y. , ShiraishiH., InoueT. Minimal catalytic domain of a group I self-splicing intron RNA. Nat. Struct. Biol. 2000; 7 :1032–1035.11062558
27. Ikawa Y. , YoshiokaW., OhkiY., ShiraishiH., InoueT. Self-splicing of the tetrahymena group I ribozyme without conserved base-triples. Genes Cells. 2001; 6 :411–420.11380619
28. Ohuchi S.J. , IkawaY., ShiraishiH., InoueT. Modular engineering of a group I intron ribozyme. Nucleic Acids Res. 2002; 30 :3473–3480.12140333
29. Lehnert V. , JaegerL., MichelF., WesthofE. New loop-loop tertiary interactions in self-splicing introns of subgroup IC and ID: a complete 3D model of the Tetrahymena thermophila ribozyme. Chem. Biol. 1996; 3 :993–1009.9000010
30. Muller U.F. Design and experimental evolution of trans-splicing group I intron ribozymes. Molecules. 2017; 22 :75.28045452
31. Guo F. , CechT.R. In vivo selection of better self-splicing introns in Escherichia coli: the role of the P1 extension helix of the Tetrahymena intron. RNA. 2002; 8 :647–658.12022231
32. Li S. , PaloM.Z., ZhangX., PintilieG., ZhangK. Snapshots of the second-step self-splicing of tetrahymena ribozyme revealed by cryo-EM. Nat. Commun. 2023; 14 :1294.36928031
33. Amini Z.N. , OlsonK.E., MullerU.F. Spliceozymes: ribozymes that remove introns from pre-mRNAs in trans. PLoS One. 2014; 9 :e101932.25014025
34. Liu C.X. , GuoS.K., NanF., XuY.F., YangL., ChenL.L. RNA circles with minimized immunogenicity as potent PKR inhibitors. Mol. Cell. 2022; 82 :420–434.34951963
35. Nelson J. , SorensenE.W., MintriS., RabideauA.E., ZhengW., BesinG., KhatwaniN., SuS.V., MiraccoE.J., IssaW.J.et al . Impact of mRNA chemistry and manufacturing process on innate immune activation. Sci. Adv. 2020; 6 :eaaz6893.32637598
36. Liu D. , ThelotF.A., PiccirilliJ.A., LiaoM., YinP. Sub-3-A cryo-EM structure of RNA enabled by engineered homomeric self-assembly. Nat. Methods. 2022; 19 :576–585.35501384
37. Zhang X. , LiS., PintilieG., PaloM.Z., ZhangK. Snapshots of the first-step self-splicing of tetrahymena ribozyme revealed by cryo-EM. Nucleic Acids Res. 2023; 51 :1317–1325.36660826
38. Abe B.T. , WesselhoeftR.A., ChenR., AndersonD.G., ChangH.Y. Circular RNA migration in agarose gel electrophoresis. Mol. Cell. 2022; 82 :1768–1777.35358469
39. Baum D.A. , SinhaJ., TestaS.M. Molecular recognition in a trans excision-splicing ribozyme: non-Watson-Crick base pairs at the 5' splice site and omegaG at the 3' splice site can play a role in determining the binding register of reaction substrates. Biochemistry. 2005; 44 :1067–1077.15654763
40. Burke J.M. , EsherickJ.S., BurfeindW.R., KingJ.L. A 3' splice site-binding sequence in the catalytic core of a group I intron. Nature. 1990; 344 :80–82.2406615
41. Michel F. , NetterP., XuM.Q., ShubD.A. Mechanism of 3' splice site selection by the catalytic core of the sunY intron of bacteriophage T4: the role of a novel base-pairing interaction in group I introns. Genes Dev. 1990; 4 :777–788.2379829
42. Price J.V. , CechT.R. Determinants of the 3' splice site for self-splicing of the tetrahymena pre-rRNA. Genes Dev. 1988; 2 :1439–1447.3209068
43. Adams P.L. , StahleyM.R., KosekA.B., WangJ., StrobelS.A. Crystal structure of a self-splicing group I intron with both exons. Nature. 2004; 430 :45–50.15175762
44. Su Z. , ZhangK., KappelK., LiS., PaloM.Z., PintilieG.D., RanganR., LuoB., WeiY., DasR.et al . Cryo-EM structures of full-length tetrahymena ribozyme at 3.1 A resolution. Nature. 2021; 596 :603–607.34381213
45. Qin S. , TangX., ChenY., ChenK., FanN., XiaoW., ZhengQ., LiG., TengY., WuM.et al . mRNA-based therapeutics: powerful and versatile tools to combat diseases. Signal Transduct. Target Ther. 2022; 7 :166.35597779
46. Stanton M.G. , Murphy-BenenatoK.E. Garner A.L. RNA Therapeutics. 2018; Cham Springer International Publishing 237–253.
47. Guo S.K. , LiuC.X., XuY.F., WangX., NanF., HuangY., LiS., NanS., LiL., KonE.et al . Therapeutic application of circular RNA aptamers in a mouse model of psoriasis. Nat. Biotechnol. 2024; 10.1038/s41587-024-02204-4.
48. Breuer J. , BarthP., NoeY., ShalamovaL., GoesmannA., WeberF., RossbachO. What goes around comes around: artificial circular RNAs bypass cellular antiviral responses. Mol. Ther. Nucleic Acids. 2022; 28 :623–635.35497503
49. Vincent H.A. , DeutscherM.P. Substrate recognition and catalysis by the exoribonuclease RNase R. J. Biol. Chem. 2006; 281 :29769–29775.16893880
50. Xiao M.S. , WiluszJ.E. An improved method for circular RNA purification using RNase R that efficiently removes linear RNAs containing G-quadruplexes or structured 3' ends. Nucleic Acids Res. 2019; 47 :8755–8769.31269210
51. Kwok C.K. , MarsicoG., BalasubramanianS. Detecting RNA G-quadruplexes (rG4s) in the transcriptome. Cold Spring Harb. Perspect. Biol. 2018; 10 :a032284.29967010
52. Michel F. , HannaM., GreenR., BartelD.P., SzostakJ.W. The guanosine binding site of the tetrahymena ribozyme. Nature. 1989; 342 :391–395.2685606
53. Doudna J.A. , CormackB.P., SzostakJ.W. RNA structure, not sequence, determines the 5' splice-site specificity of a group I intron. Proc. Natl. Acad. Sci. U.S.A. 1989; 86 :7402–7406.2678103
54. Golden B.L. , CechT.R. Conformational switches involved in orchestrating the successive steps of group I RNA splicing. Biochemistry. 1996; 35 :3754–3763.8619996
55. Bell M.A. , SinhaJ., JohnsonA.K., TestaS.M. Enhancing the second step of the trans excision-splicing reaction of a group I ribozyme by exploiting P9.0 and P10 for intermolecular recognition. Biochemistry. 2004; 43 :4323–4331.15065876
