Skip to main content An official website of the United States government Here's how you know Here's how you know Official websites use .gov A .gov website belongs to an official government organization in the United States. Secure .gov websites use HTTPS A lock ( Lock Locked padlock icon ) or https:// means you've safely connected to the .gov website. Share sensitive information only on official, secure websites. Search Log in Dashboard Publications Account settings Log out Search… Search NCBI Primary site navigation Search Logged in as: Dashboard Publications Account settings Log in Search PMC Full-Text Archive Search in PMC Journal List User Guide PERMALINK Copy As a library, NLM provides access to scientific literature. Inclusion in an NLM database does not imply endorsement of, or agreement with, the contents by NLM or the National Institutes of Health. Learn more: PMC Disclaimer | PMC Copyright Notice Chem Rev . 2026 Mar 24;126(7):3907–3956. doi: 10.1021/acs.chemrev.5c00347 Search in PMC Search in PubMed View in NLM Catalog Add to search Beyond the Sequence: Chemical and Topological Design and Innovations in mRNA Therapeutics Dangliang Liu Dangliang Liu ∇ Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139-4307, United States ‡ Broad Institute of MIT and Harvard, Cambridge, Massachusetts 02142, United States Find articles by Dangliang Liu ∇, ‡ , Hongyu Chen Hongyu Chen ∇ Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139-4307, United States ‡ Broad Institute of MIT and Harvard, Cambridge, Massachusetts 02142, United States Find articles by Hongyu Chen ∇, ‡ , Alisia Pan Alisia Pan ∇ Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139-4307, United States ‡ Broad Institute of MIT and Harvard, Cambridge, Massachusetts 02142, United States § Harvard/MIT MD-PhD Program, Boston, Massachusetts 02115, United States Find articles by Alisia Pan ∇, ‡, § , Xiao Wang Xiao Wang ∇ Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139-4307, United States ‡ Broad Institute of MIT and Harvard, Cambridge, Massachusetts 02142, United States Find articles by Xiao Wang ∇, ‡, * Author information Article notes Copyright and License information ∇ Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139-4307, United States ‡ Broad Institute of MIT and Harvard, Cambridge, Massachusetts 02142, United States § Harvard/MIT MD-PhD Program, Boston, Massachusetts 02115, United States * Email: [email protected] . Received 2025 Apr 30; Accepted 2026 Feb 13; Revised 2026 Feb 3; Collection date 2026 Apr 8. © 2026 The Authors. Published by American Chemical Society This article is licensed under CC-BY-NC-ND 4.0 PMC Copyright notice PMCID: PMC13067282 PMID: 41875901 Abstract Messenger RNA (mRNA) has rapidly emerged as a transformative therapeutic modality, exemplified by its growing applications in infectious diseases, oncology, and genetic disorders. The chemical programmability of mRNA allows researchers to modulate its function by introducing synthetic modifications across the moleculefrom the cap structure, untranslated regions (UTRs), coding sequence (CDS) to poly(A) tail and from base, backbone to ribose sugars. Beyond sequence-level design, recent advances have introduced a new dimension of control: topological engineering. Circular RNAs, branched structures, and synthetic lariat architectures are reshaping how we approach RNA stability, immunogenicity, and translation. This review surveys recent advances in the chemical and topological engineering of mRNA, emphasizing four key areas: (1) enzymatic, chemical, and hybrid methodologies that expand the repertoire of accessible mRNA modifications; (2) synthesis strategies for linear, circular, and branched mRNA topologies; (3) structure–activity relationships governing translation efficiency, decay, and immune activation; and (4) implications for next-generation mRNA-based therapeutics. By integrating chemical synthesis, synthetic biology, and RNA structural design, researchers are beginning to unlock the full therapeutic potential of engineered mRNA molecules. 1. Introduction The emergence of mRNA technology has catalyzed a paradigm shift in biomedical research, offering unprecedented avenues for novel therapeutics. The advantage of mRNA technology was exemplified during the COVID-19 pandemic, where mRNA-based vaccines outpaced traditional DNA- and protein-based platforms in development speed and scalability. Their intrinsic programmability, streamlined manufacturing, and favorable safety profileparticularly the avoidance of genomic integrationunderscore mRNA’s potential for broader clinical applications, including prophylactic vaccines, genome editing, cancer immunotherapy, and protein replacement therapy. Although the first successful in vivo translation of exogenous mRNA was achieved in 1990 using mouse models, its clinical potential remained limited by its unfavorable immunogenicity, cytotoxicity, and instability. Researchers investigated exogenous mRNA for various therapies (vaccine development, peptide/protein replacement therapy, and antitumor immunotherapy), but were unable to overcome the intrinsic shortcomings of unmodified mRNA. A pivotal breakthrough occurred in 2005 with the introduction of synthetic nucleoside modifications (e.g., m 5 C, Ψ, m 6 A, s 2 U) into mRNA via in vitro transcription (IVT). Researchers were able to significantly decrease unwanted innate immune response induced by unmodified RNAs and substantially enhance protein production, thereby enabling the application of synthetic mRNAs for intracellular protein expression without significant cytotoxicity. Drs. Katalin Karikó and Drew Weissman opened a new era for mRNA-based therapies by introducing chemical modifications to RNA molecules and were awarded the 2023 Nobel Prize in Physiology or Medicine. Building upon such pioneering work, recent efforts have greatly expanded beyond simple nucleoside substitutions introduced via IVT by exploring how chemical and structural (topological) modifications of mRNAs can regulate their function. Innovations in synthetic chemical biology now allow for precise control over RNA modification, circularization, branching, and other noncanonical topologies that modulate stability, translation, immune recognition, and cellular function. In this review, we highlight emerging trends and strategies in the chemical and topological modification and engineering of synthetic mRNA, with a focus on (1) synthetic methodologies for expanding the chemical alphabet and diversifying the structural topology of mRNA vectors; (2) the structure–activity relationships (SARs) governing synthetic mRNA function and biological performance; (3) translational applications of structurally and chemically engineered mRNAs in advanced therapeutics. Together, these advances chart a path toward more versatile, stable, and programmable mRNA-based medicines. Before proceeding to the detailed introduction, key concepts in mRNA therapeutics are summarized in Table to assist readers in navigating the field. 1. Key Concepts for mRNA Therapeutics. Category Term Description mRNA Biology 5′ Cap m 7 G connected to the mRNA via a 5′-5′ triphosphate bond that protects against degradation and initiates translation. A Cap0 structure is not further modified on the first transcribed base (m 7 GpppN), whereas additional 2′-O-methylations of the ribose on the first one or two nucleotides form Cap1 and Cap2 respectively. 5′ Untranslated Region (UTR) Noncoding segments (5′ and 3′) flanking the coding sequence that regulate mRNA stability and translation through interactions with RNA-binding proteins. Coding Sequence (CDS) The protein-encoding region of mRNA; its design and chemical modifications significantly impact translation dynamics and immunogenicity. Polyadenosine (Poly(A)) Tail A stretch of approximately 50 to 200 adenosines at the 3′ end that is essential for nuclear export, translation initiation, and mRNA stability. It interacts with poly(A) binding proteins to protect mRNA from enzymatic degradation or interacts with deadenylation complexes to trigger mRNA decay. Endosomal Escape The process by which delivered exogenous mRNA is released from endosomes, membrane-bound intracellular organelles responsible for sorting during endocytic pathways, into the cytosol; it is a major bottleneck in lipid nanoparticle (LNP) delivery as less than 5% of transfected RNA is released in the cytoplasm for active translation. Translation Efficiency (TE) Measurement of the effectiveness of protein production from mRNA, often expressed as protein output per mRNA amount over time. TE is critically influenced by initiation efficiency including ribosome recruitment efficiency, which can be improved with chemical and topological modifications. mRNA Duration/Stability The expression window depends on the persistence of intact, translation-competent mRNA transcripts in the cell, which is governed by chemical instability and enzyme decay pathways. Modifications modulate pharmacokinetics and thermodynamics of mRNA expression. Cell-type Specificity A major challenge and goal of mRNA therapeutics is preferential delivery and/or expression of transcripts while minimizing off-target expression to improve specificity. mRNA Translation The ribosome-mediated decoding of mRNA transcripts to produce protein through either cap-dependent or cap-independent translation initiation pathway. Internal Ribosome Entry Site (IRES) RNA elements most commonly found in the 5′UTR that directly recruits the 40S ribosome independent of the 5′ cap, commonly found in viral genomes and a subset of cellular mRNAs. RNA Modifications Post-transcriptional chemical alterations to RNA molecules including N 6 -methyladenosine (m 6 A), N 1 -methyladenosine (m 1 A), 5-methylcytosine (m 5 C), pseudouridine (Ψ), inosine (I), and ribose methylation (2′OMe) can endogenously modulate RNA structural stability, ribosome decoding efficiency, protein-interaction, and immune recognition pathways. Innovations in synthetic methods enable more precise control over exogenous RNA modifications that confer favorable properties to therapeutics. Synthesis & Modification Chemically modified RNA RNA, particularly therapeutic mRNA as a focus of this review, can be engineered with noncanonical nucleosides or topological structures to tune properties including immune recognition, structural stability, translation efficiency, and degradation rate. In Vitro Transcription (IVT) A cell-free method using RNA polymerase to synthesize mRNA from a DNA template. Co-transcriptional capping Natural or synthetic cap analogs can be incorporated during IVT reaction to prepare capped mRNA. Xenonucleic Acid (XNA) Synthetic nucleic acid analogues with modified sugar or backbone structures, synthesized using engineered DNA polymerases. Genetic Code Expansion Unnatural base pairs (UBPs) were introduced into DNA templates, enabling site-specific incorporation of noncanonical nucleotides into RNA during IVT. Solid Phase Oligonucleotide Synthesis Nucleoside building blocks (phosphoramidites) are sequentially added to a growing chain anchored to a solid support to chemically synthesize oligonucleotides. RNA Ligation-Enabled mRNA–oligonucleotide Assembly (LEGO) A chemoenzymatic assembly strategy that modularly builds full-length mRNAs from chemically synthesized and IVT fragments. Structure–activity Relationship (SAR) The systematic analysis of how defined structural features of a molecule or biomacromolecule determine its biological activity, enabling rational optimization of function through controlled chemical or topological modifications. Backsplicing A noncanonical RNA splicing mechanism in which a downstream 5′ splice donor is covalently joined to an upstream 3′ splice acceptor, resulting in the formation of a circular RNA molecule. RNA Topology Topological Engineering The design of nonlinear RNA architectures (circular, branched, or multitailed) to regulate mRNA function beyond sequence design. Circular RNA (circRNA) Covalently closed RNA molecules that lack 5′ and 3′ ends, making them highly resistant to exonuclease-mediated degradation. Branched RNA An RNA molecule whose backbone is not strictly linear or circular, but instead contains one or more branch points where a single RNA strand splits into two or more covalently connected arms. Lariat RNA A naturally occurring branched RNA structure formed via 2′–5′ linkage as an intermediate during mRNA splicing. Multicapped RNA An RNA molecule that contains two or more functional 5′ cap structures (e.g., m 7 G caps) within a single RNA construct, for example, by introducing branched 5′ architectures. Multitailed RNA An RNA molecule that contains two or more poly(A) tails, typically generated through branched RNA topologies in which multiple 3′ arms with poly(A) sequence. Capped circRNA (QRNA) A novel ″capped-circular″ mRNA that combines the high stability of circRNA with a branched cap structure to enable efficient cap-dependent translation, named based on visual resemblance to the letter ″Q″. Cis -regulatory Element A cis-regulatory element is a sequence, structure, or chemical feature that is covalently encoded on the same nucleic acid molecule as the gene or RNA it regulates, and whose regulatory function depends on intramolecular (same-molecule) context. Trans -acting Element Trans-acting elements are molecules that regulate a target RNA from a separate physical molecule (e.g., proteins or oligos), rather than being covalently encoded on the same RNA strand. Trans -acting Cap A trans-acting cap or trans-cap is a cap structure (or cap-mimicking moiety) that is not covalently attached to the target RNA, but instead acts in trans to promote translation initiation by interacting with cap-binding proteins or the translation machinery. Trans -acting Tail A trans-acting tail or trans-tail is a poly(A) tail or tail-like RNA element that is not covalently linked to the target RNA, but interacts in trans to enhance RNA stability or translation, typically through PABP recruitment. Therapeutic Applications Cell Reprogramming The process of converting a cell into another cell state or cell type (e.g., pluripotent or lineage-specific) through transient or sustained expression of defined proteins (e.g., transcription factor proteins) and RNAs (e.g., microRNAs), where chemically engineered mRNA or noncoding RNAs enables nonintegrating, tunable, and high-efficiency reprogramming without genetic modification. mRNA vaccine A vaccination platform in which synthetic, chemically modified mRNA encodes antigenic proteins that are translated in host cells to elicit an adaptive immune response. Enzyme/Protein Replacement Therapy (ERT/PRT) The use of mRNA to produce functional proteins or enzymes in vivo to treat diseases caused by protein loss or deficiency. Cancer Immunotherapy A therapeutic strategy that harnesses or modulates the immune system to recognize and eliminate tumor cells, where mRNA is used to encode tumor antigens, immune modulators, or engineered receptors, enabling programmable and transient immune activation with precise molecular control. Gene Editing A gene editing strategy in which synthetic mRNA is used to transiently express genome-modifying enzymes, enabling targeted DNA modification without permanent integration of the editing machinery. Chimeric Antigen Receptor (CAR) T-cell Therapy A living drug where a patient’s T cells are genetically engineered to express a synthetic receptor that recognizes an antigen, which has revolutionized the field of oncology and is demonstrating potential in treating autoimmune diseases, chronic infections, fibrosis, and age-related conditions. mRNA-based technologies would enable in vivo CAR production, minimize safety concerns for transgene integration, increase efficiency, and reduce cost. Open in a new tab 1.1. Structural and Functional Elements of Natural mRNA mRNA is a single-stranded polynucleotide that serves as the template for protein synthesis. In mammalian cells, endogenous (naturally occurring) mRNAs are transcribed from the genome in the nucleus, except mitochondrial mRNA, and subsequently exported to the cytoplasm for translation. By contrast, exogenous synthetic mRNAs are typically released in the cytosol from the delivery cargo (e.g., lipid nanoparticles or virus-like particles) via endocytosis then endosomal escape. Both endogenous and synthetic mRNAs typically comprise of five major elements: a 5′ cap structure, a 5′ untranslated region (5′ UTR), a coding sequence (CDS), a 3′ untranslated region (3′ UTR), and a polyadenosine (poly(A)) tail. Each element contributes differently to mRNA stability, localization, translation, and immunogenicity, making them key targets for chemical and topological modifications. Thus, region- and site-specific synthesis and engineering strategies are essential for modulating mRNA properties, as discussed in the following sections ( Figure A). 1. Open in a new tab Structural properties and translation of mRNA . ( A ) A typical mRNA encompasses a 5′ cap structure, 5′ untranslational region (5′ UTR), coding sequence (CDS), 3′ untranslational region (3′ UTR), and poly(A) tail. ( B ) The life cycle of synthetic mRNAs starts with endocytosis of the mRNA-vehicle complex by the cell, followed by endosomal escape or degradation. Translation of escaped mRNAs is initiated by the interaction between the 5′ m 7 G cap and cap-binding proteins, which further recruit the 40S ribosome to land on the mRNA. The 40S ribosome scans the mRNA for the start codon, followed by the recruitment of the 60S subunit to form the 80S ribosome for protein synthesis. The stop codon and termination factors terminate protein synthesis, releasing the proteins and ribosome units for the next round of translation initiation. 1.1.1. 5′ Cap Structure Conserved in nearly all eukaryotic cellular mRNAs, the m 7 G cap at the 5′ end plays a vital role in regulating the life cycle of mRNAs. An inverted guanosine methylated at the N 7 -position is connected to the first nucleotide in the RNA transcript through a 5′-5′ triphosphate bond. The cap structure without further modification on the first transcribed base (m 7 GpppN) is termed Cap0. In mammalian cells, additional 2′-O-methylations on the ribose of the first one or the first two transcribed nucleotides often occur to form Cap1 (m 7 Gppp(N m )) and Cap2 (m 7 Gppp(N m )p(N m )) structures, respectively. Recent studies also demonstrated that an additional methylation at the N 6 position can occur when the first transcribed base is adenosine (forming m 6 A m ). The cap structure plays multiple roles, including mRNA processing, nuclear export, and regulation of mRNA stability and translation by its involvement with different cap-binding proteins. Within the nucleus, the RNA cap structure is bound by a nuclear cap-binding complex (CBC) including nuclear cap-binding protein 1 (NCBP1) and cap-binding protein 2 (NCBP2), which orchestrates mRNA postsynthesis processing and nuclear-to-cytoplasmic transport. In the cytoplasm, the cap structure of mRNA can directly bind with eukaryotic initiation factor 4E (eIF4E), which further recruits eukaryotic initiation factor 4G (eIF4G) and other RNA-binding proteins to initiate translation. Importantly, m 7 G caps protect mRNA from Xrn1 exonuclease degradation from the 5′ end, resulting in longer half-lives of capped mRNAs than their uncapped counterparts. 2′-O-Methylation of the cap is also involved in the immune sensing pathway to distinguish self- and nonself-mRNAs. Specifically, exogenous uncapped or Cap0 mRNAs are recognized by the innate immune receptor RIG-I and activate the type I interferon (IFN) pathway as foreign invaders in defense against viral infections, while Cap1 and Cap2 mRNAs tend to be immune-silent. ,− 1.1.2. 5′ Untranslated Region (5′ UTR) Located between the cap and directly upstream of the start codon, the 5′ UTR plays a central role in translational regulation beyond the coding sequence. It is involved in the recruitment of translation initiation factors (eIFs) and assembly of the 43S preinitiation complex (PIC), regulating translation initiation through sequence context, secondary structures, and chemical modifications. Secondary structures can either inhibit or enhance translation, depending on their interaction with initiation factors or regulatory proteins. Complex secondary structure within the 5′ UTR may repress translation by impeding PIC assembly or masking the start codon. For instance, the iron-responsive element (IRE) in ferritin mRNA binds with iron-regulatory protein 1 (IRP1) or iron-regulatory protein 2 (IRP2) through the stem loop secondary structure to repress translation initiation, modulating iron homeostasis and cellular response to oxidative stress. − Conversely, some 5′ UTR structures can positively regulate mRNA translation: eukaryotic initiation factor 3 (eIF3) was shown to directly interact with stem-loop structures on the 5′ UTR of cell proliferation regulators such as c-JUN. Additionally, many viral genomes and a subset of cellular mRNAs contain a distinctive cis-acting RNA element at 5′ UTR known as internal ribosome entry sites (IRES), which trigger a noncanonical, cap-independent translation initiation mechanism through direct interaction with 40S ribosome. Given the increasing evidence supporting the vital role of the 5′ UTR in regulating mRNA turnover and translation, sequence engineering was applied to mRNA to optimize its 5′ UTR for enhanced translation. By combining methods including synthetic biology, massive parallel screening technology, and machine learning-based modeling, researchers identified outstanding 5′ UTR sequences that maximize mean ribosome loading from either a natural or a random sequence library and trained models for sequence prediction, thereby enhancing the properties of mRNA-based drugs. − Extensively reviewed elsewhere, 5′ UTR optimization remains a key tool in mRNA engineering. 1.1.3. Coding Sequence (CDS) Downstream of the 5′ UTR and encoding the protein of interest, the CDS affects translation efficiency based on its codon optimality, which typically matches codon usage to the tRNA pool of the host organism. Codon bias affects both translation speed and mRNA stability: optimal codons increase the rates of tRNA decoding and ribosome elongation efficiency while reducing mRNA decay, whereas rare codons may lead to ribosome stalling and degradation. , Thus, each codon in the CDS should be optimized in therapeutic mRNAs based on the codon frequency of the expression system, especially avoiding rare codons, to maximize protein production. For example, early reports demonstrated that sequence-optimized mRNAs without chemical modifications enabled the efficient production of erythropoietin (EPO) and achieved the therapeutic threshold for enzyme replacement therapy (ERT) (Section 7.5 ). Meanwhile, more recent advances took advantage of computational modeling for simultaneous mRNA secondary structure and codon optimization to induce enhanced antibody titers for the mRNA vaccine against COVID-19. These efforts showcase the importance of CDS optimization during mRNA drug design. 1.1.4. 3′ Untranslated Region (3′ UTR) The 3′ UTR contains many cis-regulatory elements to interact with trans-factors, such as RNA-binding proteins (RBPs), enzymes, and microRNAs (miRNA) to influence RNA subcellular localization, translational efficiency, and decay. Adenylate-uridylate-rich (AU-rich) elements (AREs) are the most common cis-regulatory element within 3′ UTRs, which can interact with AU-binding proteins (e.g., AUF1, HuR, Hel-N1, TTP) to activate ARE-mediated decay through rapid recruitment of degradation machinery, including deadenylation exonucleases, exosomes, and processing bodies (p-bodies). − In addition, another well-studied 3′ UTR trans-acting factor is miRNAs, the 20–22 nt regulatory RNAs that bind their complementary sequences in the 3′ UTR and mediate the degradation of target mRNA through RNA-induced silencing complexes (RISCs). Thus while a general consideration for mRNA 3′ UTR design involves avoiding miRNA binding sequences to enhance its stability, strategically introducing cell-type-specific miRNA-binding sites into the 3′ UTR of synthetic RNAs can achieve cell-type specificity in protein expression. Similar to 5′ UTR design, high-throughput technologies including massively parallel reporter assays or cellular library screening have also been applied to 3′ UTR optimization. For example, functional viral element screens and UTR library screens derived from human endogenous genes can identify regulatory RNA elements within 3′ UTRs that enhance mRNA stability, thereby contributing to the development of mRNA drugs with optimized properties. − 1.1.5. Poly(A) Tail The poly(A) tail (generally 50–200 nt) at the 3′ end is necessary for nuclear export, translation initiation, and mRNA stability. It either binds with poly(A) binding protein (PABP), which protects mRNAs from enzymatic degradation, or interacts with deadenylation complexes (such as CCR4-NOT, PAN2-PAN3, and PARN) to trigger mRNA decay. , Meanwhile, the poly(A) tail also plays an essential role in mRNA translation as it can communicate with the cap-eIF4E complex mediated by eIF4G to form the “pseudo-loop” structure to enhance translation initiation and ribosome recycling. Considering the vital role of poly(A) tail in both mRNA stability and translation, poly(A) tail modifications have been extensively explored in recent years to fine-tune the properties of mRNA (Section 5.4 ). 1.1.6. Naturally Occurring Internal Modification Beyond the sequence elements, endogenous RNAs harbor internal chemical modifications that regulate RNA processing, translation, decay, RNA-protein interactions, and immune recognition. , Naturally occurring chemical modifications of RNAsuch as 5-methylcytidine (m 5 C) and pseudouridine (Ψ)were first discovered around 1960, and over 200 distinct RNA modifications have since been identified in human rRNAs, tRNAs, and mRNAs. Common mRNA modifications include N 6 -methyladenosine (m 6 A), N 1 -methyladenosine (m 1 A), 5-methylcytosine (m 5 C), pseudouridine (Ψ), inosine (I), and ribose methylation (2′OMe). These modifications can alter structural stability, influence ribosome decoding, or modulate RNA-RBP interactions. − For instance, introduced by pseudouridine synthase (PUS) enzymes, pseudouridine (Ψ) contributes to immune response evasion of mRNA and enhanced structural stability of tRNA; cap and first m 6 A m protect mRNA from degradation, whereas internal m 6 A promotes RNA decay; and m 5 C on rRNA maintains the protein synthesis level during translation. , Given the biocompatibility and regulatory functions, these endogenous RNA modifications offer a rich resource for engineering synthetic mRNAs with improved performance and reduced immunogenicity. 1.2. Life Cycle of Synthetic mRNAs and Their Interactions with Endogenous Translation Machinery The design of synthetic mRNA therapeutics is deeply rooted in the ability to harness and integrate into the host cells’ natural translation machinery. This section outlines the intracellular journey of synthetic mRNAsfrom delivery to degradationand provides the biochemical context necessary for understanding how chemical and topological modifications exert their effects ( Figure B). 1.2.1. Cellular Uptake and Cytoplasmic Delivery The life cycle of synthetic mRNAs starts from cellular uptake. Due to their large size, negative charges, and susceptibility to nucleases, naked synthetic mRNAs cannot efficiently cross cellular membranes. Delivery vehicles such as lipid nanoparticles (LNPs) or liposomes are therefore employed to neutralize the charge of nucleic acids, to protect mRNA molecules from extracellular nucleases, and to form mRNA-vehicles for cellular uptake through receptor-mediated endocytosis. After internalization, the mRNA-vehicle complex traffics through endosomal compartments which are membrane-bound intracellular organelles responsible for sorting of internalized materials. Early endosomes maintain a weak acidic pH of ∼ 6 via proton pumps and Ras-associated binding proteins. Upon further endosomal acidification, the mRNA-vehicle complex tends to dissociate and endosomal escape releases mRNAs to reach the cytoplasm. Although the exact mechanism is poorly understood, endosomal escape has been characterized as a bottleneck for LNP-mediated therapeutics, where <5% of the total transfected RNA is released into the cytoplasm for active translation. Two general theories have been proposed to explain the endosomal escape process: (1) membrane fusion, where ionizable lipids interact with anionic lipids on the luminal side of the endosomal membrane upon endosomal acidification, inducing a nonbilayer structure that destabilize the membrane for nucleic acid payloads releasing; (2) the “Proton Sponge Effect”, where the buffering capacity of the LNP leads to an activation of proton pumps to increase a membrane potential and consequently an influx of chloride ions to balance the charge, which collectively increases the osmotic pressure inside the endosome leading to endosomal swelling and subsequent rupturing to release the nucleic acid cargo. During endosomal trafficking, exogenous mRNAs could be sensed by pattern recognition receptors (PRRs), especially RNA-sensing Toll-like receptors (TLRs) such as TLR3 (for dsRNA) and TLR7/8 (for ssRNA), which trigger innate immune activation and RNA degradation. Chemical modifications such as Ψ and 2′OMe can reduce such immunogenic responses (see Section 5 ). 1.2.2. Translation Initiation and the Canonical mRNA Translation Pathway Once released into the cytoplasm, synthetic mRNAs encounter the endogenous translational machinery to produce proteins. In eukaryotes, the canonical cap-dependent translation pathway begins with recognition of the m 7 G cap by the cap-binding protein eIF4E to form the eukaryotic initiation factor 4F heterotrimeric complex (eIF4F). eIF4F also includes two other proteins: eIF4G, a scaffold protein that interacts with PABP to form a “pseudo-loop” mRNA conformation, and eIF4A, an RNA helicase that unwinds 5′ UTR secondary structures. The eIF4F complex recruits the 43S preinitiation complex (PIC), composed of the 40S ribosomal subunit, the initiator tRNA (Met-tRNA), and several eIFs. The PIC scans the 5′ UTR of the mRNA until it recognizes the AUG start codon, initiating assembly of the complete 80S ribosome by recruiting the large ribosomal subunit (60S). Importantly, among the translation phases (initiation, elongation, and termination), PIC formation during initiation is often considered the rate-limiting step. Elongation proceeds via codon-specific decoding by aminoacyl-tRNAs, while translation terminates when eukaryotic release factors (eRFs) recognize a stop codon and disassemble the ribosome. , It is noteworthy that the rate and efficiency of translation are strongly influenced by both RNA sequence context and chemical modifications. Thus, to quantitatively evaluate the protein production capacity of mRNAs, we adopt the definition of translation efficiency (TE) of mRNAs as the amount of protein produced per mRNA molecule per unit time, which serves as a metric for describing the translatability and performance of synthetic mRNAs. 1.2.3. Cytoplasmic Stability and Degradation Pathways Synthetic mRNAs are subject to various endogenous decay pathways. The detailed mechanisms of intracellular degradation of naturally occurring mRNAs have been investigated for decades and summarized in previous reviews. , In brief, both exonuclease and endonuclease decay pathways are involved for cytoplasmic mRNA, including: (1) 3′ to 5′ Decay: initiated by deadenylation, which shortens the poly(A) tail of mRNA transcripts via complexes (e.g., PARN, PAN2/PAN3, and CCR4/CAF), followed by further enzymatic removal of nucleotides by exonucleases such as RRP44 or EXOSC10 from the 3′ end. (2) 5′ to 3′ Decay: as the m 7 G cap structure protects the mRNA from direct degradation by exonucleases, decapping by DCP2 or NUDT16 is required for mRNA 5′ to 3′ decay, followed by XRN1 degradation of the uncapped mRNA from the 5′ end. (3) Endonucleolytic cleavage: internal RNA breaks of phosphodiester bonds are catalyzed by endonucleases such as PMR1, IRE1, or SMG6. Understanding these mechanisms allows researchers to rationally design chemical or structural modifications that mitigate RNA decay (e.g., cap analogs resistant to decapping, poly(A) tail modifications that resist deadenylation) as comprehensively summarized in Section 5 . 2. Synthesis Methodology for mRNA Modifications The ability to chemically modify mRNA molecules has been critical for overcoming the limitations of RNA instability and immunogenicity for therapeutics. Since the first discovery and identification of mRNA in 1961 and the characterization of mRNA cap structures in 1975, , researchers have explored synthetic methods that enable the cell-free preparation of mRNA molecules. Yet, unmodified synthetic mRNA is unstable and triggers undesired immune responses inside human cells, limiting its therapeutic applications. The demonstration of decreased cytotoxicity and increased protein production with modified nucleosides thus highlighted the need to introduce chemical modifications and fine-tune mRNA properties for therapeutic applications. However, synthetic methodologies to access chemical modifications on mRNA molecules are still limited due to their large molecular weight and chemoenzymatic susceptibility, impeding the systematic investigation of their structure–activity relationship (SAR) and optimization of their chemical structures using traditional approaches for other types of drug molecules (e.g., small molecules, antisense oligonucleotides (ASOs), small interfering RNAs (siRNAs), and others discussed in Section 5 ). Thus, novel chemical and chemoenzymatic methods for mRNA synthesis are highly desired to facilitate the study of mRNA therapeutics. In this chapter, we summarize and review recent advances in synthesis methodologies that have expanded the accessible chemical space for next-generation mRNA drugs, each offering distinct advantages and limitations: cotranscriptional incorporation via In Vitro Transcription (IVT) (2.1) , post-transcriptional chemical/chemoenzymatic modification (2.2) , total chemical synthesis (2.3) , and chemoenzymatic synthesis (hybrid synthesis) (2.4) . 2.1. Co-transcriptional Incorporation via In Vitro Transcription (IVT) 2.1.1. IVT Basics IVT is a cell-free mRNA synthesis method that mimics cellular transcription by using RNA polymerase to polymerize a set of ribonucleoside 5′-triphosphates (NTPs) into an RNA chain in the 5′ to 3′ direction from a DNA template ( Figure A). Early attempts at IVT reactions used E. coli -derived RNA polymerase, which enabled the synthesis of poly(AU) from poly(dA:dT) DNA templates. To further increase the efficiency and scalability of IVT reactions, researchers identified bacteriophage-derived enzymes such as T7, T3, and SP6 RNA polymerases, − which have since been widely used for IVT-based mRNA production. Among commercialized RNA polymerases, T7 is the most commonly used due to its high fidelity and efficiency. However, RNA polymerase activity is sensitive to both the promoter sequence and the first transcribed nucleotide. For instance, transcription from the highly conserved T7 class III promoters produces RNAs with guanosine as the first nucleotide, while T7 class II promoters starts transcription with an adenosine. The development of polymerase-based IVT reaction not only enables highly programmable synthesis of RNA molecules with desired sequences but also provides a powerful platform technology for incorporating synthetic RNA modifications. 2. Open in a new tab Enzymatic IVT incorporation for mRNA synthesis . ( A ) In vitro transcription (IVT) enables the preparation of synthetic mRNAs based on DNA templates. ( B ) Co-transcriptional capping for preparing mRNA with cap modifications. ( C ) Co-transcriptional NTP substitution enables mRNA internal sequence modification. 2.1.2. Co-transcriptional Capping To prepare mRNA with a m 7 G cap structure, cotranscriptional capping technology was developed in 1982 to employ a dinucleotide cap analogue (i.e., m 7 GpppG) as the primer for IVT by RNA polymerase (e.g., SP6, T7) ( Figure B). However, about one-third of the RNA molecules generated this way are translationally inactive due to the reverse-incorporation of m 7 GpppG in the form of Gpppm 7 G, which cannot be recognized by eIF4E to initiate translation. To address this issue, “anti-reverse” cap analogs (ARCAs) were developed by methylating the 3′-OH position of the m 7 G nucleotide to block elongation from the m 7 G end, ensuring correct orientation for transcription elongation ( Figure B). While ARCAs improve mRNA translation activity by at least 2-fold, , they suffer from several limitations: (1) low incorporation/capping efficiency, typically from 50% to 90% depending on the structure and concentration of cap analogs during the reaction, due to competition from GTP with ARCA for transcription initiation; (2) reduced yield of the IVT reaction because a high ARCA:GTP ratio (typically 4:1 to 10:1) is required for optimal capping efficiency, constraining NTP concentration and thus synthesis efficiency; (3) limited scope of mRNA cap modifications, as ARCAs only contain the first-transcribed base and thus are incompatible with the synthesis of naturally occurring Cap1 and Cap2 structures. These limitations led to the development of trinucleotide cap analogs as transcription initiation primers, enabling efficient incorporation of m 7 GpppN 1 pN 2 (usually m 7 GpppApG) into mRNA by T7 polymerase. The trinucleotide primer takes advantage of the T7 polymerase’s preference for G for transcription initiation, allowing efficient priming by the cap analogue compared to competing nucleotides that would result in a mixed product with uncapped mRNA transcripts. Therefore, the first transcribed bases from the WT T7 promoter 5′- TAATACGACTCACTATA G -3′ need to be adjusted to 5′- TAATACGACTCACTATA AG -3′ to be compatible with the trinucleotide analog m 7 GpppApG. The novel cap analogs enable higher capping efficiency >90%, increase yield without limitations on GTP concentration, and are compatible with a broader range of chemical modifications at the cap position, especially the Cap1 structure. Some of the trinucleotide cap analogs have been commercialized (e.g., m 7 GpppA m pG as the CleanCap reagent) and are widely used for the production of mRNA vaccines or other therapeutic mRNAs. Along this line, tetranucleotide cap analogs compatible with Cap2 mRNA synthesis were also developed to further expand the scope of mRNA cap modification ( Figure B). In summary, the cotranscriptional capping technology, developed through the chemical synthesis of cap analogs, enabled the scalable production of capped mRNA directly via IVT, with chemical modifications to the m 7 G cap and the first two nucleotides. However, the limitations of this approach include: (1) individualized reaction optimization sometimes requiring exonuclease treatment given variable efficiency depending on cap analog structure and polymerase compatibility, which complicates the screening for diverse structures and limits throughput for a more systemic SAR investigation; (2) site-specific chemical modification scope restriction to the first one or two nucleotides, while modifications of the rest of mRNA chain still rely on other synthesis methodologies (see Sections 2.2–2.4 ). 2.1.3. Co-transcriptional NTP Substitution Beyond cotranscriptional capping, modified NTPs can be used as spike-ins or as a 100% substitution of natural NTPs in IVT reactions to enable cotranscriptional incorporation of RNA chemical modifications at internal sites of mRNAs ( Figure C). Canonical RNA polymerases (e.g., T7, SP6) are compatible with a range of base modifications (e.g., m 5 C, m 6 A, Ψ, m 1 Ψ, m 5 U, or s 2 U), ,, phosphate backbone modifications (e.g., phosphorothioate, boranophosphate), , and sugar modifications (e.g., 2′-fluorodeoxyribose or 2′-aminodeoxyribose). , While full substitution is possible for subtle modifications (mostly base modifications), many other modified nucleosides (especially phosphate backbone and sugar modifications) can only be substituted partially due to polymerase intolerance and a significant decrease in IVT yield. Notably, the full substitution of Ψ and its derivatives significantly reduced the immunogenicity of exogenous RNAs, thereby enabling mRNA vaccine development. , The IVT enzymatic approach has become the industry standard due to its simplicity and compatibility with CMC (Chemistry, Manufacturing, and Controls) and GMP (Good Manufacturing Practice) processes. However, limitations for this widely used technique include: (1) partial substitution of modified NTP during IVT are not site-specific, producing highly heterogeneous mRNA population and raising concerns for quality control and batch effect; (2) difficulty in interpreting the structure–activity relationship (SAR) of modifications because each modification may have variable impact on the properties of mRNA depending on the local sequence context; , (3) the narrow scope of chemical modifications limited by the intolerance of RNA polymerases for modifications that impede base-paring or backbone extension (e.g., m 1 A, m 3 C, m 6 A for base modifications, 2′-OMe, locked nucleic acids (LNAs), and 2′-MOE for ribose modifications). 2.1.4. XNA Synthesis via Engineered DNA Polymerase Beyond the naturally occurring RNA polymerases used for mRNA synthesis, researchers also repurposed and engineered DNA polymerases to use modified NTPs in synthesizing xenonucleic acid (XNA) polymer, a class of sugar-modified nucleic acids. Given the structural similarities among DNAs, RNAs, and XNAs, thermostable DNA polymerases have been mutated and repurposed to accommodate unnatural nucleotides: dideoxynucleotides for Sanger sequencing and non-natural ribonucleotides for XNA synthesis with 2′ modification. , An important example is the evolution of Thermococcus gorgonarius (Tgo, a hyperthermophilic archaeon) DNA polymerase, a polB family DNA polymerase. Using a compartmentalized self-tagging (CST) strategy, Tgo C7 (10 amino acid mutations to the original Tgo) was developed for the incorporation of locked nucleic acid (LNA) up to 72 nt. Another variant called Tgo TGK was developed by introducing two additional mutations to Tgo to further enable the successful synthesis of heavily modified mRNAs and XNAs (up to 1700 nt) with Ψ, m 5 C, 2′-F, and 2′-azido modifications. Recently, three additional mutations were introduced to decrease the steric gate of Tgo TGK to accommodate bulky 2′-OMe or 2′-MOE modifications, yielding XNA synthesis up to 800 nt. ( Figure A) Similar engineering efforts with another polB enzyme from Thermococcus kodakarensis (KOD) using a high-throughput evolving system of reverse transcription compartmentalized self-replication (RT-CSR) have led to the discovery of a KOD RTx mutant that can synthesize oligonucleotides using NTPs and 2′-OMe NTPs. Finally, a thermophilic DNA polymerase SFM4–6 derived from the polA family also showed the potential to synthesize modified RNA molecules, especially 2′-OMe- or 2′-F-modified oligonucleotides. 3. Open in a new tab Enzymatic synthesis of mRNA with engineered DNA polymerase . ( A ) Structure-guided DNA polymerase engineering generates polymerase mutants that enable the incorporation of nucleoside triphosphates and their 2′ modified derivatives. ( B ) Engineered DNA polymerases with extended substrate scope enable site-specific or region-specific synthesis of RNA molecules. The authors curated a modified x NTPs library, fluorophore-labeled primers, and ssDNA templates, and confirmed product formation using denaturing PAGE (dPAGE). (i) Modified nucleotide was introduced at a single internal site through adding a modified NTP together with three other unmodified NTPs. (ii) Multiple incorporations with four modified nucleotides achieve the synthesis of fully modified RNAs. (iii) Incorporation of a single modified nucleotide at multiple sites was achieved through careful template sequence design. (iv) Terminating sequence traversal with or without modified nucleotides was achieved using the primer extension reaction (PEX). (v) RNA with three modified nucleotides could be synthesized with engineered DNA polymerase, while the T7 RNA polymerase cannot. (vi) Full-length mRNA with a single site-specific modification was synthesized through single modified nucleotide PEX, phosphatase treatment, and enzyme denaturation, a second round of natural NTPs PEX, and DNase digestion of the template. x NTP or NTPs in each reaction scheme represent the NTP substrates used in the specific reaction. Reproduced with permission from ref. Copyright 2024 Springer Nature under [CC BY-NC-ND 4.0] [ http://creativecommons.org/licenses/by/4.0/ ]. Recently, these engineered DNA polymerases have been applied to region-specific mRNA modifications, in which an RNA primer (with or without modifications) anneals to the single-stranded DNA (ssDNA) template, and then NTPs or their modified counterparts are added into the polymerization reaction to achieve template-dependent primer extension. By careful sequence design, this method can be used to incorporate internal mRNA single-nucleotide modification through pausing and fragmented synthesis. ( Figure B) Compared with T7 RNA polymerase-based mRNA synthesis, the TGK-based method is compatible with several unnatural base modifications (e.g., A Ph TP, U Bio TP, C mBdp TP) that are too sterically demanding for T7 RNAP to incorporate. Meanwhile, mRNA site-specific modification can be achieved via one-pot fragmented synthesis, greatly expanding the scope of mRNA modification. While promising, the technology still faces several limitations when using engineered DNA polymerase for mRNA synthesis. (1) Due to the lack of strand displacement activity and primer annealing constraints, DNA polymerase-based methods require equimolar ssDNA templates and thus have limited scalability owing to the difficulties in large-scale ssDNA preparation. (2) Given its intrinsic preference for DNA synthesis, the processivity of engineered DNA polymerase for RNA synthesis is a significant issue, leading to truncated products. So far, the quality and activity of mRNA synthesized with engineered DNA polymerases are less well characterized, especially compared with those of widely used RNA polymerases (e.g., T7 polymerase). (3) Engineered DNA polymerases typically have much lower fidelity (error rate: 10 –3 ) than commonly used IVT RNA polymerase (error rate: 10 –5 ), raising potential safety concerns for therapeutic applications. 2.2. Post-transcriptional Chemical/Chemoenzymatic Modification Apart from cotranscriptional modification during IVT, post-transcriptional modification methods act on already transcribed RNA, offering precise site-specific modifications and expanding the chemical space of modifications for enhanced functional properties. 2.2.1. Cap Modification with Capping Enzymes In most eukaryotic cells, the m 7 G cap structure is introduced at the 5′ of mRNA cotranscriptionally through sequential enzymatic reactions. Recruited by RNA polymerase II to the 5′ triphosphate of growing mRNA molecules, the capping enzyme complex first hydrolyzes the 5′ γ-phosphate from nascent pre-mRNA to form the mRNA with 5′ diphosphate. It then transfers the GMP from a GTP donor to the 5′ diphosphate of mRNA. Finally, the capping enzyme complex functions as a methyltransferase to methylate the N 7 position using S-adenosyl methionine (SAM) as a donor to form the canonical Cap0 structure (m 7 GpppN, Figure A). An additional RNA cap 2′ O-methyltransferase may modify the ribose of the first and second nucleotides at the 2′ position to form Cap1 (m 7 GpppN m ) and Cap2 structures (m 7 GpppN m pN m ). ( Figure A) Following a similar pathway to eukaryotic mRNA capping, viruses that replicate in the cytoplasm of eukaryotic cells rely on their own enzymes to cap viral transcripts or endonucleolytically cleave cap structures from cellular mRNAs to avoid host recognition and antiviral immune responses. The capping enzyme of the vaccinia virus is the most well studied among the virus-derived capping machinery. , There are two viral proteins in the capping enzyme complex of vaccinia virus (VACV): (1) D1 protein that contains RNA triphosphatase, RNA guanylyltransferase, and RNA methyltransferase activities, and (2) D12 protein that stimulates the methyltransferase activity of the D1 protein. Meanwhile, VACV also contains a 2′-O-methyltransferase enzyme (VP39) that adds an additional 2′-methylation to the first nucleotide of mRNA to form the Cap1 structure. , These naturally occurring capping pathways hold potential for in vitro enzymatic mRNA capping, among which the virus capping enzymes (VCEs) have been successfully repurposed and purified for the post-transcriptional capping of an uncapped mRNA prepared through IVT, ( Figure A) enabling large-scale preparation of capped mRNA by using easily accessible substrates without the requirement for cap analogs. Different GTP analogs were synthesized and demonstrated to be compatible with VCE to introduce chemical modifications, though the capping efficiency varied significantly from 10% to 90% based on individual GTP derivatives. Resembling the method of using dinucleotide cap analog for cotranscriptional capping, VCE-based postsynthesis modification methods also suffer from a narrowed scope of chemical modification due to their limited compatibility with modified substrates. 4. Open in a new tab Cap modification with engineered capping enzymes . ( A ) Vaccinia capping enzymes for postsynthesis mRNA capping utilize (i) RNA triphosphatase (TPase) to remove the 5′ terminal phosphate; (ii) capping enzyme guanyltransferase (GTase) to hydrolyze gamma-phosphate of GTP and form a new 5′-5′ triphosphate linkage on the 5′ end of RNA; (iii) RNA methyltransferase (MTase) with an AdoMet methyl donor to form the canonical Cap0 structure; and (iv) cap 2′-O-MTase VP39, which can also act as a poly(A) polymerase processivity factor, to form Cap1 after additional methylation on the ribose. ( B ) Engineered cap modification enzymes (including trimethylguanosine synthases, CAPAM, and methyltransferases) enable the chemical derivatization of the mRNA cap structure. Beyond the application of VCE for IVT mRNA capping and modification, other evolutionarily conserved enzymes with yeast and human homologues have also been tested for site-specific mRNA cap modifications. ( Figure B) For instance, trimethylguanosine synthases (Tgs) is an endogenous enzyme that converts the canonical m 7 G caps to the 2,2,7-trimethylguanosine (TMG) in small nuclear RNAs (snRNAs) and small nucleolar RNAs (snoRNAs), as hypermethylation plays critical roles in RNA transport, splicing, and maturation. By treating the m 7 G capped mRNA with recombinant Tgs enzyme derived from either Homo sapiens (hTgs1) or Giardia lamblia (GlaTgs2), researchers achieved the introduction of alkyl groups to N 2 of guanosine to form N 2 -methyl-modified cap or N 2 -allyl-modified cap depending on the different AdoMet donor analogues applied. In a similar fashion, an engineered AdoMet synthetase can efficiently generate AdoMet analogs with different modifications (e.g., photocaged groups) as substrates for mRNA cap analogs modifications employing methyltransferases (MTase). Specifically, CAPAM (cap-specific adenosine methyltransferase) is an MTase with helical domains to specifically recognize N 6 -methylation of A m (m 6 A) in Cap1 structure to form m 6 A m . Using different AdoMet analogs as alkyl donors with CAPAM can introduce an additional alkyl group at the N 6 position of the first adenosine in the capped mRNA. , In summary, enzyme-based strategies provide site-specific modifications to the mRNA cap for subsequent functional studies. 2.2.2. mRNA 2′-OH Acylation Reactions involving the ribose 2′-OH group for RNA labeling have been explored for decades to regulate RNA properties. Acylating reagents have been developed to selectively react with 2′-OH of ribose without interfering with other functional groups in the RNA (e.g., exocyclic amines on nucleobases). ( Figure A) Effective acylating agents rely on the trade-off of reaction specificity, reaction rate, compound half-life, and chemical compatibility. For instance, 1-methyl-7-nitroisatoic anhydride (1M7) is a faster-acting compound for RNA 2′-OH labeling with reactivity similar to all four ribonucleotides compared to its parent compound NMIA ( N -Methylisatoic anhydride), speeding up experiments and reducing the risk of degradation when using this chemistry for RNA structural analysis (e.g., SHAPE-MaP). Another type of 2′-OH acylating reagent is characterized by bearing an imidazole leaving group (e.g., 2-methyl-3-furoic acid imidazolide FAI, clickable 2-methylnicotinic acid imidazole-azide reagent NAI-N3) that is not electron-deficient enough to activate the carbonyl, leading to a longer half-life and efficient acylation of RNA at superstoichiometric levels in minutes, particularly suitable for in vivo experiments requiring time for diffusion. 5NIA (5-nitroisatoic anhydride) is another long -half-life reagent with better in-cell reactivity. RNA acylation has been applied in many areas of research, including RNA labeling, RNA structural analysis, and nuclease protection studies. Given the versatility of this reagent, researchers have applied the technology to SHAPE (selective 2′-hydroxyl acylation analyzed by primer extension) in elucidating the secondary structure analysis of RNAs. 5. Open in a new tab RNA 2′-OH acylation for enhancing mRNA stability . ( A ) Schematic of the acylation reaction involving a 2′-OH and an activated carbonyl. RNA 2′-OH acylation reactions were used to investigate RNA structure and interactions, RNA functionalization and labeling, or protection of RNA from degradation. ( B ) Reversible 2′-OH acylation of mRNA enhances stability in a cloaked form, while mRNA translation can be recovered under physiological conditions through 2′-OH deprotection. Figure adapted from ref . Copyright 2023 Springer Nature. More recently, the RNA 2′-OH acylation chemistry has been explored in mRNA post-transcriptional modifications. Since the 2′-OH contributes to autolysis, blocking the 2′-OH enhances the stability of mRNA. For example, a recent study reported a reversible 2′-OH acylation method that enables the preparation of 2′-OH-cloaked mRNA using acylimidazole reagents. Subsequent treatment with water-soluble nucleophilic reagents removes the acylation adducts quantitatively, generating uncloaked mRNA and restoring its translational activity; the recovery ratio of translation largely depends on the deprotection efficiency. Thus, further mechanistic studies for the deprotection reaction may benefit broader application of this method. ( Figure B) As the first attempt to use RNA 2′-OH modification chemistry to regulate mRNA function, this work not only provides a general strategy for mRNA stabilization but also further expands the available chemical space for mRNA modifications. 2.2.3. 3′ End Chemical and Enzymatic Modification The 3′ end of mRNA, especially the poly(A) tail, is critical to translational control and RNA decay, primarily through interactions with PABP and deadenylation complexes, and thus has been an attractive target site for chemical modification and modulation of mRNA properties. The most distinct chemical signature at the 3′ end of mRNA is the 2′,3′-adenosine diol, which can be selectively oxidized to form a dialdehyde for subsequent bioconjugation reactions to improve stability and expand functionality. ( Figure A) For example, researchers treated synthetic mRNA with sodium periodate to selectively oxidize the 3′ diol, followed by condensation with hydrazines, hydroxylamines, or other nucleophiles with diverse functional groups. Subsequently, reductive amination with sodium cyanoborohydride (NaBH 3 CN) produced the labeled mRNA bearing a 3′ hexose. Additionally, this strategy can achieve specific 3′ mRNA labeling by condensing the mRNA with a click handle containing primary amine derivatives, enabling downstream labeling of fluorophores or biotin for imaging or functional studies. One caveat of this chemistry is the requirement to use uncapped mRNA or ribose-modified m 7 G cap analogs since the canonical m 7 G structure contains a 2′,3′-diol that will be nonselectively oxidized and damaged. 6. Open in a new tab mRNA 3′-OH chemical and enzymatic modification . ( A ) mRNA 3′ chemical modification through diol-oxidation and reductive amination. ( B ) Poly(A) polymerase-based enzymatic synthesis enables the spike-in of nucleotides with phosphorothioate or boranophosphate modification. ( C ) Engineered Poly(U) polymerase enables template-independent RNA synthesis at 3′ of mRNA. ( D ) pCp reagents with diverse functional groups (fluorophores, biotin, click handles) are applied for mRNA 3′ chemical modification through T4 RNA ligase-based mRNA labeling. Beyond chemical derivatization of the mRNA 3′ end, a series of enzymes, including RNA ligases and polymerases, can perform postsynthesis RNA labeling. Poly(A) polymerase (PAP) is a commercially available enzyme derived from E. coli that can recognize the 3′ end of single-stranded RNA (ssRNA) and perform template-independent synthesis of poly(A) using ATP as substrate. To explore the opportunity of using PAP for mRNA postsynthesis modification, researchers tested different modified ATP analogs as alternative substrates for poly(A) synthesis, including phosphorothioate-modified and boranophosphate-modified ATP. ( Figure B) However, partial substitution of ATP with ATPαS (Adenosine-5′-(α-thio)-triphosphate) impeded the synthesis of the poly(A) tail due to incompatibility of modified ATP analogs with PAP, leading to a significantly decreased yield when the ratio of ATP to ATPαS reached 1:1. The same issue arose in the case of boranophosphated ATP substitution. Although the PAP-based tail extension and modification provides a convenient method to introduce region-specific modification, it suffers from the intrinsic issue of generating heterogeneous products with mixed lengths of poly(A) tail. Recently, a similar method has been developed using poly(U) polymerase (PUP) mutants to perform 3′ end modification in a template-independent manner. ( Figure C) Instead of random spike-ins of NTPs, the reversible 3′-O blocking group was first installed onto the NTPs, then incorporated into 3′ of RNA by PUP one at a time. After the deblocking step, free 3′-OH was then available for the next round of 1-nucleotide extension. This sequential single-nucleotide synthesis method enables site-specific RNA modification at the 3′ end. Beyond polymerase-based mRNA 3′ end labeling, RNA ligases or nucleotidyltransferases are alternative options for introducing modifications. ( Figure D) For instance, mononucleotide 3′,5′-bisphosphates (pNps) have been demonstrated to be effective donors for T4 RNA ligase I, which couples pNps to the 3′ end of mRNA. By installing chemical modifications to the 3′ of pNp reagents, a variety of functional decorations, including fluorophores, click handles, free amines, and biotin, can be introduced to the mRNAs after the RNA ligase-mediated labeling reaction. Meanwhile, nucleotidyltransferases (e.g., CutA) catalyze the coupling of one or two additional nucleotides to the 3′ of mRNA by using NTPs as substrates, demonstrating comparable efficiency as pNps-based mRNA labeling methods. Together, these tools enable diverse 3′ end modifications for site-specific tagging and functional analyses. 2.2.4. tRNA-Modifying Enzyme for Site-Specific Modification tRNAs are the most extensively modified RNA species with over 85 known modifications installed by tRNA-modifying enzymes, modulating anticodon–codon recognition and regulating tRNA structure. Inspired by the pairs of naturally occurring tRNA modifications and their corresponding modifying enzymes, researchers have repurposed tRNA-modifying enzymes to label RNAs. For example, tRNA Ile2 -agmatidine synthetase (Tias) can recognize its counterpart tRNA Ile2 and transfer agmatine, a polyamine also present in mammalian nervous systems, to the wobble position cytosine 34 of tRNA Ile2 anticodon for deciphering AUA and distinguishing it from AUG in both bacteria and archaea. , Similar systems exist in eukaryotes, where the tRNA modifications are critical for codon recognition and specificity. By conjugating the RNA of interest with the core motif from tRNA Ile2 , diverse chemical modifications can be introduced to the RNA of Interest (ROI) by utilizing the agmatine derivatives and Tias. ( Figure A) In addition, tRNA guanine transglycosylase (TGT) derived from E. coli has been adapted for RNA site-specific modifications. It has been demonstrated that TGTs are capable of performing transglycosylation reactions, where a guanine at the wobble position of the anticodon loop is exchanged with 7-deazaguanine derivatives (PreQ1). Taking advantage of this endogenous mechanism, the 17 nt hairpin recognition motif from tRNA was inserted into the reporter mRNA (e.g., mCherry) while PreQ1 derivatives with different functional groups, such as fluorophore or biotin, were synthesized and utilized as substrates for TGT-catalyzed transglycosylation reaction. , ( Figure B) These methods provide versatile site-specific modifications based on known RNA sequence motifs, which are convenient to incorporate into the DNA template during IVT. However, such an approach relies on the introduction of structured motifs, whose folding and labeling efficiency depend on the variable sequence context of the entire mRNA. For example, the surrounding mRNA sequence context can disrupt the motif’s functional structure, thereby reducing its reactivity in labeling reactions. 7. Open in a new tab mRNA labeling with tRNA-modifying enzymes . ( A ) tRNA Ile2 -agmatidine synthetase (Tias) was used to introduce chemical modifications at Cytosine 34 (C34) of the tRNA Ile2 motif. RNA chemical derivatization is achieved through fusing the RNA of interest (ROI) with a tRNA motif and enzymatic labeling reactions (e.g., azide–alkyne click chemistry). ( B ) tRNA guanine transglycosylase (TGT) catalyzes the transglycosylation reaction between PreQ1 and the fused-tRNA motif within the ROI, which enables postsynthesis chemical labeling of RNA with biotin or various fluorophores. 2.2.5. Genetic Code Expansion for Site-Specific Modification Genetic code expansion technologies introduce unnatural base pairs (UBPs) into DNA templates, enabling site-specific incorporation of noncanonical nucleotides into RNA during IVT. Through extensive structural screening and optimization, researchers discovered several UBPs that function as third base pairs compatible with DNA replication, transcription, and mRNA translation. This achievement also led to the generation of engineered bacteria that perform DNA replication, RNA transcription using unnatural NTPs for survival and replication. , The development and broad applications of this technology have been systematically reviewed previously, , so we will focus only on applications related to mRNA modifications. Inspired by the compatibility of UBPs with transcription, including T7-based IVT, noncanonical NTPs (e.g., TPT3-TP, NaM-TP analogs), and DNA templates with UBPs were used in IVT reactions to enable site-specific introduction of functional groups to synthetic RNAs for structural, functional, and labeling studies. − For instance, click handles, including norbornene or cyclopropene moieties for inverse electron-demand Diels–Alder (iEDDA) cycloaddition, were covalently attached to the base group on the side opposite base pairing. Therefore, the click handles can be inserted into any specific sites of RNA molecules cotranscriptionally. ( Figure ) Following this line, this method has been applied to label mRNA with an internal fluorophore for intracellular mRNA visualization via in situ iEDDA. In addition, nitroxide spin label-containing RNA has been synthesized using modified TPT3 triphosphates, which enables RNA structural analysis measured by pulsed electron paramagnetic resonance (EPR) spectroscopy. In summary, genetic code expansion technology enables site-specific RNA labeling via one-step IVT, especially internal RNA modifications that could be challenging to achieve by other existing technologies. However, this method remains limited by the challenge of synthesizing NTPs containing unnatural bases, which hinders its broader application to therapeutic mRNAs. Furthermore, the discovery of novel natural base modifications that enhance incorporation efficiency and synthesis fidelity will be essential to fully achieve the potential of this emerging technology. 8. Open in a new tab Genetic code expansion for mRNA site-specific labeling . DNA templates with unnatural base pairs are used in the IVT reaction to synthesize mRNA with site-specific unnatural nucleotides by incorporating unnatural NTPs (e.g., TPT3TP analogs), which enables the installation of single or multiple unnatural base groups with functional groups (e.g., EPR probes or iEDDA handles) for mRNA structural studies or chemical labeling. 2.3. Total Chemical Synthesis of mRNA 2.3.1. Introduction to Solid-Phase Oligonucleotide Synthesis Solid-phase synthesis was first introduced to peptide synthesis in 1963, and then applied to DNA synthesis in the 1980s through the ground-breaking development of phosphoramidite chemistry. , In the typical solid-phase 3′-5′ synthesis of oligonucleotides, nucleoside building blocks (phosphoramidites) are sequentially added to a growing chain anchored to a solid support, typically controlled pore glass (CPG). Each synthesis cycle includes: (1) deblocking the 5′-dimethoxytrityl (DMTr) group on the nucleoside linked to CPG to provide a free 5′-OH, (2) coupling the free 5′-OH with the incoming 3′-phosphoramidite monomer by activator reagents, (3) capping unreacted 5′-OH groups on the CPG to prevent side reactions, (4) oxidation to convert P(III) into stable P(V) linkages. After synthesis, the desired oligonucleotides are cleaved from the CPG, while the protection groups on bases with exocyclic primary amines and phosphate backbone are then removed under basic conditions. For RNA synthesis, 2′-OH groups are protected by groups like tert-butyldinethylsilyl (TBDMS), which are deprotected by treating the oligonucleotide with hydrofluoric acid at the end of synthesis. ( Figure A) Modern optimizations of phosphoramidite chemistry, including the optimization of coupling activators, base-protecting groups, oxidation/sulfurization reagents, and instrument automation, enable DNA synthesis up to 300 nt and RNA up to 100 nt with acceptable yield and high fidelity, and have been summarized in other references. In comparison to enzymatic synthesis, chemical synthesis tolerates most of the chemical modifications through preparing phosphoramidite-derivatives with ribose, base, and phosphate backbone modifications. 9. Open in a new tab Total chemical synthesis for mRNA modification . ( A ) Solid-phase oligonucleotide synthesis technology with phosphoramidite chemistry involves the coupling and deprotection cycle, including monomer activation/coupling, failure sequence capping, oxidation, and detritylation to extend the oligonucleotide chain on a solid-phase support, followed by cleavage and deprotection to yield target products. Modifications can be introduced into synthetic RNAs by incorporating chemically modified phosphoramidite monomers (e.g., base or ribose modification) during the coupling step of solid-phase synthesis. ( B ) The combination of solid-phase oligonucleotide synthesis and chemical capping reaction enables the total chemical synthesis of minimal mRNAs (up to 110 nt). This method is compatible with chemically modified nucleotides with available phosphoramidite monomers and enables site-specific mRNA chemical modification. 2.3.2. Total Chemical Synthesis for mRNA Modification While total chemical synthesis has long been applied to therapeutic small RNAs (e.g., siRNAs, antisense oligonucleotides), its application for mRNA production is rarely explored due to two main challenges: (1) most mRNAs contain hundreds to thousands nucleotides, well beyond the typical range for chemical synthesis (∼100 nt) and (2) the unique chemical structures of mRNA (e.g., m 7 G cap structure) is usually challenging to synthesize, requiring special chemistries and multiple steps of postsynthesis modification. Pioneering work in mRNA chemical synthesis began in 1992, when researchers synthesized an m 7 G-capped 9-nt RNA oligonucleotide by combining solid-phase oligonucleotide synthesis with postsynthesis chemical capping. The 5′ monophosphate of a synthetic RNA oligonucleotide was converted into a phosphorimidazolide derivative by activating the oligonucleotide with carbonyldiimidazole, followed by a condensation reaction with m 7 GDP to provide an m 7 G-capped 9-nt oligonucleotide. Based on similar chemistry, the m 7 GDP-imidazole derivative was prepared to couple with the 5′ monophosphate of a synthetic oligonucleotide in the presence of Mn 2+ or Mg 2+ as catalysts. , Beyond imidazole-derivatives as the activated form of m 7 GDP, other capping reagents were also developed including the chloroquinoline-derivatives, which can be activated by CuCl 2 to couple with the monophosphate of RNA 5′ end. In addition, m 7 G-capped oligonucleotides have been achieved by combining solid-phase synthesis and enzymatic capping system through treating the synthetic 5′ diphosphate RNA with capping enzyme and 2′-O-methyltransferase. To facilitate high-quality purification of 5′ capped oligonucleotides, solid-phase RNA capping was achieved by treating the free 5′ monophosphate of RNA oligonucleotide attached to the CPG with m 7 GDP-imidazole capping reagents. , An alternative method employs a reversible DMTr-protected capping reagent, which is compatible with affinity-based purification of the capped oligonucleotide products after the capping reaction. Recently, the first example of translatable mRNA prepared through total chemical synthesis was reported. A series of 107 nt RNA oligonucleotides encoding FLAG- and His-tag proteins was prepared through solid-phase synthesis, followed by chemical capping reaction with m 7 GDP-imidazole derivatives. ( Figure B) This chemical method is compatible with a wide range of chemical modifications at any site of the synthetic minimal mRNA, including canonical and noncanonical nucleotides and the introduction of even unnatural structures such as PEG, thereby expanding the scope of investigation into site-specific mRNA modifications. Although still constrained by the challenges of synthesizing long RNA molecules for functional studies and the potential carryover of impurities from side reactions during solid-phase synthesis, total chemical synthesis holds great promise for producing RNA with a broad scope of site-specific chemical modifications that enzymatic synthesis cannot achieve. 2.4. Chemoenzymatic Synthesis (Hybrid Synthesis) of Modified mRNA Chemoenzymatic, or hybrid, synthesis combines the scalability of enzymatic transcription with the structural versatility of chemical synthesis. This approach overcomes the limitations of either method used in isolation: enzymatic IVT synthesis offers unlimited transcript length but narrow modification scope, while chemical synthesis enables broad modification types but is constrained by oligonucleotide length. Hybrid strategies enable efficient, modular, and site-specific assembly of mRNAs with diverse natural and unnatural modifications, facilitating the investigation of structure–activity relationships and expanding the design space for therapeutic mRNAs. 2.4.1. mRNA-oligo Conjugates for 3′ Modification Previous efforts on chemical or enzymatic labeling of the mRNA 3′ terminus enabled chemical modification at the terminal nucleotide. However, site-specific modification beyond the terminal nucleotide was challenging due to the lack of synthesis methods for mRNA internal modification. To address this, our group developed the messenger-oligonucleotide conjugated RNAs (mocRNAs) technology, which leverages both enzymatic mRNA synthesis and chemical oligonucleotide synthesis to enable modular, site-specific modifications. ( Figure A) In this strategy, a chemically synthesized 30-nt oligonucleotide containing 5′ phosphate, diverse modifications, and structural motifs was ligated to the 3′ end of an IVT mRNA using T4 RNA ligase I. This allows precise incorporation of: backbone modifications (e.g., phosphorothioates), ribose modifications (e.g., 2′-O-methyl, 2′-F), chain-terminating residues (e.g., ddC, inverted dT), and structural motifs (e.g., G-quadruplexes). This method enables precise, modular encoding of chemical modifications into RNA vectors with expanded modification sites, a feature that cannot be achieved with prior 3′ modification methods. 10. Open in a new tab Chemoenzymatic synthesis (hybrid synthesis) of modified mRNA . ( A ) Messenger-oligonucleotide conjugated RNAs (mocRNAs) are synthesized through T4 RNA ligase I-mediated enzymatic ligation between IVT mRNA and a chemically synthesized oligonucleotide, which enables the incorporation of a variety of modifications at the 3′ end of mRNA, including linkage modifications, sugar backbone modifications, chain-terminating nucleotides, or RNA motifs. Figure adapted from ref . Copyright 2022 American Chemical Society. ( B ) Ligation-enabled mRNA-oligonucleotide assembly (LEGO) enables modular assembly of multipiece oligo/RNA for site-specific mRNA modification or topological engineering. ( C ) LEGO can be used to construct a modified mRNA library with 5′ site-specific modifications, including: cap modification, base modification, linkage modification, and sugar backbone modification. Figures B and C adapted from ref . Copyright 2024 Springer Nature. ( D ) Site-specific mRNA internal modification is achieved through T4 RNA ligase II-mediated splint ligation between enzymatically synthesized mRNA fragments and chemically synthesized oligonucleotide with specific modifications. 2.4.2. Modular Multipiece mRNA Assembly by RNA LEGO The cap and the 5′ UTR have been shown to regulate both translation initiation and mRNA stability, highlighting their potential for chemical modification. However, both cotranscriptional capping and enzymatic capping are restricted to modifications tolerated by RNA polymerases and capping enzymes, and are challenging to extend the modification beyond the first two bases. To tackle this problem and further expand the modification scope of synthetic mRNAs, our group reported a modular and divergent chemo-enzymatic synthesis strategy named ligation-enabled mRNA-oligonucleotide assembly (LEGO), which allows access to potentially any modifications of interest available from chemical synthesis. ( Figure B) This approach assembles full-length mRNAs from modular components: (1) a chemically synthesized 5′ fragment (e.g., 12-nt segment of the 5′ UTR) with translation-enhancing or RNA-stabilizing modifications followed by chemical capping with various capping reagents (e.g., m 7 GDP-imidazole derivatives), (2) an IVT-synthesized internal fragment (e.g., containing Ψ, m 1 Ψ, or other functional nucleotides), and (3) a 3′ chemically synthesized oligo bearing stability-enhancing modifications and chain-terminating residues (e.g., ddC or inverted dT to block self-ligation). After preparing the internal IVT mRNA fragment, RppH is used to hydrolyze the 5′-triphosphate to a monophosphate or further to 5′ OH by calf intestinal alkaline phosphatase (CIAP), making it suitable for RtcB or T4 ligase 1 enzymatic ligation with the capped 5′ oligo. ( Figure B) The 3′ modified fragment is added via T4 RNA ligase 1 RNA ligation as described in Section 2.4.1 . In addition, we have extended this approach to engineer branched or circular mRNAs using tandem ligation and compatibility with RtcB ligase, which only circularizes fully assembled constructs, providing intrinsic quality control (see Section 3.1.3 ). Through such a modular assembly strategy, RNA LEGO supports: (1) broad chemical compatibility (caps, bases, sugars, linkages); (2) site-specific incorporation of unnatural motifs (e.g., branch points, clickable handles); (3) rapid design–build–test cycles for screening chemically modified mRNA libraries. ( Figure C) 2.4.3. mRNA Assembly by Splint Ligation Besides ssRNA-based ligation, multipiece RNA ligation can also be achieved by enzymatic oligo-splint ligation. Researchers reported the synthesis of EGFP mRNA with single-site 2′-O-methylation within the coding sequence using a three-way, one-pot splint ligation workflow. Briefly, the EGFP mRNA was split into three fragments: 3′ and 5′ unmodified fragments synthesized through IVT, and an internal RNA fragment with modifications prepared by solid-phase synthesis. Then, the three fragments were assembled through one-pot splint ligation using T4 RNA ligase II, followed by real-time gel elution for product purification. Although the synthesis efficiency and scalability were low due to limited ligation efficiency and sophisticated purification steps, this method enabled the investigation of internal mRNA modifications at single-nucleotide resolution. ( Figure D) In summary, chemoenzymatic synthesis merges the best of chemical and enzymatic platforms, allowing long transcript synthesis, modular assembly, and high chemical diversity. While the process is currently less scalable due to multiple ligation steps and purification demands, it provides an essential toolkit for programmable engineering of mRNA structure and function ( Table ). 2. Comparison of Different mRNA Synthesis/Modification Methodologies. Synthesis method Length limit Modification scope Modification site Scalability mRNA capping Reference IVT via RNA polymerase Up to 10k nt Only modifications compatible with RNAP Cap and first two bases or full substitution or random spike-in Easy to scale up Capping efficiency depends on the structure of cap analogs − ,, IVT via engineered DNA polymerase Up to 1000 nt Only modifications compatible with DNAP Any position on mRNA Restricted by the ssDNA preparation No cap ,,,,, Postsynthesis modification Based on the RNA substrates Depends on enzyme tolerance or the chemistries used Any position on synthetic mRNA Easy to scale up - ,,− ,,,,,,, Total chemical synthesis <150nt Any nucleotide with available phosphoramidite Any position on mRNA Easy to scale up up to 95% ,, Chemoenzymatic synthesis Up to 10k nt Any nucleotide with available phosphoramidite Any position on mRNA Scalable after optimization >98% capping efficiency after HPLC purification for short oligo − , Open in a new tab 3. Synthesis Methodology of RNA Topology 3.1. Synthesis of Circular RNA (circRNA) While most therapeutic mRNAs are linear, circular RNAs (circRNAs)covalently closed RNA moleculeshave emerged as promising alternatives due to their enhanced stability, resistance to exonuclease degradation, and potential for cap-independent translation. − Initially characterized as noncoding RNAs (ncRNAs), recent discoveries have shown that endogenous and synthetic circRNAs can be translated via internal ribosome entry sites (IRES) or N 6 -methyladenosine (m 6 A) modification to facilitate cap-independent translation. Endogenous circRNAs are usually generated through back-splicing of pre-mRNAs catalyzed by ribozymes or spliceosomal machinery and typically include an untranslated region (UTR) and several exons. Inspired by this natural process, researchers have developed various strategies to generate synthetic circRNAs for research and therapeutic applications. Favorable properties, such as improved stability against exonuclease degradation, enhanced thermodynamic stability, and compatibility with rolling circle amplification (RCA), have motivated the application of circRNAs in molecular devices, biosensing, and drug delivery. , Here we classify and review current strategies for synthetic circRNA production by their mechanisms and compatibility with additional chemical and topological modifications ( Table ). 3. Comparison of Different Methodologies for Circular RNA Synthesis. Method RNA length Compatibility with nucleoside modifications Scalability Compatibility with topological engineering Chemical circularization <150nt Yes Low circularization efficiency and difficult to purify theoretically compatible Backsplicing Any length No, only natural nucleotides that will not interfere with RNA secondary structures can be used High No Autocatalytic ribozyme assisted circularization short aptamer or IRES-containing RNA No, only natural nucleotides that will not interfere with RNA secondary structures can be used Low (limited to in cellulo synthesis) No Enzymatic ligation <1000nt Yes, modified nucleotides are compatible with RNA polymerase Relatively low No Chemoenzymatic synthesis Any length Yes, any modified nucleotides Scalable after optimization Yes Open in a new tab 3.1.1. Chemical Circularization The chemistry for oligonucleotide circularization and ligation has been investigated for several decades. Early attempts include using highly reactive cyanogen bromide (BrCN) together with morpholino derivatives as activators for linking two oligonucleotide strands bearing terminal 5′ hydroxyl and 3′ phosphate. , Subsequently, other condensing agents, such as ethyl-3-(3′-dimethylaminopropyl) carbodiimide (EDC), have been used to support phosphodiester bond formation, yet with variable circularization yields ranging from 20% to 95% depending on secondary structure and oligonucleotide length. The efficiency of chemical ligation for RNA oligomers was even lower than that of DNA counterparts, possibly due to steric hindrance of the 2′-hydroxyl group with neighboring phosphate groups, unfavorable N-type conformation of ribose, and side reactions of 2′,3′-cyclophosphate formation. , Circularization yields may be further reduced by undesirable intermolecular ligation and formation of 2′,5′-phosphodiester linkages that may be circumvented by using a 2′-deoxy sugar moiety at the 3′-terminal. More recent work demonstrated a novel chemical circularization strategy using 3′ amine-phosphate coupling chemistry activated by EDC and HOBt, achieving both two-step or one-pot chemical synthesis of a 126-mer circular RNA with the help of splint DNAs. With reaction yields of up to 32.4%, this work demonstrated the first example of a translatable circular RNA prepared through chemical circularization. ( Figure A) The one-pot intermolecular reaction notably prefers higher RNA concentrations at 20 μM, which may facilitate scale-up by circumventing traditional limitations of using low RNA concentration (<1 μM) to minimize undesirable competing polymerization. To date, chemical circularization is generally compatible only with the synthesis of small circRNAs (<150 nt), which cannot meet the typical length requirement for circRNAs encoding functional proteins in therapeutic applications. Additionally, multiple steps of chemical treatment may damage RNA integrity, leading to undesired byproducts. 11. Open in a new tab Synthesis methodologies for circular RNA (circRNA) . ( A ) Amine-phosphate coupling reaction-based chemical circularization enables the synthesis of circular RNAs. ( B ) Group I introns are engineered to synthesize circular RNAs via backsplicing. ( C ) Autocatalytic ribozymes are utilized to generate RNA fragments with 5′ free hydroxyl and 2′,3′-cyclic phosphate, which can be ligated by Rtcb to form the circular RNAs. ( D ) T4 RNA ligase or T4 DNA ligase-mediated ligation enables the circularization of mRNA with a homology arm or splint probe. ( E ) LEGO enables the synthesis of circular RNA through two-step ligation reactions: (1) T4 RNA ligase-mediated ligation between a synthetic oligo with a 5′ phosphate and 3′ hydroxyl group of IVT mRNA; (2) Rtcb ligase-mediated ligation between 3′ phosphate and a 5′ free hydroxyl group of mRNAs. Figure adapted from ref . Copyright 2024 Springer Nature. 3.1.2. Backsplicing for Circularization Distinct splicing mechanisms of pre-mRNA in eukaryotic cells generate two types of endogenous circRNA: exonic circRNAs and intronic circRNAs. Inspired by the natural biogenesis of circRNAs, several intron elements have been exploited for the synthesis of artificial circRNAs. The first example was a Group I intron reported in 1992 using the permuted introns and exons (PIE) strategy, which enabled the generation of circRNA both in vitro and in vivo . In practice, the desired circRNA sequence (as ‘exon’) can be placed between flanking half-intron sequences of the group I intron self-splicing system, followed by the back-splicing reaction to circularize the exon sequence. Mechanistically, the Group I intron-based method includes two transesterification reactions. The first reaction of the 3′ hydroxyl group of guanosine nucleotide reacting at the 5′ splice site leads to the release of the 3′ terminal sequence (5′-half intron). Then, the newly generated 3′ hydroxyl group attacks the 3′ splice site in the second transesterification reaction, resulting in the circularization of the engineered exogenous RNA and the release of the 3′ half intron ( Figure B). Group I introns from different species have been utilized to synthesize artificial circular RNAs for functional studies or therapeutic applications. For instance, Anabaena catalytic intron together with a strong homology arm, could facilitate back-splicing with an efficiency of up to 95%, which was further applied to the design and synthesis of circRNA vaccines by increasing the length of the circularization sequence to 5 kb and improving protein expression stability. Meanwhile, Group I intron from phage T4 thymidylate synthase (Td) was used to prepare circRNA for innate immune response evaluation and circRNA regulatory elements optimization. ,,, While PIE and similarly RNA-catalyzed methods are compatible with larger RNAs than chemical and/or enzymatic ligation strategies, they rely on large RNA catalytic motifs, which usually leave a scar sequence in the final product. Ongoing investigations into the mechanisms of group I splicing may further improve circularization efficiency and expand the modification landscape. 3.1.3. Autocatalytic Ribozyme for Circularization In Cellulo Furthermore, autocatalytic ribozymes have also been exploited to generate circRNA in cellulo . The Tornado (Twister-optimized RNA for durable overexpression) expression system has been developed to mimic the tRNA (tRNA) splicing process and achieved the synthesis of ribozyme-assisted circular RNA (racRNA). Briefly, a 5′ P3 Twister U2A and a 3′ P1 Twister undergo autocatalytic cleavage to generate free hydroxyl at the 5′ end and 2′,3′-cyclic phosphate at the 3′ end, respectively, followed by enzymatic ligation of cellular RtcB with the help of tRNA-like homology arm. , ( Figure C) These methods enable efficient synthesis of circular RNAs in cellulo so far, thus providing a powerful tool for the functional studies of endogenous circRNAs and for broadening the therapeutic applications of exogenous circRNAs. However, in terms of synthesizing therapeutic circRNAs, both backsplicing and autocalalytic ribozyme approaches suffer from incompatibility with modified nucleotides (e.g., m 1 Ψ), which alter intron or ribozyme secondary structures and sabotage splicing activity. Thus, the development of ribozymes or intron elements that are compatible with modified nucleotides will be beneficial for exploring modified circRNAs as therapeutics in the future. 3.1.4. Enzymatic Ligation-Based Circularization Several protocols have been developed for the synthesis of circRNA using different polynucleotide ligases (e.g., T4 DNA ligase, T4 RNA ligase I, and T4 RNA ligase II) that can ligate nicks in single- or double-stranded RNA constructs. These ligases catalyze the formation of phosphodiester bonds between 5′ monophosphate and 3′ hydroxyl of the RNA molecules in an ATP-dependent manner. For instance, an early report demonstrated, for the first time, the application of T4 RNA ligase to the synthesis of circRNA with any sequence by using a homology arm to bring the ends of RNA together. To further facilitate the circularization of mRNA, splint DNAs were used to preanneal with both ends of mRNA, followed by T4 RNA ligase-based ligation. ( Figure D) Recently, enzymatic ligation was used to synthesize circular RNA vaccines encoding RBD antigen for SARS-CoV-2, which demonstrated the potential of using RNA ligase-based methods for the production of therapeutic circRNAs. In comparison to the back splicing-based circularization methods, enzymatic ligation is compatible with RNA modifications, which has been proven to benefit RNA therapeutics from several aspects. However, the limitations of this method include: (1) limited scalability for large-scale construct preparation given low RNA substrate concentrations are usually necessary to minimize the potential issue of intermolecular ligation; (2) extra elements such as homology arms or splint DNAs are usually required to facilitate the ligation process by bringing the ends together, which requires careful individualized optimization to achieve decent circularization efficiency. 3.1.5. Chemoenzymatic Circularization To address the challenge of enabling RNA circularization without intron back-splicing and generalized circRNA synthesis with modified nucleotides (e.g., m 1 Ψ), our group adopted a modified RNA LEGO procedure for site-specific chemical/topological engineering of circRNA. Accordingly, tandem ligation of a chemically synthesized 5′/3′-phosphorylated oligonucleotide was performed to an IVT 5′/3′-hydroxyl mRNA using T4 RNA ligase I and RtcB ligase, where the circularization reaction was facilitated by 5′/3′ UTR-encoded homology. In addition to the compatibility with chemical modifications, including both modified nucleotides spike-in during IVT and unnatural modifications/topological elements (e.g., branch strands) for the synthetic oligonucleotides for ligation, the use of RtcB ligase added an intrinsic quality control to the circRNA synthesis workflow, where only the RNA with successful ligation of capped oligo bearing 3′ phosphate could be circularized. ( Figure E) This method enables the screening of site-specific chemical modifications in circRNA and provides a platform technology for more sophisticated circRNA engineering. 3.2. Synthesis of Branched RNAs In addition to circular topologies, branched RNA topologies have also been explored over the past few decades, particularly through studies on the biogenesis and function of lariat RNAsnaturally occurring branched RNA species formed during the splicing process. Inspired by the unique topology increasingly shown to endow stability and subsequent physiologic function, our group and others introduced the concept of topological engineering for therapeutic mRNAs as another structural dimension to regulate their properties. This section reviews current strategies for the synthesis of branched RNA constructs, organized by the diverse methodologies for introducing the branching structure ( Table ). 4. Comparison of Different Methodologies for Branched RNA Synthesis. Branching type Linkage Synthesis methods Pros Cons Branching via backbone 2′, 5′-phosphodiester bond deoxyribozymes mediated synthesis or solid phase synthesis with orthogonal protection groups Mimic the natural structure of branched and lariat RNA susceptible to endogenous debranching mechanism (e.g., Dbr1); incompatible with chemical modification (for deoxyribozymes) Branching via base modification Click handle (triazole formed between 5-octadiynyl deoxyuridine and azide) or other modifications Click chemistry (e.g., CuAAC, IEDDA) or other reaction High stability due to the unnatural linkage; compatible with modifications Require specific modifications (e.g., click handles) Branching via noncovalent hybridization Hybridization through base-pairing Direct hybridization Easy to synthesize; high programmability Less stable due to the noncovalent linkage Open in a new tab 3.2.1. Branching via Backbone Lariat RNA species form naturally during Group II introns-mediated mRNA splicing to generate a 2′,5′-branched RNA, where the 2′-hydroxyl of an internal nucleotide and the 5′-hydroxyl of another nucleotide are linked through a phosphodiester bond to form a branching strand in addition to the conventional strand through 3′,5′ linkage. Synthetic mimics of this branching pattern have been constructed using both enzymatic and chemical strategies. Using an in vitro selection assay, researchers identified several deoxyribozymes that anneal with two RNA fragments and catalyze the formation of a phosphodiester bond between the 2′-hydroxyl of an internal nucleotide on the 5′ fragment and 5′-triphosphate on the 3′ fragment. − ( Figure A) In addition to the deoxyribozymes-mediated synthesis method, solid-phase synthesis has also been exploited for the construction of branched oligonucleotides. Based on the principle of phosphoramidite chemistry (described in Section 2.3.1 ), extension of the oligonucleotide chain depends on iterative deprotecting and coupling cycles. To enable the introduction of a branching point at the 2′ position of an internal nucleotide, orthogonal protection groups for either the 2′-hydroxyl group or the 2′-amine group were developed for selective removal. After the synthesis of the stem strand, the protection group was selectively removed (2′-acetal levulinic ester for the 2′ hydroxyl group and fluorenylmethyl-oxycarbonyl for the 2′ amine) to expose the hydroxyl group/amine for branched strand synthesis. ( Figure B) Mimicking the natural structure of branched and lariat RNAs, the linkage through 2′,5′ phosphodiester bond is the substrate for endogenous lariat debranching enzyme (Dbr1), which hydrolyzes the branched RNA in cellulo . − Although ideal for probing the endogenous function of lariat RNA, branching via naturally occurring 2′,5′ phosphodiester bond suffers from intrinsic stability issues when applied to therapeutic RNA engineering. 12. Open in a new tab Synthesis of Branched RNAs . ( A ) Deoxyribozymes are utilized to catalyze the reaction between 2′-OH of an internal nucleotide with 5′ triphosphate to form the 2′-5′ branching structure. ( B ) Solid-phase synthesis enables direct synthesis of branched oligonucleotides by incorporating a brancher phosphoramidite. ( C ) Click chemistry is used to conjugate an internal 5-octadiynyl deoxyuridine with an azide group within another oligonucleotide to form a branched RNA. Figure adapted from refs , . Copyright 2024 Springer Nature. ( D ) Noncovalent hybridization is used to introduce a branched cap, a branched poly(A) tail, or regulatory elements. 3.2.2. Branching via Base Modification To overcome the instability of 2′,5′-linked branches, we and others have explored unnatural branching structures using chemically stable linkages for therapeutic mRNA engineering. Our group reported the synthesis of multitailed mRNA via chemoenzymatic assembly of a branched poly(A) oligonucleotide with IVT mRNA. To start with, a branched poly(A) tail was assembled through copper-catalyzed azide–alkyne cycloaddition (CuAAC) between an internal 5-octadiynyl deoxyuridine (OU) containing oligonucleotide and a 5′ azide containing oligonucleotide. Alkyne handles were introduced internally into chemically synthesized oligonucleotides using base-modified phosphoramidites that serve as branching points. Following a similar fashion, multicap mRNAs were synthesized using the same 5-octadiynyl deoxyuridine (OU) for the introduction of the branching point. Click chemistry was used to assemble the capped oligo with an internal OU as the stem strand and another capped oligo with a 3′-azide as the branched strand. The synthetic oligonucleotide with a branched cap was subsequently enzymatically ligated to the IVT mRNA to construct multicap mRNA. Furthermore, our group also applied the branching strategy to the synthesis of circular RNA with a cap structure to leverage the cap-dependent mechanism for efficient circular RNA translation initiation. ( Figure C) These unnatural branches are resistant to enzymatic debranching and are structurally compatible with a wide range of chemical and topological modifications, offering an expanded toolkit for mRNA design and engineering. 3.2.3. Branching via Hybridization In addition to the covalent branching strategy, a noncovalent approach can be leveraged by using base-pair hybridization. ( Figure D) This strategy relies on the design of complementary oligonucleotide sequences engineered into the RNA construct, which can stably hybridize to form functional branch-like structures. For example, our group recently reported that an intermolecular trans -acting, capped oligonucleotide hybridizing to the circRNA 5′ UTR with 20 nt complementary regions enhanced circRNA translation. Meanwhile, the hybridization branching method was recently applied to boost mRNA expression by tethering a poly(A) tail mimetic or regulatory elements to the 3′ UTR of a specifically endogenous target mRNA. , In total, the noncovalent strategy provides an extra regulatory mechanism to tune the expression of endogenous and exogenous mRNAs with a relatively short oligo component. However, noncovalent branching can dissociate under physiological conditions or be removed by RNA helicases, making it less effective than covalent branching. Despite these challenges, noncovalent hybridization remains a promising tool for introducing transient branching or enhancing RNA function in therapeutic settings ( Table ). 4. Purification Methods of Synthetic mRNA The purity of mRNA products is crucial for their activity and cytotoxicity, which will ultimately influence the final therapeutic outcome. The impurities in the mRNA products usually come from three sources: the leftover substrates, unavoidable byproducts, and unwanted side products from the preparative reaction. During in vitro mRNA synthesis, components such as DNA plasmids, RNA polymerase, nucleotide starting materials, or enzymes for mRNA modification can induce pro-inflammatory cytokines if not removed. Meanwhile, IVT reactions usually generate byproducts such as truncated transcripts, uncapped mRNA, or double-stranded RNA (dsRNA), most of which have been demonstrated to activate innate immune response pathways and inhibit translation. Therefore, rigorous purification is essential for both research applications and therapeutic-grade mRNA manufacturing. This section reviews key purification methods, emphasizing their strengths, limitations, and relevance for chemically or topologically modified RNAs. 4.1. Precipitation Purification The most commonly used method for mRNA in a laboratory setting is to precipitate the mRNA using monovalent cation-containing buffers (e.g., sodium acetate, ammonium acetate, or lithium chloride) with ethanol or isopropanol. This method removes most leftovers, such as NTP starting materials and enzymes. But it does not effectively remove RNA byproducts, including truncated fragments and dsRNAs, which negatively affect mRNA function. 4.2. Chromatography Purification Among chromatography-based methods, high-performance liquid chromatography (HPLC) is the gold standard for mRNA purification, effectively removing RNA byproducts. Reverse-phase HPLC (RP-HPLC) has been widely utilized in mRNA purification. In the RP-HPLC, the negatively charged phosphate backbone of IVT mRNAs pairs with cationic species in the mobile phase, usually quaternary ammonium compounds or amine-containing compounds (e.g., triethylamine, diethylamine, etc.), to make the mRNAs more lipophilic, allowing them to interact with the stationary phase of the reverse-phase chromatography column. Based on this mechanism, mRNAs of varying lengths exhibit different retention times depending on their lipophilicity, enabling the separation of target mRNA products from truncated RNAs or dsRNAs. According to the pioneering work on the application of mRNA for protein production, mRNA products purified through this method showed significantly lower levels of dsRNA that induce type I interferons (IFNs) or other proinflammatory cytokines, leading to 10- to 1000-fold greater translation levels in primary cells. , Additionally, HPLC purification facilitates the preparation of circRNA with minimal immunogenicity by removing unreacted linear RNAs and free introns. In light of the importance of purification to minimize undesirable immunostimulation, other chromatography-based methods have been developed, including size-exclusion HPLC (SEC), ion-exchange HPLC (IEC), affinity chromatography using oligo(dT) containing resins, and cellulose chromatography for dsRNA removal. − 4.3. RNA-Encoded Purification Handles In addition to method optimization for HPLC, structural modification of oligonucleotides or mRNAs has also been exploited to facilitate their purification. For instance, a cleavable DMTr-containing cap analog was used to separate the capped oligonucleotide from uncapped starting material based on the hydrophobicity of the DMTr group; the pNp reagent or 3′ chemical labeling reaction were used to introduce the fluorophores (e.g., Cy5) to the 3′ of mRNA, which usually induces a significant retention time shift on HPLC and facilitates mRNA purification due to the hydrophobic property of most fluorophores. , Hydrophobic photocaged tag-modified cap analogs were also developed to separate capped mRNA from uncapped mRNA by RP-HPLC. After HPLC purification, UV-treatment was used for photocleavage to recover the canonical m 7 G cap structure. This method enables 100% capping efficiency after HPLC purification with versatility applicable to 600 nt to 4000 nt mRNAs. Additionally, a trinucleotide mRNA cap analog bearing N 6 -benzylated at the first adenosine position was reported recently, demonstrating that the hydrophobic benzyl group as a purification handle also confers enhanced translational properties. ( Figure ) These methods facilitate the purification of mRNA and remove byproducts that can lead to cytotoxicity, which is especially useful when sophisticated chemical modifications are involved. 13. Open in a new tab Chemical modification facilitates the purification of mRNAs . ( A ) A trinucleotide cap analog (m 7 GpppBn 6 ApG) was used for cotranscriptional capping during IVT. The benzyl group on adenosine added extra hydrophobicity to the mRNA molecule and helped separate capped and uncapped mRNA on the reverse-phase HPLC column. Reproduced from ref . Copyright 2024 American Chemical Society. ( B ) A hydrophobic tag containing a tert -butyl (tBu) group in a 2-nitrobenzyl (Nb) photocaging molecule was attached to the cap analog. The capped mRNA product was purified by reverse-phase HPLC, followed by cleavage of the hydrophobic tag using UV treatment. Figure adapted from ref published under a Creative Commons Attribution 4.0 International License [ http://creativecommons.org/licenses/by/4.0/ ]. 4.4. Enzymatic Purification Methods Researchers are also utilizing RNase specific for the reaction byproducts as another cost-effective method for mRNA purification, even for constructs with complex modifications and topologies. One such example is the use of RNase III enzymes to remove dsRNA from the IVT product. Previous results demonstrated that RNase III treatment enhanced the protein production of modified mRNA transfected into T cells, leading to improved performance of CAR-T therapeutics. Another widely used enzyme for therapeutic RNA purification is RNase R, which specifically degrades ssRNA after the circularization reaction, yielding circRNA with higher purity. ,, More recently, the poly(A) polymerase (PAP) has also been applied for circRNA purification, where the enzyme appends a poly(A) tail on residual linear RNA impurities to allow removal of those by oligo(dT) capturing. In summary, high-purity mRNA is a prerequisite for safe and effective therapeutic application. Chromatographic techniques like RP-HPLC remain the benchmark, but novel purification strategiesespecially enzyme-aided and tag-encoded approachesare offering greater flexibility, selectivity, and scalability. 5. Structure–Activity Relationship of Chemical Modifications The therapeutic potential of mRNA depends not only on its sequence but also on its chemical composition and structural features. Chemical modifications affect nearly every aspect of mRNA behaviorfrom stability and translational efficiency to immunogenicity and pharmacokinetics. The structures used for mRNA modification derive from several resources: First, endogenous RNA modifications offer an ideal resource for modifying synthetic mRNAs; Second, unnatural modifications have proven crucial for further broadening the scope and success of oligonucleotide-based drugs, including ASOs, siRNAs, and aptamers. These modifications, such as phosphorothiolate, 2′-methoxyethyl (2′-MOE), 2′-Fluoro, phosphorodiamidate morpholino oligomer (PMO), and locked nucleic acid (LNA), were applied to enhance the stability and tune base-pairing affinity of therapeutic oligonucleotides. − Finally, the structure-guided molecular design was also applied to develop novel mRNA modifications based on knowledge of mRNA-protein interactions that regulate translation. − Although, in theory, there is a vast chemical space to be explored for RNA design, most of these approaches have not been applied to protein-encoding RNAs due to limited chemical accessibility and exploration. In this chapter, we will summarize the current progress on studies of the structure–activity relationship (SAR) for mRNA modifications, enabled by various novel synthesis methodologies. These efforts can be categorized by four regions on mRNA molecules: cap modification, coding sequence modification or full substitution, UTR modification, and tail modification. ( Figure ) 14. Open in a new tab Landscape of mRNA chemical modifications . The modifications in different regions of mRNA are summarized, including cap modification, UTR modification, CDS modification, and tail modification. 5.1. mRNA Cap Modification The m 7 G cap regulates both mRNA stability and translation, making it an important candidate for chemical modification. Thus, by using the cotranscriptional capping technology, researchers were able to introduce a variety of chemical modifications to the mRNA cap by preparing cap analog derivatives. All the reported synthetic mRNAs with cap modification are summarized in Tables –, together with their biochemical properties. For instance, dinucleotide cap analog derivatives, including modified m 7 G, triphosphate group, and the first nucleotide, were used to evaluate the influence of diverse modifications on mRNA properties. The results showed that 2′/3′- O -methylation, locked nucleic acid (LNA) analogs, or other functional groups (e.g., azide or propargyl groups) of m 7 G that favor a 3′-endo sugar pucker usually increase translation activity by enhancing interaction with eIF4E at translation initiation. ,− However, the effect of guanosine N 7 modification (e.g., benzyl group) was controversial as different reports reached opposite conclusions. ,, ( Table ) These controversial results likely arise from differences in synthesis methods, where varying reaction treatments may affect the quality of the final mRNA product, or from distinct sequence contexts that influence the performance of chemical modifications. 5. SAR Study of mRNA Cap Using Dinucleotide Cap Analogs. Entry Modification Capping efficiency (%) eIF4E binding K AS x10 –6 (M –1 ) Decapping Relative translation efficiency Ref m7G cap modification 1 m 7 G ppp G 69 12.6 - 1.00 2 Bn 7 G ppp G 79 14.6 - 1.87 (Rel. to 1 ) 3 Et 7 G ppp G 81 3.1 - 0.68 (Rel. to 1 ) 4 m 2 2,7 G ppp G 91 18.4 - 1.23 (Rel. to 1 ) 5 m 3 2,2,7 G ppp G 66 ND - 0.37 (Rel. to 1 ) 6 A ppp G - - - 0.008 (Rel. to 1 ) 7 G ppp G - - - 0.593 (Rel. to 1 ) 8 m 2 3,7 G ppp G - - - 1.00 9 Ant-m 7 G ppp G - - - - 10 Mant-m 7 G ppp G - - - - 11 N 3 -m 7 G ppp G 85 - half-life:15 min 3.7 (Rel. to 1 ) , 12 N 3 -m 7 G pppp G 72 - half-life:15 min 3.3 (Rel. to 1 ) 13 2′-N 3 -m 7 G pp s p (D1) G 80 - Resistant 7.5 (Rel. to 1 ) 14 2′-N 3 -m 7 G pp s p (D2) G 77 - Resistant 6.9 (Rel. to 1 ) 15 3′-N 3 -m 7 G pp s p (D1) G 82 - Resistant 5.9 (Rel. to 1 ) 16 3′-N 3 -m 7 G pp s p (D2) G 78 - Resistant 6.8 (Rel. to 1 ) 17 Lock Nucleic Acid-m 7 G ppp G 50 - Resistant ∼2.8 (Rel. to 1 ) 18 2′-N-m 7 G ppp G - 5.6 - 0.9 (Rel. to 1 ) 19 2′-N-Biotin-m 7 G ppp G - 3.8 - 1.0 (Rel. to 1 ) 20 Bn 7 G ppp G - K D : 217 nM Hydrolyzed 5.2 (Rel. to 1 ) 21 2-MeBn 7 G ppp G - K D : 1910 nM Resistant 1.3 (Rel. to 1 ) 22 3-MeBn 7 G ppp G - K D : 319 nM Resistant 6.4 (Rel. to 1 ) 23 4-MeBn 7 G ppp G - K D : 482 nM Hydrolyzed 4.4 (Rel. to 1 ) 24 4-iPrBn 7 G ppp G - K D : 534 nM Resistant 1.5 (Rel. to 1 ) 25 3,5-di-MeBn 7 G ppp G - K D : 646 nM Resistant 4.5 (Rel. to 1 ) 26 4-F-Bn 7 G ppp G - K D : 261 nM Resistant 8.7 (Rel. to 1 ) 27 4-Cl-Bn 7 G ppp G - K D : 221 nM Resistant 20.3 (Rel. to 1 ) 28 4-Br-Bn 7 G ppp G - K D : 172 nM Hydrolyzed 9.9 (Rel. to 1 ) 29 2,4-di-F-Bn 7 G ppp G - K D : 598 nM Resistant 5.9 (Rel. to 1 ) 30 3,4-di-F-Bn 7 G ppp G - K D : 489 nM Resistant 6.0 (Rel. to 1 ) 31 4-CF 3 –Bn 7 G ppp G - K D : 293 nM Resistant 32 4-COOH-Bn 7 G ppp G - K D : > 5000 nM Resistant 0.2 (Rel. to 1 ) 33 4-NO 2 –Bn 7 G ppp G - K D : 367 nM Resistant 7.1 (Rel. to 1 ) 34 α-Naphm-Bn 7 G ppp G - K D : 3040 nM Resistant 35 β-Naphm-Bn 7 G ppp G - K D : 1060 nM Resistant 1.0 (Rel. to 1 ) 36 Bn-m 2 2,7 G ppp G - K D : 171 nM Resistant 13.2 (Rel. to 1 ) 37 4-Cl-Bn-m 2 2,7 G ppp G - K D : 132 nM Resistant 11.2 (Rel. to 1 ) Triphosphate modification 38 Bn 7 G pppp G 76 128.2 - 1.98 (Rel. to 1 ) 39 Bn 7 m 3′‑O G pppp G 64 70.7 - 2.87 (Rel. to 1 ) 40 m 2 2,7 G pppp G 92 107.0 - 1.29 (Rel. to 1 ) 41 m 3 2,2,7 G pppp G 70 - - 0.52 (Rel. to 1 ) 42 Bn 7 m 2 G pppp G 81 188.9 - 2.55 (Rel. to 1 ) 43 m 7 G pppp m 7 G 74 47.0 - 3.14 (Rel. to 1 ) 44 m 7 G pppp m 7 G - - - 0.176 (Rel. to 1 ) 45 m 7 G ppp s G - 23.6 Hydrolyzed - 46 m 7 G pp s p G - 45.0 Hydrolyzed - 47 m 7 G p s pp (D1) G - 30.8 Resistant - 48 m 7 G p s pp (D2) G - 10.3 Resistant - 49 m 2 2,7 G pp s p (D1) G - - - 3.884 (Rel. to 1 ) 50 m 2 2,7 G pp s p (D2) G - - - 4.042 (Rel. to 1 ) 51 m 2 2,7 G pp s p (D2) G 79 19.3 Resistant 1.8 (Rel. to 4 ) 52 m 2 3,7 G ppCH 2 p G 85 4.4 Resistant - 53 m 2 2,7 G pp Se p (D1) G 59 38.5 half-life:15 min 0.6 (Rel. to 4 ) 54 m 2 2,7 G pp Se p (D2) G 57 19.0 Resistant 1.4 (Rel. to 4 ) 55 m 2 2,7 G pp BH3 p (D1) G 63 39.4 Resistant 0.5 (Rel. to 4 ) 56 m 2 2,7 G pp BH3 p (D2) G 73 13.2 Resistant 1.0 (Rel. to 4 ) 57 m 7 G pp BH3 p m 7 G 79 11.1 Resistant 1.7 (Rel. to 4 ) 58 m 2 2,7 G ppNHp G 87 10.2 Resistant 0.5 (Rel. to 4 ) 59 m 2 2,7 G pNHpp G 88 18.5 Hydrolyzed 0.5 (Rel. to 4 ) 60 m 7 G pp BH3 p (D1) G - - Resistant - 61 m 7 G pp BH3 p (D2) G - - Resistant - 62 m 7 G p BH3 pp (D1) G - - Hydrolyzed - 63 m 7 G p BH3 pp (D2) G - - Resistant - 64 m 7 G pp BH3 p m 7 G - - Resistant 0.83 (Rel. to 8 ) 65 m 2 2,7 G pp BH3 p (D1) G - - - 2.25 (Rel. to 8 ) 66 m 2 2,7 G pp BH3 p (D2) G - - - 1.66 (Rel. to 8 ) 67 m 2 2,7 G pp s p (D1) G - 42.1 - - 68 m 2 2,7 G pp s p (D2) G - 18.3 - 1.91 (Rel. to 1 ) 69 m 2 2,7 G pppp G - - - 0.11 (Rel. to 70 ) 70 m 2 2,7 G pp s p (D1) G - - - 1.00 71 m 2 2,7 G pp s p (D2) G - - - 0.64 (Rel. to 70 ) 72 m 2 2,7 G pp s pp (D1/D2 mix) G - - - 0.20 (Rel. to 70 ) 73 m 2 2,7 G ppp s p (D1/D2 mix) G - - - 0.28 (Rel. to 70 ) 74 m 2 2,7 G pppp s (D1) G - - - 0.62 (Rel. to 70 ) 75 m 2 2,7 G pppp s (D2) G - - - 0.61 (Rel. to 70 ) 76 m 2 2,7 G pp s p s (D1/D2 mix) G - - - 1.60 (Rel. to 70 ) 77 m 2 2,7 G pp s p s (D3) G - - - 1.20 (Rel. to 70 ) 78 m 2 2,7 G pp s p s (D4) G - - - 1.50 (Rel. to 70 ) 79 m 2 2,7 G pp s p s p (D1/D2 mix) G - - - 1.56 (Rel. to 70 ) 80 m 2 2,7 G pp s p s p (D3/D4 mix) G - - - 0.40 (Rel. to 70 ) 81 m 2 2,7 G ppp s p s (D1/D2 mix) G - - - 1.79 (Rel. to 70 ) 82 m 2 2,7 G ppp s p s (D3/D4 mix) G - - - 1.57 (Rel. to 70 ) 83 m 2 2,7 G ppp-triazole-C 2 H 4 NHpG G 41 24.7 - 0.16 (Rel. to 8 ) 84 m 2 2,7 G ppp-triazole-C 2 H 4 NHppG G 38 82.9 - 0.45 (Rel. to 8 ) 85 m 2 2,7 G ppp-triazole-C 2 H 4 OpG G 42 49.9 - 0.66 (Rel. to 8 ) 86 m 2 2,7 G ppp-triazole-C 2 H 4 OppG G 30 87.8 - 0.51 (Rel. to 8 ) 87 m 2 2,7 G pp-triazole-C 2 H 4 OppG G 38 24.1 - 0.30 (Rel. to 8 ) 88 m 7 G triazole-ppp G 80 2.24 - 0.11 (Rel. to 8 ) 89 m 7 G triazole-pppp G 77 8.6 - 0.15 (Rel. to 1 ) 90 m 7 G ppS G - 0.82 Hydrolyzed - 91 m 7 G pppS G 82 14.1 Hydrolyzed 0.9 (Rel. to 1 ) 92 m 7 G Spp G - 1.47 Resistant - 93 m 7 G Sppp G 86 19.8 Resistant 1.08 (Rel. to 1 ) 94 m 7 G SppCH 2 p G 82 7.54 Resistant 0.56 (Rel. to 1 ) 95 m 2 2,7 G Sppp G 70 18.2 Resistant 1.73 (Rel. to 1 ) 96 m 2 2,7 G pppS G 76 10.6 Hydrolyzed 2.23 (Rel. to 1 ) 97 m 7 G CH 2 ppS G 88 5.87 Resistant 0.56 (Rel. to 1 ) 98 m 7 G SpppS G 84 10.5 Resistant 1.09 (Rel. to 1 ) 99 m 7 G pp s pS (D1) G 88 41.0 Hydrolyzed 1.22 (Rel. to 1 ) 100 m 7 G pp s pS (D2) G 95 23.8 Hydrolyzed 1.46 (Rel. to 1 ) 101 m 7 G Spp s p (D1) G 98 25.7 Resistant 1.07 (Rel. to 1 ) 102 m 7 G Spp s p (D2) G 95 18.6 Resistant 1.50 (Rel. to 1 ) 103 m 7 G Spp s pS (D1) G 88 33.2 Resistant 1.18 (Rel. to 1 ) 104 m 7 G Spp s pS (D2) G 79 17.3 Resistant 1.20 (Rel. to 1 ) First nucleotide modification 105 m 7 G ppp 2′ dG 81 - - 0.46 (Rel. to 1 ) 106 m 7 G ppp 2′ OMe-G 86 8.0 - 0.08 (Rel. to 1 ) 107 m 7 G ppp m 7 G 73 3.7 - 2.66 (Rel. to 1 ) 108 m 2 2,7 G ppp 6S G - 16.7 - 0.45 (Rel. to 1 ) 109 m 2 2,7 G pp s p (D1) 6S G - 54.6 - 0.60 (Rel. to 1 ) 110 m 2 2,7 G pp s p (D2) 6S G - 27.0 - 0.68 (Rel. to 1 ) Open in a new tab a Translation activity evaluated in rabbit reticulocyte system or in vitro translation system. b Translation activity evaluated in cellulo . 8. SAR Study of mRNA Cap Using Chemoenzymatic Synthesis . Entry Modification Capping efficiency (%) eIF4E binding K D (nM) Decapping with hDcp2/half-life Relative translation efficiency Cap modification 1 m 7 G ppp G 100 - - 1.00 2 Bn 7 G ppp A 100 - - 0.75 3 ClBn 7 G ppp A 100 - - 0.93 4 ClBnOEt 7 G ppp A 100 - - 0.71 5 LNAm 7 G ppp A 100 - 15 min 4.53 First base modification 6 m 7 G ppp A 100 2019 5 min 1.81 7 m 7 G ppp U 100 - - 0.85 8 m 7 G ppp C 100 - - 1.28 9 m 7 G ppp I 100 - - 2.50 10 m 7 G ppp m 6 A 100 - - 3.01 11 m 7 G ppp A PS G 100 - - 0.61 12 m 7 G ppp 2F-A 100 - - 0.81 13 m 7 G ppp L -A 100 - - 1.14 14 m 7 G ppp dA 100 - - 1.61 15 m 7 G ppp 2OMe-A 100 - 5 min 2.64 16 m 7 G ppp 2MOE-A 100 - - 3.38 17 m 7 G ppp LNA-A 100 1708 >8h 4.77 18 LNAm 7 G ppp LNA-A 100 1425 >8h 8.63 Multiple bases modification 19 m 7 G ppp A PS G PS A PS G PS A PS A PS 100 - - 1.42 20 m 7 G ppp dA dG dA dG dA dA 100 - - 2.41 21 m 7 G ppp 2OMe*6 100 2446 10 min 6.91 22 m 7 G ppp 2MOE*6 100 - - 5.39 23 m 7 G ppp LNA*6 100 - - 4.77 24 LNAm 7 G ppp LNA*6 100 - - 4.25 25 LNAm 7 G ppp 2OMe*6 100 1570 15 min 7.49 Open in a new tab a Translation activity evaluated with Hela cell 24 h post transfection. While the practicality of initial nucleotide modifications using dinucleotide cap analogs was limited by low capping efficiency, the development of trinucleotide or tetranucleotide cap analogs enabled the IVT synthesis of mRNA with Cap1, Cap2, or base-modified nucleotides (e.g., m 6 A, Bn 6 A) to enhance mRNA translation. ,, ( Table ) In contrast, cap modifications utilizing virus capping enzymes to modify first base (e.g., m 6 A or propargyl- 6 A) revealed that the first base modification negatively impacted the translation. ,, ( Table ). These contradictory findings may stem from two possible reasons: (1) differences in synthesis workflows, resulting in variability in modification efficiency, mRNA integrity, and downstream purification quality; and (2) cell-type-specific effects that yield divergent translational responses across experimental systems. Beyond the m 7 G cap and first nucleotide modification, cap analogs with modified triphosphate were also prepared. In the endogenous mRNA degradation pathway, the DcpS and Dcp2 enzymes are responsible for mRNA decapping, which in turn promotes exonuclease-mediated degradation. DcpS recognizes dinucleotide/trinucleotide cap analogs or short capping oligonucleotides and cleaves the triphosphate between the γ and β position; whereas the Dcp1/2 complex usually cleaves longer transcripts between the β and α position. To block the potential degradation by the decapping enzyme and thus increase the stability of synthetic mRNA, caps with tetraphosphate, substitutions at beta-phosphate position (e.g., BH 3 , Se, NH, S), or substituents on the nitrogenous base (e.g., triazole) demonstrated that triphosphate modifications could increase the half-life of cap analogs significantly and contribute to enhanced mRNA translation. ,,− Interestingly, such substitution introduced an additional chiral center in the triphosphate chain, which showed distinct degradation kinetics and translation outcomes. Further structural studies revealed that two phosphate isomers showed different binding kinetics with eIF4E, which partially explains the observed differences in translational properties. , 6. SAR Study of mRNA Cap Using Trinucleotide or Tetranucleotide Cap Analogs. Entry Modification Capping efficiency (%) eIF4E binding K AS x10 –6 (M -1 ) Decapping Relative translation efficiency Ref m7G modification 1 m 7 G ppp A G - 89 26.6 - 1.0 b , 2 Locked nucleic acid m 7 G ppp 2′-OMe A G - 53 - - ∼5 (Rel. to 1 ) 3 3′-propargyl-m 7 G ppp 2′-OMe A G - - - - 1.3 (Rel. to 1 ) Triphosphate modification 4 m 7 G p s pp (S p ) 2′-OMe A G - - - Resistant ∼0.7 (Rel. to 14 ) 5 m 7 G p s pp (R p ) 2′-OMe A G - - - Resistant ∼0.8 (Rel. to 14 ) 6 m 7 G ppp 2′-OMe A G - - - - 5.93 7 m 7 G ppp-triazole- A G - - - - 0.21 (Rel. to 6 ) 8 m 7 G ppp-triazole- 2′-OMe A G - - - - 0.43 (Rel. to 6 ) 9 m 7 G ppp-CH 2 –triazole- A G - - - - 0.22 (Rel. to 6 ) 10 m 7 G ppp-CH 2 –triazole- 2′-OMe A G - - - - 0.42 (Rel. to 6 ) 11 m 7 G ppp-C 2 H 4 -triazole- A G - - - - 0.25 (Rel. to 6 ) 12 m 7 G ppp-C 2 H 4 -triazole- 2′-OMe A G - - - - 0.21 (Rel. to 6 ) 13 m 7 G ppp-triazole-C 2 H 4 - 2′-OMe A G - - - - 4.79 (Rel. to 6 ) Nucleotides modification 14 m 7 G ppp 2′-OMe A G - 90 45.6 nM Hydrolyzed 1.21 (Rel. to 1 ) , 15 m 7 G ppp m 6 A G - 78 35.2 nM Hydrolyzed 0.9 (Rel. to 1 ) 16 m 7 G ppp m 6 Am G - 77 32.8 nM Hydrolyzed 1.56 (Rel. to 1 ) 17 m 7 G ppp C G - 60 25.1 nM Resistant 0.73 (Rel. to 1 ) 18 m 7 G ppp 2′-OMe C G - 54 29.9 nM Resistant 0.97 (Rel. to 1 ) 19 m 7 G ppp G G - 80 22.7 nM Resistant 0.34 (Rel. to 1 ) 20 m 7 G ppp 2′-OMe G G - 86 22.8 nM Resistant 0.41 (Rel. to 1 ) 21 m 7 G ppp U G - 56 - Resistant 0.45 (Rel. to 1 ) 22 m 7 G ppp 2′-OMe U G - 56 - Resistant 0.91 (Rel. to 1 ) 23 m 7 G ppp Bn 6 Am G - 80 23.9 nM Hydrolyzed ∼3.0 (Rel. to 14 ) 24 m 7 G ppp 2′-OMe A 2′-OMe G G - - Hydrolyzed ∼1.0 (Rel. to 1 ) 25 m 7 G ppp A 2′-OMe G G - - Hydrolyzed ∼1.8 (Rel. to 1 ) 26 m 7 G ppp m 6 Am 2′-OMe G G - - Hydrolyzed ∼2.0 (Rel. to 1 ) 27 m 7 G ppp m 6 A 2′-OMe G G - - Hydrolyzed ∼4.0 (Rel. to 1 ) Open in a new tab a Translation activity evaluated in cellulo . 7. SAR Study of mRNA Cap Using Virus Capping Enzymes. Entry Modification Relative translation efficiency Ref 1 m 7 G ppp G 1.0 2 N 2 -Me-m 7 G ppp G ∼0.5 (Rel. to 1 ) 3 m 7 G ppp 2′-OMe A ∼1.6 (Rel. to 1 ) 4 m 7 G ppp m 6 Am ∼0.2 (Rel. to 1 ) 5 m 7 G ppp N 6 -propargyl-Am ∼1.0 (Rel. to 1 ) 6 Bn 7 G ppp G ∼0.5 (Rel. to 1 ) 7 Allyl- 7 G ppp G <0.1 (Rel. to 1 ) 8 Azido- 7 G ppp G <0.1 (Rel. to 1 ) 9 Vinylbenzyl- 7 G ppp G <0.1 (Rel. to 1 ) 10 4-Cl-Bn 7 G ppp G 1.3 (Rel. to 1 ) 11 4-CN-Bn 7 G ppp G 0.8 (Rel. to 1 ) 12 Pyridine- 7 G ppp G 0.4 (Rel. to 1 ) 13 4-NO 2 –Bn 7 G ppp G - 14 4-Br-Bn 7 G ppp G - 15 3,5–2-CF 3 –Bn 7 G ppp G - Open in a new tab a Translation activity evaluated in rabbit reticulocyte system or in vitro translation system. b Translation activity evaluated in cellulo . Although cotranscriptional capping and post-transcriptional enzymatic cap modifications enable the preparation of a series of mRNA with different cap or nucleotide modifications, several issues with these methods prevent a comprehensive investigation of mRNA SAR at the 5′ end. First, different cap analogs usually have distinct capping efficiency and correct capping orientation rate (for dinucleotide cap analogs), so that the translation outcomes are the combined effect of capping efficiency and the modification itself, which makes the interpretation of specific modification challenging and may contribute to the opposite results obtained from different modification strategies mentioned above. Second, current cap modification strategies are restricted to the first two bases of mRNA, and the effects of extended modifications at the 5′ end have not elucidated. Given that the m 7 G cap and the 5′ UTR are closely involved in the translation initiation process, further expanding the chemical space for mRNA 5′ end modification will provide a more comprehensive understanding of mRNA SAR. Recently, our group utilized the ligation-enabled mRNA-oligonucleotide assembly (LEGO) technology to systematically interrogate chemical modifications SAR at the 5′ end of mRNA. ( Table ) Through chemical synthesis of the capped oligonucleotide and enzymatic ligation with IVT mRNA, > 98% capping efficiency could be achieved for all the constructs without sequence bias. A variety of modified nucleotides (e.g., m 6 A, 2′-OMe) can be chemically incorporated into the capped oligonucleotides without restrictions from enzymatic tolerance and with enhanced mRNA translation. Meanwhile, the effects of other novel modifications have also been identified, including base modification (e.g., inosine), sugar backbone modification (2′-FA, LA, LNA, 2′-MOE, dA), and phosphate linkage modification (e.g., phosphorothioate) for the first six nucleotides, which for the first time demonstrated the feasibility and benefits of modifying mRNA beyond the first two bases. Based on combinatorial screening of modifications at different sites, a ∼ 10-fold increase in translation efficiency was observed compared to the unmodified construct. Mechanistically, chemical modifications such as LNAm 7 G and LNA for the first nucleotide improve translation initiation by increasing the binding affinity of the mRNA with cap-binding proteins eIF4E and downstream eIF4G. Additionally, modifications, including LNA as the first nucleotide or six repeated 2′-OMe, protect mRNA from decapping, as demonstrated by an increased half-life of the modified oligonucleotide in the hDcp2 capping experiment. In summary, the LEGO technology unlocked the chemical space at the mRNA 5′ end and can be potentially applied to explore more diverse chemical modifications. 5.2. CDS Modification As the major component of mRNA molecules, the coding sequence carries the genetic information for translation and significantly influences the biochemical properties of mRNA biochemical properties. Although the concept of using mRNA as a therapeutic modality has been explored for several decades, the immunogenicity of synthetic mRNA has led to significant cytotoxicity and translation inhibition, which largely restricts its efficiency for therapeutic purposes. The revolutionary work for this field was achieved in 2005 when mRNA base modifications were discovered to alleviate the immunogenicity triggered by synthetic mRNA. Following this line, researchers explored more diverse modifications for the purpose of generating mRNA with desired properties. Among all methods for synthesizing mRNA with nucleotide modifications, enzymatic IVT incorporation, whether full or partial substitution, was the most convenient and prevalent. By utilizing this method, researchers prepared a variety of chemically modified mRNAs, including m 5 C, s 2 U, m 5 U, m 6 A, m 1 Ψ, Ψ, 5moU, phosphorothioated NTP substitutions, which allow the functional interrogation of these modifications. Pseudouridine (Ψ) was the first chemical modification of mRNA identified to significantly increase protein production. Based on the results from in cellulo luciferase reporter assay, mRNA full substitution of U with Ψ enhances protein synthesis by 5 to 9 fold in comparison to unmodified mRNA in several cell lines. In 2005, m 1 Ψ, another natural modification found in 18S rRNA, was introduced to mRNAs. m 1 Ψ outperformed Ψ in terms of protein production both in vitro and in vivo . Mechanistically, previous studies have identified several possible explanations for the enhanced translation, which can be grouped into two main categories: modulation of the innate immune response and regulation of translation dynamics. First, incorporation of m 1 Ψ or Ψ extensively regulates the innate immune response induced by exogenous mRNAs, thus alleviating translational inhibition. (1) m 1 Ψ incorporation reduces the synthesis of immunogenic antisense RNA and dsRNAs during IVT. , (2) The incorporation of m 1 Ψ alters the secondary structure of mRNA due to the differences in base-pairing strength with uridine, which minimizes interactions with immune receptors such as TLR3 or RIG-I. , (3) The interaction of mRNA with single-stranded RNA immune receptors (e.g., TLR7, PKR) was changed by m 1 Ψ modification, whose altered hydrogen bonding face and steric “bump” disrupts the mRNA-protein interaction. The reported data demonstrated that decreased interaction with immune-sensing receptors alleviates activation of downstream cytokine signaling pathways and decreases the phosphorylation level of the translation initiation factor 2-alpha (eIF-2α), thereby improving translation. ,− (4) A recent study also discovered that exonucleases do not adequately process pseudouridine-containing RNAs to generate TLR-agonistic ligands, thus evading immune detection. (5) m 1 Ψ was recently discovered to help RNAs evade immune surveillance through reduced binding with proton-sensing TRIM25, which is an RNA-binding E3 ubiquitin ligase inducing mRNA degradation. Second, m 1 Ψ and Ψ have also been shown to directly modulate the decoding kinetics during translation beyond merely interfering with innate immune sensing, though with contradictory results. For instance, incorporation of m 1 Ψ has been reported to dramatically alter translation by inducing ribosome pausing and increasing ribosome density on mRNA. Although further evidence is required, researchers have proposed that enhanced ribosome loading may facilitate more permissive initiation, either through ribosome recycling on the same mRNA or de novo ribosome recruitment. In contrast, using an in vitro translation system, other researchers found that Ψ within the coding sequence can modestly influence translation speed and decoding, but promotes the synthesis of multiple peptide products from a single mRNA in a context-dependent manner. More recently, the same group reported that m 1 Ψ does not significantly alter the rate of amino acid incorporation by cognate tRNAs but may subtly affect the fidelity of incorporation. Moreover, while pseudourine modifications confer desirable properties to mRNA constructs, challenges and limitations remain. More recently, researchers discovered that incorporation of m 1 Ψ into mRNA results in +1 ribosomal frameshifting due to the m 1 Ψ-induced ribosome stalling during IVT mRNA translation. These results highlighted the potential safety issues associated with CDS-modified mRNAs induced by the heterogeneity of protein products. Overall, the precise regulatory roles of m 1 Ψ and Ψ in translation dynamics remain poorly understood, partly due to difficulties in fully disentangling the decoding process from cellular immune sensing mechanisms, especially in cellulo studies. Beyond the application of m 1 Ψ or Ψ, other mRNA base modifications have also been extensively explored. Full substitution with modified NTPs during IVT was the most convenient method to introduce chemical modifications to CDS. For instance, 5-methoxyuridine (5moU) substitution enhances protein production from mRNA encoding the Cas9 protein and thrombopoietin (TPO); , 5-methyl-C (m 5 C) full substitution was found to decrease the immunogenicity of mRNA and enhance protein production. In some cases, the combination of m 5 C and m 1 Ψ can further increase the translation efficiency of mRNA than their single-nucleotide-substitution counterparts. , However, full substitution of adenosine with N 6 -methyladenosine (m 6 A) or uridine with 2-thiouridine (s 2 U) was shown to decrease translation efficiency significantly, although m 6 A modification could also reduce immunogenicity of mRNA. In contrast, 5% substitution of the m 6 A can slightly enhance translation. A subsequent mechanistic study using single-molecule fluorescence revealed that m 6 A modification acts as a barrier for tRNA accommodation, thereby disrupting translation elongation. Furthermore, N 4 -acetylcytidine (ac 4 C), a naturally occurring mRNA modification catalyzed by the acetyltransferase NAT10, has recently been incorporated into synthetic mRNAs through full substitution during in vitro transcription (IVT). While the ac 4 C modification promotes translation and stability of endogenous mRNAs, a recent study demonstrated that substituting cytidine with ac 4 C resulted in comparable protein levels and reduced inflammatory gene expression in immune cells. , While full substitution offers mRNA constructs with enhanced translation and reduced immunogenicity, a more recent study showed that incorporating modifications at different positions results in vastly different translation performance. Recently, DNA polymerase-based RNA synthesis methods enabled site- and region-specific mRNA modification using modified NTP analogs. Using an in vitro translation system with an IRES-containing mRNA, researchers demonstrated that full substitution of C with m 5 C completely abolished mRNA translation because base modification altered the IRES secondary structure and reduced its interaction with RNA-binding proteins. This contrasts with substitution in the coding sequence, where translation is slightly reduced. Moreover, site-specific m 5 C modifications at either the initial or middle region of CDS led to 2–3 fold enhancement of mRNA translation, which highlights the importance of introducing modifications in a more controllable and accurate manner. Unlike base modifications, which generally facilitate translation through altered interactions with the initiation machinery, 2′-OH modification that blocks hydrolysis in the coding region was recently developed using several synthetic methods to increase RNA stability. To this end, researchers employed a reversible 2′-OH acylation reagent that effectively protects RNAs from both thermal and enzymatic degradation. Functional demonstrations using sgRNAs or mRNAs (e.g., GFP) showed a significant increase in RNA half-life. Furthermore, cloaked and nontranslatable mRNA was deprotected in cellulo to release functional mRNA, ultimately leading to higher total protein production due to increased stability. Recently, chemoenzymatic synthesis was used to investigate site-specific 2′-OMe modification within the mRNA coding sequence, particularly at the codon immediately downstream of the start codon. The results demonstrated that, beyond minimizing hydrolytic degradation, 2′-OMe influences IRES-dependent mRNA translation in a position-dependent manner (the second-nucleotide methylation of the codon abolishes translation, while the third-nucleotide methylation increases protein production by 2-fold). Mechanistically, a recent single-molecule fluorescence study investigated the influence of 2′-OMe on mRNA decoding kinetics, showing that 2′-OMe within coding regions disrupts key steps in codon reading during cognate tRNA selection. These results further highlighted the necessity of introducing chemical modification in a site-specific manner to maximize the effects for boosting translation, which requires more systemic SAR study in the future. Phosphate backbone phosphorothioate modification within CDS has been shown to regulate mRNA properties. Through single or multiple phosphorothioated NTP substitutions, backbone-modified mRNAs were prepared. The results showed that single-base substitutions significantly increase translation efficiency, whereas multiple-base substitutions, especially triple and quadruple substitutions, decrease translation efficiency. Mechanistic study revealed that phosphorothioate modification increases ribosome loading on mRNA, suggesting an increased translation initiation rate. In summary, coding sequence modification plays a vital role in regulating stability, translatability, and immunogenicity of mRNA. Although significant achievements have been made over the past few decades, our knowledge of SARs of synthetic mRNA internal chemical space is still limited, especially in a region or context-specific manner. Future applications of synthetic methods combining chemical and enzymatic synthesis to construct a more diverse internal modification library will be beneficial to further the chemical space of mRNA. 5.3. UTR Modification The untranslational region (UTR) regulates both translation and stability through interactions with RNA-binding proteins. Given the importance of the UTR, several synthetic methods have been applied to prepare mRNA with modified UTRs. For instance, mRNAs with site-specific m 6 A modification in the 5′ UTR were prepared by enzymatic IVT incorporation of m 6 ATP and an intentional UTR design that left only adenosine for substitution. This method revealed that the m 6 A modification in the 5′ UTR enables cap-independent translation of mRNA as a mechanism of mRNA translation under physiologic stresses. Additionally, researchers combined enzymatic IVT incorporation and T4 RNA ligase 2-based splint ligation to synthesize mRNA with phosphorothioate modification restricted to the 5′ UTR. In this way, they were able to compare the effect of phosphorothioate modification in a region-specific manner. The results demonstrated that phosphorothioated ATP and CTP increase the mRNA translation initiation rate in an in vitro translation system. In contrast, the whole sequence substitution with phosphorothioated ATP and CTP led to decreased translation efficiency. Taking advantage of chemenzymatic synthesis, our group recently reported the synthesis of mRNAs with 5′ UTR modifications up to the first six nucleotides. Reporter mRNA screening enabled the identification of several modifications that enhance both stability and translation efficiency, such as 2′OMe × 6 or LNA × 6. ( Table ) Additionally, genetic code expansion technology has also been used to synthesize mRNA with site-specific modifications. A recent study reported a method to incorporate an unnatural base into the 3′ UTR using the expanded genetic code, which was well tolerated by the translation machinery and enabled site-specific labeling of mRNA. These results showed that the effects of chemical modifications vary significantly across different regions of the mRNA and highlighted the importance of introducing modifications in a more region-selective manner within the UTR. Given the current need for more efficient therapeutic mRNA, the development of synthesis methods that enable region-specific modification of each mRNA construct component will be necessary to further enhance efficiency and allow more therapeutic scenarios in the future. 5.4. Poly(A) Tail Modification The 3′ poly(A) tail of mRNA regulates translation through interactions with RNA-binding proteins such as PABP and affects mRNA stability through interactions with deadenylation complexes and exonucleases. Several synthesis methodologies were developed to modify the poly(A) tail and modulate mRNA properties. First, 3′ chemical derivatization was performed by selective 3′-diol cleavage using sodium periodate, followed by reductive amination to form hexose nucleotide derivatives resistant to the exonucleases degradation. Activity tests demonstrated that this terminal chemical modification increased mRNA stability, thereby increasing protein production. By using enzymatic modification with polyA polymerase, an azide-containing ATP analog was incorporated for 3′ terminal modification. Further click reactions with biotin-alkyne form an exonuclease-resistant unnatural linkage that both protects mRNA from degradation and increases protein production by 2–3 fold. Additionally, PAP was used to enzymatically incorporate phosphate-modified ATP analogs, including ATPαS and ATPαBH 3 . The results demonstrated that phosphate modification significantly increases the stability of the poly(A) tail. However, partial ATPαS incorporation showed marginal impact on protein expression, while ATPαBH 3 incorporation decreased protein production significantly, suggesting that boranophosphate modification was less compatible with mRNA expression in vivo than phosphorothioate. To further explore the chemical space, our group used chemoenzymatic synthesis to investigate a series of chemical modifications on the poly(A) tail with a broader structural scope and well-defined structure. Different elements, including modified internucleotide linkage, pentose sugar modification, structural motif, and chain-terminating nucleotide, were enzymatically ligated to the 3′ end of mRNA and screened for their translation activity. The results showed that terminal phosphorothioate linkage increases the total protein production by 3–4 fold. It is noteworthy that poly(A) tail modification regulates more of mRNA stability than translation initiation. Ribose modification with 2′MOE showed the most significant effect in terms of total protein production (4–6 fold enhancement). In addition, a structured DNA quadruplex (G4) at the 3′ terminus increases protein production, while an unstructured ssDNA tail does not, presumably due to the end protection offered by the structured sequence. Overall, introducing nuclease-resistant chemical modifications to the 3′ end provides a general method for improving stability and generating mRNA with longer expression duration. 6. Structure Activity Relationship of RNA Topology The majority of the endogenous functional mRNAs are linear, which consist of both 5′ and 3′ termini, and internal nucleotides sequentially linked with each other through canonical 5′-3′ diphosphate ester bonds. Beyond classic mRNAs, RNAs with other nonlinear topologies have also been identified within the cell, including lariat RNAs as intermediate byproducts from mRNA splicing and circular RNAs acting as both regulatory RNAs and protein-encoding RNAs. Recently, given the increasing knowledge about the functions of naturally occurring RNA species with various topologies, topological engineering has also been applied to the design of therapeutic mRNAs, adding another dimension for therapeutic mRNA optimization beyond sequence design and chemical modifications. ( Figure ) 15. Open in a new tab Topological engineering of therapeutic mRNA . ( A ) The mRNA with multiple branched polyA tail preserves multimeric poly(A)-PABPC1 interactions for prolonged translation and provides resistance to exonuclease degradation through combining the 3′ chemical modifications. Figure 15A adapted from ref . Copyright 2024 Springer Nature. ( B ) The branched topology at the 5′ end of mRNA enables the introduction of a dual cap structure, which enhances translation initiation by increasing ribosome loading and translation efficiency. ( C ) Circular RNA (circRNA) shows prolonged duration through blocking the exonuclease degradation and 5′ phosphate-mediated immunogenicity. The cap structure is covalently introduced to the circular RNA to construct the QRNA. QRNA shows prolonged duration due to its circular topology and enhanced translation efficiency via the cap-dependent translation initiation mechanism. Figures B and C adapted from ref . Copyright 2024 Springer Nature. 6.1. Branched mRNA The most representative endogenous branched mRNA is lariat RNA. Although the biological significance of lariat RNA is less understood, the branch structures suggest a new possibility for RNA topology. Recently, our group introduced the concept of topological engineering to the field of therapeutic mRNA by exploring synthetic mRNAs with various topologies. 6.1.1. Multitail mRNA The introduction of topological engineering to the 3′ end of mRNA was explored as a proof-of-concept. Previous data have shown that introducing site-specific exonuclease-resistant modification at poly(A) tails increases mRNA stability and consequently, protein production, possibly by maintaining poly(A)-PABPC1 interactions. Given that PABPC1 functions as a multimeric protein complex, as revealed by structural studies, our group constructed an mRNA with multiple poly(A) tails via a branched topology, which preserved multimeric poly(A)-PABPC1 interactions for prolonged translation. ( Figure A) By using the chemoenzymatic synthesis method, branched poly(A) tails with exonucleases-resistant chemical modifications were assembled via click chemistry with a normal 5′ to 3′ directionality, followed by enzymatic ligation with the IVT mRNA to form the mRNA construct with multiple poly(A) tails. Luciferase reporter assays in cellulo demonstrated that having three branches and synergistic incorporation of 2′- O -methoxyethyl (2′ MOE) and phosphorothioate (PS) on both the stem and branched poly(A) oligos significantly increase mRNA stability. Mechanistically, modified poly(A) tails can form multimeric interactions with PABPC1 as a natural linear poly(A) tail, and both chemical and topological modifications effectively protect poly(A) from degradation to preserve functional tails for prolonged protein expression. The stabilized Cas9 mRNA construct with branched poly(A) tails was applied to multiplex genome editing of Pcsk9 and Angptl3 in vivo and showed substantially increased genome editing efficiency at lower mRNA dosage. 6.1.2. Multicap mRNA In addition to introducing multiple poly(A) tails to increase mRNA stability, our group recently reported a strategy to enhance mRNA translation efficiency by installing a branched multiple cap structure at the 5′ end. Resembling the construction of multitail mRNA, synthetic and capped oligonucleotides with chemical modifications were assembled through click chemistry and then ligated with the IVT mRNA containing 5′ monophosphate. ( Figure B) Activity screening results demonstrated that a branched dual cap structure increases mRNA translation efficiency by 2-fold through enhanced interactions with translation initiation factors, such as eIF4E. Subsequent topological screening of the length of branch and stem strands revealed an optimal topology for the 5′ end structure that maximizes translation efficiency. Interestingly, mRNA with only a cap on the branch (and not on the stem) showed comparable translation activity to capped linear mRNA, indicating that the unnatural branch linkage at 5′ end of mRNA is well tolerated by the cellular translation machinery. Furthermore, the combination of an optimal branched topology and chemical modifications resulted in mRNA with a 15-fold higher protein production level than unmodified mRNA at 24 h post transfection. Functional demonstrations indicated that chemically modified dual-capped mRNA enhances SARS-CoV-2 mRNA vaccine efficacy and plasma hEPO expression in protein replacement therapeutic setting. In summary, these examples demonstrated that branched mRNA topology is compatible with the endogenous translation machinery and can be applied to regulate both mRNA translation efficiency and stability in combination with chemical modifications, opening a new avenue for therapeutic mRNA engineering. 6.2. circRNA and QRNA 6.2.1. circRNA Circular RNA (circRNA) is another topology of endogenous RNAs. They are generated by 3′ exonucleolytic cleavage of lariat splicing byproducts or a noncanonical splicing event called backsplicing. Backsplicing produces covalently closed RNA molecules without 5′ and 3′ ends. Since its discovery 40 years ago, this noncanonical RNA species has been extensively explored regarding its biogenesis and properties. Functional studies demonstrated that circRNAs play an important role as regulatory elements: acting as miRNA sponges to fine-tune mRNA translation; acting as protein sponges to enhance the functions of RNA-binding proteins (RBPs); and acting as scaffolds to mediate complex formation between specific enzymes and substrates. Details for the characterization and mechanistic studies have been comprehensively reviewed previously. , More recently, researchers identified a class of endogenous circRNAs, including circ-ZNF609, circMbl, circFBXW7, circPINTexon2, and circ-SHPRH, that are translatable and produce functional proteins. − Contrary to the canonical mRNAs with 5′ cap and 3′ tail that undergo cap-dependent translation, the translation of circRNA is mainly driven by two mechanisms, including internal ribosome entry sites (IRESs) driven translation or m 6 A modification driven translation. , ( Figure C) It has also been found that circRNAs are more stable than their linear counterparts due to the lack of exonuclease-dependent decay. In addition, circularization of RNA helps with attenuating immune response through bypassing cellular RNA sensors such as RIG-I and Toll-like receptors (TLRs) or leveraging the internal m 6 A modifications to avoid nonself detection. , Given the favorable properties of circRNAs, researchers explored the possibility of developing circRNA-based vectors for therapeutic protein expression. For instance, researchers engineered the Group I catalytic intron system to efficiently produce circRNA via backsplicing, achieving circularization of sequences up to 5kb. After comparing several IRES elements, the reporter assay demonstrated that the optimized and purified circRNA outperformed the modified linear mRNA with the same sequence in terms of protein production and expression duration. The following mechanistic characterization revealed that the immunogenicity and protein expression of circRNA depend on purity; and small amounts of contaminating linear RNA can trigger a cellular immune response, highlighting the importance of product purification during circRNA preparation. To further increase the translatability of circRNAs, a high-throughput screening was performed to systematically discover RNA sequences that can direct circRNA translation and identified the 18S rRNA complementarity or structured RNA sequences as functional elements important for driving circRNA translation. Recently, the same group optimized various elements within the circRNA, including vector topology, 5′ and 3′ untranslated regions, IRESs, and synthetic aptamers that can recruit translation machinery, generating the optimal circRNA with more durable translation in vivo. The progress of circRNA optimization recently led to the development of the first circRNA-based vaccine for SARS-CoV-2 spike protein, which enables higher and more durable antigen production. Although the processing method optimization and sequence engineering contribute to the enhancement of circRNA translatability by hundreds fold, current circRNAs still suffer from several limitations: (1) restricted by the synthesis method using intron or ribozyme, circRNAs are not compatible with base modifications including m 1 Ψ which has been proven to increase the efficacy of therapeutic mRNAs; (2) it was not feasible to incorporate site-specific chemical modifications such as m 6 A, which are favorable for circRNA translation; (3) the translation efficiency of circRNAs is still much lower than linear mRNA, especially those with optimized cap structure as well as 5′/3′ untranslated regions sequence, due to the inefficiency of cap-independent translation mechanism. 6.2.2. Capped circRNA (QRNA) To tackle these issues and take advantage of the favorable properties of circRNA, our group recently developed “QRNA” by introducing branched cap structures to circRNA, which simultaneously maintains the high stability of circRNA and removes the IRES by harnessing the cap-dependent translation initiation mechanism to enhance its translatability. We denote such capped-circular mRNA as QRNA, after the construct’s visual resemblance to the letter “Q”. ( Figure C) A panel of Nano luciferase (Nluc) QRNAs were constructed and screened, bearing various modified caps and beneficial chemical modifications such as site-specific m 6 A or PS modifications in the 5′ UTR. m 1 Ψ-modified QRNA with m 7 G or LNAm 7 G cap resulted in 27- to 144-fold enhancement of translation efficiency in cellulo and 30- to 59-fold enhancement in vivo compared to the unmodified circRNA with an IRES prepared by backsplicing. In summary, circular RNA, as a complement to traditional linear mRNA, offers new opportunities to improve the properties of RNA-based therapeutics, especially stability and immunogenicity, thereby enhancing protein production both in vitro and in vivo . Additionally, further topological engineering of circRNA results in the development of QRNA, which provides superior translation efficiency and stability simultaneously. 6.3. Trans-Acting Elements and Distal Cis-Acting Elements 6.3.1. Cap Proximal Translation Driven by Hybridization The efficient translation induced by the branched cap demonstrates the tolerance of the endogenous translation machinery to noncanonical cap structures and the great flexibility of the translation initiation process. ( Figure A) Inspired by noncanonical cap-dependent translation, our group further explored other topologies of synthetic mRNAs. First, given that the 3′ end of mRNA is usually in close proximity with the 5′ end through RNA-protein interaction according to the “close-loop” model of translation initiation, our group reported that topological inversion of the natural 5′-cap/3′-poly(A) feature to a synthetic 3′-cap/5′-poly(A) could also induce translation, which demonstrated that a distal cis -acting cap element could effectively drive translation through the canonical eIF4E-dependent mechanism. ( Figure C) Second, the trans -acting cap element was also effective for driving mRNA translation where a capped oligonucleotide hybridized to a complementary sequence upstream of the ORF in the circRNA could induce expression of the downstream ORF. ( Figure B) Furthermore, when the branched capped oligonucleotide (with m 7 G or its modified derivatives) was placed internally between two ORFs, we demonstrated that it could drive the translation of both upstream and downstream ORFs either through direct 3D proximity of the branched cap with upstream UTR and start codon or through 1D ribosome 3′ to 5′ back scanning. ( Figure C–D) 16. Open in a new tab Different modes of cap proximal translation . ( A ) Covalent and branched cap structure is well-tolerated by the translation machinery and initiates translation through “slot-in” mode. ( B ) Noncovalent trans-acting cap hybridization to either circRNA or linear mRNA can initiate translation. ( C ) Noncovalent cis-acting cap drives translation via homology that brings the 3′ cap to the start codon. ( D ) Cap structure could drive the translation of both proximal upstream and downstream ORFs within a circular RNA. ( E ) Tethering of a poly(A) tail mimetic to endogenous mRNA through hybridization selectively enhances protein production. ( F ) Hybridization of translation-activating RNAs (taRNAs) to the 3′ UTR of endogenous mRNA enhances translation by recruiting the translation machinery. Figures A-D adapted from ref . Copyright 2024 Springer Nature. In summary, these results suggested a general “cap-proximal” mechanism: the 5′ cap and UTR sequences connected to mRNA through the natural phosphodiester backbone, an unnatural covalent linkage, or hybridization can initiate translation. ( Figure ) Such a mechanism not only illustrates the general translation process but also inspires future synthetic mRNA structural design with broader chemical scope and topology. 6.3.2. Translation Enhancement Driven by Hybridization of Poly(A) Tail Mimetics or Translation Regulating Elements Beyond regulating the 5′ end of mRNA, trans-acting elements have also been employed for enhancing translation intermolecularly through hybridizing a poly(A) mimetic to the 3′ end of mRNA. By polyadenylating short RNA sequences antisense to the 3′ UTR of specific endogenous mRNA, a recent report demonstrated that the “booster poly(A) mimetic” could selectively and significantly enhance mRNA expression both in vitro and in vivo . This result indicated that the branched poly(A) tail structure through hybridization is well-tolerated by the translation machinery and sufficient to mimic the function of poly(A) tail. ( Figure E) Additionally, a translation-activating RNA (taRNA) was developed to bind the specific endogenous mRNA of interest and directly upregulate its translation. taRNA is a bifunctional molecule with a guide sequence for targeting 3′ UTR and domain selected from viral or mammalian IRESs for recruiting translation machinery. ( Figure F) 7. Therapeutic and Scientific Applications Given their increased stability, higher translation efficiency, decreased immunogenicity, and programmability, chemical modifications and topological engineering of mRNAs provide powerful tools for both basic biology studies and disease treatment. Below, we provide examples from fields that attract the most attention for mRNA therapeutics, including cell reprogramming and stem cell engineering, vaccines for infectious diseases, cancer immunotherapy, gene editing, and enzyme/protein replacement therapy. ( Figure ) 17. Open in a new tab Therapeutic and scientific applications of modified mRNAs . Chemically and topologically modified mRNAs have been applied to a variety of therapies, including cell reprogramming and stem cell engineering, gene editing, vaccines for infectious diseases, cancer immunotherapy, and enzyme/protein replacement therapy. 7.1. Cell Reprogramming and Stem Cell Engineering The ability to restore the pluripotency of somatic cells through coexpression of reprogramming factors has revolutionized cell biology, providing a powerful tool and new opportunities for regenerative medicine. In 2006, pioneering work showed that the cell type can be altered by transfecting exogenous transcription factors, called Yamanaka factors, to generate induced pluripotent stem cells (iPSCs) that resemble the differentiation potential of embryonic stem cells (ESCs). Considering the great benefit of iPSC in the investigation of developmental biology and for developing therapies, researchers continued to explore methodologies to enable more efficient cell reprogramming. Several vectors, including virus-based integration or gene expression, transposon-based genomic integration, and plasmid-based transfection, were used to deliver transcription factors (TFs) that drive cell differentiation, but suffered from low induction efficiency (typically 0.001% to 0.1%). The emergence of mRNA technology has provided new opportunities for cell reprogramming, enabling efficient protein expression while avoiding the risk of genome integration. The proof-of-concept work on using mRNA to generate iPSCs was reported in 2010 by utilizing chemically modified mRNA with an ARCA cap analog and an m 5 C/Ψ substitution. mRNAs encoding the Yamanaka factors ( KLF4 , c-MYC , OCT4 , and SOX2 ) drove efficient protein synthesis 6 h post-transfection in human primary fibroblast cells without significant cytotoxicity. However, due to the instability of mRNA and the requirement for long-lasting transcription factor expression for successful cell-state alteration, a daily transfection protocol was necessary to maintain an effective level of TFs for 18 days, yielding an induction efficiency of up to 4.4%. Although mRNA transfection has been shown to be the most efficient method for generating iPS cells to date, the requirement for multiple rounds of transfection remains labor-intensive and prohibitive for scale-up. To further increase the efficiency of iPSC induction, researchers showed that self-amplifying mRNA encoding Yamanaka factors with a Cap1 structure, synthesized using 2′-O-Methyltransferase, enables long-lasting TF expression and reduces the transfection frequency to once every 3 days. Together with miRNA transfection and optimized culture conditions, chemically modified mRNAs with ARCA cap analog and m 5 C/Ψ substitution led to an induction efficiency of 90% in a recent work. In summary, the application of chemical modification significantly enhances the stability and translation efficiency of mRNAs, leading to more efficient production of TFs and higher iPSC induction efficiency. Beyond iPSC induction, mRNA technology has also been applied in stem cell differentiation and direct cell transdifferentiation. For instance, mRNA encoding MYOD with ARCA cap and m 5 C/Ψ drives the transdifferentiation from human foreskin fibroblasts to myoblasts; using the same chemical modification, synthetic mRNA encoding PDX1 promotes the differentiation of hESCs to insulin-producing cells; the similar strategy has also been utilized to induce the differentiation from human iPSC to neurons by transfecting mRNAs encoding transcription factors including NEUROG1, NEUROG2, NEUROG3, NEUROD1 and NEUROD2. More comprehensive examples of using chemically modified mRNA for cell reprogramming have been reviewed previously. 7.2. Vaccine for Infectious Diseases Vaccines that stimulate the immune system have been proven highly effective against various infectious diseases caused by bacteria or viruses. In addition to traditional vectors for vaccine development, including inactivated, live-attenuated, and subunit or recombinant vaccines, mRNA-based vaccines have been demonstrated to be effective for infectious diseases in clinical trials during the COVID-19 pandemic. The most successful mRNA vaccines are the FDA-approved SARS-CoV-2 vaccines developed by Moderna (Spikevax) and BioNTech/Pfizer (BNT-162), both of which encode the full-length SARS-CoV-2 spike glycoprotein. To enhance the properties of these vaccines, the trinucleotide Cap1 analog was utilized, and uridine-5′-triphosphate (UTP) was fully substituted with N 1 -methylpseudouridine-5′-triphosphate (m 1 ΨTP) during in vitro transcription. The chemical modification alleviates the innate immune response triggered by exogenous mRNA molecules, thereby enhancing translatability and stability. In vivo administration of modified mRNA-LNP complex results in higher antigen expression levels and induces higher antibody titers and more potent T-helper-1 cell (Th1)-mediated immunity. The rapid development of SARS-CoV-2 mRNA vaccines provided the first proof-of-concept study in real-world clinical settings, and the encouraging results also benefited the investigation of vaccines for other viruses. For instance, researchers prepared the chemically modified mRNA with Cap1 structure and m 1 ΨTP full substitution encoding premembrane and envelope (prM–E) glycoproteins of ZIKV H/PF/2013 to develop a vaccine against Zika virus (ZIKV). An in vivo study demonstrated that a single low dose of intradermal immunization with mRNA-LNP elicited potent and durable neutralizing antibody responses in both mice and nonhuman primates, protecting rhesus macaques against ZIKV challenge at 5 weeks after immunization. Utilizing a similar strategy, clinical trial data was reported for mRNA vaccines against influenza viruses including H10N8 and H7N9. Chemically modified mRNAs encoding the full-length, membrane-bound form of the hemagglutinin (HA) glycoprotein from the H10N8 influenza strain or the H7N9 influenza strain were intramuscularly delivered. The results suggested that, although the mRNA influenza viruses vaccines induced robust humoral immune responses, no significant cell-mediated responses were detected. Furthermore, by leveraging the programmability and multiplexing capabilities of mRNA preparation, researchers developed a multivalent nucleoside-modified mRNA vaccine against all known influenza viruses through intramuscular administration of an mRNA-LNP complex encoding hemagglutinin antigens from all 20 known influenza A virus subtypes and influenza B virus lineages. The data from the mouse model indicated that multivalent mRNA vaccines successfully induce antibodies against multiple antigens and provide protection against variable viruses, demonstrating the great potential of developing multivalent vaccines using mRNA technology. , More recently, our group reported an in vivo evaluation of chemically and topologically modified mRNA encoding the SARS-CoV-2 spike protein. Through chemical modification screening for cap analogs and 5′ UTR, we identified the optimal combination of 5′ modifications (LNAm 7 G-LNA-2′-OMe × 5) that can enhance protein production of mRNA by up to 30-fold in vivo in combination with dual-cap topology and 3′ tail modification. Using our optimized mRNA construct with enhanced translatability and stability, we achieved a markedly stronger RBD-specific humoral response, reaching up to a 17.1-fold increase in antibody titers after the first dose and a 3.7-fold increase after the booster dose, and a substantially more robust T-helper cell response (a 4.0-fold increase in IFNγ-producing CD8 + T cells and a 4.2-fold increase in IFNγ-producing CD4 + T cells). These findings underscore the critical role of improved antigen expression in boosting the efficacy of mRNA-based vaccination strategies. Meanwhile, enhanced antigen production also allows lower mRNA dosage during vaccination, thereby alleviating manufacturing costs and minimizing potential side effects or toxicity induced by the mRNA-LNP complex. 7.3. Cancer Immunotherapy Along with the success and promising results of mRNA-based vaccines for infectious diseases, the application of mRNA-based technology in cancer treatment has gained increasing attention over the past few years. Several ongoing clinical trials have demonstrated the feasibility and effectiveness of using therapeutic mRNAs with different types of cancers, including melanoma, glioblastoma, breast, colorectal, and lung cancer. In the field of mRNA-based cancer immunotherapy, there are currently two major directions being intensively investigated: mRNA cancer vaccine and mRNA-based therapeutic protein production. We will summarize the current progress of chemically modified mRNAs in the development of next-generation cancer immunotherapy in this review. 7.3.1. mRNA Cancer Vaccine mRNA cancer vaccines are designed to induce or boost the active immune response against tumor cells or tissues. They target tumor-associated or tumor-specific antigens (TAAs or TSAs), leading to tumor cell clearance and long-lasting therapeutic responses through immune memory. The design of cancer vaccines involves the following steps: (1) identification of tumor-specific mutations or nonconforming sequences; (2) prediction of antigen epitopes specific for tumor cells; (3) design and preparation of mRNA constructs/libraries encoding TAAs or TSAs for vaccination. Resembling the mechanism of mRNA vaccines for infectious diseases, mRNA-based cancer vaccines are taken up by immune cells near the injection site after administration, especially by dendritic cells that are the most efficient antigen-presenting cells (APCs). After cellular uptake, proteins produced from transfected mRNA are subsequently subjected to post-translational modifications. Thereafter, the cellular degradation pathways generate antigen fragments directed for antigen presentation via the major histocompatibility complex (MHC) class I. After recognizing the antigens displayed by APCs, the T cells are activated and ultimately induce apoptosis in cancer cells by secreting molecules such as perforin and granzyme. The first clinically proven mRNA-based cancer vaccine is the mRNA-loaded dendritic cell (DC) vaccine. This innovative treatment involves extracting dendritic cells, differentiating and maturing them in vitro , transfecting them with mRNA using electroporation, and then reinfusing the mRNA-based DC vaccine back into patients. ( Figure A) This strategy has proven effective against several tumor types in clinical trials, including breast cancer, melanoma, ovarian cancer, and glioblastoma. Additionally, in vivo mRNA cancer vaccines are another type of cancer immunotherapy actively being tested in clinical trials due to their lower cost, more convenient manufacturing, and better scalability. A liposomal RNA vaccine was designed targeting four nonmutated TAAs prevalent in melanoma. Natural nucleotide triphosphates and the β-S-ARCA(D1) cap analogue are used for in vitro transcription synthesis of mRNA vaccine. Clinical trial results showed that therapeutic responses in melanoma patients came with strong CD4+ and CD8+ T cell immunity against the vaccine antigens. This suggests that nonmutant shared tumor antigens can be effective targets for mRNA-based cancer vaccination. 18. Open in a new tab Application of modified mRNAs for immunotherapy . ( A ) mRNA-based cancer vaccines are developed by identifying potential antigens from tumor samples using next-generation sequencing (NGS), followed by the design and formulation of mRNA constructs encoding tumor-specific or tumor-associated antigens (TSA/TAA). Vaccination is achieved either through direct administration of the mRNA or by reinfusion of antigen-loaded dendritic cells. ( B ) mRNA-based therapeutic protein production was applied for immunotherapy by in vivo expression of antitumor antibodies, immune regulators, or CAR constructs. Furthermore, personalized mRNA cancer vaccines based on TSAs provide enhanced vaccination efficiency compared to TAAs-based vaccines, as they are designed according to the specific mutations of individual patients. To be more specific, genomic information is acquired from patients’ tumor samples through next-generation sequencing. This allows researchers to identify patient-specific somatic mutations by comparing the tumor data with healthy tissues. Neoantigens for MHC epitopes are predicted using algorithms that guide the design of mRNA-based personalized cancer vaccines. For example, a recent study reported the first-in-human application of individualized mutanome vaccines for patients with melanoma. Using the mRNA with natural nucleotide triphosphates and the β-S-ARCA(D1) cap analogue encoding personalized TSAs, researchers showed that all patients developed T cell responses against multiple vaccine neo-epitopes, confirming that vaccine was able to trigger an immune attack against tumors and demonstrating the potential of mRNA-based personalized cancer vaccines. While promising, a key limitation of personalized mRNA cancer vaccines is the need to obtain sequencing data and design individualized constructs for each patient, which increases costs and limits accessibility. In addition, accurately predicting tumor-specific antigens from sequencing data remains challenging and will require the development of more advanced algorithms. Although several pioneering works have showcased the feasibility of mRNA-based antigen vaccines, the translatability and stability of mRNA constructs encoding antigens have not been systematically investigated or optimized. Using the COVID-19 mRNA vaccine as an example, our group demonstrated that improving mRNA translation efficiency and stability for vaccine development significantly enhanced immune responses. It is foreseeable that further engineering of the mRNA constructs could benefit the design of next-generation mRNA-based cancer vaccines. 7.3.2. mRNA-Based Therapeutic Protein Production In comparison to cancer vaccines that induce a systemic and active immune response by expressing TSAs or TAAs, mRNA-based therapeutic protein production introduces immunotherapeutic or immunomodulatory molecules, such as anticancer antibodies or immunostimulatory cytokines, to regulate the immune response against cancer cells. For instance, mRNA therapeutics have been applied to develop cell therapies, such as CAR-T cells, for cancer immunotherapy. Resembling traditional CAR-T therapy, mRNA encoding the CAR constructs is transfected into T cells isolated from patients, followed by reinfusion of CAR-expressing T cells. mRNA-based CAR-T production offers higher efficiency and raises fewer safety concerns than virus transduction or DNA-based transfection. Higher CAR expression correlates with increased potency in anticancer effects and stronger activation-induced cell death (AICD). More recently, researchers demonstrated the concept of in vivo CAR-T production to reduce cardiac fibrosis and restore cardiac function. Through administrating chemically modified mRNA with a Cap1 structure and fully substituted m 1 ΨTP in T-cell-targeting lipid nanoparticles, CAR can be effectively expressed in T lymphocytes, generating antifibrotic CAR-T cells in situ . ( Figure B) This pioneering work showcased the unique advantages of mRNA technology in the development of CAR-T cell-based therapies, which significantly decrease the cost and simplify the overall manufacturing process. Meanwhile, mRNA-encoded antibodies represent another category of cancer immunotherapy by expressing monoclonal antibodies that target and eliminate cancer cells. ( Figure B) For example, nucleoside-modified mRNA encoding bispecific T-cell-engaging (TCE) antibodies was developed for endogenous antibody synthesis that eliminated advanced tumors as effectively as the corresponding purified antibody. In addition, a recent study developed a cocktail of mRNA-encoded cytokines and T cell costimulators such as IL-23, IL-36γ, and OX40L for anticancer therapy. ( Figure B) When administered intratumorally, this triplet mRNA mixture with chemical modifications recruits immune cells into tumors, converting a “cold tumor” into a “hot tumor” and enabling effective tumor destruction. Impressively, mouse models demonstrated immunity from tumor rechallenge and synergistic effects with existing immune checkpoint inhibition. The study demonstrated an effective, translatable therapy for different tumor cell types by regulating the tumor microenvironment. Contrary to active immunization, such as cancer vaccines, which activate immunity by expressing antigens, passive immunization, including antibody or immunomodulatory proteins, usually requires a relatively high dose of mRNA or repeated dosing to reach the therapeutic threshold and requires a more efficient delivery approach targeting specific tissues. Thus, it is beneficial to apply chemically modified mRNA constructs with enhanced stability and translation efficiency to achieve better therapeutic outcomes at lower cost with reduced toxicity. 7.4. Gene Editing Gene editing has attracted tremendous attention due to its immense therapeutic and scientific potential. During the past few decades, several generations of endonucleases have been exploited and engineered to recognize and cut DNA at specific locations, including meganucleases, zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeats (CRIPSR)/CRISPR-associated nucleases (Cas). , Among these technologies, the CRISPR-Cas system has emerged as the dominant gene editing tool due to its excellent programmability and robustness. The CRISPR-Cas9 system consists of two essential components: the Cas9 enzyme, which acts like molecular scissors to cleave the DNA by generating a double-stranded break (DBS), and guide RNA (gRNA) that guides the Cas9 enzyme to a predefined genomic site through RNA-DNA hybridization. , Although powerful for a variety of applications and therapies, many indications require precise DNA editing without inducing double-stranded breaks. Thus, second-generation genome editing tools have been developed based on the CRISPR-Cas system, including base editors, prime editors, and CRISPR-based epigenetic editors. Base editor modified the CRISPR-Cas9 system through utilizing a Cas9-nickase fused to a base editing enzyme capable of altering individual bases. In contrast, prime editor fuses the Cas9-nickase to a reverse transcriptase to insert genetic information into the genome copying the template sequence within prime editing guide RNA (pegRNA). CRISPRon/off systems function through fusing the Cas protein with transcriptional regulators that modulate gene expression at the epigenetic level. Since the development of these genome-editing tools, several vectors have been exploited to achieve intracellular delivery, including DNA plasmids, ribonucleoprotein (RNP) complexes, adeno-associated virus (AAV), and mRNA-LNP. Among these methods, mRNA-LNP delivery has been intensively investigated due to its high protein expression efficiency, low toxicity and immunogenicity, low risk of insertional mutagenesis, and minimal restriction on cargo size. , Recent work demonstrated the great potential of chemically modified mRNA for delivering gene-editing tools, resulting in enhanced editing efficiency. For example, chemically modified mRNA encoding Cas9 (m 1 Ψ) and gRNA targeting the mouse transthyretin (Ttr) gene was encapsulated into LNPs and injected into a mouse model for the treatment of transthyretin amyloidosis with a dosage of 5 mg/kg. In vivo results demonstrated that a single administration of CRISPR-Cas9 LNP results in significant knockout of the mouse transthyretin (Ttr) gene in the liver, with >97% reduction in serum protein level that persisted over 12 months. Recently, our group synthesized chemically and topologically optimized mRNA constructs with branched poly(A) tails encoding Cas9. Through in vivo administration of modified Cas9 mRNA/gRNA-LNP complex targeting Pcsk9 and Angptl3 in mouse liver, we achieved 13.2-fold higher Cas9 protein expression than the control mRNA. mRNA with a chemically modified, branched poly(A) tail showed more efficient multiplexed genome editing with minimal mRNA dosage (0.7 mg/kg), leading to a higher editing rate (3.3-fold for Pcsk9 editing percentage and 9.0-fold Angptl3 editing percentage) and 71% reduction in serum Pcsk9 and 66% reduction in Angptl3 levels. This example demonstrated that mRNA constructs with longer half-lives significantly benefit multiplexed gene-editing efficiency. Modified mRNAs have also been applied to other genome-editing tools, such as base editors. Researchers discovered that full substitution of uridine with 5-methoxyuridine (5moU) increases the expression level of the mRNA-encoded adenine base editor (ABE), leading to higher A-to-G conversion rates in vitro . Using the chemically modified ABE mRNA, they corrected a splice-site mutation in Tyrosinemia I mouse model with liver damage, harboring a homozygous G•C to A•T point mutation at the last nucleotide of exon 8 in the Fah gene. Additionally, CD117/LNP–mRNA complex was developed, targeting the stem cell factor receptor (CD117) on hematopoietic stem cells (HSCs). Using chemically modified mRNA with Cap1 trinucleotide analogue and m 1 ΨTP encoding either a CRISPR-Cas9 adenine base editor or the pro-apoptotic BH3-only gene PUMA (p53 up-regulated modulator of apoptosis), the researchers corrected a disease mutation in HSCs or depleted HSCs through nongenotoxic conditioning, respectively. Meanwhile, the stability of guide RNA (gRNA) or pegRNA and the other essential components in the gene editing system also significantly influence the overall editing efficiency. Thus, systematic investigations into gRNA or pegRNA modification have been conducted extensively to regulate their stability or interactions with the Cas enzyme/genome, which have been previously reviewed elsewhere. − Furthermore, the kinetics of the editor protein and gRNA/pegRNA are usually nonsynchronous because the concentration peak of the cellular editor protein usually lags behind that of cellular gRNA/pegRNA, which compromises overall editing efficiency. Therefore, it is foreseeable that modified mRNA constructs with higher stability and faster protein production rates will benefit future applications of mRNA-based genome editing. 7.5. Enzyme/Protein Replacement Therapy Enzymes/protein replacement therapy is another promising application of mRNA technology and has been intensively investigated to treat a variety of diseases by producing functional proteins or enzymes in vivo to compensate for or restore the disease phenotype caused by protein loss-of-function or underproduction. The cargo of protein replacement therapy includes hormones, metabolic enzymes, soluble factors, etc. In contrast to other therapeutic applications, such as vaccine or cancer immunotherapy, mRNA-based protein replacement therapy has several additional desired features: (1) prolonged mRNA stability and high protein expression level to reach the higher therapeutic threshold; (2) minimal immunogenicity for optimal protein expression outcome with lower cytotoxicity; (3) redosability to maintain the functional concentration of therapeutic proteins; (4) cell type/tissue-specific expression to avoid the side effects. Thus, in order to meet these requirements and enable a vast array of protein replacement therapies, next-generation mRNA constructs with optimal duration and translation efficiency have been intensively exploited to unlock the promise of mRNA therapeutics. Chemical modifications have broadened the therapeutic effects of mRNAs for ERT. For example, erythropoietin (EPO) is a hormone produced primarily by the kidneys that stimulates the bone marrow to produce red blood cells. Several studies have exploited chemically modified mRNA encoding EPO for in vivo hormone replacement therapy. A pioneering research demonstrated that a single injection of high-performance liquid chromatography (HPLC)-purified, in vitro -transcribed EPO mRNAs containing the modified nucleoside pseudouridine elevated serum EPO level within 6 h in vivo and such level was maintained for 4 days, which is 10–100-fold higher than canonical mRNAs containing uridine. Another study discovered that modified mRNA with m 7 Gppp Bn6 A m pG cap analogue encoding human EPO showed 3-fold increase in protein expression in vivo 24 h post administration. More recently, our group reported the application of RNA LEGO for chemically and topologically optimized mRNA constructs encoding human EPO for hormone replacement therapy. The results demonstrated that on top of m 1 Ψ full substitution, chemically modified dual-cap in combination with tail modification further enhanced the total plasma hEPO level by 8-fold, and induced significant therapeutic effects on the percentage of reticulocytes in vivo . Metabolic diseases have also been an attractive target for mRNA-based therapies. Isolated methylmalonic acidemia/aciduria (MMA) is a devastating metabolic disorder caused by full or partial deficiency of methylmalonyl-CoA mutase (MUT), which is a vitamin B12-dependent mitochondrial enzyme mediating the intracellular metabolism of amino acids and fatty acids. A chemically modified mRNA-LNP complex encoding methylmalonyl-CoA mutase (MUT) with a 5-methoxy-UTP and Cap1 structure was administered to a mouse model with MMA, and the results demonstrated that a single intravenous dose of hMUT mRNA increased hepatic MUT expression, enhanced propionate oxidation, and lowered plasma and tissue methylmalonic acid levels. Similarly, deficiency of the mitochondrial enzyme, propionyl-CoA carboxylase (PCC), composed of PCCA and PCCB subunits, leads to propionic acidemia/aciduria (PA), which is an ultrarare but life-threatening metabolic disorder. mRNAs with m 1 Ψ modification encoding both human PCCA (hPCCA) and PCCB (hPCCB) were developed to restore the expression of functional PCC enzymes in the liver and reduce primary disease-associated toxins in PA mice model. This pioneering work led to the first clinical trial of mRNA-based enzyme replacement therapy, and the preliminary analysis showed that mRNA dosing was well tolerated by patients and alleviated exacerbated PA symptoms. , Furthermore, modified mRNAs with Cap1 and m 1 Ψ substitution have also been applied to encode soluble factors, such as coagulation factors, to treat hemophilia A (HemA) or hemophilia B (HemB). For instance, factor VIII (FVIII) protein-encoding mRNA was developed to treat hemophilia A and the factor IX mRNA was used for hemophilia B treatment. , These pioneering works have demonstrated the great potential of utilizing chemically modified mRNA for protein replacement therapies. We note that there are still unmet challenges. (1) The inherently limited translation efficiency of mRNA in vivo , combined with the need for elevated protein expression above a therapeutic threshold, often necessitates high dosing levels that can trigger toxicity. (2) The short half-life of mRNA typically requires frequent redosing of mRNA-LNP formulations, which makes clinical setting difficult and may cumulatively provoke immune activation, especially in chronic treatment settings, potentially compromising long-term therapeutic efficacy. (3) Targeted delivery or tissue-specific expression methods, especially for tissues beyond the liver, are necessary to expand the therapeutic window and avoid off-target effects. Based on these considerations, further investigation into chemical modifications to generate next-generation mRNA constructs with optimal duration, translation efficiency, low immunogenicity, and enhanced cell-type specificity will enable mRNA therapeutics to address more challenging indications. 8. Challenges and Perspectives Chemical modification plays a vital role in the development of mRNA therapeutics and enables substantial improvement in mRNA properties. Beyond the SARS-CoV-2 vaccine, the potential of mRNA technology is even greater, as more and more new therapies have been investigated over the past few years. It is foreseeable that mRNA-based therapies will meet various medical needs beyond vaccines, cell reprogramming, cancer immunotherapy, genome editing, and protein replacement therapy. To advance next-generation mRNA therapeutics, several key challenges need to be addressed: (1) Maximizing mRNA pharmacokinetic window: mRNA-based therapy enables more efficient protein expression and fewer safety concerns in comparison to other modalities like recombinant proteins, DNA-based methods, or virus-based methods. However, the mRNA molecule suffers from chemical instability and enzymatic degradation. Thus, therapeutic applications that require sustained protein concentrations above the therapeutic threshold will demand multiple dosages. Although this is of less concern for mRNA-based vaccines, RNA’s short half-life will limit clinical implementation in the more challenging applications such as protein replacement therapies requiring multiple magnitudes more efficient production. Thus, further optimization of the pharmacokinetic properties of mRNA molecules will be necessary by introducing chemical modifications in combination with sequence engineering. (2) Enhancing the translation efficiency by expanding the chemical space of mRNA: pioneering works have demonstrated the great potential of fine-tuning the chemical structure of mRNA for a better translation performance. However, due to the limitations on synthesis methodologies, current chemical engineering focuses on the end structures of mRNA (the 5′ and 3′ ends) or full substitution with modified nucleotides, leaving a huge chemical space within mRNA molecules unexplored. More advanced synthesis methods that enable the programmable introduction of chemical modifications in a site-specific or region-specific manner will be necessary to further elucidate the structure–activity relationships of functional mRNAs and guide the design of next-generation mRNA constructs. (3) Cell-type specificity and targeted delivery: beyond the stability and translatability of mRNA constructs, specificity also plays an important role in the development of mRNA-based therapies. We need to consider this to maximize therapeutic effects while minimizing side effects by avoiding off-target protein expression. Previous reports provided valuable information for developing cell-type specific mRNA delivery or creating regulatory methods with tissue or cell-type specificity, such as the introduction of miRNA responsive element to the mRNA, RNA editing-based mRNA activity switch, − riboswitch elements for RNA sensing, untranslational region optimization or development of targeted delivery vehicles. , Developing new platform technologies and combining these methods with specific therapies will expand the therapeutic window for mRNA therapeutics and address more unmet clinical needs. (4) Improving endosomal escape: despite decades of developments in the LNP field, the limited release of mRNA payloads from the endosome to the cytoplasm remains an often overlooked bottleneck for the current LNP-mediated therapeutics. Although several hypotheses have been proposed regarding the mechanism of LNP endosomal escape, such as the lipid-membrane infusion theory or the proton sponge effect, the exact mechanism remains unknown. Thus, an in-depth investigation into the endosomal escape mechanism will accelerate our understanding of the mechanism and facilitate the development of new methods improving mRNA cytoplasmic release. (5) Multiplexed mRNA therapeutics: as demonstrated in vaccines for infectious diseases and cancer, mRNA cocktails encoding multiple antigens can help maximize therapeutic immune effects. Multiplexed mRNA cocktails encoding multiple TFs will also be critical to the future development of stem cell therapy and regenerative medicine. Furthermore, in contrast to rare diseases, common diseases often have polygenic backgrounds and involve complex biological pathways, requiring potential multigene targeting. Therefore, mRNA molecules encoding multiple cargos that can be administered simultaneously are desired to achieve synergistic effects, and chemically and topologically modified mRNA will be critical for multiplexing capacity. (6) Scalable manufacture and distribution: although chemical modifications have significantly improved various properties of mRNA, the introduction of modifications usually involves multiple steps of small molecule synthesis or oligonucleotide chemistries, which increases the cost of mRNA manufacturing, complicates the CMC process, and restricts the large-scale material preparation. Thus, it is valuable to develop synthesis and purification methods that enable scalable material preparation compatible with desired chemical modifications. Furthermore, cold-chain (−80 °C) is always necessary for the distribution of mRNA products due to their intrinsic chemical instability, which limits accessibility and increases costs. Although different LNP formulation strategies have been developed to enhance the storage stability of mRNA-LNP complexes, further investigation and optimization of mRNA chemical structures would provide an alternative method to increase their structural stability, thus enabling more efficient distribution and broader clinical applications of mRNA therapeutics, impacting real patient care. 9. Conclusion mRNA has evolved from a naturally occurring biomolecule to a programmable, multifunctional platform for treating disease. The past two decades have witnessed remarkable progress in both the chemical modification and topological engineering of mRNAs, enabling control over translation efficiency, stability, immunogenicity, and spatial-temporal regulation in living systems. On the chemical front, advances in enzymatic, chemical, and hybrid synthesis strategies have vastly expanded the accessible modification space across all domains of the mRNA molecule (including the cap, UTRs, coding region, poly(A) tail, and backbone). These modifications not only improve expression and stability but also reduce immunogenicity and open new functional modalities for targeted control, imaging, and delivery. On the topological front, mRNA is no longer confined to a linear format. The emergence of circular, branched, and hybrid RNA architectures (e.g., QRNA)enabled by innovations in ligation chemistry, ribozyme design, and chemoenzymatic assemblyintroduces new dimensions of regulatory control. Circular and branched mRNAs offer exceptional stability and translation control, while synthetic topologies enable programmable switches and logic gates for next-generation RNA therapeutics. Yet, despite these advances, the therapeutic outcome of chemically modified and topologically engineered mRNAs remains context-dependent, influenced by RNA sequence and structure, the cellular and tissue environment, and delivery strategy. Addressing these complexities will require integrated approaches that combine high-throughput experimental platforms, data-driven modeling, and structure–activity relationship (SAR) analysis at scale. Looking forward, we envision a future in which therapeutic mRNAs are designed with precision medicinal chemistry akin to small-molecule drugs or engineered proteins: assembled from well characterized modules, tailored to specific delivery and expression goals, and manufactured through streamlined, scalable workflows. In this emerging paradigm, RNA is not just a protein-synthesis template but a programmable platformmodular, tunable, and multifunctionalpoised to redefine the landscape of medicine. Acknowledgments We would like to thank Seth Furniss, Thatcher Lee, Peizhe Ren, and Wendy Wang (MIT and Broad Institute) for the help with manuscript editing and proofreading. X.W. acknowledges the support from the Ono Pharma Breakthrough Science Initiative Award, Merkin Institute Fellowship, Klarman Cell Observatory, Packard Fellowship, Escaping Velocity Award, Sloan Research Fellowship, Dimon Foundation and NIH DP2 New Innovator Award (1DP2GM146245). Biographies Dangliang Liu is a Ph.D. candidate in the Department of Chemistry at Massachusetts Institute of Technology. He received his B.S. in pharmaceutical science (2020) and M.S. in chemical biology (2022) at Peking University with Prof. Suwei Dong. His research interests focus on exploring the chemical space of mRNA and advancing its therapeutic potential. Hongyu Chen is a postdoctoral researcher at the Broad Institute of MIT and Harvard. He received his B.A. in Chemistry (2021) from Boston University with Prof. John Porco and Ph.D. in Chemistry (2025) from Massachusetts Institute of Technology. His research is focused on utilizing synthetic organic chemistry within the context of RNA modification to improve its therapeutic potential. Alisia Pan is a M.D.-Ph.D. student at Harvard Medical School and the Department of Chemistry at Massachusetts Institute of Technology. She received her B.S. in Chemistry (2023) from Yale University with thesis research advised by Prof. Anna Pyle. Her research interests focus on improving cell-type specificity of mRNA therapeutics for translational applications. Xiao Wang is an Associate Professor at MIT Department of Chemistry and Core Institute Member at the Broad Institute of MIT and Harvard. Dr. Wang’s lab is focused developing a molecular toolbox of in situ sequencing methods to study the RNA lifecycle and gene regulation mechanisms in context of tissue structure and function with both single-cell and spatial resolutions, and expanding the alphabet and topology of synthetic mRNA for basic research and RNA therapeutics to explore the mechanism and treatment of human diseases through RNA-centered approaches. She received her B.S. degrees in chemistry and molecular engineering (2010) from Peking University with undergraduate research in Prof. Jian Pei’s lab and a Ph.D. in chemistry from University of Chicago (2015) with Prof. Chuan He, followed by postdoctoral training (2019) as a Life Science Research Fellow at Stanford University in bioengineering and neuroscience with Prof. Karl Deisseroth. †. D.L. and H.C. contributed equally to this work. X.W. conceived the topic and proposed the structure of this review. D.L., H.C., A.P., and X. W. produced the figures and prepared the manuscript. CRediT: Dangliang Liu writing - original draft, writing - review & editing; Hongyu Chen writing - original draft, writing - review & editing; Alisia Pan writing - review & editing; Xiao Wang conceptualization, supervision, writing - original draft, writing - review & editing. The authors declare the following competing financial interest(s): X.W. is a consultant, equity holder and scientific cofounder of Stellaromics and Convergence Bio. Published as part of Chemical Reviews special issue “Synthetic Biology”. References Sahin U., Karikó K., Türeci Ö.. mRNA-Based Therapeutics Developing a New Class of Drugs. Nat. Rev. Drug Discov. 2014;13(10):759–780. doi: 10.1038/nrd4278. [ DOI ] [ PubMed ] [ Google Scholar ] Hogan M. J., Pardi N.. mRNA Vaccines in the COVID-19 Pandemic and Beyond. Annual Review of Medicine. 2022;73(1):17–39. doi: 10.1146/annurev-med-042420-112725. [ DOI ] [ PubMed ] [ Google Scholar ] Xiao Y., Tang Z., Huang X., Chen W., Zhou J., Liu H., Liu C., Kong N., Tao W.. Emerging mRNA Technologies: Delivery Strategies and Biomedical Applications. Chem. Soc. Rev. 2022;51(10):3828–3845. doi: 10.1039/D1CS00617G. [ DOI ] [ PubMed ] [ Google Scholar ] Wolff J. A., Malone R. W., Williams P., Chong W., Acsadi G., Jani A., Felgner P. L.. Direct Gene Transfer into Mouse Muscle in Vivo. Science. 1990;247(4949):1465–1468. doi: 10.1126/science.1690918. [ DOI ] [ PubMed ] [ Google Scholar ] Karikó K., Buckstein M., Ni H., Weissman D.. Suppression of RNA Recognition by Toll-like Receptors: The Impact of Nucleoside Modification and the Evolutionary Origin of RNA. Immunity. 2005;23(2):165–175. doi: 10.1016/j.immuni.2005.06.008. [ DOI ] [ PubMed ] [ Google Scholar ] Rohner E., Yang R., Foo K. S., Goedel A., Chien K. R.. Unlocking the Promise of mRNA Therapeutics. Nat. Biotechnol. 2022;40(11):1586–1600. doi: 10.1038/s41587-022-01491-z. [ DOI ] [ PubMed ] [ Google Scholar ] Mauer J., Luo X., Blanjoie A., Jiao X., Grozhik A. V., Patil D. P., Linder B., Pickering B. F., Vasseur J.-J., Chen Q., Gross S. S., Elemento O., Debart F., Kiledjian M., Jaffrey S. R.. Reversible Methylation of m6Am in the 5′ Cap Controls mRNA Stability. Nature. 2017;541(7637):371–375. doi: 10.1038/nature21022. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Gebhardt A., Habjan M., Benda C., Meiler A., Haas D. A., Hein M. Y., Mann A., Mann M., Habermann B., Pichlmair A.. mRNA Export through an Additional Cap-Binding Complex Consisting of NCBP1 and NCBP3. Nat. Commun. 2015;6(1):8192. doi: 10.1038/ncomms9192. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Aitken C. E., Lorsch J. R.. A Mechanistic Overview of Translation Initiation in Eukaryotes. Nat. Struct Mol. Biol. 2012;19(6):568–576. doi: 10.1038/nsmb.2303. [ DOI ] [ PubMed ] [ Google Scholar ] Abbas Y. M., Laudenbach B. T., Martínez-Montero S., Cencic R., Habjan M., Pichlmair A., Damha M. J., Pelletier J., Nagar B.. Structure of Human IFIT1 with Capped RNA Reveals Adaptable mRNA Binding and Mechanisms for Sensing N1 and N2 Ribose 2′-O Methylations. Proc. Natl. Acad. Sci. U.S.A. 2017;114(11):2106–2115. doi: 10.1073/pnas.1612444114. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Schuberth-Wagner C., Ludwig J., Bruder A. K., Herzner A.-M., Zillinger T., Goldeck M., Schmidt T., Schmid-Burgk J. L., Kerber R., Wolter S., Stümpel J.-P., Roth A., Bartok E., Drosten C., Coch C., Hornung V., Barchet W., Kümmerer B. M., Hartmann G., Schlee M.. A Conserved Histidine in the RNA Sensor RIG-I Controls Immune Tolerance to N1–2′O-Methylated Self RNA. Immunity. 2015;43(1):41–51. doi: 10.1016/j.immuni.2015.06.015. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Ramanathan A., Robb G. B., Chan S.-H.. mRNA Capping: Biological Functions and Applications. Nucleic Acids Res. 2016;44(16):7511–7526. doi: 10.1093/nar/gkw551. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Babendure J. R., Babendure J. L., Ding J.-H., Tsien R. Y.. Control of Mammalian Translation by mRNA Structure near Caps. RNA. 2006;12(5):851–861. doi: 10.1261/rna.2309906. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Mignone F., Gissi C., Liuni S., Pesole G.. Untranslated Regions of mRNAs. Genome Biol. 2002;3(3):reviews0004.1. doi: 10.1186/gb-2002-3-3-reviews0004. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Muckenthaler M., Gray N. K., Hentze M. W.. IRP-1 Binding to Ferritin mRNA Prevents the Recruitment of the Small Ribosomal Subunit by the Cap-Binding Complex eIF4F. Mol. Cell. 1998;2(3):383–388. doi: 10.1016/S1097-2765(00)80282-8. [ DOI ] [ PubMed ] [ Google Scholar ] Hentze M. W., Caughman S. W., Rouault T. A., Barriocanal J. G., Dancis A., Harford J. B., Klausner R. D.. Identification of the Iron-Responsive Element for the Translational Regulation of Human Ferritin mRNA. Science. 1987;238(4833):1570–1573. doi: 10.1126/science.3685996. [ DOI ] [ PubMed ] [ Google Scholar ] Cairo G., Recalcati S.. Iron-Regulatory Proteins: Molecular Biology and Pathophysiological Implications. Expert Rev. Mol. Med. 2007;9(33):1–13. doi: 10.1017/S1462399407000531. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Lee A. S. Y., Kranzusch P. J., Cate J. H. D.. eIF3 Targets Cell-Proliferation Messenger RNAs for Translational Activation or Repression. Nature. 2015;522(7554):111–114. doi: 10.1038/nature14267. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Yang Y., Wang Z.. IRES-Mediated Cap-Independent Translation, a Path Leading to Hidden Proteome. Journal of Molecular Cell Biology. 2019;11(10):911–919. doi: 10.1093/jmcb/mjz091. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Leppek K., Das R., Barna M.. Functional 5′ UTR mRNA Structures in Eukaryotic Translation Regulation and How to Find Them. Nat. Rev. Mol. Cell Biol. 2018;19(3):158–174. doi: 10.1038/nrm.2017.103. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Ding W., Cheng J., Guo D., Mao L., Li J., Lu L., Zhang Y., Yang J., Jiang H.. Engineering the 5′ UTR-Mediated Regulation of Protein Abundance in Yeast Using Nucleotide Sequence Activity Relationships. ACS Synth. Biol. 2018;7(12):2709–2714. doi: 10.1021/acssynbio.8b00127. [ DOI ] [ PubMed ] [ Google Scholar ] Sample P. J., Wang B., Reid D. W., Presnyak V., McFadyen I. J., Morris D. R., Seelig G.. Human 5′ UTR Design and Variant Effect Prediction from a Massively Parallel Translation Assay. Nat. Biotechnol. 2019;37(7):803–809. doi: 10.1038/s41587-019-0164-5. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Cao J., Novoa E. M., Zhang Z., Chen W. C. W., Liu D., Choi G. C. G., Wong A. S. L., Wehrspaun C., Kellis M., Lu T. K.. High-Throughput 5′ UTR Engineering for Enhanced Protein Production in Non-Viral Gene Therapies. Nat. Commun. 2021;12(1):4138. doi: 10.1038/s41467-021-24436-7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Castillo-Hair S., Fedak S., Wang B., Linder J., Havens K., Certo M., Seelig G.. Optimizing 5′UTRs for mRNA-Delivered Gene Editing Using Deep Learning. Nat. Commun. 2024;15(1):5284. doi: 10.1038/s41467-024-49508-2. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Metkar M., Pepin C. S., Moore M. J.. Tailor Made: The Art of Therapeutic mRNA Design. Nat. Rev. Drug Discov. 2024;23:67–83. doi: 10.1038/s41573-023-00827-x. [ DOI ] [ PubMed ] [ Google Scholar ] Presnyak V., Alhusaini N., Chen Y.-H., Martin S., Morris N., Kline N., Olson S., Weinberg D., Baker K. E., Graveley B. R., Coller J.. Codon Optimality Is a Major Determinant of mRNA Stability. Cell. 2015;160(6):1111–1124. doi: 10.1016/j.cell.2015.02.029. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Hanson G., Coller J.. Codon Optimality, Bias and Usage in Translation and mRNA Decay. Nat. Rev. Mol. Cell Biol. 2018;19(1):20–30. doi: 10.1038/nrm.2017.91. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Thess A., Grund S., Mui B. L., Hope M. J., Baumhof P., Fotin-Mleczek M., Schlake T.. Sequence-Engineered mRNA Without Chemical Nucleoside Modifications Enables an Effective Protein Therapy in Large Animals. Molecular Therapy. 2015;23(9):1456–1464. doi: 10.1038/mt.2015.103. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhang H., Zhang L., Lin A., Xu C., Li Z., Liu K., Liu B., Ma X., Zhao F., Jiang H., Chen C., Shen H., Li H., Mathews D. H., Zhang Y., Huang L.. Algorithm for Optimized mRNA Design Improves Stability and Immunogenicity. Nature. 2023;621(7978):396–403. doi: 10.1038/s41586-023-06127-z. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kuersten S., Goodwin E. B.. The Power of the 3′ UTR: Translational Control and Development. Nat. Rev. Genet. 2003;4(8):626–637. doi: 10.1038/nrg1125. [ DOI ] [ PubMed ] [ Google Scholar ] Otsuka H., Fukao A., Funakami Y., Duncan K. E., Fujiwara T.. Emerging Evidence of Translational Control by AU-Rich Element-Binding Proteins. Front. Genet. 2019;10:332. doi: 10.3389/fgene.2019.00332. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Barreau C.. AU-Rich Elements and Associated Factors: Are There Unifying Principles? Nucleic Acids Res. 2005;33(22):7138–7150. doi: 10.1093/nar/gki1012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Dai W., Zhang G., Makeyev E. V.. RNA-Binding Protein HuR Autoregulates Its Expression by Promoting Alternative Polyadenylation Site Usage. Nucleic Acids Res. 2012;40(2):787–800. doi: 10.1093/nar/gkr783. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Shang R., Lee S., Senavirathne G., Lai E. C.. microRNAs in Action: Biogenesis, Function and Regulation. Nat. Rev. Genet. 2023;24(12):816–833. doi: 10.1038/s41576-023-00611-y. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhao W., Pollack J. L., Blagev D. P., Zaitlen N., McManus M. T., Erle D. J.. Massively Parallel Functional Annotation of 3′ Untranslated Regions. Nat. Biotechnol. 2014;32(4):387–391. doi: 10.1038/nbt.2851. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Orlandini Von Niessen A. G., Poleganov M. A., Rechner C., Plaschke A., Kranz L. M., Fesser S., Diken M., Löwer M., Vallazza B., Beissert T., Bukur V., Kuhn A. N., Türeci Ö., Sahin U.. Improving mRNA-Based Therapeutic Gene Delivery by Expression-Augmenting 3′ UTRs Identified by Cellular Library Screening. Molecular Therapy. 2019;27(4):824–836. doi: 10.1016/j.ymthe.2018.12.011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Seo J. J., Jung S.-J., Yang J., Choi D.-E., Kim V. N.. Functional Viromic Screens Uncover Regulatory RNA Elements. Cell. 2023;186(15):3291–3306. doi: 10.1016/j.cell.2023.06.007. [ DOI ] [ PubMed ] [ Google Scholar ] Mugridge J. S., Coller J., Gross J. D.. Structural and Molecular Mechanisms for the Control of Eukaryotic 5′-3′ mRNA Decay. Nat. Struct Mol. Biol. 2018;25(12):1077–1085. doi: 10.1038/s41594-018-0164-z. [ DOI ] [ PubMed ] [ Google Scholar ] Schoenberg D. R., Maquat L. E.. Regulation of Cytoplasmic mRNA Decay. Nat. Rev. Genet. 2012;13(4):246–259. doi: 10.1038/nrg3160. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Passmore L. A., Coller J.. Roles of mRNA Poly(A) Tails in Regulation of Eukaryotic Gene Expression. Nat. Rev. Mol. Cell Biol. 2022;23(2):93–106. doi: 10.1038/s41580-021-00417-y. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Delaunay S., Helm M., Frye M.. RNA Modifications in Physiology and Disease: Towards Clinical Applications. Nat. Rev. Genet. 2024;25(2):104–122. doi: 10.1038/s41576-023-00645-2. [ DOI ] [ PubMed ] [ Google Scholar ] Cui L., Ma R., Cai J., Guo C., Chen Z., Yao L., Wang Y., Fan R., Wang X., Shi Y.. RNA Modifications: Importance in Immune Cell Biology and Related Diseases. Sig Transduct Target Ther. 2022;7(1):334. doi: 10.1038/s41392-022-01175-9. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Sloan K. E., Warda A. S., Sharma S., Entian K.-D., Lafontaine D. L. J., Bohnsack M. T.. Tuning the Ribosome: The Influence of rRNA Modification on Eukaryotic Ribosome Biogenesis and Function. RNA Biology. 2017;14(9):1138–1152. doi: 10.1080/15476286.2016.1259781. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Shi H., Chai P., Jia R., Fan X.. Novel Insight into the Regulatory Roles of Diverse RNA Modifications: Re-Defining the Bridge between Transcription and Translation. Mol. Cancer. 2020;19(1):78. doi: 10.1186/s12943-020-01194-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Suzuki T.. The Expanding World of tRNA Modifications and Their Disease Relevance. Nat. Rev. Mol. Cell Biol. 2021;22(6):375–392. doi: 10.1038/s41580-021-00342-0. [ DOI ] [ PubMed ] [ Google Scholar ] Frye M., Jaffrey S. R., Pan T., Rechavi G., Suzuki T.. RNA Modifications: What Have We Learned and Where Are We Headed? Nat. Rev. Genet. 2016;17(6):365–372. doi: 10.1038/nrg.2016.47. [ DOI ] [ PubMed ] [ Google Scholar ] Hou X., Zaks T., Langer R., Dong Y.. Lipid Nanoparticles for mRNA Delivery. Nat. Rev. Mater. 2021;6(12):1078–1094. doi: 10.1038/s41578-021-00358-0. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chatterjee S., Kon E., Sharma P., Peer D.. Endosomal Escape: A Bottleneck for LNP-Mediated Therapeutics. Proc. Natl. Acad. Sci. U.S.A. 2024;121(11):e2307800120. doi: 10.1073/pnas.2307800120. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Gilleron J., Querbes W., Zeigerer A., Borodovsky A., Marsico G., Schubert U., Manygoats K., Seifert S., Andree C., Stöter M., Epstein-Barash H., Zhang L., Koteliansky V., Fitzgerald K., Fava E., Bickle M., Kalaidzidis Y., Akinc A., Maier M., Zerial M.. Image-Based Analysis of Lipid Nanoparticle-Mediated siRNA Delivery, Intracellular Trafficking and Endosomal Escape. Nat. Biotechnol. 2013;31(7):638–646. doi: 10.1038/nbt.2612. [ DOI ] [ PubMed ] [ Google Scholar ] Liu G., Gack M. U.. Distinct and Orchestrated Functions of RNA Sensors in Innate Immunity. Immunity. 2020;53(1):26–42. doi: 10.1016/j.immuni.2020.03.017. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wang J., Shin B.-S., Alvarado C., Kim J.-R., Bohlen J., Dever T. E., Puglisi J. D.. Rapid 40S Scanning and Its Regulation by mRNA Structure during Eukaryotic Translation Initiation. Cell. 2022;185(24):4474–4487. doi: 10.1016/j.cell.2022.10.005. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jackson R. J., Hellen C. U. T., Pestova T. V.. The Mechanism of Eukaryotic Translation Initiation and Principles of Its Regulation. Nat. Rev. Mol. Cell Biol. 2010;11(2):113–127. doi: 10.1038/nrm2838. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] von der Haar T., Gross J. D., Wagner G., McCarthy J. E. G.. The mRNA Cap-Binding Protein eIF4E in Post-Transcriptional Gene Expression. Nat. Struct Mol. Biol. 2004;11(6):503–511. doi: 10.1038/nsmb779. [ DOI ] [ PubMed ] [ Google Scholar ] Garneau N. L., Wilusz J., Wilusz C. J.. The Highways and Byways of mRNA Decay. Nat. Rev. Mol. Cell Biol. 2007;8(2):113–126. doi: 10.1038/nrm2104. [ DOI ] [ PubMed ] [ Google Scholar ] Brenner S., Jacob F., Meselson M.. An Unstable Intermediate Carrying Information from Genes to Ribosomes for Protein Synthesis. Nature. 1961;190(4776):576–581. doi: 10.1038/190576a0. [ DOI ] [ PubMed ] [ Google Scholar ] Furuichi Y., Miura K.-I.. A Blocked Structure at the 5′ Terminus of mRNA from Cytoplasmic Polyhedrosis Virus. Nature. 1975;253(5490):374–375. doi: 10.1038/253374a0. [ DOI ] [ PubMed ] [ Google Scholar ] Muthukrishnan S., Both G. W., Furuichi Y., Shatkin A. J.. 5′-Terminal 7-Methylguanosine in Eukaryotic mRNA Is Required for Translation. Nature. 1975;255(5503):33–37. doi: 10.1038/255033a0. [ DOI ] [ PubMed ] [ Google Scholar ] Krieg P. A., Melton D. A.. Functional Messenger RNAs Are Produced by SP6 in Vitro Transcription of Cloned cDNAs. Nucleic Acids Res. 1984;12(18):7057–7070. doi: 10.1093/nar/12.18.7057. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jorgensen E. D., Durbin R. K., Risman S. S., McAllister W. T.. Specific Contacts between the Bacteriophage T3, T7, and SP6 RNA Polymerases and Their Promoters. J. Biol. Chem. 1991;266(1):645–651. doi: 10.1016/S0021-9258(18)52483-2. [ DOI ] [ PubMed ] [ Google Scholar ] Imburgio D., Rong M., Ma K., McAllister W. T.. Studies of Promoter Recognition and Start Site Selection by T7 RNA Polymerase Using a Comprehensive Collection of Promoter Variants. Biochemistry. 2000;39(34):10419–10430. doi: 10.1021/bi000365w. [ DOI ] [ PubMed ] [ Google Scholar ] Coleman T. M.. Superior 5′ Homogeneity of RNA from ATP-Initiated Transcription under the T7 2.5 Promoter. Nucleic Acids Res. 2004;32(1):14e–114. doi: 10.1093/nar/gnh007. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Pasquinelli A. E., Dahlberg J. E., Lund E.. Reverse 5′ Caps in RNAs Made in Vitro by Phage RNA Polymerases. RNA. 1995;1(9):957–967. [ PMC free article ] [ PubMed ] [ Google Scholar ] Stepinski J., Waddell C., Stolarski R., Darzynkiewicz E., Rhoads R. E.. Synthesis and Properties of mRNAs Containing the Novel “Anti-Reverse” Cap Analogs 7-Methyl(3′-O-Methyl)GpppG and 7-Methyl (3′-Deoxy)GpppG. RNA. 2001;7(10):1486–1495. [ PMC free article ] [ PubMed ] [ Google Scholar ] Jemielity J., Fowler T., Zuberek J., Stepinski J., Lewdorowicz M., Niedzwiecka A., Stolarski R., Darzynkiewicz E., Rhoads R. E.. Novel “Anti-Reverse” Cap Analogs with Superior Translational Properties. RNA. 2003;9(9):1108–1122. doi: 10.1261/rna.5430403. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Ishikawa M., Murai R., Hagiwara H., Hoshino T., Suyama K.. Preparation of Eukaryotic mRNA Having Differently Methylated Adenosine at the 5′-Terminus and the Effect of the Methyl Group in Translation. Nucleic Acids Symp. Ser. 2009;53(1):129–130. doi: 10.1093/nass/nrp065. [ DOI ] [ PubMed ] [ Google Scholar ] Vaidyanathan S., Azizian K. T., Haque A. K. M. A., Henderson J. M., Hendel A., Shore S., Antony J. S., Hogrefe R. I., Kormann M. S. D., Porteus M. H., McCaffrey A. P.. Uridine Depletion and Chemical Modification Increase Cas9 mRNA Activity and Reduce Immunogenicity without HPLC Purification. Molecular Therapy - Nucleic Acids. 2018;12:530–542. doi: 10.1016/j.omtn.2018.06.010. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Drazkowska K., Tomecki R., Warminski M., Baran N., Cysewski D., Depaix A., Kasprzyk R., Kowalska J., Jemielity J., Sikorski P. J.. 2′- O -Methylation of the Second Transcribed Nucleotide within the mRNA 5′ Cap Impacts the Protein Production Level in a Cell-Specific Manner and Contributes to RNA Immune Evasion. Nucleic Acids Res. 2022;50(16):9051–9071. doi: 10.1093/nar/gkac722. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Senthilvelan A., Vonderfecht T., Shanmugasundaram M., Pal I., Potter J., Kore A. R.. Trinucleotide Cap Analogue Bearing a Locked Nucleic Acid Moiety: Synthesis, mRNA Modification, and Translation for Therapeutic Applications. Org. Lett. 2021;23(11):4133–4136. doi: 10.1021/acs.orglett.1c01037. [ DOI ] [ PubMed ] [ Google Scholar ] Anderson B. R., Muramatsu H., Nallagatla S. R., Bevilacqua P. C., Sansing L. H., Weissman D., Karikó K.. Incorporation of Pseudouridine into mRNA Enhances Translation by Diminishing PKR Activation. Nucleic Acids Res. 2010;38(17):5884–5892. doi: 10.1093/nar/gkq347. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Uchida S., Kataoka K., Itaka K.. Screening of mRNA Chemical Modification to Maximize Protein Expression with Reduced Immunogenicity. Pharmaceutics. 2015;7(3):137–151. doi: 10.3390/pharmaceutics7030137. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kawaguchi D., Kodama A., Abe N., Takebuchi K., Hashiya F., Tomoike F., Nakamoto K., Kimura Y., Shimizu Y., Abe H.. Phosphorothioate Modification of mRNA Accelerates the Rate of Translation Initiation to Provide More Efficient Protein Synthesis. Angew. Chem., Int. Ed. 2020;59(40):17403–17407. doi: 10.1002/anie.202007111. [ DOI ] [ PubMed ] [ Google Scholar ] Strzelecka D., Smietanski M., Sikorski P. J., Warminski M., Kowalska J., Jemielity J.. Phosphodiester Modifications in mRNA Poly(A) Tail Prevent Deadenylation without Compromising Protein Expression. RNA. 2020;26(12):1815–1837. doi: 10.1261/rna.077099.120. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhu B., Hernandez A., Tan M., Wollenhaupt J., Tabor S., Richardson C. C.. Synthesis of 2′-Fluoro RNA by Syn5 RNA Polymerase. Nucleic Acids Res. 2015;43(14):e94. doi: 10.1093/nar/gkv367. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Aurup H., Williams D. M., Eckstein F.. 2’-Fluoro and 2-Amino-2’-Deoxynucleoside 5′-Triphosphates as Substrates for T7 RNA Polymerase. Biochemistry. 1992;31(40):9636–9641. doi: 10.1021/bi00155a016. [ DOI ] [ PubMed ] [ Google Scholar ] Morais P., Adachi H., Yu Y.-T.. The Critical Contribution of Pseudouridine to mRNA COVID-19 Vaccines. Front. Cell Dev. Biol. 2021;9:789427. doi: 10.3389/fcell.2021.789427. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Basu I., Gorai B., Chandran T., Maiti P. K., Hussain T.. Selection of Start Codon during mRNA Scanning in Eukaryotic Translation Initiation. Commun. Biol. 2022;5(1):587. doi: 10.1038/s42003-022-03534-2. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Astatke M., Ng K., Grindley N. D. F., Joyce C. M.. A Single Side Chain Prevents Escherichia Coli DNA Polymerase I (Klenow Fragment) from Incorporating Ribonucleotides. Proc. Natl. Acad. Sci. U.S.A. 1998;95(7):3402–3407. doi: 10.1073/pnas.95.7.3402. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Patel P. H., Loeb L. A.. Multiple Amino Acid Substitutions Allow DNA Polymerases to Synthesize RNA. J. Biol. Chem. 2000;275(51):40266–40272. doi: 10.1074/jbc.M005757200. [ DOI ] [ PubMed ] [ Google Scholar ] Pinheiro V. B., Taylor A. I., Cozens C., Abramov M., Renders M., Zhang S., Chaput J. C., Wengel J., Peak-Chew S.-Y., McLaughlin S. H., Herdewijn P., Holliger P.. Synthetic Genetic Polymers Capable of Heredity and Evolution. Science. 2012;336(6079):341–344. doi: 10.1126/science.1217622. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Cozens C., Pinheiro V. B., Vaisman A., Woodgate R., Holliger P.. A Short Adaptive Path from DNA to RNA Polymerases. Proc. Natl. Acad. Sci. U.S.A. 2012;109(21):8067–8072. doi: 10.1073/pnas.1120964109. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Freund N., Taylor A. I., Arangundy-Franklin S., Subramanian N., Peak-Chew S.-Y., Whitaker A. M., Freudenthal B. D., Abramov M., Herdewijn P., Holliger P.. A Two-Residue Nascent-Strand Steric Gate Controls Synthesis of 2′-O-Methyl- and 2′-O-(2-Methoxyethyl)-RNA. Nat. Chem. 2023;15:91–100. doi: 10.1038/s41557-022-01050-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Ellefson J. W., Gollihar J., Shroff R., Shivram H., Iyer V. R., Ellington A. D.. Synthetic Evolutionary Origin of a Proofreading Reverse Transcriptase. Science. 2016;352(6293):1590–1593. doi: 10.1126/science.aaf5409. [ DOI ] [ PubMed ] [ Google Scholar ] Chen T., Hongdilokkul N., Liu Z., Adhikary R., Tsuen S. S., Romesberg F. E.. Evolution of Thermophilic DNA Polymerases for the Recognition and Amplification of C2′-Modified DNA. Nature Chem. 2016;8(6):556–562. doi: 10.1038/nchem.2493. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Brunderová M., Havlíček V., Matyašovský J., Pohl R., PoštováSlavětínská L., Krömer M., Hocek M.. Expedient Production of Site Specifically Nucleobase-Labelled or Hypermodified RNA with Engineered Thermophilic DNA Polymerases. Nat. Commun. 2024;15(1):3054. doi: 10.1038/s41467-024-47444-9. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Potapov V., Krudup S., Maguire S., Unlu I., Guan S., Buss J. A., Smail B. A., Van Eeuwen T., Taylor M. S., Burns K. H., Ong J. L., Trachman R. J.. Discrete Measurements of RNA Polymerase and Reverse Transcriptase Fidelity Reveal Evolutionary Tuning. RNA. 2024;30(9):1246–1258. doi: 10.1261/rna.080002.124. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Cho E.-J., Takagi T., Moore C. R., Buratowski S.. mRNA Capping Enzyme Is Recruited to the Transcription Complex by Phosphorylation of the RNA Polymerase II Carboxy-Terminal Domain. Genes Dev. 1997;11(24):3319–3326. doi: 10.1101/gad.11.24.3319. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Decroly E., Ferron F., Lescar J., Canard B.. Conventional and Unconventional Mechanisms for Capping Viral mRNA. Nat. Rev. Microbiol. 2012;10(1):51–65. doi: 10.1038/nrmicro2675. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Shuman S., Hurwitz J.. Mechanism of mRNA Capping by Vaccinia Virus Guanylyltransferase: Characterization of an Enzyme-Guanylate Intermediate. Proc. Natl. Acad. Sci. U.S.A. 1981;78(1):187–191. doi: 10.1073/pnas.78.1.187. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kyrieleis O. J. P., Chang J., de la Peña M., Shuman S., Cusack S.. Crystal Structure of Vaccinia Virus mRNA Capping Enzyme Provides Insights into the Mechanism and Evolution of the Capping Apparatus. Structure. 2014;22(3):452–465. doi: 10.1016/j.str.2013.12.014. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Barbosa E., Moss B.. mRNA(Nucleoside-2’-)-Methyltransferase from Vaccinia Virus. Purification and Physical Properties. J. Biol. Chem. 1978;253(21):7692–7697. doi: 10.1016/S0021-9258(17)34425-3. [ DOI ] [ PubMed ] [ Google Scholar ] Hodel A. E., Gershon P. D., Quiocho F. A.. Structural Basis for Sequence-Nonspecific Recognition of 5′-Capped mRNA by a Cap-Modifying Enzyme. Mol. Cell. 1998;1(3):443–447. doi: 10.1016/S1097-2765(00)80044-1. [ DOI ] [ PubMed ] [ Google Scholar ] Fuchs A.-L., Neu A., Sprangers R.. A General Method for Rapid and Cost-Efficient Large-Scale Production of 5′ Capped RNA. RNA. 2016;22(9):1454–1466. doi: 10.1261/rna.056614.116. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Corbett K. S., Edwards D. K., Leist S. R., Abiona O. M., Boyoglu-Barnum S., Gillespie R. A., Himansu S., Schäfer A., Ziwawo C. T., DiPiazza A. T., Dinnon K. H., Elbashir S. M., Shaw C. A., Woods A., Fritch E. J., Martinez D. R., Bock K. W., Minai M., Nagata B. M., Hutchinson G. B., Wu K., Henry C., Bahl K., Garcia-Dominguez D., Ma L., Renzi I., Kong W.-P., Schmidt S. D., Wang L., Zhang Y., Phung E., Chang L. A., Loomis R. J., Altaras N. E., Narayanan E., Metkar M., Presnyak V., Liu C., Louder M. K., Shi W., Leung K., Yang E. S., West A., Gully K. L., Stevens L. J., Wang N., Wrapp D., Doria-Rose N. A., Stewart-Jones G., Bennett H., Alvarado G. S., Nason M. C., Ruckwardt T. J., McLellan J. S., Denison M. R., Chappell J. D., Moore I. N., Morabito K. M., Mascola J. R., Baric R. S., Carfi A., Graham B. S.. SARS-CoV-2 mRNA Vaccine Design Enabled by Prototype Pathogen Preparedness. Nature. 2020;586(7830):567–571. doi: 10.1038/s41586-020-2622-0. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Ohno H., Akamine S., Mochizuki M., Hayashi K., Akichika S., Suzuki T., Saito H.. Versatile Strategy Using Vaccinia Virus-Capping Enzyme to Synthesize Functional 5′ Cap-Modified mRNAs. Nucleic Acids Res. 2023;51(6):e34. doi: 10.1093/nar/gkad019. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Hausmann S., Zheng S., Costanzo M., Brost R. L., Garcin D., Boone C., Shuman S., Schwer B.. Genetic and Biochemical Analysis of Yeast and Human Cap Trimethylguanosine Synthase. J. Biol. Chem. 2008;283(46):31706–31718. doi: 10.1074/jbc.M806127200. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Schulz D., Holstein J. M., Rentmeister A.. A Chemo-Enzymatic Approach for Site-Specific Modification of the RNA Cap. Angew. Chem. Int. Ed. 2013;52(30):7874–7878. doi: 10.1002/anie.201302874. [ DOI ] [ PubMed ] [ Google Scholar ] Michailidou F., Klöcker N., Cornelissen N. V., Singh R. K., Peters A., Ovcharenko A., Kümmel D., Rentmeister A.. Engineered SAM Synthetases for Enzymatic Generation of AdoMet Analogs with Photocaging Groups and Reversible DNA Modification in Cascade Reactions. Angew. Chem. Int. Ed. 2021;60(1):480–485. doi: 10.1002/anie.202012623. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Akichika S., Hirano S., Shichino Y., Suzuki T., Nishimasu H., Ishitani R., Sugita A., Hirose Y., Iwasaki S., Nureki O., Suzuki T.. Cap-Specific Terminal N 6 -Methylation of RNA by an RNA Polymerase II-Associated Methyltransferase. Science. 2019;363(6423):eaav0080. doi: 10.1126/science.aav0080. [ DOI ] [ PubMed ] [ Google Scholar ] van Dülmen M., Muthmann N., Rentmeister A.. Chemo-Enzymatic Modification of the 5′ Cap Maintains Translation and Increases Immunogenic Properties of mRNA. Angew. Chem. Int. Ed. 2021;60(24):13280–13286. doi: 10.1002/anie.202100352. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Bollu A., Peters A., Rentmeister A.. Chemo-Enzymatic Modification of the 5′ Cap To Study mRNAs. Acc. Chem. Res. 2022;55(9):1249–1261. doi: 10.1021/acs.accounts.2c00059. [ DOI ] [ PubMed ] [ Google Scholar ] Velema W. A., Kool E. T.. The Chemistry and Applications of RNA 2′-OH Acylation. Nat. Rev. Chem. 2020;4(1):22–37. doi: 10.1038/s41570-019-0147-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Mortimer S. A., Weeks K. M.. A Fast-Acting Reagent for Accurate Analysis of RNA Secondary and Tertiary Structure by SHAPE Chemistry. J. Am. Chem. Soc. 2007;129(14):4144–4145. doi: 10.1021/ja0704028. [ DOI ] [ PubMed ] [ Google Scholar ] Park H. S., Kietrys A. M., Kool E. T.. Simple Alkanoyl Acylating Agents for Reversible RNA Functionalization and Control. Chem. Commun. 2019;55(35):5135–5138. doi: 10.1039/C9CC01598A. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Spitale R. C., Crisalli P., Flynn R. A., Torre E. A., Kool E. T., Chang H. Y.. RNA SHAPE Analysis in Living Cells. Nat. Chem. Biol. 2013;9(1):18–20. doi: 10.1038/nchembio.1131. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Busan S., Weidmann C. A., Sengupta A., Weeks K. M.. Guidelines for SHAPE Reagent Choice and Detection Strategy for RNA Structure Probing Studies. Biochemistry. 2019;58(23):2655–2664. doi: 10.1021/acs.biochem.8b01218. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Weeks K. M., Mauger D. M.. Exploring RNA Structural Codes with SHAPE Chemistry. Acc. Chem. Res. 2011;44(12):1280–1291. doi: 10.1021/ar200051h. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Fang L., Xiao L., Jun Y. W., Onishi Y., Kool E. T.. Reversible 2′-OH Acylation Enhances RNA Stability. Nat. Chem. 2023;15(9):1296–1305. doi: 10.1038/s41557-023-01246-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Gampe C., White A. C. S., Siva S., Zécri F., Diener J.. 3′-Modification Stabilizes mRNA and Increases Translation in Cells. Bioorg. Med. Chem. Lett. 2018;28(14):2451–2453. doi: 10.1016/j.bmcl.2018.06.008. [ DOI ] [ PubMed ] [ Google Scholar ] Mamot A., Sikorski P. J., Siekierska A., De Witte P., Kowalska J., Jemielity J.. Ethylenediamine Derivatives Efficiently React with Oxidized RNA 3′ Ends Providing Access to Mono and Dually Labelled RNA Probes for Enzymatic Assays and in Vivo Translation. Nucleic Acids Res. 2022;50(1):e3. doi: 10.1093/nar/gkab867. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Balbo P. B., Bohm A.. Mechanism of Poly(A) Polymerase: Structure of the Enzyme-MgATP-RNA Ternary Complex and Kinetic Analysis. Structure. 2007;15(9):1117–1131. doi: 10.1016/j.str.2007.07.010. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wiegand D. J., Rittichier J., Meyer E., Lee H., Conway N. J., Ahlstedt D., Yurtsever Z., Rainone D., Kuru E., Church G. M.. Template-Independent Enzymatic Synthesis of RNA Oligonucleotides. Nat. Biotechnol. 2025;43:762–772. doi: 10.1038/s41587-024-02244-w. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] England T. E., Uhlenbeck O. C.. 3′-Terminal Labelling of RNA with T4 RNA Ligase. Nature. 1978;275(5680):560–561. doi: 10.1038/275560a0. [ DOI ] [ PubMed ] [ Google Scholar ] Tomecki R., Kobylecki K., Drazkowska K., Hyjek-Skladanowska M., Dziembowski A.. Reproducible and Efficient New Method of RNA 3′-End Labelling by CutA Nucleotidyltransferase-Mediated CC-Tailing. RNA Biology. 2021;18(sup2):623–639. doi: 10.1080/15476286.2021.1999104. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Terasaka N., Kimura S., Osawa T., Numata T., Suzuki T.. Biogenesis of 2-Agmatinylcytidine Catalyzed by the Dual Protein and RNA Kinase TiaS. Nat. Struct Mol. Biol. 2011;18(11):1268–1274. doi: 10.1038/nsmb.2121. [ DOI ] [ PubMed ] [ Google Scholar ] Mandal D., Köhrer C., Su D., Russell S. P., Krivos K., Castleberry C. M., Blum P., Limbach P. A., Söll D., RajBhandary U. L.. Agmatidine, a Modified Cytidine in the Anticodon of Archaeal tRNAIle, Base Pairs with Adenosine but Not with Guanosine. Proc. Natl. Acad. Sci. U.S.A. 2010;107(7):2872–2877. doi: 10.1073/pnas.0914869107. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Li F., Dong J., Hu X., Gong W., Li J., Shen J., Tian H., Wang J.. A Covalent Approach for Site-Specific RNA Labeling in Mammalian Cells. Angew. Chem. 2015;127(15):4680–4685. doi: 10.1002/ange.201410433. [ DOI ] [ PubMed ] [ Google Scholar ] Garcia G. A., Kittendorf J. D.. Transglycosylation: A Mechanism for RNA Modification (and Editing?) Bioorganic Chemistry. 2005;33(3):229–251. doi: 10.1016/j.bioorg.2005.01.001. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Alexander S. C., Busby K. N., Cole C. M., Zhou C. Y., Devaraj N. K.. Site-Specific Covalent Labeling of RNA by Enzymatic Transglycosylation. J. Am. Chem. Soc. 2015;137(40):12756–12759. doi: 10.1021/jacs.5b07286. [ DOI ] [ PubMed ] [ Google Scholar ] Busby, K. N. ; Devaraj, N. K. . Enzymatic Covalent Labeling of RNA with RNA Transglycosylation at Guanosine (RNA-TAG). In Methods in Enzymology; Elsevier, 2020; Vol. 641, pp 373–399. 10.1016/bs.mie.2020.03.009. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kimoto M., Hirao I.. Genetic Alphabet Expansion Technology by Creating Unnatural Base Pairs. Chem. Soc. Rev. 2020;49(21):7602–7626. doi: 10.1039/D0CS00457J. [ DOI ] [ PubMed ] [ Google Scholar ] Malyshev D. A., Dhami K., Lavergne T., Chen T., Dai N., Foster J. M., Corrêa I. R., Romesberg F. E.. A Semi-Synthetic Organism with an Expanded Genetic Alphabet. Nature. 2014;509(7500):385–388. doi: 10.1038/nature13314. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhang Y., Ptacin J. L., Fischer E. C., Aerni H. R., Caffaro C. E., San Jose K., Feldman A. W., Turner C. R., Romesberg F. E.. A Semi-Synthetic Organism That Stores and Retrieves Increased Genetic Information. Nature. 2017;551(7682):644–647. doi: 10.1038/nature24659. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Feldman A. W., Romesberg F. E.. Expansion of the Genetic Alphabet: A Chemist’s Approach to Synthetic Biology. Acc. Chem. Res. 2018;51(2):394–403. doi: 10.1021/acs.accounts.7b00403. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Domnick C., Eggert F., Kath-Schorr S.. Site-Specific Enzymatic Introduction of a Norbornene Modified Unnatural Base into RNA and Application in Post-Transcriptional Labeling. Chem. Commun. 2015;51(39):8253–8256. doi: 10.1039/C5CC01765C. [ DOI ] [ PubMed ] [ Google Scholar ] Domnick C., Eggert F., Wuebben C., Bornewasser L., Hagelueken G., Schiemann O., Kath-Schorr S.. EPR Distance Measurements on Long Non-coding RNAs Empowered by Genetic Alphabet Expansion Transcription. Angew. Chem., Int. Ed. 2020;59(20):7891–7896. doi: 10.1002/anie.201916447. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Bornewasser L., Domnick C., Kath-Schorr S.. Stronger Together for In-Cell Translation: Natural and Unnatural Base Modified mRNA. Chem. Sci. 2022;13(17):4753–4761. doi: 10.1039/D2SC00670G. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Hoffmann E. S., De Pascali M. C., Neu L., Domnick C., Soldà A., Kath-Schorr S.. Reverse Transcription as Key Step in RNA in Vitro Evolution with Unnatural Base Pairs. RSC Chem. Biol. 2024;5(6):556–566. doi: 10.1039/D4CB00084F. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Le A. V., Hartman M. C. T.. Improved Synthesis of the Unnatural Base NaM, and Evaluation of Its Orthogonality in in Vitro Transcription and Translation. RSC Chem. Biol. 2024;5(11):1111–1121. doi: 10.1039/D4CB00121D. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Bornewasser L., Domnick C., Kath-Schorr S.. Stronger Together for In-Cell Translation: Natural and Unnatural Base Modified mRNA. Chem. Sci. 2022;13(17):4753–4761. doi: 10.1039/D2SC00670G. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Merrifield R. B.. Solid Phase Peptide Synthesis. I. The Synthesis of a Tetrapeptide. J. Am. Chem. Soc. 1963;85(14):2149–2154. doi: 10.1021/ja00897a025. [ DOI ] [ Google Scholar ] Matteucci M. D., Caruthers M. H.. Synthesis of Deoxyoligonucleotides on a Polymer Support. J. Am. Chem. Soc. 1981;103(11):3185–3191. doi: 10.1021/ja00401a041. [ DOI ] [ PubMed ] [ Google Scholar ] Beaucage S. L., Caruthers M. H.. Deoxynucleoside PhosphoramiditesA New Class of Key Intermediates for Deoxypolynucleotide Synthesis. Tetrahedron Lett. 1981;22(20):1859–1862. doi: 10.1016/S0040-4039(01)90461-7. [ DOI ] [ Google Scholar ] Obexer R., Nassir M., Moody E. R., Baran P. S., Lovelock S. L.. Modern Approaches to Therapeutic Oligonucleotide Manufacturing. Science. 2024;384(6692):eadl4015. doi: 10.1126/science.adl4015. [ DOI ] [ PubMed ] [ Google Scholar ] Shi Y., Zhen X., Zhang Y., Li Y., Koo S., Saiding Q., Kong N., Liu G., Chen W., Tao W.. Chemically Modified Platforms for Better RNA Therapeutics. Chem. Rev. 2024;124(3):929–1033. doi: 10.1021/acs.chemrev.3c00611. [ DOI ] [ PubMed ] [ Google Scholar ] Iwase R., Maeda M., Fujiwara T., Sekine M., Hata T., Miura K.. Molecular Design of a Eukaryotic Messenger RNA and Its Chemical Synthesis. Nucleic Acids Res. 1992;20(7):1643–1648. doi: 10.1093/nar/20.7.1643. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Sawai H., Wakai H., Nakamura-Ozaki A.. Synthesis and Reactions of Nucleoside 5‘-Diphosphate Imidazolide. A Nonenzymatic Capping Agent for 5‘-Monophosphorylated Oligoribonucleotides in Aqueous Solution. J. Org. Chem. 1999;64(16):5836–5840. doi: 10.1021/jo990286u. [ DOI ] [ Google Scholar ] Piecyk K., Davis R. E., Jankowska-Anyszka M.. 5′-Terminal Chemical Capping of Spliced Leader RNAs. Tetrahedron Lett. 2012;53(36):4843–4847. doi: 10.1016/j.tetlet.2012.06.127. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Koukhareva I. I., Lebedev A. V.. Chemical Route to the Capped RNAs. Nucleosides, Nucleotides & Nucleic Acids. 2004;23(10):1667–1680. doi: 10.1081/NCN-200031492. [ DOI ] [ PubMed ] [ Google Scholar ] Nagata S., Hamasaki T., Uetake K., Masuda H., Takagaki K., Oka N., Wada T., Ohgi T., Yano J.. Synthesis and Biological Activity of Artificial mRNA Prepared with Novel Phosphorylating Reagents. Nucleic Acids Res. 2010;38(21):7845–7857. doi: 10.1093/nar/gkq638. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jemielity J., Heinonen P., Lönnberg H., Darzynkiewicz E.. A NOVEL APPROACH TO SOLID PHASE CHEMICAL SYNTHESIS OF OLIGONUCLEOTIDE mRNA CAP ANALOGS. Nucleosides, Nucleotides & Nucleic Acids. 2005;24(5–7):601–605. doi: 10.1081/NCN-200061922. [ DOI ] [ PubMed ] [ Google Scholar ] Leiter J., Reichert D., Rentmeister A., Micura R.. Practical Synthesis of Cap-4 RNA. ChemBioChem. 2020;21(1–2):265–271. doi: 10.1002/cbic.201900590. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Veliath E., Gaffney B. L., Jones R. A.. Synthesis of Capped RNA Using a DMT Group as a Purification Handle. Nucleosides, Nucleotides and Nucleic Acids. 2014;33(1):40–52. doi: 10.1080/15257770.2013.864417. [ DOI ] [ PubMed ] [ Google Scholar ] Abe N., Imaeda A., Inagaki M., Li Z., Kawaguchi D., Onda K., Nakashima Y., Uchida S., Hashiya F., Kimura Y., Abe H.. Complete Chemical Synthesis of Minimal Messenger RNA by Efficient Chemical Capping Reaction. ACS Chem. Biol. 2022;17(6):1308–1314. doi: 10.1021/acschembio.1c00996. [ DOI ] [ PubMed ] [ Google Scholar ] Aditham A., Shi H., Guo J., Zeng H., Zhou Y., Wade S. D., Huang J., Liu J., Wang X.. Chemically Modified mocRNAs for Highly Efficient Protein Expression in Mammalian Cells. ACS Chem. Biol. 2022;17:3352. doi: 10.1021/acschembio.1c00569. [ DOI ] [ PubMed ] [ Google Scholar ] Chen H., Liu D., Aditham A., Guo J., Huang J., Kostas F., Maher K., Friedrich M. J., Xavier R. J., Zhang F., Wang X.. Chemical and Topological Design of Multi-Capped mRNA and Capped Circular RNA for Translation Augmentation. Nat. Biotechnol. 2025;43(7):1128–1143. doi: 10.1038/s41587-024-02393-y. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Hertler J., Slama K., Schober B., Özrendeci Z., Marchand V., Motorin Y., Helm M.. Synthesis of Point-Modified mRNA. Nucleic Acids Res. 2022;50:e115. doi: 10.1093/nar/gkac719. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Contreras R., Cheroutre H., Degrave W., Fiers W.. Simple, Efficient in Vitro Synthesis of Capped RNA Useful for Direct Expression of Cloned Eukaryoti Genes. Nucleic Acids Res. 1982;10(20):6353–6362. doi: 10.1093/nar/10.20.6353. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Houlihan G., Arangundy-Franklin S., Holliger P.. Exploring the Chemistry of Genetic Information Storage and Propagation through Polymerase Engineering. Acc. Chem. Res. 2017;50(4):1079–1087. doi: 10.1021/acs.accounts.7b00056. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Moody E. R., Obexer R., Nickl F., Spiess R., Lovelock S. L.. An Enzyme Cascade Enables Production of Therapeutic Oligonucleotides in a Single Operation. Science. 2023;380(6650):1150–1154. doi: 10.1126/science.add5892. [ DOI ] [ PubMed ] [ Google Scholar ] Wiegand D. J., Rittichier J., Meyer E., Lee H., Conway N. J., Ahlstedt D., Yurtsever Z., Rainone D., Kuru E., Church G. M.. Template-Independent Enzymatic Synthesis of RNA Oligonucleotides. Nat. Biotechnol. 2025;43:762–772. doi: 10.1038/s41587-024-02244-w. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Li F., Dong J., Hu X., Gong W., Li J., Shen J., Tian H., Wang J.. A Covalent Approach for Site-Specific RNA Labeling in Mammalian Cells. Angew. Chem. Int. Ed. 2015;54(15):4597–4602. doi: 10.1002/anie.201410433. [ DOI ] [ PubMed ] [ Google Scholar ] Chen H., Liu D., Guo J., Aditham A., Zhou Y., Tian J., Luo S., Ren J., Hsu A., Huang J., Kostas F., Wu M., Liu D. R., Wang X.. Branched Chemically Modified Poly(A) Tails Enhance the Translation Capacity of mRNA. Nat. Biotechnol. 2025;43:194–203. doi: 10.1038/s41587-024-02174-7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen Y. G., Kim M. V., Chen X., Batista P. J., Aoyama S., Wilusz J. E., Iwasaki A., Chang H. Y.. Sensing Self and Foreign Circular RNAs by Intron Identity. Mol. Cell. 2017;67(2):228–238. doi: 10.1016/j.molcel.2017.05.022. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wesselhoeft R. A., Kowalski P. S., Anderson D. G.. Engineering Circular RNA for Potent and Stable Translation in Eukaryotic Cells. Nat. Commun. 2018;9(1):2629. doi: 10.1038/s41467-018-05096-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen Y. G., Chen R., Ahmad S., Verma R., Kasturi S. P., Amaya L., Broughton J. P., Kim J., Cadena C., Pulendran B., Hur S., Chang H. Y.. N6-Methyladenosine Modification Controls Circular RNA Immunity. Mol. Cell. 2019;76(1):96–109. doi: 10.1016/j.molcel.2019.07.016. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wen S., Qadir J., Yang B. B.. Circular RNA Translation: Novel Protein Isoforms and Clinical Significance. Trends in Molecular Medicine. 2022;28(5):405–420. doi: 10.1016/j.molmed.2022.03.003. [ DOI ] [ PubMed ] [ Google Scholar ] Ali M. M., Li F., Zhang Z., Zhang K., Kang D.-K., Ankrum J. A., Le X. C., Zhao W.. Rolling Circle Amplification: A Versatile Tool for Chemical Biology, Materials Science and Medicine. Chem. Soc. Rev. 2014;43(10):3324. doi: 10.1039/c3cs60439j. [ DOI ] [ PubMed ] [ Google Scholar ] Li J., Mohammed-Elsabagh M., Paczkowski F., Li Y.. Circular Nucleic Acids: Discovery, Functions and Applications. ChemBioChem. 2020;21(11):1547–1566. doi: 10.1002/cbic.202000003. [ DOI ] [ PubMed ] [ Google Scholar ] Sokolova N. I., Ashirbekova D. T., Dolinnaya N. G., Shabarova Z. A.. Chemical Reactions within DNA Duplexes Cyanogen Bromide as an Effective Oligodeoxyribonucleotide Coupling Agent. FEBS Lett. 1988;232(1):153–155. doi: 10.1016/0014-5793(88)80406-X. [ DOI ] [ PubMed ] [ Google Scholar ] Wang S., Kool E. T.. Circular RNA Oligonucleotides. Synthesis, Nucleic Acid Binding Properties, and a Comparison with Circular DNAs. Nucleic Acids Res. 1994;22(12):2326–2333. doi: 10.1093/nar/22.12.2326. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] DOlinnaya N. G., Blumenfeld M., Merenkova I. N., Oretskaya T. S., Krynetskaya N., Ivanovskaya M. G., Vasseur M., Shabarova Z. A.. Oligonucleotide Circularization by Template-Directed Chemical Ligation. Nucleic Acids Res. 1993;21(23):5403–5407. doi: 10.1093/nar/21.23.5403. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Fedorova O. A., Gottikh M. B., Oretskaya T. S., Shabarova Z. A.. Cyanogen Bromide-Induced Chemical Ligation: Mechanism and Optimization of the Reaction Conditions. Nucleosides and Nucleotides. 1996;15(6):1137–1147. doi: 10.1080/07328319608007382. [ DOI ] [ Google Scholar ] Petkovic S., Müller S.. RNA Circularization Strategies in Vivo and in Vitro. Nucleic Acids Res. 2015;43(4):2454–2465. doi: 10.1093/nar/gkv045. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Nakamoto K., Abe N., Tsuji G., Kimura Y., Tomoike F., Shimizu Y., Abe H.. Chemically Synthesized Circular RNAs with Phosphoramidate Linkages Enable Rolling Circle Translation. Chem. Commun. 2020;56(46):6217–6220. doi: 10.1039/D0CC02140G. [ DOI ] [ PubMed ] [ Google Scholar ] Puttaraju M., Been M.. Group I Permuted Intron-Exon (PIE) Sequences Self-Splice to Produce Circular Exons. Nucleic Acids Res. 1992;20(20):5357–5364. doi: 10.1093/nar/20.20.5357. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Qu L., Yi Z., Shen Y., Lin L., Chen F., Xu Y., Wu Z., Tang H., Zhang X., Tian F., Wang C., Xiao X., Dong X., Guo L., Lu S., Yang C., Tang C., Yang Y., Yu W., Wang J., Zhou Y., Huang Q., Yisimayi A., Liu S., Huang W., Cao Y., Wang Y., Zhou Z., Peng X., Wang J., Xie X. S., Wei W.. Circular RNA Vaccines against SARS-CoV-2 and Emerging Variants. Cell. 2022;185(10):1728–1744. doi: 10.1016/j.cell.2022.03.044. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Liu C.-X., Guo S.-K., Nan F., Xu Y.-F., Yang L., Chen L.-L.. RNA Circles with Minimized Immunogenicity as Potent PKR Inhibitors. Mol. Cell. 2022;82(2):420–434. doi: 10.1016/j.molcel.2021.11.019. [ DOI ] [ PubMed ] [ Google Scholar ] Chen R., Wang S. K., Belk J. A., Amaya L., Li Z., Cardenas A., Abe B. T., Chen C.-K., Wender P. A., Chang H. Y.. Engineering Circular RNA for Enhanced Protein Production. Nat. Biotechnol. 2023;41:262–272. doi: 10.1038/s41587-022-01393-0. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Liao K.-C., Eshaghi M., Hong Z., Saw T. Y., Lim J. A. J., Han J., Aw J. G. A., Tan K. Y., Yap A., Gao X., Cheng Y. A., Lim S. Y., Cheang Y. Z. N., Saron W. A. A., Rathore A. P. S., Zhang L., Shunmuganathan B., Gupta R., Tan S. L. I., Qian X., Purushotorman K., Subramaniam N., Vardy L. A., Macary P. A., John A., Yang Y. Y., Alonso S., Song H., Huber R. G., Wan Y.. Characterization of Group I Introns in Generating Circular RNAs as Vaccines. Nucleic Acids Res. 2025;53(4):gkaf089. doi: 10.1093/nar/gkaf089. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Litke J. L., Jaffrey S. R.. Highly Efficient Expression of Circular RNA Aptamers in Cells Using Autocatalytic Transcripts. Nat. Biotechnol. 2019;37(6):667–675. doi: 10.1038/s41587-019-0090-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Unti M. J., Jaffrey S. R.. Highly Efficient Cellular Expression of Circular mRNA Enables Prolonged Protein Expression. Cell Chemical Biology. 2024;31(1):163–176. doi: 10.1016/j.chembiol.2023.09.015. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Miller E. S., Kutter E., Mosig G., Arisaka F., Kunisawa T., Rüger W.. Bacteriophage T4 Genome. Microbiol Mol. Biol. Rev. 2003;67(1):86–156. doi: 10.1128/MMBR.67.1.86-156.2003. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Beadudry D., Perreault J.-P.. An Efficient Strategy for the Synthesis of Circular RNA Molecules. Nucleic Acids Res. 1995;23(15):3064–3066. doi: 10.1093/nar/23.15.3064. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kim Y.-S., Kim D.-H., An D., Lim Y., Seo Y.-J., Kim H. K., Kang H.-Y.. The RNA Ligation Method Using Modified Splint DNAs Significantly Improves the Efficiency of Circular RNA Synthesis. Animal Cells and Systems. 2023;27(1):208–218. doi: 10.1080/19768354.2023.2265165. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Talhouarne G. J. S., Gall J. G.. Lariat Intronic RNAs in the Cytoplasm of Vertebrate Cells. Proc. Natl. Acad. Sci. U.S.A. 2018;115(34):E7970-E7977. doi: 10.1073/pnas.1808816115. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wang Y., Silverman S. K.. Deoxyribozymes That Synthesize Branched and Lariat RNA. J. Am. Chem. Soc. 2003;125(23):6880–6881. doi: 10.1021/ja035150z. [ DOI ] [ PubMed ] [ Google Scholar ] Wang Y., Silverman S. K.. A General Two-Step Strategy to Synthesize Lariat RNAs. RNA. 2006;12(2):313–321. doi: 10.1261/rna.2259406. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wang Y., Silverman S. K.. Efficient One-Step Synthesis of Biologically Related Lariat RNAs by a Deoxyribozyme. Angew. Chem. Int. Ed. 2005;44(36):5863–5866. doi: 10.1002/anie.200501643. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Tago N., Katolik A., Clark N. E., Montemayor E. J., Seio K., Sekine M., Hart P. J., Damha M. J.. Design, Synthesis, and Properties of Phosphoramidate 2′,5′-Linked Branched RNA: Toward the Rational Design of Inhibitors of the RNA Lariat Debranching Enzyme. J. Org. Chem. 2015;80(20):10108–10118. doi: 10.1021/acs.joc.5b01719. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Carriero S., Damha M. J.. Solid-Phase Synthesis of Branched Oligonucleotides. CP Nucleic Acid Chemistry. 2002;9(1):4. doi: 10.1002/0471142700.nc0414s09. (14.1–4.14.32) [ DOI ] [ PubMed ] [ Google Scholar ] Katolik A., Johnsson R., Montemayor E., Lackey J. G., Hart P. J., Damha M. J.. Regiospecific Solid-Phase Synthesis of Branched Oligoribonucleotides That Mimic Intronic Lariat RNA Intermediates. J. Org. Chem. 2014;79(3):963–975. doi: 10.1021/jo4024182. [ DOI ] [ PubMed ] [ Google Scholar ] Torkzaban B., Zhu Y., Lopez C., Alexander J. M., Ma J., Sun Y., Maschhoff K. R., Hu W., Jacob M. H., Lin D., Mao H.-Q., Martin S., Coller J.. Use of Polyadenosine Tail Mimetics to Enhance mRNA Expression from Genes Associated with Haploinsufficiency Disorders. Molecular Therapy Nucleic Acids. 2025;36(1):102453. doi: 10.1016/j.omtn.2025.102453. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Cao Y., Liu H., Lu S. S., Jones K. A., Govind A. P., Jeyifous O., Simmons C. Q., Tabatabaei N., Green W. N., Holder Jimmy. L., Tahmasebi S., George A. L., Dickinson B. C.. RNA-Based Translation Activators for Targeted Gene Upregulation. Nat. Commun. 2023;14(1):6827. doi: 10.1038/s41467-023-42252-z. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Bitounis D., Jacquinet E., Rogers M. A., Amiji M. M.. Strategies to Reduce the Risks of mRNA Drug and Vaccine Toxicity. Nat. Rev. Drug Discov. 2024;23(4):281–300. doi: 10.1038/s41573-023-00859-3. [ DOI ] [ PubMed ] [ Google Scholar ] Karikó K., Muramatsu H., Ludwig J., Weissman D.. Generating the Optimal mRNA for Therapy: HPLC Purification Eliminates Immune Activation and Improves Translation of Nucleoside-Modified, Protein-Encoding mRNA. Nucleic Acids Res. 2011;39(21):e142. doi: 10.1093/nar/gkr695. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Pardi N., Tuyishime S., Muramatsu H., Kariko K., Mui B. L., Tam Y. K., Madden T. D., Hope M. J., Weissman D.. Expression Kinetics of Nucleoside-Modified mRNA Delivered in Lipid Nanoparticles to Mice by Various Routes. J. Controlled Release. 2015;217:345–351. doi: 10.1016/j.jconrel.2015.08.007. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Aviv H., Leder P.. Purification of Biologically Active Globin Messenger RNA by Chromatography on Oligothymidylic Acid-Cellulose. Proc. Natl. Acad. Sci. U.S.A. 1972;69(6):1408–1412. doi: 10.1073/pnas.69.6.1408. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Lukavsky P. J., Puglisi J. D.. Large-Scale Preparation and Purification of Polyacrylamide-Free RNA Oligonucleotides. RNA. 2004;10(5):889–893. doi: 10.1261/rna.5264804. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Baiersdörfer M., Boros G., Muramatsu H., Mahiny A., Vlatkovic I., Sahin U., Karikó K.. A Facile Method for the Removal of dsRNA Contaminant from In Vitro-Transcribed mRNA. Molecular Therapy - Nucleic Acids. 2019;15:26–35. doi: 10.1016/j.omtn.2019.02.018. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Beck J. D., Reidenbach D., Salomon N., Sahin U., Türeci Ö., Vormehr M., Kranz L. M.. mRNA Therapeutics in Cancer Immunotherapy. Mol. Cancer. 2021;20(1):69. doi: 10.1186/s12943-021-01348-0. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Depaix A., Mlynarska-Cieslak A., Warminski M., Sikorski P. J., Jemielity J., Kowalska J.. RNA Ligation for Mono and Dually Labeled RNAs. Chem.Eur. J. 2021;27(47):12190–12197. doi: 10.1002/chem.202101909. [ DOI ] [ PubMed ] [ Google Scholar ] Inagaki M., Abe N., Li Z., Nakashima Y., Acharyya S., Ogawa K., Kawaguchi D., Hiraoka H., Banno A., Meng Z., Tada M., Ishida T., Lyu P., Kokubo K., Murase H., Hashiya F., Kimura Y., Uchida S., Abe H.. Cap Analogs with a Hydrophobic Photocleavable Tag Enable Facile Purification of Fully Capped mRNA with Various Cap Structures. Nat. Commun. 2023;14(1):2657. doi: 10.1038/s41467-023-38244-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Foster J. B., Choudhari N., Perazzelli J., Storm J., Hofmann T. J., Jain P., Storm P. B., Pardi N., Weissman D., Waanders A. J., Grupp S. A., Karikó K., Resnick A. C., Barrett D. M.. Purification of mRNA Encoding Chimeric Antigen Receptor Is Critical for Generation of a Robust T-Cell Response. Hum. Gene Ther. 2019;30(2):168–178. doi: 10.1089/hum.2018.145. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Xiao M.-S., Wilusz J. 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(16):8755–8769. doi: 10.1093/nar/gkz576. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhang J., Chen S., Yang J., Zhao F.. Accurate Quantification of Circular RNAs Identifies Extensive Circular Isoform Switching Events. Nat. Commun. 2020;11(1):90. doi: 10.1038/s41467-019-13840-9. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wei L., Zhang M., Zhang H., Zhao H., Mei Y.. A Poly(A) Polymerase and Oligo(dT)-Dependent Method for the Purification of Engineering Circular RNAs That Enhances Protein Expression in Eukaryotic Cells. Bioengineering. 2024 doi: 10.1101/2024.12.27.630476. [ DOI ] [ Google Scholar ] Selvam C., Mutisya D., Prakash S., Ranganna K., Thilagavathi R.. Therapeutic Potential of Chemically Modified Si RNA: Recent Trends. Chem. Biol. Drug Des. 2017;90(5):665–678. doi: 10.1111/cbdd.12993. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Hu B., Zhong L., Weng Y., Peng L., Huang Y., Zhao Y., Liang X.-J.. Therapeutic siRNA: State of the Art. Sig Transduct Target Ther. 2020;5(1):101. doi: 10.1038/s41392-020-0207-x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Patutina O. A., Gaponova S. K., Sen’kova A. V., Savin I. A., Gladkikh D. V., Burakova E. A., Fokina A. A., Maslov M. A., Shmendel’ E. V., Wood M. J. A., Vlassov V. V., Altman S., Stetsenko D. A., Zenkova M. A.. Mesyl Phosphoramidate Backbone Modified Antisense Oligonucleotides Targeting miR-21 with Enhanced in Vivo Therapeutic Potency. Proc. Natl. Acad. Sci. U.S.A. 2020;117(51):32370–32379. doi: 10.1073/pnas.2016158117. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jia Y., Chiu T.-L., Amin E. A., Polunovsky V., Bitterman P. B., Wagner C. R.. Design, Synthesis and Evaluation of Analogs of Initiation Factor 4E (eIF4E) Cap-Binding Antagonist Bn7-GMP. Eur. J. Med. Chem. 2010;45(4):1304–1313. doi: 10.1016/j.ejmech.2009.11.054. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Šponer J., Bussi G., Krepl M., Banáš P., Bottaro S., Cunha R. A., Gil-Ley A., Pinamonti G., Poblete S., Jurečka P., Walter N. G., Otyepka M.. RNA Structural Dynamics As Captured by Molecular Simulations: A Comprehensive Overview. Chem. Rev. 2018;118(8):4177–4338. doi: 10.1021/acs.chemrev.7b00427. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Lama D., Verma C. S.. Deciphering the Mechanistic Effects of eIF4E Phosphorylation on mRNA-cap Recognition. Protein Sci. 2020;29(6):1373–1386. doi: 10.1002/pro.3798. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Grudzien E., Stepinski J., Jankowska-Anyszka M., Stolarski R., Darzynkiewicz E., Rhoads R. E.. Novel Cap Analogs for in Vitro Synthesis of mRNAs with High Translational Efficiency. RNA. 2004;10(9):1479–1487. doi: 10.1261/rna.7380904. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Su W., Slepenkov S., Grudzien-Nogalska E., Kowalska J., Kulis M., Zuberek J., Lukaszewicz M., Darzynkiewicz E., Jemielity J., Rhoads R. E.. Translation, Stability, and Resistance to Decapping of mRNAs Containing Caps Substituted in the Triphosphate Chain with BH 3, Se, and NH. RNA. 2011;17(5):978–988. doi: 10.1261/rna.2430711. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kore A. R., Shanmugasundaram M., Charles I., Vlassov A. V., Barta T. J.. Locked Nucleic Acid (LNA)-Modified Dinucleotide mRNA Cap Analogue: Synthesis, Enzymatic Incorporation, and Utilization. J. Am. Chem. Soc. 2009;131(18):6364–6365. doi: 10.1021/ja901655p. [ DOI ] [ PubMed ] [ Google Scholar ] Mamot A., Sikorski P. J., Warminski M., Kowalska J., Jemielity J.. Azido-Functionalized 5′ Cap Analogues for the Preparation of Translationally Active mRNAs Suitable for Fluorescent Labeling in Living Cells. Angew. Chem. Int. Ed. 2017;56(49):15628–15632. doi: 10.1002/anie.201709052. [ DOI ] [ PubMed ] [ Google Scholar ] Senthilvelan A., Vonderfecht T., Shanmugasundaram M., Potter J., Kore A. R.. Click-iT Trinucleotide Cap Analog: Synthesis, mRNA Translation, and Detection. Bioorg. Med. Chem. 2023;77:117128. doi: 10.1016/j.bmc.2022.117128. [ DOI ] [ PubMed ] [ Google Scholar ] Wojcik R., Baranowski M. R., Markiewicz L., Kubacka D., Bednarczyk M., Baran N., Wojtczak A., Sikorski P. J., Zuberek J., Kowalska J., Jemielity J.. Novel N7-Arylmethyl Substituted Dinucleotide mRNA 5′ Cap Analogs: Synthesis and Evaluation as Modulators of Translation. Pharmaceutics. 2021;13(11):1941. doi: 10.3390/pharmaceutics13111941. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Sikorski P. J., Warminski M., Kubacka D., Ratajczak T., Nowis D., Kowalska J., Jemielity J.. The Identity and Methylation Status of the First Transcribed Nucleotide in Eukaryotic mRNA 5′ Cap Modulates Protein Expression in Living Cells. Nucleic Acids Res. 2020;48(4):1607–1626. doi: 10.1093/nar/gkaa032. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Cornelissen N. V., Mineikaitė R., Erguven M., Muthmann N., Peters A., Bartels A., Rentmeister A.. Post-Synthetic Benzylation of the mRNA 5′ Cap via Enzymatic Cascade Reactions. Chem. Sci. 2023;14(39):10962–10970. doi: 10.1039/D3SC03822J. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kowalska J., Lewdorowicz M., Zuberek J., Grudzien-Nogalska E., Bojarska E., Stepinski J., Rhoads R. E., Darzynkiewicz E., Davis R. E., Jemielity J.. Synthesis and Characterization of mRNA Cap Analogs Containing Phosphorothioate Substitutions That Bind Tightly to eIF4E and Are Resistant to the Decapping Pyrophosphatase DcpS. RNA. 2008;14(6):1119–1131. doi: 10.1261/rna.990208. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kuhn A. N., Diken M., Kreiter S., Selmi A., Kowalska J., Jemielity J., Darzynkiewicz E., Huber C., Türeci Ö., Sahin U.. Phosphorothioate Cap Analogs Increase Stability and Translational Efficiency of RNA Vaccines in Immature Dendritic Cells and Induce Superior Immune Responses in Vivo. Gene Ther. 2010;17(8):961–971. doi: 10.1038/gt.2010.52. [ DOI ] [ PubMed ] [ Google Scholar ] Kowalska J., Wypijewska del Nogal A., Darzynkiewicz Z. M., Buck J., Nicola C., Kuhn A. N., Lukaszewicz M., Zuberek J., Strenkowska M., Ziemniak M., Maciejczyk M., Bojarska E., Rhoads R. E., Darzynkiewicz E., Sahin U., Jemielity J.. Synthesis, Properties, and Biological Activity of Boranophosphate Analogs of the mRNA Cap: Versatile Tools for Manipulation of Therapeutically Relevant Cap-Dependent Processes. Nucleic Acids Res. 2014;42(16):10245–10264. doi: 10.1093/nar/gku757. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Strenkowska M., Grzela R., Majewski M., Wnek K., Kowalska J., Lukaszewicz M., Zuberek J., Darzynkiewicz E., Kuhn A. N., Sahin U., Jemielity J.. Cap Analogs Modified with 1,2-Dithiodiphosphate Moiety Protect mRNA from Decapping and Enhance Its Translational Potential. Nucleic Acids Res. 2016:gkw896. doi: 10.1093/nar/gkw896. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Walczak S., Sikorski P. J., Kasprzyk R., Kowalska J., Jemielity J.. Exploring the Potential of Phosphotriazole 5′ mRNA Cap Analogues as Efficient Translation Initiators. Org. Biomol. Chem. 2018;16(36):6741–6748. doi: 10.1039/C8OB01720D. [ DOI ] [ PubMed ] [ Google Scholar ] Wojtczak B. A., Sikorski P. J., Fac-Dabrowska K., Nowicka A., Warminski M., Kubacka D., Nowak E., Nowotny M., Kowalska J., Jemielity J.. 5′-Phosphorothiolate Dinucleotide Cap Analogues: Reagents for Messenger RNA Modification and Potent Small-Molecular Inhibitors of Decapping Enzymes. J. Am. Chem. Soc. 2018;140(18):5987–5999. doi: 10.1021/jacs.8b02597. [ DOI ] [ PubMed ] [ Google Scholar ] Kozarski M., Drazkowska K., Bednarczyk M., Warminski M., Jemielity J., Kowalska J.. Towards Superior mRNA Caps Accessible by Click Chemistry: Synthesis and Translational Properties of Triazole-Bearing Oligonucleotide Cap Analogs. RSC Adv. 2023;13(19):12809–12824. doi: 10.1039/D3RA00026E. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Warminski M., Kowalska J., Nowak E., Kubacka D., Tibble R., Kasprzyk R., Sikorski P. J., Gross J. D., Nowotny M., Jemielity J.. Structural Insights into the Interaction of Clinically Relevant Phosphorothioate mRNA Cap Analogs with Translation Initiation Factor 4E Reveal Stabilization via Electrostatic Thio-Effect. ACS Chem. Biol. 2021;16(2):334–343. doi: 10.1021/acschembio.0c00864. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhang H.-J., Ociepa M., Nassir M., Zheng B., Lewicki S. A., Salmaso V., Baburi H., Nagel J., Mirza S., Bueschbell B., Al-Hroub H., Perzanowska O., Lin Z., Schmidt M. A., Eastgate M. D., Jacobson K. A., Müller C. E., Kowalska J., Jemielity J., Baran P. S.. Stereocontrolled Access to Thioisosteres of Nucleoside Di- and Triphosphates. Nat. Chem. 2024;16:249–258. doi: 10.1038/s41557-023-01347-2. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Gunawardana D., Domashevskiy A. V., Gayler K. R., Goss D. J.. Efficient Preparation and Properties of mRNAs Containing a Fluorescent Cap Analog: Anthraniloyl-m7 GpppG. Translation. 2015;3(1):e988538. doi: 10.4161/21690731.2014.988538. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jemielity J., Lukaszewicz M., Kowalska J., Czarnecki J., Zuberek J., Darzynkiewicz E.. Synthesis of Biotin Labelled Cap Analogue - Incorporable into mRNA Transcripts and Promoting Cap-Dependent Translation. Org. Biomol. Chem. 2012;10(43):8570. doi: 10.1039/c2ob26060c. [ DOI ] [ PubMed ] [ Google Scholar ] Nowakowska M., Kowalska J., Martin F., d’Orchymont A., Zuberek J., Lukaszewicz M., Darzynkiewicz E., Jemielity J.. Cap Analogs Containing 6-Thioguanosine - Reagents for the Synthesis of mRNAs Selectively Photo-Crosslinkable with Cap-Binding Biomolecules. Org. Biomol. Chem. 2014;12(27):4841–4847. doi: 10.1039/C4OB00059E. [ DOI ] [ PubMed ] [ Google Scholar ] Warminski M., Trepkowska E., Smietanski M., Sikorski P. J., Baranowski M. R., Bednarczyk M., Kedzierska H., Majewski B., Mamot A., Papiernik D., Popielec A., Serwa R. A., Shimanski B. A., Sklepkiewicz P., Sklucka M., Sokolowska O., Spiewla T., Toczydlowska-Socha D., Warminska Z., Wolosewicz K., Zuberek J., Mugridge J. S., Nowis D., Golab J., Jemielity J., Kowalska J.. Trinucleotide mRNA Cap Analogue N 6-Benzylated at the Site of Posttranscriptional m6 A m Mark Facilitates mRNA Purification and Confers Superior Translational Properties In Vitro and In Vivo. J. Am. Chem. Soc. 2024;146(12):8149–8163. doi: 10.1021/jacs.3c12629. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen H., Liu D., Aditham A., Guo J., Huang J., Kostas F., Maher K., Friedrich M. J., Xavier R. J., Zhang F., Wang X.. Chemical and Topological Design of Multicapped mRNA and Capped Circular RNA to Augment Translation. Nat. Biotechnol. 2025;43:1128–1143. doi: 10.1038/s41587-024-02393-y. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Karikó K., Muramatsu H., Welsh F. A., Ludwig J., Kato H., Akira S., Weissman D.. Incorporation of Pseudouridine Into mRNA Yields Superior Nonimmunogenic Vector With Increased Translational Capacity and Biological Stability. Molecular Therapy. 2008;16(11):1833–1840. doi: 10.1038/mt.2008.200. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Andries O., Mc Cafferty S., De Smedt S. C., Weiss R., Sanders N. N., Kitada T.. N1-Methylpseudouridine-Incorporated mRNA Outperforms Pseudouridine-Incorporated mRNA by Providing Enhanced Protein Expression and Reduced Immunogenicity in Mammalian Cell Lines and Mice. J. Controlled Release. 2015;217:337–344. doi: 10.1016/j.jconrel.2015.08.051. [ DOI ] [ PubMed ] [ Google Scholar ] Durbin A. F., Wang C., Marcotrigiano J., Gehrke L.. RNAs Containing Modified Nucleotides Fail To Trigger RIG-I Conformational Changes for Innate Immune Signaling. mBio. 2016;7(5):e00833-16. doi: 10.1128/mBio.00833-16. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Mauger D. M., Cabral B. J., Presnyak V., Su S. V., Reid D. W., Goodman B., Link K., Khatwani N., Reynders J., Moore M. J., McFadyen I. J.. mRNA Structure Regulates Protein Expression through Changes in Functional Half-Life. Proc. Natl. Acad. Sci. U.S.A. 2019;116(48):24075–24083. doi: 10.1073/pnas.1908052116. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Svitkin Y. V., Cheng Y. M., Chakraborty T., Presnyak V., John M., Sonenberg N.. N1-Methyl-Pseudouridine in mRNA Enhances Translation through eIF2α-Dependent and Independent Mechanisms by Increasing Ribosome Density. Nucleic Acids Res. 2017;45(10):6023–6036. doi: 10.1093/nar/gkx135. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Nelson J., Sorensen E. W., Mintri S., Rabideau A. E., Zheng W., Besin G., Khatwani N., Su S. V., Miracco E. J., Issa W. J., Hoge S., Stanton M. G., Joyal J. L.. Impact of mRNA Chemistry and Manufacturing Process on Innate Immune Activation. Sci. Adv. 2020;6(26):eaaz6893. doi: 10.1126/sciadv.aaz6893. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Nance K. D., Meier J. L.. Modifications in an Emergency: The Role of N1-Methylpseudouridine in COVID-19 Vaccines. ACS Cent. Sci. 2021;7(5):748–756. doi: 10.1021/acscentsci.1c00197. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Monroe J., Eyler D. E., Mitchell L., Deb I., Bojanowski A., Srinivas P., Dunham C. M., Roy B., Frank A. T., Koutmou K. S.. N1-Methylpseudouridine and Pseudouridine Modifications Modulate mRNA Decoding during Translation. Nat. Commun. 2024;15(1):8119. doi: 10.1038/s41467-024-51301-0. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Bérouti M., Wagner M., Greulich W., Piseddu I., Gärtig J., Hansbauer L., Müller-Hermes C., Heiss M., Pichler A., Tölke A. J., Witte G., Hopfner K.-P., Anz D., Sattler M., Carell T., Hornung V.. Pseudouridine RNA Avoids Immune Detection through Impaired Endolysosomal Processing and TLR Engagement. Cell. 2025;188(18):4880. doi: 10.1016/j.cell.2025.05.032. [ DOI ] [ PubMed ] [ Google Scholar ] Kim M., Pyo Y., Hyun S.-I., Jeong M., Choi Y., Kim V. N.. Exogenous RNA Surveillance by Proton-Sensing TRIM25. Science. 2025;388(6742):eads4539. doi: 10.1126/science.ads4539. [ DOI ] [ PubMed ] [ Google Scholar ] Eyler D. E., Franco M. K., Batool Z., Wu M. Z., Dubuke M. L., Dobosz-Bartoszek M., Jones J. D., Polikanov Y. S., Roy B., Koutmou K. S.. Pseudouridinylation of mRNA Coding Sequences Alters Translation. Proc. Natl. Acad. Sci. U.S.A. 2019;116(46):23068–23074. doi: 10.1073/pnas.1821754116. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Mulroney T. E., Pöyry T., Yam-Puc J. C., Rust M., Harvey R. F., Kalmar L., Horner E., Booth L., Ferreira A. P., Stoneley M., Sawarkar R., Mentzer A. J., Lilley K. S., Smales C. M., Von Der Haar T., Turtle L., Dunachie S., Klenerman P., Thaventhiran J. E. D., Willis A. E.. N1-Methylpseudouridylation of mRNA Causes + 1 Ribosomal Frameshifting. Nature. 2024;625(7993):189–194. doi: 10.1038/s41586-023-06800-3. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jiang T., Henderson J. M., Coote K., Cheng Y., Valley H. C., Zhang X.-O., Wang Q., Rhym L. H., Cao Y., Newby G. A., Bihler H., Mense M., Weng Z., Anderson D. G., McCaffrey A. P., Liu D. R., Xue W.. Chemical Modifications of Adenine Base Editor mRNA and Guide RNA Expand Its Application Scope. Nat. Commun. 2020;11(1):1979. doi: 10.1038/s41467-020-15892-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhang Y., Xi X., Yu H., Yang L., Lin J., Yang W., Liu J., Fan X., Xu Y.. Chemically Modified In-Vitro-Transcribed mRNA Encoding Thrombopoietin Stimulates Thrombopoiesis in Mice. Molecular Therapy - Nucleic Acids. 2022;29:657–671. doi: 10.1016/j.omtn.2022.08.017. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Choi J., Ieong K.-W., Demirci H., Chen J., Petrov A., Prabhakar A., O’Leary S. E., Dominissini D., Rechavi G., Soltis S. M., Ehrenberg M., Puglisi J. D.. N6-Methyladenosine in mRNA Disrupts tRNA Selection and Translation-Elongation Dynamics. Nat. Struct Mol. Biol. 2016;23(2):110–115. doi: 10.1038/nsmb.3148. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Arango D., Sturgill D., Alhusaini N., Dillman A. A., Sweet T. J., Hanson G., Hosogane M., Sinclair W. R., Nanan K. K., Mandler M. D., Fox S. D., Zengeya T. T., Andresson T., Meier J. L., Coller J., Oberdoerffer S.. Acetylation of Cytidine in mRNA Promotes Translation Efficiency. Cell. 2018;175(7):1872–1886. doi: 10.1016/j.cell.2018.10.030. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Nance K. D., Gamage S. T., Alam M. M., Yang A., Levy M. J., Link C. N., Florens L., Washburn M. P., Gu S., Oppenheim J. J., Meier J. L.. Cytidine Acetylation Yields a Hypoinflammatory Synthetic Messenger RNA. Cell Chemical Biology. 2022;29(2):312–320. doi: 10.1016/j.chembiol.2021.07.003. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Choi J., Indrisiunaite G., DeMirci H., Ieong K.-W., Wang J., Petrov A., Prabhakar A., Rechavi G., Dominissini D., He C., Ehrenberg M., Puglisi J. D.. 2′-O-Methylation in mRNA Disrupts tRNA Decoding during Translation Elongation. Nat. Struct Mol. Biol. 2018;25(3):208–216. doi: 10.1038/s41594-018-0030-z. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kawaguchi D., Kodama A., Abe N., Takebuchi K., Hashiya F., Tomoike F., Nakamoto K., Kimura Y., Shimizu Y., Abe H.. Phosphorothioate Modification of mRNA Accelerates the Rate of Translation Initiation to Provide More Efficient Protein Synthesis. Angew. Chem. Int. Ed. 2020;59(40):17403–17407. doi: 10.1002/anie.202007111. [ DOI ] [ PubMed ] [ Google Scholar ] Meyer K. D., Patil D. P., Zhou J., Zinoviev A., Skabkin M. A., Elemento O., Pestova T. V., Qian S.-B., Jaffrey S. R.. 5′ UTR m6A Promotes Cap-Independent Translation. Cell. 2015;163(4):999–1010. doi: 10.1016/j.cell.2015.10.012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Anhäuser L., Hüwel S., Zobel T., Rentmeister A.. Multiple Covalent Fluorescence Labeling of Eukaryotic mRNA at the Poly(A) Tail Enhances Translation and Can Be Performed in Living Cells. Nucleic Acids Res. 2019;47(7):e42. doi: 10.1093/nar/gkz084. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kristensen L. S., Andersen M. S., Stagsted L. V. W., Ebbesen K. K., Hansen T. B., Kjems J.. The Biogenesis, Biology and Characterization of Circular RNAs. Nat. Rev. Genet. 2019;20(11):675–691. doi: 10.1038/s41576-019-0158-7. [ DOI ] [ PubMed ] [ Google Scholar ] Liu C.-X., Chen L.-L.. Circular RNAs: Characterization, Cellular Roles, and Applications. Cell. 2022;185(12):2016–2034. doi: 10.1016/j.cell.2022.04.021. [ DOI ] [ PubMed ] [ Google Scholar ] Legnini I., Di Timoteo G., Rossi F., Morlando M., Briganti F., Sthandier O., Fatica A., Santini T., Andronache A., Wade M., Laneve P., Rajewsky N., Bozzoni I.. Circ-ZNF609 Is a Circular RNA That Can Be Translated and Functions in Myogenesis. Mol. Cell. 2017;66(1):22–37. doi: 10.1016/j.molcel.2017.02.017. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Pamudurti N. R., Bartok O., Jens M., Ashwal-Fluss R., Stottmeister C., Ruhe L., Hanan M., Wyler E., Perez-Hernandez D., Ramberger E., Shenzis S., Samson M., Dittmar G., Landthaler M., Chekulaeva M., Rajewsky N., Kadener S.. Translation of CircRNAs. Mol. Cell. 2017;66(1):9–21. doi: 10.1016/j.molcel.2017.02.021. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Yang Y., Gao X., Zhang M., Yan S., Sun C., Xiao F., Huang N., Yang X., Zhao K., Zhou H., Huang S., Xie B., Zhang N.. Novel Role of FBXW7 Circular RNA in Repressing Glioma Tumorigenesis. JNCI: Journal of the National Cancer Institute. 2018;110(3):304–315. doi: 10.1093/jnci/djx166. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhang M., Zhao K., Xu X., Yang Y., Yan S., Wei P., Liu H., Xu J., Xiao F., Zhou H., Yang X., Huang N., Liu J., He K., Xie K., Zhang G., Huang S., Zhang N.. A Peptide Encoded by Circular Form of LINC-PINT Suppresses Oncogenic Transcriptional Elongation in Glioblastoma. Nat. Commun. 2018;9(1):4475. doi: 10.1038/s41467-018-06862-2. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Yang Y., Fan X., Mao M., Song X., Wu P., Zhang Y., Jin Y., Yang Y., Chen L.-L., Wang Y., Wong C. C., Xiao X., Wang Z.. Extensive Translation of Circular RNAs Driven by N6-Methyladenosine. Cell Res. 2017;27(5):626–641. doi: 10.1038/cr.2017.31. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Fan X., Yang Y., Chen C., Wang Z.. Pervasive Translation of Circular RNAs Driven by Short IRES-like Elements. Nat. Commun. 2022;13(1):3751. doi: 10.1038/s41467-022-31327-y. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wesselhoeft R. A., Kowalski P. S., Parker-Hale F. C., Huang Y., Bisaria N., Anderson D. G.. RNA Circularization Diminishes Immunogenicity and Can. Extend Translation Duration In Vivo. Mol. Cell. 2019;74(3):508–520. doi: 10.1016/j.molcel.2019.02.015. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen C.-K., Cheng R., Demeter J., Chen J., Weingarten-Gabbay S., Jiang L., Snyder M. P., Weissman J. S., Segal E., Jackson P. K., Chang H. Y.. Structured Elements Drive Extensive Circular RNA Translation. Mol. Cell. 2021;81(20):4300–4318. doi: 10.1016/j.molcel.2021.07.042. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen Y. G., Chen R., Ahmad S., Verma R., Kasturi S. P., Amaya L., Broughton J. P., Kim J., Cadena C., Pulendran B., Hur S., Chang H. Y.. N6-Methyladenosine Modification Controls Circular RNA Immunity. Mol. Cell. 2019;76(1):96–109. doi: 10.1016/j.molcel.2019.07.016. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Takahashi K., Yamanaka S.. Induction of Pluripotent Stem Cells from Mouse Embryonic and Adult Fibroblast Cultures by Defined Factors. Cell. 2006;126(4):663–676. doi: 10.1016/j.cell.2006.07.024. [ DOI ] [ PubMed ] [ Google Scholar ] Robinton D. A., Daley G. Q.. The Promise of Induced Pluripotent Stem Cells in Research and Therapy. Nature. 2012;481(7381):295–305. doi: 10.1038/nature10761. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Warren L., Manos P. D., Ahfeldt T., Loh Y.-H., Li H., Lau F., Ebina W., Mandal P. K., Smith Z. D., Meissner A., Daley G. Q., Brack A. S., Collins J. J., Cowan C., Schlaeger T. M., Rossi D. J.. Highly Efficient Reprogramming to Pluripotency and Directed Differentiation of Human Cells with Synthetic Modified mRNA. Cell Stem Cell. 2010;7(5):618–630. doi: 10.1016/j.stem.2010.08.012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Yoshioka N., Gros E., Li H.-R., Kumar S., Deacon D. C., Maron C., Muotri A. R., Chi N. C., Fu X.-D., Yu B. D., Dowdy S. F.. Efficient Generation of Human iPSCs by a Synthetic Self-Replicative RNA. Cell Stem Cell. 2013;13(2):246–254. doi: 10.1016/j.stem.2013.06.001. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kogut I., McCarthy S. M., Pavlova M., Astling D. P., Chen X., Jakimenko A., Jones K. L., Getahun A., Cambier J. C., Pasmooij A. M. G., Jonkman M. F., Roop D. R., Bilousova G.. High-Efficiency RNA-Based Reprogramming of Human Primary Fibroblasts. Nat. Commun. 2018;9(1):745. doi: 10.1038/s41467-018-03190-3. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Preskey D., Allison T. F., Jones M., Mamchaoui K., Unger C.. Synthetically Modified mRNA for Efficient and Fast Human iPS Cell Generation and Direct Transdifferentiation to Myoblasts. Biochem. Biophys. Res. Commun. 2016;473(3):743–751. doi: 10.1016/j.bbrc.2015.09.102. [ DOI ] [ PubMed ] [ Google Scholar ] Wang X., Yu L., Ding Y., Guo X., Yuan Y., Li D.. Gene Manipulation of Human Embryonic Stem Cells by In Vitro-Synthesized mRNA for Gene Therapy. CGT. 2015;15(4):428–435. doi: 10.2174/1566523215666150515144533. [ DOI ] [ PubMed ] [ Google Scholar ] Goparaju S. K., Kohda K., Ibata K., Soma A., Nakatake Y., Akiyama T., Wakabayashi S., Matsushita M., Sakota M., Kimura H., Yuzaki M., Ko S. B. H., Ko M. S. H.. Rapid Differentiation of Human Pluripotent Stem Cells into Functional Neurons by mRNAs Encoding Transcription Factors. Sci. Rep. 2017;7(1):42367. doi: 10.1038/srep42367. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Wang A. Y. L.. Application of Modified mRNA in Somatic Reprogramming to Pluripotency and Directed Conversion of Cell Fate. IJMS. 2021;22(15):8148. doi: 10.3390/ijms22158148. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Vogel A. B., Kanevsky I., Che Y., Swanson K. A., Muik A., Vormehr M., Kranz L. M., Walzer K. C., Hein S., Güler A., Loschko J., Maddur M. S., Ota-Setlik A., Tompkins K., Cole J., Lui B. G., Ziegenhals T., Plaschke A., Eisel D., Dany S. C., Fesser S., Erbar S., Bates F., Schneider D., Jesionek B., Sänger B., Wallisch A.-K., Feuchter Y., Junginger H., Krumm S. A., Heinen A. P., Adams-Quack P., Schlereth J., Schille S., Kröner C., De La Caridad Güimil Garcia R., Hiller T., Fischer L., Sellers R. S., Choudhary S., Gonzalez O., Vascotto F., Gutman M. R., Fontenot J. A., Hall-Ursone S., Brasky K., Griffor M. C., Han S., Su A. A. H., Lees J. A., Nedoma N. L., Mashalidis E. H., Sahasrabudhe P. V., Tan C. Y., Pavliakova D., Singh G., Fontes-Garfias C., Pride M., Scully I. L., Ciolino T., Obregon J., Gazi M., Carrion R., Alfson K. J., Kalina W. V., Kaushal D., Shi P.-Y., Klamp T., Rosenbaum C., Kuhn A. N., Türeci Ö., Dormitzer P. R., Jansen K. U., Sahin U.. BNT162b Vaccines Protect Rhesus Macaques from SARS-CoV-2. Nature. 2021;592(7853):283–289. doi: 10.1038/s41586-021-03275-y. [ DOI ] [ PubMed ] [ Google Scholar ] Pardi N., Hogan M. J., Pelc R. S., Muramatsu H., Andersen H., DeMaso C. R., Dowd K. A., Sutherland L. L., Scearce R. M., Parks R., Wagner W., Granados A., Greenhouse J., Walker M., Willis E., Yu J.-S., McGee C. E., Sempowski G. D., Mui B. L., Tam Y. K., Huang Y.-J., Vanlandingham D., Holmes V. M., Balachandran H., Sahu S., Lifton M., Higgs S., Hensley S. E., Madden T. D., Hope M. J., Karikó K., Santra S., Graham B. S., Lewis M. G., Pierson T. C., Haynes B. F., Weissman D.. Zika Virus Protection by a Single Low-Dose Nucleoside-Modified mRNA Vaccination. Nature. 2017;543(7644):248–251. doi: 10.1038/nature21428. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Arevalo C. P., Bolton M. J., Le Sage V., Ye N., Furey C., Muramatsu H., Alameh M.-G., Pardi N., Drapeau E. M., Parkhouse K., Garretson T., Morris J. S., Moncla L. H., Tam Y. K., Fan S. H. Y., Lakdawala S. S., Weissman D., Hensley S. E.. A Multivalent Nucleoside-Modified mRNA Vaccine against All Known Influenza Virus Subtypes. Science. 2022;378(6622):899–904. doi: 10.1126/science.abm0271. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Feldman R. A., Fuhr R., Smolenov I., Ribeiro A. M., Panther L., Watson M., Senn J. J., Smith M., Almarsson Ö., Pujar H. S., Laska M. E., Thompson J., Zaks T., Ciaramella G.. mRNA Vaccines against H10N8 and H7N9 Influenza Viruses of Pandemic Potential Are Immunogenic and Well Tolerated in Healthy Adults in Phase 1 Randomized Clinical Trials. Vaccine. 2019;37(25):3326–3334. doi: 10.1016/j.vaccine.2019.04.074. [ DOI ] [ PubMed ] [ Google Scholar ] Kong B., Kim Y., Kim E. H., Suk J. S., Yang Y.. mRNA: A Promising Platform for Cancer Immunotherapy. Adv. Drug Delivery Rev. 2023;199:114993. doi: 10.1016/j.addr.2023.114993. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Carroll R. G., June C. H.. Programming the Next Generation of Dendritic Cells. Molecular Therapy. 2007;15(5):846–848. doi: 10.1038/sj.mt.6300166. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Sahin U., Oehm P., Derhovanessian E., Jabulowsky R. A., Vormehr M., Gold M., Maurus D., Schwarck-Kokarakis D., Kuhn A. N., Omokoko T., Kranz L. M., Diken M., Kreiter S., Haas H., Attig S., Rae R., Cuk K., Kemmer-Brück A., Breitkreuz A., Tolliver C., Caspar J., Quinkhardt J., Hebich L., Stein M., Hohberger A., Vogler I., Liebig I., Renken S., Sikorski J., Leierer M., Müller V., Mitzel-Rink H., Miederer M., Huber C., Grabbe S., Utikal J., Pinter A., Kaufmann R., Hassel J. C., Loquai C., Türeci Ö.. An RNA Vaccine Drives Immunity in Checkpoint-Inhibitor-Treated Melanoma. Nature. 2020;585(7823):107–112. doi: 10.1038/s41586-020-2537-9. [ DOI ] [ PubMed ] [ Google Scholar ] Borden E. S., Buetow K. H., Wilson M. A., Hastings K. T.. Cancer Neoantigens: Challenges and Future Directions for Prediction, Prioritization, and Validation. Front. Oncol. 2022;12:836821. doi: 10.3389/fonc.2022.836821. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Sahin U., Derhovanessian E., Miller M., Kloke B.-P., Simon P., Löwer M., Bukur V., Tadmor A. D., Luxemburger U., Schrörs B., Omokoko T., Vormehr M., Albrecht C., Paruzynski A., Kuhn A. N., Buck J., Heesch S., Schreeb K. H., Müller F., Ortseifer I., Vogler I., Godehardt E., Attig S., Rae R., Breitkreuz A., Tolliver C., Suchan M., Martic G., Hohberger A., Sorn P., Diekmann J., Ciesla J., Waksmann O., Brück A.-K., Witt M., Zillgen M., Rothermel A., Kasemann B., Langer D., Bolte S., Diken M., Kreiter S., Nemecek R., Gebhardt C., Grabbe S., Höller C., Utikal J., Huber C., Loquai C., Türeci Ö.. Personalized RNA Mutanome Vaccines Mobilize Poly-Specific Therapeutic Immunity against Cancer. Nature. 2017;547(7662):222–226. doi: 10.1038/nature23003. [ DOI ] [ PubMed ] [ Google Scholar ] Wu J., Wu W., Zhou B., Li B.. Chimeric Antigen Receptor Therapy Meets mRNA Technology. Trends Biotechnol. 2024;42(2):228–240. doi: 10.1016/j.tibtech.2023.08.005. [ DOI ] [ PubMed ] [ Google Scholar ] Almåsbak H., Rian E., Hoel H. J., Pulè M., Wälchli S., Kvalheim G., Gaudernack G., Rasmussen A.-M.. Transiently Redirected T Cells for Adoptive Transfer. Cytotherapy. 2011;13(5):629–640. doi: 10.3109/14653249.2010.542461. [ DOI ] [ PubMed ] [ Google Scholar ] Rurik J. G., Tombácz I., Yadegari A., Méndez Fernández P. O., Shewale S. V., Li L., Kimura T., Soliman O. Y., Papp T. E., Tam Y. K., Mui B. L., Albelda S. M., Puré E., June C. H., Aghajanian H., Weissman D., Parhiz H., Epstein J. A.. CAR T Cells Produced in Vivo to Treat Cardiac Injury. Science. 2022;375(6576):91–96. doi: 10.1126/science.abm0594. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Stadler C. R., Bähr-Mahmud H., Celik L., Hebich B., Roth A. S., Roth R. P., Karikó K., Türeci Ö., Sahin U.. Elimination of Large Tumors in Mice by mRNA-Encoded Bispecific Antibodies. Nat. Med. 2017;23(7):815–817. doi: 10.1038/nm.4356. [ DOI ] [ PubMed ] [ Google Scholar ] Hewitt S. L., Bai A., Bailey D., Ichikawa K., Zielinski J., Karp R., Apte A., Arnold K., Zacharek S. J., Iliou M. S., Bhatt K., Garnaas M., Musenge F., Davis A., Khatwani N., Su S. V., MacLean G., Farlow S. J., Burke K., Frederick J. P.. Durable Anticancer Immunity from Intratumoral Administration of IL-23, IL-36γ, and OX40L mRNAs. Sci. Transl. Med. 2019;11(477):eaat9143. doi: 10.1126/scitranslmed.aat9143. [ DOI ] [ PubMed ] [ Google Scholar ] Li H., Yang Y., Hong W., Huang M., Wu M., Zhao X.. Applications of Genome Editing Technology in the Targeted Therapy of Human Diseases: Mechanisms, Advances and Prospects. Sig Transduct Target Ther. 2020;5(1):1. doi: 10.1038/s41392-019-0089-y. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Pacesa M., Pelea O., Jinek M.. Past, Present, and Future of CRISPR Genome Editing Technologies. Cell. 2024;187(5):1076–1100. doi: 10.1016/j.cell.2024.01.042. [ DOI ] [ PubMed ] [ Google Scholar ] Cong L., Ran F. A., Cox D., Lin S., Barretto R., Habib N., Hsu P. D., Wu X., Jiang W., Marraffini L. A., Zhang F.. Multiplex Genome Engineering Using CRISPR/Cas Systems. Science. 2013;339(6121):819–823. doi: 10.1126/science.1231143. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Popovitz J., Sharma R., Hoshyar R., Soo Kim B., Murthy N., Lee K.. Gene Editing Therapeutics Based on mRNA Delivery. Adv. Drug Delivery Rev. 2023;200:115026. doi: 10.1016/j.addr.2023.115026. [ DOI ] [ PubMed ] [ Google Scholar ] Finn J. D., Smith A. R., Patel M. C., Shaw L., Youniss M. R., Van Heteren J., Dirstine T., Ciullo C., Lescarbeau R., Seitzer J., Shah R. R., Shah A., Ling D., Growe J., Pink M., Rohde E., Wood K. M., Salomon W. E., Harrington W. F., Dombrowski C., Strapps W. R., Chang Y., Morrissey D. V.. A Single Administration of CRISPR/Cas9 Lipid Nanoparticles Achieves Robust and Persistent In Vivo Genome Editing. Cell Reports. 2018;22(9):2227–2235. doi: 10.1016/j.celrep.2018.02.014. [ DOI ] [ PubMed ] [ Google Scholar ] Breda L., Papp T. E., Triebwasser M. P., Yadegari A., Fedorky M. T., Tanaka N., Abdulmalik O., Pavani G., Wang Y., Grupp S. A., Chou S. T., Ni H., Mui B. L., Tam Y. K., Weissman D., Rivella S., Parhiz H.. In Vivo Hematopoietic Stem Cell Modification by mRNA Delivery. Science. 2023;381(6656):436–443. doi: 10.1126/science.ade6967. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Ryan D. E., Diamant-Levi T., Steinfeld I., Taussig D., Visal-Shah S., Thakker S., Lunstad B. D., Kaiser R. J., McCaffrey R., Ortiz M., Townsend J., Welch W. R. W., Singh M., Curry B., Dellinger D. J., Bruhn L.. Phosphonoacetate Modifications Enhance the Stability and Editing Yields of Guide RNAs for Cas9 Editors. Biochemistry. 2023;62(24):3512–3520. doi: 10.1021/acs.biochem.1c00768. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen Z., Kelly K., Cheng H., Dong X., Hedger A. K., Li L., Sontheimer E. J., Watts J. K.. In Vivo Prime Editing by Lipid Nanoparticle Co-Delivery of Chemically Modified pegRNA and Prime Editor mRNA. GEN Biotechnology. 2023;2(6):490–502. doi: 10.1089/genbio.2023.0045. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen Q., Zhang Y., Yin H.. Recent Advances in Chemical Modifications of Guide RNA, mRNA and Donor Template for CRISPR-Mediated Genome Editing. Adv. Drug Delivery Rev. 2021;168:246–258. doi: 10.1016/j.addr.2020.10.014. [ DOI ] [ PubMed ] [ Google Scholar ] Karikó K., Muramatsu H., Keller J. M., Weissman D.. Increased Erythropoiesis in Mice Injected With Submicrogram Quantities of Pseudouridine-Containing mRNA Encoding Erythropoietin. Molecular Therapy. 2012;20(5):948–953. doi: 10.1038/mt.2012.7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] An D., Schneller J. L., Frassetto A., Liang S., Zhu X., Park J.-S., Theisen M., Hong S.-J., Zhou J., Rajendran R., Levy B., Howell R., Besin G., Presnyak V., Sabnis S., Murphy-Benenato K. E., Kumarasinghe E. S., Salerno T., Mihai C., Lukacs C. M., Chandler R. J., Guey L. T., Venditti C. P., Martini P. G. V.. Systemic Messenger RNA Therapy as a Treatment for Methylmalonic Acidemia. Cell Reports. 2017;21(12):3548–3558. doi: 10.1016/j.celrep.2017.11.081. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jiang L., Park J.-S., Yin L., Laureano R., Jacquinet E., Yang J., Liang S., Frassetto A., Zhuo J., Yan X., Zhu X., Fortucci S., Hoar K., Mihai C., Tunkey C., Presnyak V., Benenato K. E., Lukacs C. M., Martini P. G. V., Guey L. T.. Dual mRNA Therapy Restores Metabolic Function in Long-Term Studies in Mice with Propionic Acidemia. Nat. Commun. 2020;11(1):5339. doi: 10.1038/s41467-020-19156-3. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Koeberl D., Schulze A., Sondheimer N., Lipshutz G. S., Geberhiwot T., Li L., Saini R., Luo J., Sikirica V., Jin L., Liang M., Leuchars M., Grunewald S.. Interim Analyses of a First-in-Human Phase 1/2 mRNA Trial for Propionic Acidaemia. Nature. 2024;628(8009):872–877. doi: 10.1038/s41586-024-07266-7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Ramaswamy S., Tonnu N., Tachikawa K., Limphong P., Vega J. B., Karmali P. P., Chivukula P., Verma I. M.. Systemic Delivery of Factor IX Messenger RNA for Protein Replacement Therapy. Proc. Natl. Acad. Sci. U.S.A. 2017;114(10):E1941. doi: 10.1073/pnas.1619653114. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Chen C.-Y., Tran D. M., Cavedon A., Cai X., Rajendran R., Lyle M. J., Martini P. G. V., Miao C. H.. Treatment of Hemophilia A Using Factor VIII Messenger RNA Lipid Nanoparticles. Molecular Therapy - Nucleic Acids. 2020;20:534–544. doi: 10.1016/j.omtn.2020.03.015. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Fujita Y., Hirosawa M., Hayashi K., Hatani T., Yoshida Y., Yamamoto T., Saito H.. A Versatile and Robust Cell Purification System with an RNA-Only Circuit Composed of microRNA-Responsive ON and OFF Switches. Sci. Adv. 2022;8(1):eabj1793. doi: 10.1126/sciadv.abj1793. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Jiang K., Koob J., Chen X. D., Krajeski R. N., Zhang Y., Volf V., Zhou W., Sgrizzi S. R., Villiger L., Gootenberg J. S., Chen F., Abudayyeh O. O.. Programmable Eukaryotic Protein Synthesis with RNA Sensors by Harnessing ADAR. Nat. Biotechnol. 2023;41(5):698–707. doi: 10.1038/s41587-022-01534-5. [ DOI ] [ PubMed ] [ Google Scholar ] Qian Y., Li J., Zhao S., Matthews E. A., Adoff M., Zhong W., An X., Yeo M., Park C., Yang X., Wang B.-S., Southwell D. G., Huang Z. J.. Programmable RNA Sensing for Cell Monitoring and Manipulation. Nature. 2022;610(7933):713–721. doi: 10.1038/s41586-022-05280-1. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Kaseniit K. E., Katz N., Kolber N. S., Call C. C., Wengier D. L., Cody W. B., Sattely E. S., Gao X. J.. Modular, Programmable RNA Sensing Using ADAR Editing in Living Cells. Nat. Biotechnol. 2023;41(4):482–487. doi: 10.1038/s41587-022-01493-x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Zhao E. M.. RNA-Responsive Elements for Eukaryotic Translational Control. Nat. Biotechnol. 2022;40:539–545. doi: 10.1038/s41587-021-01068-2. [ DOI ] [ PubMed ] [ Google Scholar ] Cheng Q., Wei T., Farbiak L., Johnson L. T., Dilliard S. A., Siegwart D. J.. Selective Organ Targeting (SORT) Nanoparticles for Tissue-Specific mRNA Delivery and CRISPR-Cas Gene Editing. Nat. Nanotechnol. 2020;15(4):313–320. doi: 10.1038/s41565-020-0669-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Yin D.. Dendritic-Cell-Targeting Virus-like Particles as Potent mRNA Vaccine Carriers. Nature Biomedical Engineering. 2025;9:185–200. doi: 10.1038/s41551-024-01208-4. [ DOI ] [ PubMed ] [ Google Scholar ] Kim B., Hosn R. R., Remba T., Yun D., Li N., Abraham W., Melo M. B., Cortes M., Li B., Zhang Y., Dong Y., Irvine D. J.. Optimization of Storage Conditions for Lipid Nanoparticle-Formulated Self-Replicating RNA Vaccines. J. Controlled Release. 2023;353:241–253. doi: 10.1016/j.jconrel.2022.11.022. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Articles from Chemical Reviews are provided here courtesy of American Chemical Society ACTIONS View on publisher site PDF (19.9 MB) Cite Collections Permalink PERMALINK Copy RESOURCES Similar articles Cited by other articles Links to NCBI Databases Cite Copy Download .nbib .nbib Format: AMA APA MLA NLM Add to Collections Create a new collection Add to an existing collection Name your collection * Choose a collection Unable to load your collection due to an error Please try again Add Cancel Follow NCBI NCBI on X (formerly known as Twitter) NCBI on Facebook NCBI on LinkedIn NCBI on GitHub NCBI RSS feed Connect with NLM NLM on X (formerly known as Twitter) NLM on Facebook NLM on YouTube National Library of Medicine 8600 Rockville Pike Bethesda, MD 20894 Web Policies FOIA HHS Vulnerability Disclosure Help Accessibility Careers NLM NIH HHS USA.gov Back to Top