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Expanded genetic code

The expanded genetic code refers to the extension of the canonical genetic code, which normally encodes 20 standard amino acids using 64 triplets of nucleotides (codons), by incorporating non-canonical or unnatural amino acids (ncAAs) into proteins at specific sites through engineered translational machinery.^[1] This is achieved primarily via orthogonal aminoacyl-tRNA synthetase (aaRS)/tRNA pairs that recognize unique codons, such as the amber stop codon (UAG), without interfering with the host's native translation system, allowing site-specific insertion of ncAAs with novel chemical properties like fluorescence, photocrosslinking, or bioorthogonal handles.^[2] Pioneered in the late 20th century, the concept builds on early demonstrations of tRNA-mediated amino acid reassignment in cell-free systems dating back to 1962, but significant in vivo breakthroughs occurred in the 1980s and 1990s through the work of Peter G. Schultz and colleagues at the Scripps Research Institute.^[3] In 1989, Schultz's group first incorporated ncAAs using amber suppression in vitro,^[4] followed by the development of orthogonal pairs derived from archaeal organisms, such as the tyrosyl-tRNA synthetase (TyrRS) from Methanocaldococcus jannaschii and the pyrrolysyl-tRNA synthetase (PylRS) from Methanosarcina mazei.^[5] By 2001, Wang et al. achieved the first genetic encoding of an ncAA (O-methyl-L-tyrosine) in living Escherichia coli, marking the birth of in vivo genetic code expansion and enabling the incorporation of over 13 distinct ncAAs by 2003.^[1] Subsequent innovations, including quadruplet codon decoding and directed evolution of synthetases via methods like phage-assisted continuous evolution (PACE), have expanded the repertoire to more than 400 ncAAs across diverse organisms, from bacteria to mammalian cells and even mice.^[3] Key methods for expansion include amber suppression, where the UAG codon is reassigned via suppressor tRNAs, and advanced strategies like frame-shift suppression with quadruplet codons or genome recoding to free up additional codons in engineered strains such as Syn61Δ3 E. coli.^[2] Orthogonal systems ensure fidelity, with PylRS variants particularly versatile for charging diverse ncAAs, including backbone-modified ones, as demonstrated in recent evolutions by the Chin and Badran labs in 2024.^[3] In vitro approaches, such as cell-free translation systems (e.g., PURExpress), complement in vivo methods for rapid prototyping and incorporation of unstable ncAAs.^[3] Applications of the expanded genetic code span chemical biology, protein engineering, and therapeutics, enabling precise studies of protein structure, dynamics, and interactions through site-specific labeling or modifications.^[1] For instance, ncAAs like p-acetyl-L-phenylalanine facilitate bioorthogonal chemistry for protein conjugation, while photocaged amino acids allow light-controlled protein activation.^[5] In medicine, expanded codes have produced optimized biologics, including antibody-drug conjugates like ARX788 (in clinical trials NCT04829604), achieving yields up to 5 g/L in mammalian systems.^[2] Synthetic biology leverages these tools for creating proteins with enhanced enzymatic activity or novel functions, such as multiplexed ncAA incorporation (up to four per protein) for complex post-translational mimicry.^[3] Despite progress, challenges persist, including limited codon availability in the universal code, reduced translation efficiency in higher eukaryotes, and the need for orthogonal pairs that evade cellular toxicity or immune responses.^[2] Ongoing research focuses on ribosome engineering, ncAA biosynthesis pathways, and semi-synthetic organisms with unnatural base pairs to further broaden the code's scope.^[3]

Overview

Introduction

The expanded genetic code refers to the engineered modification of the canonical genetic code, which normally encodes 20 standard amino acids, to enable the site-specific incorporation of non-standard amino acids (nsAAs) into proteins during translation. This is achieved through the development of orthogonal translation components, such as engineered tRNA-aminoacyl tRNA synthetase (aaRS) pairs, that recognize unique codons and charge tRNAs with nsAAs without interfering with the host organism's native protein synthesis machinery.^[6] The approach allows for precise control over protein composition, introducing chemical functionalities not found in natural biology, such as photocrosslinkers, fluorescent probes, or bioorthogonal handles.^[6] A pivotal historical milestone occurred in 1989, when Peter G. Schultz and colleagues demonstrated the first site-specific incorporation of an unnatural amino acid into a protein using an amber suppressor tRNA chemically acylated with a non-standard residue, such as p-fluorophenylalanine, in an in vitro E. coli translation system.^[4] This method laid the foundation for subsequent in vivo expansions, including the 2001 achievement of incorporating O-methyl-L-tyrosine in living E. coli via evolved orthogonal aaRS/tRNA pairs. These advances built on earlier suppressor tRNA technologies but shifted toward genetically encoded systems for broader applicability.^[6] The expansion of the genetic code has profound implications for biotechnology, medicine, and synthetic biology, enabling the creation of proteins with enhanced therapeutic properties, such as antibody-drug conjugates for targeted cancer treatment (e.g., ARX788 for HER2-positive breast cancer) and tools for studying protein dynamics in vivo.^[6] By facilitating the integration of over 200 distinct nsAAs in various organisms, including bacteria, yeast, and mammals, this technology supports innovations in drug discovery, biosensors, and materials science while minimizing disruption to endogenous translation.^[6]

Non-Standard Amino Acids

Non-standard amino acids (nsAAs) encompass a diverse class of building blocks beyond the canonical 20 amino acids, broadly classified into naturally occurring rare variants and unnatural, synthetically engineered ones. Naturally occurring nsAAs include selenocysteine (Sec) and pyrrolysine (Pyl), which are incorporated into proteins in specific organisms despite not being part of the standard genetic code. Selenocysteine, the 21st amino acid, features a selenol group in place of the thiol in cysteine and is found in enzymes across prokaryotes, eukaryotes, and archaea, where it participates in redox reactions. Pyrrolysine, the 22nd, contains a pyrroline ring and is utilized in methanogenic archaea and certain bacteria for methylamine metabolism. In contrast, unnatural nsAAs are chemically synthesized to introduce novel functionalities, such as p-acetyl-L-phenylalanine (pAcPhe), which bears a ketone group enabling biorthogonal conjugation reactions for site-specific protein labeling without interfering with native biology.^[7]^[8]^[9] These nsAAs expand the chemical repertoire of proteins by imparting specialized properties that enhance structural, functional, or analytical capabilities. For instance, photocrosslinking is facilitated by nsAAs like p-benzoyl-L-phenylalanine (pBpa), which generates a reactive diradical upon UV irradiation to form covalent bonds with nearby residues or biomolecules, aiding in the mapping of protein interactions. Fluorescence properties can be introduced via nsAAs such as p-azido-L-phenylalanine (pAzF), whose azide moiety allows post-incorporation attachment of fluorophores through click chemistry, enabling real-time visualization of protein dynamics. Metal chelation is another key attribute, exemplified by nsAAs with bipyridine or nitrilotriacetic acid side chains that bind transition metals, supporting applications in catalysis or imaging. These properties allow precise modulation of protein behavior while maintaining compatibility with cellular machinery.^[10]^[11]^[12] Synthesis of nsAAs typically involves chemical routes tailored to their modified structures, with variants of the Strecker synthesis being particularly versatile for producing D-amino acids and other stereoisomers. The classical Strecker reaction condenses an aldehyde, ammonia, and cyanide to form α-amino nitriles, which are hydrolyzed to amino acids; adaptations incorporate chiral catalysts or modified precursors to yield enantiopure unnatural variants, such as fluorinated or azido-substituted D-phenylalanines. Additionally, nsAAs can serve as post-translational modification mimics, replicating effects like phosphorylation (e.g., via O-phosphoserine analogs) or ubiquitination through designed side chains that emulate native PTM chemistry without requiring enzymatic processing. To date, over 200 distinct nsAAs have been successfully incorporated into recombinant proteins, as cataloged in databases like iNClusive (over 466 as of 2023), demonstrating the breadth of this expansion.^[13]^[14]^[15]^[16]^[17]^[18] Representative examples include fluorinated amino acids like (2,3,4,5,6-pentafluorophenyl)glycine, which enhance protein thermal stability by strengthening hydrophobic cores through fluorine's electronegativity and low polarizability. Such nsAAs are briefly incorporated via codon suppression techniques to site-specifically diversify proteomes.^[13]^[14]^[15]^[16]^[17]

Codon Reassignment Methods

Amber Codon Suppression

Amber codon suppression utilizes the amber stop codon (UAG or TAG) to enable site-specific incorporation of non-standard amino acids (nsAAs) into proteins during translation. In this approach, an orthogonal transfer RNA (tRNA) with its anticodon mutated to CUA is engineered to recognize the UAG codon, while an orthogonal aminoacyl-tRNA synthetase (aaRS) specifically charges this tRNA with the desired nsAA. This suppresses translation termination by competing with the release factor 1 (RF1), allowing the ribosome to insert the nsAA at the UAG site and continue polypeptide elongation. The orthogonality of the tRNA/aaRS pair ensures minimal cross-reactivity with endogenous components, preserving the fidelity of the standard genetic code elsewhere.^[19] The efficiency of amber suppression is influenced by several factors, primarily the competition between the suppressor tRNA and RF1 at UAG codons, which often results in truncated proteins or reduced yields. Suppression efficiency can be enhanced by overexpressing the orthogonal tRNA or optimizing the aaRS for better charging, but a significant improvement comes from using RF1-deficient Escherichia coli strains, such as the genomically recoded Syn61ΔprfA, where amber suppression yields increase by up to 250% for multi-site incorporations. In these strains, the absence of RF1 eliminates termination at UAG, though RF2 still handles other stop codons, maintaining cell viability.^[20] Pioneering work in amber suppression began with the 1989 demonstration of site-specific incorporation of p-fluorophenylalanine into a protein using chemically aminoacylated suppressor tRNA in an in vitro translation system, marking the first use of this method for unnatural amino acid insertion. This in vitro proof-of-concept evolved into in vivo applications in the 1990s and 2000s, with the development of orthogonal pairs derived from archaeal or eukaryotic sources expressed in bacterial hosts. By the 2010s, amber suppression had enabled the incorporation of over 100 distinct nsAAs, including photocrosslinkers, fluorescent probes, and post-translationally modified residues, into recombinant proteins in various organisms.^[19] Despite its versatility, amber suppression faces limitations due to off-target suppression at the approximately 300 natural UAG stop codons in the E. coli genome, which can disrupt essential protein termination and reduce cellular fitness. This issue is particularly pronounced in RF1-deficient strains, where unintended read-through may occur more frequently. To mitigate such effects and expand coding capacity, strategies like quadruplet codon decoding have been developed as alternatives or complements, allowing nsAA incorporation without relying on stop codon reassignment.^[21]

Rare Sense Codon Reassignment

Rare sense codon reassignment involves repurposing low-frequency sense codons, such as AGG or AGA for arginine in Escherichia coli, to encode non-standard amino acids (nsAAs) in organisms engineered to minimize interference with native translation. The strategy entails depleting the natural tRNAs that decode these rare codons through genome recoding—replacing all instances of the target codon with synonymous alternatives—and subsequently introducing an orthogonal tRNA/synthetase pair specific for the desired nsAA. This approach leverages the low usage of rare codons (e.g., AGG occurs only about 1,400 times in the E. coli genome, primarily in non-essential genes) to avoid disrupting proteome function while expanding the code.^[22]^[23] A seminal example is the reassignment of the AGG codon to incorporate the non-canonical amino acid L-homoarginine in E. coli. Researchers engineered a variant of the pyrrolysyl-tRNA synthetase (HarRS) to charge tRNA^Pyl_CCU with L-homoarginine, while eliminating AGG from 38 sites in 32 essential genes via synonymous substitutions. Mass spectrometry confirmed incorporation of L-homoarginine at AGG sites in model proteins, achieving detectable levels in the proteome (approximately 0.4% relative abundance) under optimized conditions, demonstrating efficient suppression of the depleted codon. Similar strategies have been applied to other nsAAs, though efficiencies vary based on synthetase affinity and cellular context.^[22] This method offers key advantages over amber (UAG) suppression, as it avoids competition with release factor 1 (RF1), which can limit incorporation efficiency at stop codons to 20-50% in standard strains. By targeting sense codons in recoded backgrounds, rare sense reassignment enables the simultaneous incorporation of multiple distinct nsAAs into a single protein using different reassigned codons, facilitating complex protein engineering without the toxicity or yield penalties associated with stop codon readthrough. Orthogonal tRNA/synthetase pairs, such as those derived from archaea, ensure specificity and minimal cross-reactivity with canonical translation.^[22]^[23] Despite these benefits, challenges persist, primarily the need for extensive genome-wide recoding to fully eliminate natural codon usage and prevent mistranslation or growth defects. For instance, the 2016 design and partial assembly of a recoded E. coli strain involved synonymous replacement of all 62,214 instances of seven codons (including rare sense codons for arginine, leucine, and serine, plus UAG) across the 4.3 Mb genome, reducing the code to 57 codons and enabling safer reassignment, though full assembly required iterative synthesis and validation of essential genes. This effort culminated in 2025 with the realization of a fully viable 57-codon E. coli strain, further enhancing the scalability of rare sense codon reassignment. Partial recoding, as in early AGG studies, limits scalability due to residual endogenous tRNA activity, and achieving high-fidelity incorporation (>90%) often demands further optimization of orthogonal components.^[24]^[22]^[25]

Quadruplet Codons

Quadruplet codons represent an approach to expand the genetic code by introducing four-base codon sequences that trigger a +1 ribosomal frameshift, enabling the decoding of non-standard amino acids (nsAAs) without repurposing the standard 64 triplet codons. Engineered transfer RNAs (tRNAs) feature extended anticodon loops, typically seven or eight nucleotides long, which pair with the quadruplet codon in the ribosomal A-site, facilitating a four-nucleotide translocation during translation. This mechanism operates through models such as the "yardstick" model, where the tRNA anticodon fully base-pairs with the four-base codon, or the "slippery" model involving partial slippage and realignment, with recent refinements suggesting a four-base interaction in the P-site post-translocation.^[26]^[27] The development of quadruplet codon decoding began in the 1990s with in vitro demonstrations using synthetic mRNAs and mutant tRNAs to incorporate amino acids at four-base sites like AGGU or CGGG. In vivo applications emerged in the early 2000s, with seminal work by Moore and colleagues engineering a suppressor tRNA to decode an expanded amber codon (UAGA) in Escherichia coli, achieving functional protein expression. Further advancements in the 2010s involved orthogonal tRNA-synthetase pairs, such as the pyrrolysyl-tRNA synthetase (PylRS)/tRNA^Pyl^ system, optimized for quadruplet anticodons like CUA G, which improved charging specificity for nsAAs. Efficiency, initially below 5% relative to triplet decoding, reached 20-50% by the mid-2010s through tRNA body modifications and anticodon stem-loop optimizations that reduced ribosomal slippage.^[28]^[29] This strategy theoretically expands the codon repertoire to 256 possibilities (4^4), allowing simultaneous incorporation of multiple nsAAs at distinct sites without interference from the natural code, as demonstrated in proteins for fluorescence resonance energy transfer (FRET) studies and nuclear magnetic resonance (NMR) spectroscopy. Applications include multiplexed protein labeling for structural biology and the creation of logic-gated circuits in synthetic biology, where quadruplet codons control protein function in response to environmental cues. In therapeutic contexts, quadruplet decoding has enabled the production of vaccine antigens with nsAAs mimicking pathogen modifications, enhancing immunogenicity.^[26]^[30]^[31] Despite these advances, quadruplet decoding suffers from lower fidelity and efficiency compared to triplet suppression, primarily due to ribosomal slippage causing frameshift errors or competition with endogenous triplet tRNAs, often yielding only 1-25% of wild-type protein levels for multi-site incorporations. Cross-decoding of embedded triplets can further reduce orthogonality, limiting the number of usable quadruplet codons to around 20-30. Recent progress in eukaryotic systems, such as Caenorhabditis elegans, has achieved stable quadruplet frameshifting with 30-50% efficiency using hybrid PylRS/tRNA variants, enabling applications like photocaged nsAA incorporation for optogenetic control of neuronal proteins. These eukaryotic adaptations address prior prokaryotic biases and pave the way for broader in vivo expansion.^[32]^[33]^[34]

Orthogonal Translation Components

Engineered tRNA/Synthetase Pairs

The principle of orthogonality in engineered tRNA/aminoacyl-tRNA synthetase (aaRS) pairs relies on importing components from archaea or eukaryotes into bacterial hosts, where they do not interact with the endogenous translation machinery. A seminal example is the tyrosyl-tRNA synthetase (TyrRS) and its cognate tRNA^Tyr from Methanocaldococcus jannaschii, which exhibit minimal cross-reactivity with Escherichia coli aaRSs and tRNAs due to differences in identity elements such as the anticodon and acceptor stem. This pair, first demonstrated in 2001, allows selective charging of non-canonical amino acids (ncAAs) without disrupting canonical protein synthesis.^[35] Engineering these pairs for ncAA specificity typically involves directed evolution using positive and negative selection schemes in vivo. Libraries of aaRS mutants are generated through error-prone PCR or DNA shuffling, then screened for activation of the target ncAA (positive selection, often via suppression of an amber codon in a reporter gene like chloramphenicol acetyltransferase) while eliminating charging of the 20 standard amino acids (negative selection, using a toxic reporter like barnase). A key illustration is the evolution of an M. jannaschii TyrRS variant specific for p-acetyl-L-phenylalanine (pAcPhe), achieved through iterative selection yielding mutants with altered active-site residues that accommodate the ketone group. This approach has been widely adopted, enabling high-fidelity charging where optimized pairs achieve >90% efficiency for the target ncAA and <1% misacylation of canonical amino acids.^[36] Prominent examples include variants derived from M. jannaschii TyrRS, which have been tailored for over 50 aromatic and phenylalanine derivatives. Another major class stems from the pyrrolysyl-tRNA synthetase (PylRS) and tRNA^Pyl pair from Methanosarcina species, naturally orthogonal in E. coli and engineered for pyrrolysine analogs like N^ε-acetyl-L-lysine or cyclopropynylated lysine through mutations in the aaRS binding pocket. Recent evolutions of PylRS by the Chin and Badran groups in 2024 have further expanded its versatility to charge diverse ncAAs, including backbone-modified ones.^[3] By the 2020s, dozens of such orthogonal pairs had been developed across various aaRS scaffolds, including leucyl- and tryptophanyl-tRNA synthetases, facilitating diverse ncAA incorporations with robust orthogonality and minimal off-target effects.^[37]^[38]^[39]

Orthogonal Sets in Model Organisms

Orthogonal tRNA/aminoacyl-tRNA synthetase (aaRS) pairs, originally derived from archaea or bacteria, have been successfully implemented in bacterial model organisms such as Escherichia coli to enable the incorporation of non-canonical amino acids (ncAAs). A prominent example is the pyrrolysyl-tRNA synthetase (PylRS) and its cognate tRNA^Pyl_CUA from Methanosarcina mazei, which was imported into E. coli to genetically encode pyrrolysine and later engineered variants for diverse ncAAs like N^ε-acetyl-L-lysine.^[40] This system demonstrates high orthogonality in bacteria, minimizing cross-reactivity with endogenous translation machinery, and has been used to produce proteins with up to 70% suppression efficiency at amber codons in optimized strains.^[41] However, overexpression of these exogenous components can lead to cellular toxicity due to interference with native aaRS activities or resource competition, which is often mitigated through inducible expression systems, such as arabinose-inducible promoters, to control levels during protein synthesis.^[42] In eukaryotic model organisms, adapting orthogonal sets presents unique challenges, including the nuclear export of engineered tRNAs and competition from host synthetases, which can reduce charging efficiency and lead to misacylation. In Saccharomyces cerevisiae, early efforts in the 2000s faced issues with tRNA nuclear retention and poor cytoplasmic functionality, but by 2010, optimized PylRS/tRNA^Pyl_CUA pairs achieved functional expression through modifications enhancing tRNA export signals and synthetase stability, enabling ncAA incorporation at amber sites with moderate yields.^[43] Similarly, in mammalian cells like HEK293, initial implementations in the early 2000s using archaeal or bacterial orthogonal pairs encountered low orthogonality due to host editing mechanisms and tRNA instability, but engineered variants improved fidelity by the mid-2000s, allowing site-specific ncAA insertion in proteins expressed via transient transfection.^[44] Comparisons across model organisms highlight differences in implementation efficiency, often attributed to variations in translation machinery and codon usage biases. In E. coli, orthogonal systems typically achieve 60-80% amber suppression efficiency, benefiting from streamlined bacterial ribosomes and minimal tRNA competition, whereas in HEK293 cells, efficiencies range from 20-50% due to eukaryotic regulatory layers and higher background suppression. Codon bias adjustments, such as optimizing tRNA anticodon loops to match host preferences or using codon-optimized genes, further enhance performance; for instance, in yeast, tailoring tRNA sequences to S. cerevisiae codon usage increased ncAA incorporation by 2-3 fold compared to unmodified prokaryotic tRNAs.^[45] As of 2024, advances have focused on stable genomic integration of orthogonal sets to enable long-term, heritable expression without reliance on plasmids, reducing toxicity and improving consistency. In E. coli, chromosomally integrated PylRS/tRNA^Pyl_CUA systems created auxotrophic strains with altered genetic codes, achieving sustained ncAA-dependent growth and up to 90% incorporation fidelity.^[46] In mammalian models, transgenic integration of engineered aaRS/tRNA pairs into mouse genomes has supported efficient ncAA labeling in vivo, paving the way for applications in multicellular organisms.^[47]

Specialized Ribosome Systems

o-Ribosome

The o-ribosome, or orthogonal ribosome, represents a engineered ribosomal system designed to selectively translate orthogonal messenger RNAs (o-mRNAs) bearing altered Shine-Dalgarno (SD) sequences, thereby dedicating translation resources to the synthesis of proteins incorporating non-standard amino acids (nsAAs) without interfering with the host cell's proteome.^[48] This system achieves specificity through mutations in the anti-Shine-Dalgarno (ASD) sequence of the 16S ribosomal RNA (rRNA), which is the complement of the canonical SD sequence (AGGAGG) and located at positions 1535–1541 in Escherichia coli 16S rRNA.^[48] By altering the ASD (originally CCUCCU), the o-ribosome preferentially binds and initiates translation on o-mRNAs with complementary mutated SD sequences, such as those changed at one or more positions from the wild-type GGAGG, enabling up to 100-fold greater selectivity for o-mRNAs compared to natural mRNAs.^[48]^[49] The foundational design was developed in 2005 by Rackham and Chin using a dual positive-negative selection strategy in E. coli, where libraries of mutated 16S rRNA variants (targeting all seven nucleotides in the ASD region) were screened against corresponding o-mRNA libraries to identify orthogonal pairs that confer chloramphenicol resistance (positive selection via CAT reporter) while avoiding 5-fluorouracil toxicity (negative selection via UPRT).^[48] This approach generated a network of at least four orthogonal ribosome-o-mRNA pairs that operate independently of the wild-type translation machinery, demonstrating the system's capacity for parallel, non-competing translation pathways.^[48] In 2007, the Chin laboratory further evolved the o-ribosome (termed ribo-Xm in the study) specifically for enhanced genetic code expansion, introducing the key mutation A1191G in the 16S rRNA to improve amber (TAG) codon suppression efficiency when paired with orthogonal amber suppressor tRNAs.^[49] This evolution was achieved through directed selection for improved decoding of o-mRNAs containing amber codons, resulting in a ribosome that boosts nsAA incorporation yields: from approximately 20% to over 60% full-length protein for constructs with a single amber codon, and from less than 1% to over 20% for those with two amber codons.^[49] The A1191G mutation is thought to reduce interactions with release factor 1 at the amber codon, thereby favoring suppressor tRNA accommodation and enabling the production of proteins with multiple nsAAs.^[49] Subsequent variants of the o-ribosome have incorporated additional 16S rRNA mutations to refine initiation efficiency and orthogonality, such as combined ASD alterations with helix 44 modifications, allowing for more robust translation of recoded genes in model organisms like E. coli. These improvements maintain the core selectivity of the original design while supporting applications in multi-site nsAA incorporation for proteins up to hundreds of residues long.^[49]

Ribo-X

The Ribo-X system represents an advanced orthogonal ribosome engineered to boost the efficiency of non-standard amino acid (nsAA) incorporation in the expanded genetic code within Escherichia coli. Introduced in 2007 by the Chin laboratory (Wang et al.), it evolves from the foundational orthogonal ribosome by applying directed evolution to select variants that preferentially translate orthogonal mRNAs containing amber (TAG) stop codons, enabling precise nsAA insertion without competing with the host translation machinery. This development addresses limitations in earlier systems, where low suppression efficiency led to truncated proteins and reduced yields.^[50] At its core, Ribo-X operates through mutations acquired during selection that diminish the ribosome's affinity for release factor 1 (RF1), the endogenous factor that terminates translation at amber codons. This alteration allows the orthogonal amber suppressor tRNA, charged with an nsAA by a paired orthogonal aminoacyl-tRNA synthetase, to outcompete RF1 more effectively during decoding. Consequently, Ribo-X selectively engages orthogonal mRNAs—those with altered Shine-Dalgarno sequences that evade native ribosomes—while minimizing off-target translation. In vitro assays confirm Ribo-X's reduced RF1 binding, supporting its role in favoring productive suppression over termination.^[50]^[51] Ribo-X offers substantial advantages in performance, elevating nsAA incorporation yields to over 60% for proteins with a single amber codon and over 20% for those with two, compared to approximately 20% and less than 1% with unmodified orthogonal ribosomes. These gains stem from lower background activity on native mRNAs and higher fidelity in suppression, achieving overall efficiencies exceeding 80% in optimized E. coli strains when paired with refined orthogonal components. The system's orthogonality also mitigates toxicity from leaky expression, as it confines activity to designated mRNAs.^[50] In applications, Ribo-X facilitates tunable expression of nsAA-containing proteins, particularly those prone to toxicity when truncated or harboring reactive groups like azides or alkynes. By integrating with inducible promoters for rRNA expression, it enables temporal control of translation in vivo, allowing researchers to synchronize nsAA incorporation with cellular growth phases. This has proven valuable for producing modified superfolder GFP or calmodulin variants with photocrosslinkers, advancing studies in protein dynamics, therapeutic design, and synthetic biology.^[50]

Ribo-Q

The Ribo-Q ribosome is an engineered orthogonal ribosome variant designed to facilitate the incorporation of non-standard amino acids (nsAAs) by efficiently decoding quadruplet codons. Developed through directed evolution, it features specific mutations in the 16S rRNA that alter the decoding center to favor binding of quadruplet-decoding tRNAs, thereby reducing frameshifting errors and enhancing translation fidelity for expanded genetic codes. Introduced in 2010, this system builds on prior orthogonal ribosome designs by targeting residues in the A-site to improve selectivity for modified tRNAs.^[52] Key mutations in the 16S rRNA of Ribo-Q include A1196G, often combined with alterations at nearby positions such as C1195A or C1195T and A1197G, depending on the specific variant (e.g., Ribo-Q1 with A1196G and A1197G). These changes, identified via saturation mutagenesis of libraries derived from earlier ribosome variants, preferentially accommodate extended anticodons of quadruplet tRNAs while maintaining compatibility with standard triplet decoding when needed. The design ensures orthogonality, allowing Ribo-Q to translate orthogonal mRNAs containing quadruplet codons without significant interference from host ribosomes.^[52]^[53] In terms of functionality, Ribo-Q enhances frameshift suppression during translation of quadruplet codons, such as CGGG or CUA G, by stabilizing the interaction between the ribosome's A-site and cognate quadruplet tRNAs. This selectivity arises from the reconfigured A-site geometry, which discriminates against standard triplet tRNAs and promotes accurate nsAA insertion at defined sites. When paired with orthogonal tRNA/synthetase systems, Ribo-Q enables the site-specific incorporation of multiple distinct nsAAs in a single protein, supporting applications in protein engineering.^[52] Performance evaluations demonstrate that Ribo-Q achieves incorporation yields of 50-70% for nsAAs at quadruplet codons, a substantial improvement over unmodified orthogonal ribosomes, which often suffer from lower efficiency due to frameshifting. It is fully compatible with orthogonal synthetases, such as pyrrolysyl-tRNA synthetase variants, allowing seamless integration into existing genetic code expansion workflows. These yields were measured in E. coli expression systems using reporter proteins like superfolder green fluorescent protein.^[52] Ribo-Q evolved from the o-Ribosome, an earlier orthogonal system, through iterative selection to specifically support quadruplet codon decoding and broader expansion of the genetic code. This derivation involved introducing the A-site mutations into o-Ribosome scaffolds to optimize for four-base codon recognition, resulting in a more versatile platform for synthetic biology.^[52]

Ribosome Stapling

Ribosome stapling is a specialized engineering approach to generate orthogonal ribosomes by covalently linking their small (30S) and large (50S) subunits via an RNA staple, enabling selective and efficient translation of orthogonal mRNAs for expanded genetic code applications in Escherichia coli. Introduced in 2015, this method addresses limitations in subunit association for orthogonal systems by fusing a circularly permuted 23S rRNA into the 16S rRNA at helix 44 and helix 101, connected by a flexible RNA hinge derived from the J5/J5a region of the Tetrahymena group I intron. The resulting ~4500-nucleotide fused rRNA assembles into a functional ribosome that preferentially recognizes orthogonal Shine-Dalgarno (oSD) sequences on target mRNAs, isolating translation from endogenous ribosomes and facilitating the incorporation of non-standard amino acids (nsAAs) through amber suppression.^[54] To enhance control over assembly and purity, later implementations incorporate an MS2 stem-loop tag into the stapled orthogonal rRNA (o-rRNA), allowing affinity purification via binding to the MS2 coat protein. This tethering step increases the local concentration of stapled ribosomes relative to native ones, minimizing cross-assembly and enabling high-fidelity orthogonal translation. The purified stapled ribosomes are then used to translate o-mRNAs containing amber codons, where orthogonal tRNA/synthetase pairs deliver nsAAs, such as p-acetylphenylalanine, with reduced competition from release factor 1 (RF1). This setup supports localized translation by confining activity to engineered mRNAs, improving overall system orthogonality.^[55] The primary benefit of ribosome stapling lies in its ability to boost nsAA incorporation efficiency by enforcing subunit pairing and insulating the ribosome from cellular competitors, achieving translation activities comparable to non-stapled orthogonal ribosomes. For instance, the optimized O-d2d8 variant supports amber suppression and cellular growth at 30-40% of wild-type rates initially, with directed evolution yielding variants that reach 81% growth efficiency while incorporating nsAAs at sites previously limited by RF1 competition. This enhancement is particularly valuable for challenging sequences, such as polyproline stretches, where stapled ribosomes enable elongation without auxiliary factors like EF-P, demonstrating up to 10-fold improvements in polymerization of non-canonical building blocks.^[55] Variants of the technique vary in RNA staple linker length to fine-tune subunit geometry and activity, with shorter staples like O-d0d0 promoting tight association but lower flexibility, and longer ones like O-d2d8 balancing assembly efficiency and translational fidelity. Evolved iterations, such as O-d2d8(5), further expand capabilities by incorporating β-amino acids or other non-proteinogenic monomers via amber suppression, supporting the synthesis of novel biopolymers. While photocaged variants for spatiotemporal control have been explored in related tethering systems, direct application to stapled ribosomes remains under development to enable light-activated translation initiation.^[55] Despite these advantages, ribosome stapling faces limitations, including reduced initial rRNA expression (~25% of wild-type levels) due to folding challenges in the extended transcript and potential steric hindrance during subunit docking. In crowded cellular environments, the bulky fused structure may exacerbate assembly inefficiencies or interfere with polysome formation, necessitating strain optimizations like RF1 depletion to maximize nsAA yields. These constraints highlight the need for further engineering to approach full wild-type performance in vivo.^[54]

Engineered Peptidyl Transferase Center

The peptidyl transferase center (PTC) of the bacterial ribosome, formed primarily by the 23S rRNA in the large subunit, catalyzes the formation of peptide bonds between amino acids during protein synthesis. Engineering the PTC through targeted mutations in this rRNA region enables the incorporation of non-standard amino acids (nsAAs), such as D-amino acids, by alleviating steric clashes and altering the geometry of the active site to accommodate atypical substrates. These modifications expand the genetic code beyond the standard L-amino acids, facilitating the synthesis of proteins with novel properties, though they often require orthogonal ribosome systems to minimize interference with native translation. Initial efforts to engineer the PTC emerged in the early 2000s, focusing on mutations to enhance compatibility with D-amino acids. Researchers overexpressed modified 23S rRNA from multicopy plasmids in Escherichia coli, introducing alterations in the PTC loop (nucleotides 2447–2450) and helix 89 (nucleotides 2457–2462), such as substitutions converting 2447GAUA2450 to 2447UUAC2450 or 2447GAUAA2450 paired with changes in helix 89 like U2460G. These mutant ribosomes incorporated D-methionine and D-phenylalanine into model proteins like dihydrofolate reductase with efficiencies of 11–23% relative to L-amino acid counterparts, while retaining up to 45% suppression activity compared to wild-type levels. The mechanism involves local conformational changes in the PTC that reduce discrimination against the reversed chirality of D-aminoacyl-tRNAs, allowing peptide bond formation without major disruption to the catalytic core. Subsequent refinements, such as additional mutations in the same regions, improved D-amino acid tolerance in cell-free systems, yielding functional proteins with up to 50% activity retention.^[56] Further engineering has targeted PTC variants for other ncAAs, including β-amino acids, using selection methods like β-puromycin. For example, selected ribosomes enable incorporation of select β-amino acids, such as methyl-β-alanine isomers, at relative efficiencies up to ~20% in reporter proteins. Ongoing computational design and selections continue to refine PTC mutations (e.g., in helices H73 and H75) to improve general performance and support diverse nsAAs in orthogonal systems, though efficiencies for sterically hindered residues like α-aminoisobutyric acid (Aib) remain low (typically 3–15%). These designs support non-ribosomal chemistries, like altered amide bond angles, but carry risks of global translation impairment, including reduced cell fitness and off-target effects, necessitating selective expression of mutant rRNAs to limit dominant-negative impacts on endogenous ribosomes.^[57]^[58] These engineered PTCs continue to support ncAA incorporation in contemporary genetic code expansion workflows, as reviewed in 2024.^[47]

Applications

Biomedical and Therapeutic Uses

The expanded genetic code enables the site-specific incorporation of non-standard amino acids (nsAAs) into therapeutic proteins, enhancing their pharmacological properties for biomedical applications. By introducing chemically reactive nsAAs, such as p-acetylphenylalanine, researchers can perform precise modifications like PEGylation, which extends protein half-life and reduces immunogenicity compared to random conjugation methods.^[59] For instance, site-specific PEGylation of cytokines has been used to improve bioavailability; similarly, PEGylated interleukin-2 (IL-2) engineered via genetic code expansion selectively activates regulatory T cells, showing promise in autoimmune disease models by sustaining immunosuppression without broad immune activation.^[60]^[61] Photocaging groups incorporated as nsAAs allow spatiotemporal control of protein function, facilitating light-activated therapeutics. These photocaged nsAAs, such as nitrobenzyl-lysine, block activity until ultraviolet or visible light uncages them, enabling on-demand drug release or enzyme activation in targeted tissues.^[62] In therapeutic contexts, photocaged nanobodies have been generated using genetic code expansion to inhibit proteins in vivo upon illumination, offering potential for precise modulation in cancer or neurological treatments. For example, incorporation of photocaged glutamic acid into key residues permits optical control of protein signaling pathways, advancing light-responsive biologics.^[63] Clinical applications of expanded code-modified proteins include antibody-drug conjugates (ADCs) and fluorinated therapeutics. In the 2010s, ADCs with site-specifically incorporated nsAAs, such as p-acetylphenylalanine for payload attachment, entered clinical trials, improving homogeneity and efficacy over traditional cysteine-based conjugates; ARX788, an anti-HER2 ADC from Ambrx, has progressed to Phase III trials (as of 2025) for HER2-positive breast cancer, demonstrating improved progression-free survival in pivotal studies.^[64]^[65]^[66] Fluorinated nsAAs, like 3-fluoro-tyrosine, enhance protein stability and metabolic resistance; recent developments in 2023 involved their use in engineered therapeutics to improve pharmacokinetics, though no direct FDA approvals for expanded code-derived fluorinated proteins were reported that year, building on prior preclinical successes.^[47]^[67] As of 2025, no therapeutics produced via expanded genetic code have been FDA-approved, though several are in late-stage clinical trials. In gene therapy, the expanded code facilitates site-specific nsAA incorporation into viral vector capsids, enhancing targeting and reducing off-target effects. Adeno-associated virus (AAV) vectors modified with azide-bearing nsAAs in capsid proteins allow click chemistry conjugation of targeting ligands, improving tropism for specific cell types like neurons or hepatocytes.^[68] This approach has been applied to AAV2 variants, where nsAA insertion during production enables precise functionalization without disrupting assembly, advancing therapies for genetic disorders.^[69]^[70] A key challenge in translating nsAA-containing therapeutics to humans is potential immunogenicity, as noncanonical structures may elicit anti-drug antibodies. Studies indicate that certain nsAAs, particularly those with bulky or charged side chains, can alter T-cell epitope presentation, potentially triggering immune responses, though conservative nsAAs mimicking natural ones show minimal reactivity.^[71]^[72] Strategies like selecting low-immunogenic nsAAs and preclinical screening in humanized models are essential to mitigate this risk.^[73]

Industrial and Synthetic Biology Uses

In industrial and synthetic biology, the expanded genetic code enables the incorporation of non-standard amino acids (nsAAs) into enzymes to enhance their stability and performance as biocatalysts. For instance, nsAAs such as fluorinated or cyclopropyl derivatives can rigidify protein structures, improving thermostability for processes requiring high temperatures, like biofuel production or chemical synthesis. A notable example involves engineering a stereoselective cyclopropanation biocatalyst with genetically encoded staples formed via nsAAs, which increased its melting temperature by up to 30°C while maintaining catalytic efficiency, allowing sustained activity under industrial conditions.^[74] Similarly, keratinase variants incorporating ncAAs like p-bromophenylalanine (pBpF) exhibited an 8.2-fold longer half-life at 60°C, facilitating robust degradation of keratin waste in leather processing.^[75] The use of nsAAs has also advanced the design of protein-based materials through self-assembly and click chemistry. Proteins engineered with azide- or alkyne-bearing nsAAs, such as p-azidophenylalanine, enable site-specific crosslinking via copper-free click reactions to form stable hydrogels with tunable mechanical properties for applications in tissue engineering scaffolds.^[76] In the 2020s, these approaches have contributed to bioink development, where self-assembling peptides modified with nsAAs improve printability and biocompatibility in 3D bioprinting, yielding constructs with shear-thinning behavior suitable for extrusion-based fabrication of complex structures.^[77] Fluorescent nsAAs integrated via genetic code expansion have transformed biosensors for real-time industrial monitoring. For example, fluorogenic nsAAs inserted into protein scaffolds enable sensitive detection of metabolites in fermentation broths.^[78] This allows precise control of bioprocesses, optimizing yields in microbial production of chemicals or biofuels. Scale-up efforts leverage engineered Escherichia coli strains to achieve high-density fermentation for nsAA-containing proteins, with reported yields reaching hundreds of mg/L in optimized cultures. These advancements support industrial viability.^[79]

Future Directions

Recoded Synthetic Genomes

Recoded synthetic genomes represent a transformative approach to expanding the genetic code by systematically redesigning the DNA sequence of an entire organism through synonymous codon compression. This method involves replacing multiple synonymous codons with a reduced set, thereby eliminating or reassigning specific codons across the genome to create "blank" codons available for encoding non-standard amino acids (nsAAs) without interfering with native protein synthesis. Unlike targeted suppression techniques, this global recoding ensures that freed codons are absent from essential genes, enabling orthogonal tRNA/aminoacyl-tRNA synthetase pairs to incorporate nsAAs proteome-wide with minimal off-target effects. A seminal milestone in bacterial recoding was the development of the genomically recoded Escherichia coli strain C321.ΔA, in which all 321 instances of the TAG (amber) stop codon were replaced with the synonymous TAA stop codon, and the prfA gene encoding release factor 1 (RF1) was deleted. This freed the TAG codon for reassignment to nsAAs, allowing efficient amber suppression without RF1-mediated termination and demonstrating enhanced viral resistance due to disrupted horizontal gene transfer. Building on this, advanced compression efforts have further reduced the codon repertoire; for instance, a 2025 study engineered an E. coli variant with a 57-codon genetic code by compressing four serine codons, two alanine codons, and the amber stop into three serine, two alanine, and one stop codon equivalents, enabling the reallocation of seven codons for potential nsAA encoding while maintaining viability through adaptive evolution.^[80] In eukaryotic systems, the 2020s Synthetic Yeast Genome Project (Sc2.0 and proposed Sc3.0) has advanced recoding by synonymously refactoring open reading frames to minimize codon diversity, with designs freeing multiple codons through systematic replacement of synonymous sets, paving the way for multi-nsAA incorporation in complex pathways.^[81] These efforts highlight the potential to reassign over 50 codons for nsAAs in highly compressed genomes, vastly expanding proteomic diversity. The primary benefits of recoded genomes include achieving global orthogonality, where nsAAs can be encoded without suppression artifacts such as ribosomal stalling or premature termination in native transcripts, and improved biosafety through genetic isolation from natural organisms. This approach avoids the toxicity and inefficiency of partial suppression systems by ensuring freed codons are never encountered in the host genome, facilitating high-fidelity multi-site nsAA incorporation for applications like novel therapeutics. However, significant technical hurdles persist, including reduced cell fitness from altered codon bias, which disrupts translation speed, mRNA stability, and proteome balance, often necessitating extensive adaptive laboratory evolution to restore growth rates comparable to wild-type strains. Ethical concerns also arise regarding the environmental release of such organisms, given their potential to outcompete natural variants or evade standard antibiotics, alongside the immense engineering challenges of synthesizing and assembling multi-megabase recoded chromosomes without introducing deleterious mutations.

Expanded Genetic Alphabet

The expanded genetic alphabet refers to the incorporation of synthetic nucleobases into DNA and RNA, forming unnatural base pairs (UBPs) that function orthogonally to the natural A-T and G-C pairs, thereby increasing the informational density of the genetic code. This approach contrasts with codon reassignment by directly augmenting the nucleotide repertoire, allowing for a larger number of unique codons. Pioneering work by Floyd Romesberg's laboratory at The Scripps Research Institute developed the d5SICS-dNaM UBP, where d5SICS (6-methyl-8-(6'-N-benzamidomethyl-isoxazol-3'-yl)-2'-deoxyadenosine) and dNaM (2-fluoro-6-(2''-N-benzamidomethyl-pyridin-5'-yl)-2'-deoxyguanosine) rely on hydrophobic and shape-complementary interactions for selective pairing, rather than hydrogen bonding. This pair was first demonstrated to replicate efficiently in vitro using evolved polymerases, such as those derived from Taq DNA polymerase, achieving replication fidelities comparable to natural bases. In 2014, Romesberg and colleagues achieved a landmark by engineering an Escherichia coli strain harboring a plasmid with the d5SICS-dNaM UBP, marking the first semi-synthetic organism (SSO) with a stably expanded six-letter genetic alphabet in vivo, where the unnatural nucleotides were retained through multiple rounds of cell division with negligible loss.^[82] Integration of UBPs into functional genetic systems involves their transcription into RNA containing unnatural bases, which can then form codons recognized by orthogonal tRNAs charged with non-standard amino acids (nsAAs). In a 2017 advancement, the same group engineered an SSO where an unnatural codon derived from the d5SICS-dNaM pair was transcribed and decoded by an orthogonal tRNA-synthetase pair, enabling site-specific incorporation of nsAAs like α-hydroxyl-naphthalene-1-acetic acid into green fluorescent protein expressed in E. coli. This process required orthogonal tRNAs modified to carry the unnatural anticodon and paired with an engineered aminoacyl-tRNA synthetase specific for the nsAA, ensuring minimal cross-reactivity with the natural code. Such systems expand the proteome's chemical diversity, allowing proteins with novel properties, such as enhanced fluorescence or chemical reactivity. Progress in the 2020s has focused on scaling to an eight-letter alphabet, exemplified by the hachimoji system developed by Romesberg and colleagues, which incorporates two additional UBPs (P-Z and S-B, where P is 2-amino-8-(2-thienyl)-purine and Z is 6-amino-5-nitropyridin-2-one, paired with S as 6-methyl-8-(6-methylisoxazol-3-yl)-purine and B as 2-nitro-6-(2-thienyl)-pyridine). This system supports stable duplex formation, replication, and transcription in vitro, with potential for up to 512 unique codons (8³), vastly exceeding the natural 64. While full in vivo implementation of the eight-letter code in E. coli remains under development, partial successes include E. coli RNA polymerase-mediated transcription of six- and eight-letter templates with high selectivity (greater than 99.9% for unnatural base incorporation).^[83] With a six-letter alphabet, the theoretical expansion yields 216 total codons, providing 152 additional codons beyond the natural set for encoding new nsAAs or functions. Despite these advances, limitations persist, particularly in replication fidelity, which for UBPs like d5SICS-dNaM averages around 99% in vivo—slightly lower than the near-100% for natural pairs—leading to gradual erosion of unnatural nucleotides over generations without selective pressure. Toxicity from unnatural nucleoside triphosphates and the need for evolved polymerases or transporters to maintain cellular uptake further challenge scalability. Ongoing efforts aim to improve these metrics through computational design and directed evolution to enable robust, heritable expansion.^[84]^[85]

Selective Pressure Incorporation

Selective Pressure Incorporation (SPI) is a method for residue-specific incorporation of non-standard amino acids (nsAAs) into recombinant proteins expressed in vivo, relying on the natural mistranslation machinery of auxotrophic host cells. The protocol involves culturing auxotrophic bacterial strains, such as Escherichia coli mutants deficient in biosynthesis of a specific canonical amino acid (e.g., methionine or proline auxotrophs), in chemically defined minimal media that lacks the essential amino acid but is supplemented with a structurally similar nsAA analog.^[86] As the cells deplete any residual natural amino acid and experience amino acid starvation, the endogenous aminoacyl-tRNA synthetases mischarge cognate tRNAs with the nsAA, enabling its global incorporation at all codon positions corresponding to the targeted natural amino acid during protein translation, resulting in functional proteins if the analog maintains structural compatibility.^[86] This pressure-driven selection ensures survival and growth only through utilization of the provided nsAA, typically yielding recombinant proteins with high purity after purification.^[16] The technique traces its origins to the 1970s, with early demonstrations using auxotrophic strains to incorporate D-amino acids into proteins and study stereochemical effects on folding and function, as shown in model enzymes like beta-galactosidase.^[87] Over time, SPI has been extended to other nsAA analogs, including fluorinated or phosphorylated variants, with reported incorporation yields reaching up to 100% for simple, single-domain proteins under optimized conditions.^[86] As of 2025, advances in SPI include residue-specific ncAA exchange in auxotrophic hosts for global proteome engineering and antimicrobial peptide design.^[88] A primary advantage of SPI is that it circumvents the need for genetic engineering of tRNAs, synthetases, or the host ribosome, allowing straightforward adaptation of standard expression systems for nsAA production.^[16] This makes it particularly suitable for global, residue-specific incorporation across the entire proteome or targeted recombinant proteins, facilitating studies on protein stability, enzyme kinetics, or antimicrobial peptide design without complex orthogonal components.^[86] Despite these benefits, SPI is constrained to nsAAs that serve as effective analogs recognizable by host synthetases, limiting its scope to chemically similar variants rather than highly diverse or unnatural structures.^[16] Furthermore, the proteome-wide replacement can induce off-target effects, such as misfolding of essential cellular proteins or reduced host viability, necessitating careful selection of auxotrophs and growth conditions to minimize toxicity.^[86]

In Vitro Protein Synthesis

In vitro protein synthesis enables the translation of expanded genetic codes using cell-free systems, which provide a controlled environment for incorporating non-standard amino acids (nsAAs) without the constraints of living cells. These platforms typically rely on supplemented extracts or reconstituted components to support orthogonal translation machinery, allowing rapid prototyping and modification of proteins with nsAAs at specified codons. By decoupling synthesis from cellular metabolism, such systems facilitate direct addition of orthogonal tRNA/synthetase pairs and nsAAs, often targeting amber (TAG) stop codons suppressed by engineered suppressors.^[89] Key systems include the PURExpress platform, a commercially available reconstituted E. coli-based system with purified translation factors, which can be augmented with orthogonal tRNA/synthetase pairs for nsAA incorporation. For instance, the pyrrolysyl-tRNA synthetase (PylRS)/tRNA^CUA_Pyl pair enables site-specific integration of nsAAs like N^ε-acetyl-lysine into target proteins. Similarly, crude E. coli lysates derived from release factor 1 (RF1)-deficient strains, such as the genomically recoded C321.ΔA, support high-fidelity nsAA incorporation when supplemented with orthogonal components and nsAAs, achieving yields of 0.9–1.7 mg/mL for superfolder GFP variants bearing p-azido-L-phenylalanine or similar ncAAs.^[90] These setups bypass the need for cellular uptake or toxicity concerns associated with nsAAs, enabling high-throughput screening of protein variants. Advantages of these cell-free approaches include accelerated iteration cycles for genetic code expansion, as reactions can be optimized in hours rather than days, and the absence of cellular barriers allows direct testing of potentially cytotoxic nsAAs or synthetases. In the 2020s, integration with microfluidics has further enhanced throughput, with droplet-based or continuous-flow formats yielding nsAA-modified proteins at scales of several mg/L per reaction while minimizing reagent consumption.^[91] For example, continuous-exchange cell-free (CECF) formats, which use dialysis to replenish energy sources and remove byproducts, have enabled the production of proteins with multiple nsAAs, such as dual incorporation of acetyl-lysine and thioacetyl-lysine into histone H3 tails, supporting studies of post-translational mimicry.^[89] Despite these advances, limitations persist, including short active translation windows—typically 4–20 hours—due to energy depletion and factor instability, which restrict overall productivity compared to in vivo systems. Scalability also poses challenges, as batch reactions remain dominant for preparative scales, while microfluidic setups, though efficient for screening, struggle with transitioning to gram-level production without specialized infrastructure. Ongoing refinements, such as energy-regenerating modules, aim to address these issues for broader adoption in synthetic biology. As of 2025, cell-free systems have advanced with automated biofoundries for scalable ncAA protein production and integration with AI for drug discovery.^[92]^[93]

Chemical Protein Synthesis

Chemical protein synthesis provides a non-biological approach to incorporating non-standard amino acids (nsAAs) into proteins by assembling synthetic peptide fragments, enabling precise control over sequence and modifications independent of ribosomal machinery.^[94] This method is particularly valuable for the expanded genetic code, as it allows the integration of nsAAs—such as fluorophores, photocrosslinkers, or post-translationally modified residues—directly into defined positions during fragment synthesis.^[95] Unlike biological systems, chemical synthesis circumvents limitations of cellular translation, offering flexibility for studying protein function with unnatural building blocks.^[96] The cornerstone technique is native chemical ligation (NCL), developed in the 1990s, which chemoselectively joins unprotected peptide segments in aqueous solution to form a native amide bond.^[97] In NCL, a C-terminal thioester on one peptide fragment reacts with an N-terminal cysteine on the second fragment, proceeding via a transient thioester intermediate that rearranges to yield the ligated product without racemization or side reactions.^[94] This process is highly efficient for peptides up to approximately 50 amino acids per fragment and has been demonstrated to work with all 20 proteinogenic amino acids, readily extending to nsAAs incorporated during solid-phase peptide synthesis.^[95] For instance, nsAAs like β-turn mimics or metal-binding residues can be placed at ligation sites or elsewhere, facilitating the creation of modified proteins for biophysical studies. A key variant, expressed protein ligation (EPL), combines chemical and recombinant methods to extend NCL to larger proteins by generating recombinant thioester-tagged polypeptides via intein-mediated cleavage.^[98] Developed in 1998, EPL allows ligation of a synthetic peptide containing nsAAs to the C-terminus of an expressed protein domain, enabling semisynthesis of full-length proteins with site-specific modifications.^[99] This approach has been instrumental for introducing nsAAs in contexts requiring post-translational additions, such as ubiquitination, where NCL joins ubiquitin thioesters to target protein cysteines, yielding natively linked polyubiquitin chains for functional assays.^[100] Advantages of these techniques include atomic-level precision in nsAA placement and the ability to perform ligations under mild conditions that preserve protein folding, contrasting with global incorporation methods.^[101] They enable the synthesis of proteins with homogeneous modifications, such as singly ubiquitinated histones, which are challenging to produce biologically.^[102] In the 2020s, advances in automated flow chemistry have enhanced scalability, allowing rapid synthesis and sequential NCL of peptide fragments to assemble proteins exceeding 100 amino acids, such as antibody fragments and enzymes, in hours rather than days.^[103] These developments have broadened applications in probing nsAA effects on protein stability and interactions. As of 2025, recent NCL enhancements include traceless multi-segment ligations for complex protein assemblies and improved strategies for N-terminal modifications with ncAAs.^[104]^[105]

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Jun 23, 2024 · Due to its high availability, purity, printability, and biocompatibility, it is often used as a bioink in bioprinting applications [28, 29].
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Rapid discovery and evolution of nanosensors containing ... - Nature
Sep 5, 2024 · Here we report on a platform that streamlines biosensor discovery and unlocks directed nanosensor evolution through genetically encodable fluorogenic amino ...
[74]
Decoupling Protein Production from Cell Growth Enhances the Site ...
Nov 5, 2020 · MmOP is orthogonal in E. coli, yeast, and mammalian cells, which means it does not interfere with the aminoacyl-tRNA synthetases nor tRNAs of ...
[75]
Sc3.0: revamping and minimizing the yeast genome
Aug 13, 2020 · Here, we present a proposal to design and synthesize the next version of the synthetic yeast genome, dubbed Sc3.0.Sc3. 0 Genome Design · Genome Minimization By... · Sc3. 0 Benefits And...
[76]
Efficient and sequence-independent replication of DNA containing a ...
Jul 6, 2012 · We recently developed a class of candidate unnatural base pairs, exemplified by the pair formed between d5SICS and dNaM. Here, we examine the ...Sign Up For Pnas Alerts · Results And Discussion · Pcr Selection And...
[77]
A unified Watson-Crick geometry drives transcription of six-letter ...
Dec 12, 2023 · Here, we show that E. coli RNAP selectively recognizes unnatural nucleobases in a six-letter expanded genetic system.
[78]
A semisynthetic organism engineered for the stable expansion of the ...
The genetic alphabet encodes all biological information, but it is limited to four letters that form two base pairs. To expand the alphabet, we developed ...
[79]
Expansion of the Genetic Alphabet: A Chemist's Approach to ...
Dec 2, 2017 · The most fundamental approach to this goal is the creation of a fifth and sixth nucleotide that pair to form a third, unnatural base pair (UBP).Abstract · Conspectus · Figure 2
[80]
Incorporation of non-standard amino acids into proteins - PMC - NIH
Feb 21, 2019 · In this review, we focus on the recent achievements aimed at overcoming the challenges of NSAA incorporation and the emerging applications of proteins ...
[81]
The Application of Cell-Free Protein Synthesis in Genetic Code ...
Mar 19, 2019 · An overview of genetic code expansion. The evolved aminoacyl-tRNA synthetase (AARS) charges the tRNA with the non-canonical amino acid (ncAA).
[82]
Native Chemical Ligation of Peptides and Proteins - PMC - NIH
Native chemical ligation (NCL) enables the assembly of long peptides and proteins from their cognate peptide building blocks (Camarero & Muir, 2001; P. Dawson, ...
[83]
Protein synthesis by native chemical ligation - PNAS
The efficacy of all 20 naturally occurring amino acids in native chemical ligation reactions has been demonstrated by synthesis and analysis of defined mixtures ...
[84]
[PDF] SYNTHESIS OF NATIVE PROTEINS BY CHEMICAL LIGATION ...
A remarkable feature of the native chemical ligation of unprotected peptide segments is the absence of racemization in the coupling reaction. Detailed studies ...
[85]
Synthesis of Proteins by Native Chemical Ligation - Science
A simple technique has been devised that allows the direct synthesis of native backbone proteins of moderate size.
[86]
Expressed protein ligation: A general method for protein engineering
A protein semisynthesis method—expressed protein ligation—is described that involves the chemoselective addition of a peptide to a recombinant protein.
[87]
Expressed protein ligation: a general method for protein engineering
Jun 9, 1998 · Expressed protein ligation is a simple and powerful new method in protein engineering to introduce sequences of unnatural amino acids, posttranslational ...
[88]
Chemical methods for protein site-specific ubiquitination - PMC
Feb 25, 2021 · Native chemical ligation (NCL), characterized by the ligation of a N-terminal cysteine-containing peptide and a second peptide possessing a C- ...
[89]
Semisynthesis of Proteins by Expressed Protein Ligation
Jul 1, 2003 · Expressed protein ligation (EPL) is a protein engineering approach that allows recombinant and synthetic polypeptides to be chemoselectively and ...
[90]
Semisynthesis of Ubiquitinated Histone H2B via Native Chemical ...
Apr 19, 2022 · The ubiquitylation of histone H2B at Lys (K) 120 is known to be associated with methylation of H3K4 and H3K79, transcription elongation, ...
[91]
Synthesis of proteins by automated flow chemistry - Science
May 29, 2020 · Hartrampf et al. used an automated chemistry platform to optimize fast-flow peptide synthesis and were able to produce fully synthetic single-domain proteins.
[92]
Enhancing Native Chemical Ligation for Challenging ... - NIH
In this review, we focus on recent developments aimed at enhancing and expanding native chemical ligation for challenging protein syntheses.