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DNA synthesis

DNA synthesis is the process by which deoxyribonucleic acid (DNA), the molecule that carries genetic instructions in living organisms, is produced to duplicate genetic information or create custom sequences.^[1] In biological contexts, it primarily refers to DNA replication, a fundamental mechanism occurring during the S phase of the cell cycle that ensures each daughter cell receives an identical copy of the genome before division.^[1] This semi-conservative process involves unwinding the double helix and using each strand as a template to synthesize complementary new strands, resulting in two complete DNA molecules from one.^[2] Beyond natural replication, DNA synthesis also encompasses artificial methods in biotechnology, where short oligonucleotides are chemically assembled into longer DNA constructs for applications in synthetic biology and gene engineering.^[3] The biological replication of DNA requires four key components: nucleotide substrates in the form of deoxyribonucleoside triphosphates (dNTPs), a single-stranded DNA template, a short RNA or DNA primer to initiate synthesis, and a suite of enzymes within the replisome complex.^[2] Synthesis proceeds bidirectionally from origins of replication, forming replication forks where the leading strand is synthesized continuously in the 5' to 3' direction, while the lagging strand is produced discontinuously as short Okazaki fragments that are later joined by DNA ligase.^[2] Central to this process is DNA polymerase, which catalyzes the addition of nucleotides complementary to the template—adenine (A) pairs with thymine (T), and guanine (G) with cytosine (C)—while incorporating proofreading mechanisms to achieve an error rate as low as 1 in 10^7 base pairs.^[2] Other essential enzymes include helicase, which unwinds the DNA helix; primase, which synthesizes the RNA primers; and topoisomerase, which relieves torsional stress ahead of the fork.^[2] This highly coordinated replication, taking about 8 hours in human cells to copy approximately 6 billion base pairs, is crucial for growth, repair, and the faithful transmission of genetic material across generations.^[1] In laboratory settings, chemical DNA synthesis enables the de novo creation of DNA without biological templates, revolutionizing fields like genomics and synthetic biology.^[3] The dominant method, solid-phase phosphoramidite chemistry, involves a cyclic four-step process—deprotection, coupling of protected nucleoside phosphoramidites, capping of unreacted chains, and oxidation—repeated on a solid support to build oligonucleotides up to 100-200 nucleotides long with error rates below 0.5%.^[3] These short synthons are then assembled into larger genes or circuits using techniques such as polymerase cycling assembly (PCA) or Gibson assembly, which exploit overlapping sequences and enzymatic ligation to construct double-stranded DNA up to several kilobases.^[3] Advances in this technology, including microarray-based parallel synthesis, have drastically reduced costs to as low as $0.0001 per base and enabled high-throughput production, facilitating innovations like custom gene synthesis for therapeutics, metabolic engineering, and genome editing.^[3] Despite challenges like sequence-dependent errors and scalability for very long DNA, ongoing improvements in fidelity and automation continue to bridge the gap between reading (sequencing) and writing (synthesis) genomes.^[4]

Natural Mechanisms of DNA Synthesis

DNA Replication

DNA replication is a semi-conservative process in which each strand of the parental DNA double helix serves as a template for the synthesis of a new complementary strand, ensuring accurate duplication of the genome during the cell cycle. This model was proposed by James Watson and Francis Crick in 1953 based on the structure of the DNA double helix. It was experimentally confirmed in 1958 by Matthew Meselson and Franklin Stahl through density-gradient centrifugation experiments using isotopically labeled DNA in Escherichia coli, demonstrating that after one round of replication, the DNA molecules had an intermediate density, and after two rounds, they separated into hybrid and light forms. In prokaryotes like E. coli, replication initiates at a single origin of replication, oriC, where the DnaA protein binds to specific sequences and recruits the DnaB helicase (in complex with DnaC) to unwind the double helix by breaking hydrogen bonds, forming a replication bubble. Primase (DnaG) then synthesizes short RNA primers complementary to the single-stranded DNA templates, enabling DNA polymerase III holoenzyme—the primary replicative polymerase—to begin elongation in the 5' to 3' direction. During elongation, synthesis occurs continuously on the leading strand, while the lagging strand is synthesized discontinuously as short Okazaki fragments (typically 1000–2000 nucleotides long), each requiring a new RNA primer. DNA polymerase I removes the RNA primers using its 5' to 3' exonuclease activity and replaces them with DNA via its polymerase activity, while DNA ligase seals the resulting nicks by forming phosphodiester bonds. Termination occurs when replication forks meet at the terminus region, facilitated by the Tus protein binding to Ter sites to block helicase progression. The core polymerization reaction catalyzed by DNA polymerase is:

\text{dNTP} + (\text{DNA})_n \rightarrow (\text{DNA})_{n+1} + \text{PP}_\text{i}

where a deoxyribonucleoside triphosphate (dNTP) is added to the growing 3' end, releasing pyrophosphate (PPi).^[5] In eukaryotes, replication initiates at thousands of origins per chromosome (up to 100,000 across the human genome) to accommodate larger genomes, with origins activated in a cell cycle-timed manner influenced by chromatin structure, resulting in a slower replication rate of about 50 nucleotides per second compared to prokaryotes. The linear chromosomes pose an end-replication problem, where the lagging strand at telomeres cannot be fully completed by standard polymerases; this is addressed by telomerase, a reverse transcriptase that extends the 3' overhang using an RNA template, allowing subsequent priming and completion. Overall fidelity is maintained by the 3' to 5' exonuclease proofreading activity of replicative polymerases, which removes mismatched nucleotides, achieving an error rate of approximately 1 in 10^9 base pairs incorporated when combined with post-replication mismatch repair.^[6]^[7]

Reverse Transcription

Reverse transcription is the process by which DNA is synthesized from an RNA template, a discovery made independently by Howard Temin and David Baltimore in 1970 through their identification of an RNA-dependent DNA polymerase in retroviruses, which challenged the prevailing central dogma of molecular biology that genetic information flows unidirectionally from DNA to RNA. For this groundbreaking work, along with Renato Dulbecco's contributions to tumor virology, Temin and Baltimore shared the 1975 Nobel Prize in Physiology or Medicine. The key enzyme, reverse transcriptase (RT), functions as an RNA-dependent DNA polymerase that synthesizes complementary DNA (cDNA) strands using deoxyribonucleoside triphosphates (dNTPs) as substrates, and it also possesses intrinsic RNase H activity that degrades the RNA strand within RNA-DNA hybrids to facilitate subsequent synthesis steps.^[8] In retroviruses, RT is packaged within the viral particle and initiates synthesis shortly after infection.^[9] The process begins with the priming of viral RNA by a host tRNA annealed to the primer binding site (PBS) near the 5' end, leading to the synthesis of a minus-strand DNA from the 3' end of the RNA template.^[9] RNase H activity partially degrades the RNA template, exposing a cytosine-rich segment of the minus-strand DNA that anneals to the complementary 3' end of the RNA via repeated regions (R) and unique sequences (US), enabling a strand transfer.^[9] Minus-strand synthesis then continues to the 5' end of the RNA, after which RNase H fully removes the remaining RNA, except for the polypurine tract (PPT), which serves as a primer for plus-strand DNA synthesis.^[9] A second strand transfer occurs using the terminal repeats (R-US), allowing completion of the double-stranded DNA product, which includes long terminal repeats (LTRs) at both ends and is subsequently integrated into the host genome as a provirus by the viral integrase.^[9] Reverse transcriptase exhibits lower fidelity compared to cellular DNA polymerases, lacking 3'→5' exonuclease proofreading activity, with an error rate of approximately 1 in 10^4 nucleotides incorporated, contributing to high mutation rates in retroviruses that enable immune evasion and drug resistance. In biology, reverse transcription plays a central role in the retroviral life cycle, such as in HIV where the proviral DNA integrates and directs viral gene expression and replication.^[9] It also underlies the propagation of endogenous retroviruses (ERVs), ancient viral sequences comprising about 8% of the human genome that can influence gene regulation and evolution. Additionally, telomerase, a specialized ribonucleoprotein, employs a reverse transcriptase subunit (TERT) to extend telomeres using an internal RNA template (TERC), maintaining chromosomal ends during cell division in stem and cancer cells. The core reaction catalyzed by reverse transcriptase is represented as:

\text{dNTP} + (\text{RNA})_m \rightarrow (\text{DNA})_n + \text{PP}_i

where the enzyme incorporates deoxyribonucleotides opposite the RNA template without proofreading in most retroviral RTs.^[8]

DNA Repair Synthesis

DNA repair synthesis is a critical process in maintaining genomic integrity, involving the targeted polymerization of DNA to fill gaps created after the excision of damaged or erroneous segments. This localized synthesis occurs in several excision repair pathways, including base excision repair (BER), nucleotide excision repair (NER), and mismatch repair (MMR), where DNA polymerases incorporate deoxyribonucleotide triphosphates (dNTPs) in a 5' to 3' direction using the undamaged complementary strand as a template. Unlike full genome replication, repair synthesis is activated in response to specific lesions and typically involves short to moderate patch lengths, ensuring precise restoration without extensive chromosomal disruption.^[10] In BER, which addresses small, non-helix-distorting base lesions such as oxidative damage (e.g., 8-oxoguanine) or alkylation, the process begins with damage recognition by DNA glycosylases that excise the faulty base, creating an abasic (AP) site. AP endonuclease (APE1) then cleaves the phosphodiester backbone at this site, generating a single-nucleotide gap in short-patch BER (1 nt) or a longer gap in long-patch BER (2-10 nt). DNA polymerase β (Pol β) primarily fills short-patch gaps with high fidelity, lacking intrinsic proofreading but benefiting from low error rates (~10^{-5} per nucleotide), while Pol δ or ε handles longer patches in conjunction with flap endonuclease 1 (FEN1) to remove displaced flaps and DNA ligase to seal the nick. For example, oxidative base damage from reactive oxygen species is repaired via this pathway, preventing mutations that could lead to cellular dysfunction.^[11]^[12] NER targets bulky, helix-distorting adducts, such as UV-induced cyclobutane pyrimidine dimers (e.g., thymine dimers), excising a 24-32 nucleotide oligonucleotide segment containing the lesion through coordinated action of xeroderma pigmentosum proteins (XPA-XPG). The resulting gap is filled by Pol δ or ε, which incorporate nucleotides with proofreading activity for high fidelity (error rates <10^{-6}), assisted by replication factor C (RFC), proliferating cell nuclear antigen (PCNA), FEN1 for flap processing, and ligase I for joining. This pathway is essential for removing UV-induced damage, with synthesis ensuring accurate templating to avoid photoproduct persistence.^[10]^[13]^[14] MMR corrects replication errors, such as base-base mismatches or small insertion/deletion loops, primarily in eukaryotes through recognition by MutSα (MSH2-MSH6) or MutSβ (MSH2-MSH3) complexes, followed by recruitment of MutLα (MLH1-PMS2). Exonuclease 1 (EXO1) excises the erroneous strand up to 100-1000 nucleotides in a PCNA- and RFC-dependent manner, creating a gap filled by Pol δ, which employs 3'→5' exonuclease proofreading for exceptional fidelity (enhancing overall replication accuracy 100-1000-fold). FEN1 and ligase complete the process, with this pathway integrating with replication forks to post-replicatively correct errors and suppress recombination between divergent sequences.^[15]^[16]

In Vitro Enzymatic DNA Synthesis

Polymerase Chain Reaction (PCR)

The polymerase chain reaction (PCR) is an in vitro technique for exponentially amplifying specific segments of DNA, revolutionizing molecular biology by enabling the production of large quantities of target DNA from minute starting amounts without relying on living cells. Invented by Kary Mullis in 1983 while working at Cetus Corporation, PCR was first described in a 1985 publication that demonstrated its application in amplifying beta-globin sequences for sickle cell anemia diagnosis. Mullis received the Nobel Prize in Chemistry in 1993 for this innovation, which facilitated molecular cloning and other genetic analyses previously limited by the need for bacterial propagation.^[17] The core components of a PCR reaction include template DNA containing the target sequence, two oligonucleotide primers that flank the region to be amplified (typically 18-22 nucleotides long), deoxynucleotide triphosphates (dNTPs) as building blocks, and a thermostable DNA polymerase. The polymerase most commonly used is Taq, isolated from the thermophilic bacterium Thermus aquaticus, which maintains activity at high temperatures and has an optimal extension temperature of around 72°C.^[18] These components are assembled in a buffer solution and subjected to repeated thermal cycles in a thermocycler, mimicking the strand separation aspect of natural DNA replication but in a controlled, artificial environment.^[19] Each PCR cycle consists of three main steps: denaturation at 94-98°C to separate the double-stranded DNA template into single strands, annealing at 50-65°C where primers hybridize to their complementary sequences on the template, and extension at 72°C where Taq polymerase synthesizes new DNA strands by adding dNTPs to the 3' ends of the primers. Typically, 20-40 cycles are performed, leading to exponential amplification where the number of target DNA copies approximately doubles with each cycle under ideal conditions, yielding N = N_0 \times 2^n after n cycles, or more generally N = N_0 \times (1 + E)^n with efficiency E approaching 1 for optimized reactions.^[20]^[21] PCR has broad applications, including preparing DNA inserts for cloning into vectors, generating amplicons for Sanger sequencing, and forensic analysis such as short tandem repeat (STR) profiling to identify individuals from crime scene samples. Despite its utility, PCR can suffer from limitations like non-specific amplification due to primer misbinding at lower temperatures and formation of primer dimers, which compete with target amplification and produce extraneous products. These issues are often mitigated through optimization strategies, such as hot-start PCR, where the polymerase is initially inactivated (e.g., via antibodies or chemical modifications) and activated only during the first denaturation step to prevent premature activity.^[22]^[23] Taq polymerase lacks 3'→5' exonuclease proofreading activity, resulting in a fidelity of approximately 1 error per 10^4 to 10^5 nucleotides incorporated, which can introduce mutations during amplification of longer targets.^[24]^[25]

Reverse Transcription PCR (RT-PCR)

Reverse transcription polymerase chain reaction (RT-PCR) is a laboratory technique that combines reverse transcription of RNA to complementary DNA (cDNA) with subsequent PCR amplification, enabling the analysis of RNA templates such as messenger RNA (mRNA). Developed in the late 1980s, RT-PCR built upon the 1970 discovery of reverse transcriptase by Howard Temin and David Baltimore, which demonstrated RNA-templated DNA synthesis in retroviruses. The seminal demonstration of RT-PCR occurred in 1988, when Frohman et al. introduced a method to rapidly produce full-length cDNAs from rare transcripts using a single gene-specific primer, marking a key advancement for amplifying low-abundance RNAs.^[26]^[27] RT-PCR can be performed in two main formats: two-step or one-step protocols. In the two-step format, reverse transcription is conducted separately from PCR amplification, allowing independent optimization of each stage and the use of flexible priming strategies; this approach generates a stable cDNA pool that can be stored and used for multiple downstream PCR reactions.^[28] In contrast, the one-step format integrates reverse transcription and PCR in a single reaction tube using gene-specific primers, reducing handling steps, minimizing contamination risk, and simplifying high-throughput workflows, though it limits flexibility for multi-gene analysis.^[28] Both formats typically employ retroviral-derived reverse transcriptase, such as Moloney murine leukemia virus (MMLV) reverse transcriptase, which exhibits RNA-dependent DNA polymerase activity, alongside thermostable DNA polymerases like Taq for the amplification phase.^[29]^[28] The core process begins with cDNA synthesis from mRNA templates, initiated by reverse transcriptase using primers such as oligo(dT) to target the poly(A) tail, random hexamers for unbiased coverage across the transcriptome, or gene-specific primers for targeted transcripts.^[30] This step converts single-stranded RNA into double-stranded cDNA, which then serves as the template for standard PCR amplification using gene-specific primers, dNTPs, and a thermostable DNA polymerase to generate exponential copies of the target sequence.^[30] Key applications of RT-PCR include quantitative real-time reverse transcription PCR (RT-qPCR), which monitors amplification in real time to quantify gene expression levels. RT-qPCR employs fluorescent dyes like SYBR Green, which intercalates with double-stranded DNA to provide non-specific detection, or probe-based systems such as TaqMan, where a fluorophore-quencher-labeled probe enables specific target identification via hydrolysis during extension, yielding cycle threshold (Ct) values that inversely correlate with initial template abundance.^[31] In diagnostics, RT-PCR excels at viral load detection, as exemplified by its widespread use in quantifying SARS-CoV-2 RNA in clinical samples during the COVID-19 pandemic, where assays targeting the viral nucleocapsid or spike genes achieve high sensitivity for early infection diagnosis. Additionally, RT-PCR facilitates studies of alternative splicing by amplifying and distinguishing splice variants through isoform-specific primers, revealing regulatory mechanisms in gene expression.^[32] RT-PCR offers high sensitivity, capable of detecting low-abundance transcripts down to single-molecule levels in optimized assays, making it invaluable for analyzing rare RNAs in limited samples.^[33] However, its reliance on RNA substrates introduces risks from RNase contamination, which can degrade templates and lead to false negatives, necessitating rigorous RNase-free protocols and inhibitors. For relative quantification in RT-qPCR, the Livak method (also known as the 2^{-\Delta\Delta C_t} method) is widely used to normalize target gene expression against a reference gene and control sample. The formula calculates fold change as $2^{-\Delta\Delta C_t}, where \Delta C_t = C_{t,\text{target}} - C_{t,\text{reference}} represents the difference in Ct values between the target and reference genes, and \Delta\Delta C_t is the difference between \Delta C_t values of the sample and calibrator. This assumes near-100% PCR efficiency for both amplicons and provides a straightforward measure of relative expression changes.^[34] Unlike standard PCR, which amplifies directly from DNA templates, RT-PCR addresses RNA instability by incorporating a dedicated reverse transcription step with enzymes like MMLV reverse transcriptase, which lacks RNase H activity in engineered variants to preserve RNA templates during cDNA synthesis, enabling reliable analysis of transient RNA populations.^[29]^[35]

Error-Prone PCR

Error-prone PCR is a variant of the standard polymerase chain reaction (PCR) intentionally designed to incorporate random mutations into amplified DNA sequences, thereby generating diverse genetic libraries for directed evolution studies. Unlike conventional PCR, which aims for high-fidelity amplification, error-prone PCR exploits the inherent error rate of DNA polymerases under suboptimal conditions to introduce point mutations, insertions, and deletions at controllable frequencies. This technique emerged in the early 1990s as a key tool in protein engineering, with the foundational method described by Cadwell and Joyce in 1992, who demonstrated randomization of genes through biased PCR conditions using Taq polymerase. The development of error-prone PCR in the 1990s facilitated breakthroughs in directed evolution, notably by Frances H. Arnold, who applied it to evolve enzymes such as subtilisin E for enhanced thermostability and activity in organic solvents, achieving up to 100-fold improvements in targeted properties through iterative rounds of mutagenesis and screening. Similarly, Greg Winter and colleagues utilized error-prone PCR for antibody engineering, enabling affinity maturation of human antibody fragments displayed on phage, which increased binding affinities by orders of magnitude for therapeutic applications. These early applications underscored the method's utility in mimicking natural evolution in vitro to optimize protein function. Mutations in error-prone PCR primarily consist of transitions and transversions, with occasional insertions or deletions arising from polymerase slippage, resulting in a spectrum that can be biased toward specific nucleotide changes depending on reaction parameters. The mutation rate is tunable, typically achieving 1–5 mutations per kilobase, by adjusting factors such as the number of PCR cycles (often 20–30), which amplifies cumulative errors since each cycle introduces base-pairing mistakes at a low rate (e.g., 10^{-4} to 10^{-3} per base per cycle). The overall frequency approximates the product of per-cycle error rate and cycle number, allowing libraries with 10^6–10^9 variants for subsequent selection.^[36] Common methods to induce errors include using unbalanced dNTP concentrations (e.g., elevating dGTP and dCTP to favor G-C transitions while limiting dATP and dTTP), incorporating Mn^{2+} ions (0.05–0.5 mM) to chelate Mg^{2+} and destabilize base pairing, employing low-fidelity polymerases like Mutazyme II, or introducing nucleotide biases in primers to target specific regions. These approaches ensure even distribution of mutations across the gene while minimizing excessive frameshifts that could inactivate most variants.^[36] In applications, error-prone PCR serves as random mutagenesis for optimizing enzymes, such as enhancing the thermostability of lipases or proteases for industrial biocatalysis, where variants with improved performance under extreme conditions are screened from libraries. It is also pivotal in antibody affinity maturation, generating scFv or Fab fragments with higher antigen specificity for drug discovery. Mutant libraries created via error-prone PCR are often combined with recombination techniques like DNA shuffling—developed by Stemmer in 1994—to explore larger sequence spaces by reassembling beneficial mutations, amplifying diversity beyond simple point changes and yielding synergistic improvements in protein properties.^[36]

Chemical DNA Synthesis

Oligonucleotide Synthesis

Oligonucleotide synthesis refers to the chemical production of short, single-stranded DNA fragments, typically ranging from 10 to 200 nucleotides in length, using solid-phase phosphoramidite chemistry without the need for templates or enzymes. This method, pioneered in the early 1980s, enables the automated assembly of custom DNA sequences essential for applications in molecular biology, such as probes and primers. The process builds oligonucleotides in the 3' to 5' direction on a solid support, allowing for iterative addition of protected nucleoside monomers.^[37] The standard phosphoramidite approach involves four cyclic steps performed on an automated synthesizer. First, deprotection removes the 5'-dimethoxytrityl (DMT) protecting group from the growing chain using an acid, such as 3% trichloroacetic acid in dichloromethane, exposing the 5'-hydroxyl for the next addition. Second, coupling introduces a protected nucleoside phosphoramidite monomer (for adenine, cytosine, guanine, or thymine) activated by tetrazole, forming a phosphite triester linkage with approximately 98-99% efficiency per step. Third, capping acetylates any unreacted 5'-hydroxyl groups using acetic anhydride and N-methylimidazole to prevent further extension of failure sequences. Finally, oxidation converts the unstable phosphite triester to a stable phosphate triester using iodine in water or pyridine. These cycles repeat until the desired length is achieved, after which the oligonucleotide is cleaved from the support (e.g., controlled-pore glass beads) and deprotected under basic conditions.^[38] Nucleoside phosphoramidites serve as the key building blocks, featuring β-cyanoethyl-protected phosphates and base-specific protecting groups (e.g., benzoyl for adenine and cytosine, isobutyryl for guanine) to ensure selective reactivity. The overall crude yield for a 50-mer oligonucleotide is approximately 37% when assuming a 98% coupling efficiency per step, calculated as:

\text{[Yield](/page/Yield)} = (0.98)^n

where n is the number of nucleotides added. This exponential decay highlights the method's sensitivity to stepwise efficiency.^[39] Post-synthesis purification typically employs reverse-phase high-performance liquid chromatography (HPLC) or polyacrylamide gel electrophoresis (PAGE) to isolate full-length products from truncated failures. Modified oligonucleotides, such as those conjugated with biotin for affinity capture or fluorophores for detection, are readily synthesized by incorporating functionalized phosphoramidites during assembly. Despite its efficiency, phosphoramidite synthesis has limitations, including cumulative error rates from incomplete couplings and side reactions like depurination, which predominantly result in deletions and restrict practical lengths to around 100 nucleotides. Costs range from $0.10 to $0.50 per base, depending on scale and modifications, making it economical for short sequences but less viable for longer constructs.^[3]

Gene Synthesis and Assembly

Gene synthesis and assembly involves the hierarchical construction of long DNA sequences, typically exceeding 1 kb, from shorter synthetic oligonucleotides to produce custom genes de novo. This process enables the creation of novel genetic constructs without relying on natural templates, facilitating precise engineering for various biological applications. The field originated with the pioneering work of Har Gobind Khorana's group, who in 1977 achieved the total chemical synthesis of a functional Escherichia coli tyrosine suppressor tRNA gene, marking the first de novo assembly of a biologically active gene through ligation of chemically synthesized oligonucleotides.50484-1/fulltext) This breakthrough laid the foundation for modern techniques, which evolved in the 2000s with the advent of high-throughput commercial services, such as those pioneered by GeneArt starting in 2000, enabling scalable production of synthetic genes through automated assembly pipelines.^[40] Several enzymatic methods have been developed to assemble overlapping oligonucleotides into full-length genes, improving efficiency and reducing errors compared to early ligation-based approaches. Overlap extension PCR, introduced in 1989, utilizes thermocycling to anneal and extend overlapping oligonucleotides, allowing iterative assembly of gene fragments into complete sequences.^[41] Gibson assembly, described in 2009, performs isothermal joining in a single reaction through the coordinated action of a 5' exonuclease for end resection, a polymerase for gap filling, and a ligase for sealing nicks, accommodating up to several hundred kilobases with 20-40 bp overlaps for optimal efficiency.^[42] Golden Gate assembly, developed in 2008, employs type IIS restriction enzymes that cut outside their recognition sites to enable scarless, directional ligation of multiple fragments in a one-pot reaction, minimizing unwanted recombinations and supporting modular designs.^[43] Assembly efficiency in these methods generally increases with overlap lengths of 20-40 bp, balancing specificity and yield.^[44] Gene design for synthesis prioritizes features that enhance expression and compatibility with assembly methods, such as codon optimization to match host organism preferences, thereby improving translation efficiency without altering the protein sequence. Sequences are also engineered to avoid unwanted restriction sites that could interfere with cloning or assembly, particularly in methods like Golden Gate. Error correction is integral, as chemical oligonucleotide synthesis introduces mutations at rates of 1 in 100-300 bases; post-assembly strategies include selective hybridization to biotinylated probes for enriching correct sequences or high-throughput sequencing to verify and select error-free clones.00237-2) Applications of gene synthesis span synthetic biology and biotechnology, exemplified by the 2010 assembly of a 1.08 Mb synthetic genome for Mycoplasma mycoides by Craig Venter's team, which was transplanted into a recipient cell to create the first self-replicating synthetic bacterium, demonstrating the potential for genome-scale engineering.^[45] In vaccine development, synthetic genes serve as templates for in vitro transcription of mRNA encoding antigens, as seen in the rapid design and production of SARS-CoV-2 spike protein genes for COVID-19 vaccines.^[46] Metabolic engineering benefits from custom gene synthesis to introduce or optimize pathways, such as assembling polyketide synthase genes in yeast for biofuel production. Current capabilities support assembly of genes up to 50-100 kb, with commercial providers offering sequence-verified constructs delivered in weeks. Costs have declined to approximately $0.07-0.20 per base pair as of 2025, driven by high-throughput oligonucleotide production via microarray printing followed by automated assembly.^[47]^[48] This scalability enables widespread use in research and industry, though longer assemblies require iterative hierarchical strategies to maintain fidelity.

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M-MuLV reverse transcriptase: Selected properties and improved ...
Nov 22, 2021 · This review will give a brief description of the structure, thermal stability, processivity, and fidelity, focusing on improving M-MuLV RT for practical usage.
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A primer to directed evolution: current methodologies and future ...
Jan 27, 2023 · (b) In error-prone PCR, random mutations are introduced by a low fidelity polymerase, resulting in linear DNA fragments with point mutations, ...
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Deoxyoligonucleotide synthesis via the phosphoramidite method
Deoxyoligonucleotide synthesis via the phosphoramidite method. Gene Amplif Anal. 1983:3:1-26. Authors. M H Caruthers , S L Beaucage, C Becker, J W Efcavitch ...Missing: DNA | Show results with:DNA
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Technical Brief – Synthesis of Long Oligonucleotides - Glen Research
While an average coupling efficiency of 98.0% would theoretically yield 68% full-length product for a 20mer, it would only yield 13% full-length product for a ...
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The gene makers - SCI – Where Science Meets Business
GeneArt in Germany, the world's largest manufacturer of genes, has seen its production capacity double each year since it entered the market in 2000. 'The gene ...
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Site-directed mutagenesis by overlap extension using the ...
Apr 15, 1989 · Overlap extension represents a new approach to genetic engineering. Complementary oligodeoxyribo-nucleotide (oligo) primers and the polymerase chain reaction ...
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Enzymatic assembly of DNA molecules up to several hundred ...
Apr 12, 2009 · We describe an isothermal, single-reaction method for assembling multiple overlapping DNA molecules by the concerted action of a 5′ exonuclease, a DNA ...
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https://www.thermofisher.com/blog/life-in-the-lab/gibson-assembly-101-expert-cloning-tips-you-need-to-know/
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Gibson Assembly 101: Expert Cloning Tips You Need to Know
May 29, 2025 · The proper overlap length for Gibson Assembly typically ranges from 20 to 40 base pairs. This length is sufficient to ensure that the fragments ...What is Gibson assembly? · Gibson Assembly protocol · Gibson Assembly...
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Creation of a Bacterial Cell Controlled by a Chemically Synthesized ...
May 20, 2010 · We report the design, synthesis, and assembly of the 1.08–mega–base pair Mycoplasma mycoides JCVI-syn1.0 genome starting from digitized genome sequence ...
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mRNA vaccines for infectious diseases: principles, delivery ... - Nature
Aug 25, 2021 · Principles of mRNA design and synthesis. mRNA vaccines comprise synthetic mRNA molecules that direct the production of the antigen that will ...
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2025 Pricing Announcement - Elegen
Elegen now ships US-made, cell-free, ENFINIA™ standard complexity linear DNA as low as $0.15 / base pair and high-complexity plasmids from $0.20/ base pair.
[49]
Genes Overview | Twist Bioscience
From 7¢ (USD) per base pair*. ok, 2 - 4 ... We make use of Twist Bioscience's high success rate and fast turnover times of gene synthesis and custom cloning.