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Cell-free system

A cell-free system is an in vitro biochemical platform that utilizes cellular extracts or purified components to replicate and study biological processes, such as protein synthesis, metabolic pathways, and gene expression, in the absence of intact living cells.^[1] These systems provide a controlled environment for harnessing cellular machinery like ribosomes, enzymes, and transcription factors without the barriers of cell walls or membranes.^[2] The origins of cell-free systems date to 1897, when Eduard Buchner demonstrated alcoholic fermentation using cell-free yeast extracts, establishing that enzymes could catalyze reactions independently of viable cells.^[2] Significant progress in protein synthesis occurred in the mid-20th century; in 1950, Paul Zamecnik and colleagues showed amino acid incorporation into proteins using rat liver extracts, and by 1961, Marshall Nirenberg and Heinrich Matthaei employed Escherichia coli extracts to decode the genetic code, linking messenger RNA to polypeptide synthesis.^[3] Over decades, refinements in extract preparation—such as sonication and dialysis—have boosted yields from micrograms to milligrams of protein per milliliter.^[3] Cell-free systems are classified into two primary types: extract-based, which rely on crude lysates from prokaryotic (E. coli) or eukaryotic sources (e.g., rabbit reticulocytes, wheat germ, or insect cells) to provide a complex mix of endogenous components, and purified or reconstituted systems, exemplified by the PURE system developed in 2001, which assembles defined recombinant enzymes, ribosomes, and substrates for customizable reactions.^[3] Extract-based systems offer high activity and scalability but may include inhibitors, while purified systems enable precise modifications, such as incorporating non-canonical amino acids.^[1] Both formats support batch, continuous-exchange, or microfluidic setups to sustain reactions for hours to days.^[1] Key applications span fundamental research and biotechnology, including rapid prototyping of genetic circuits, high-throughput protein engineering, and production of therapeutics like antibodies or toxic proteins that are challenging in living cells.^[1] Advantages include accelerated timelines (hours versus days for cell-based methods), inherent biocontainment, and flexibility for adding unnatural components or harsh conditions without toxicity concerns.^[3] Emerging uses involve biosensors for diagnostics (e.g., detecting pathogens like SARS-CoV-2), decentralized manufacturing via lyophilized kits, and synthetic biology for building artificial cells or metabolic cascades producing biofuels and pharmaceuticals.^[1] Recent innovations, such as energy-regenerating modules and integration with nanomaterials, promise broader industrial adoption.^[4]

Fundamentals

Definition and principles

A cell-free system is an in vitro biochemical platform that utilizes cell lysates, crude extracts, or purified components to replicate essential cellular processes such as transcription, translation, and metabolic reactions outside of intact living cells.^[5] These systems are prepared by isolating subcellular fractions through techniques like centrifugation, which remove cell membranes while retaining cytosolic and organelle components necessary for biological activity.^[3] This approach enables the study and engineering of biomolecular processes in a controlled, open environment without the constraints of cellular barriers or homeostasis.^[6] The core principles of cell-free systems rely on assembling subcellular fractions—such as ribosomes, enzymes, and cofactors—to mimic the machinery of living cells and drive reactions like protein synthesis.^[3] These platforms can support both prokaryotic processes, often derived from bacterial extracts like those from Escherichia coli, and eukaryotic ones, using sources such as rabbit reticulocyte lysates or wheat germ extracts.^[1] Key terminology includes "cell extract," referring to crude lysates containing a mixture of cellular components, and "in vitro reconstitution," which involves assembling purified macromolecules to form minimal systems.^[7] Basic components of cell-free systems include nucleic acid templates (DNA or RNA) that provide genetic instructions for transcription and translation, along with amino acids or nucleotides as building blocks for biopolymer synthesis.^[5] Energy sources such as ATP, GTP, and regeneration substrates like phosphoenolpyruvate (PEP) sustain these reactions, while salts and magnesium ions (Mg²⁺) maintain ionic conditions for enzymatic function.^[3] In protein synthesis, energy is primarily derived from the hydrolysis of ATP and GTP, which powers ribosomal translocation and tRNA charging; for instance, during translation elongation, GTP hydrolysis by elongation factors facilitates aminoacyl-tRNA delivery and peptide bond formation.^[5] These hydrolysis reactions can be represented as:

\text{ATP} + \text{H}_2\text{O} \rightarrow \text{ADP} + \text{P}_i + \text{energy}

\text{GTP} + \text{H}_2\text{O} \rightarrow \text{GDP} + \text{P}_i + \text{energy}

where P_i denotes inorganic phosphate, releasing energy to drive the non-spontaneous steps of translation.^[3]

Historical development

The origins of cell-free systems trace back to the late 19th century, when German chemist Eduard Buchner demonstrated that cell extracts could perform biochemical reactions without intact living cells. In 1897, Buchner prepared a yeast extract by grinding yeast cells with sand and filtering the mixture, showing that this acellular preparation could convert sugar into alcohol and carbon dioxide through fermentation, mimicking the process in living yeast.^[8]^[9] This work established the concept of enzymes as non-living catalysts capable of driving metabolic reactions independently of cellular integrity, challenging the prevailing vitalist views of the time. For his discovery of cell-free fermentation, Buchner was awarded the Nobel Prize in Chemistry in 1907.^[8] Advancements in the mid-20th century extended cell-free systems to protein synthesis, facilitated by techniques like ultracentrifugation that enabled the isolation of key cellular components such as ribosomes and soluble factors. In the 1950s, pioneering experiments demonstrated amino acid incorporation into proteins using bacterial extracts, laying the groundwork for mechanistic studies.^[10] A landmark achievement came in 1961 with Marshall Nirenberg and Heinrich Matthaei's use of an Escherichia coli S30 extract—a supernatant obtained after centrifugation at 30,000 × g—to synthesize proteins in vitro. By adding synthetic polyuridylic acid (poly-U) RNA to the extract, they observed the specific incorporation of phenylalanine into a polypeptide chain, producing polyphenylalanine and confirming that the triplet codon UUU encodes phenylalanine in the genetic code.^[11] This experiment not only validated the triplet nature of the genetic code but also highlighted the utility of cell-free systems for decoding RNA-directed protein synthesis.^[11] During the 1960s and 1970s, cell-free systems were adapted for eukaryotic translation, with rabbit reticulocyte lysates emerging as a key tool for studying hemoglobin synthesis and mRNA-dependent protein production. Initial demonstrations in the late 1950s and early 1960s showed that reticulocyte extracts could incorporate radioactive amino acids into hemoglobin, revealing regulatory mechanisms like heme control of globin synthesis.86041-6/fulltext) By the 1970s, refinements allowed the addition of exogenous mRNAs to direct the synthesis of specific proteins, making these lysates a standard for eukaryotic in vitro translation studies.33116-2/pdf)^[12] The late 20th century saw a shift toward more defined and controllable cell-free platforms, bridging traditional extracts to synthetic biology applications. In the 1980s and 1990s, efforts focused on optimizing extract-based systems for higher fidelity and yield, setting the stage for fully reconstituted approaches. This culminated in 2001 with Takuya Ueda's development of the PURE (protein synthesis using recombinant elements) system, a purified, recombinant-based cell-free translation platform that assembles all essential components—ribosomes, translation factors, aminoacyl-tRNA synthetases, and energy sources—without crude extracts, enabling precise control over protein production at rates up to 160 μg/ml/h.^[13]

Types

Cell extract-based systems

Cell extract-based systems utilize crude or semi-purified lysates derived from whole cells, providing a complex mixture of endogenous components that support protein synthesis without the need for intact cellular membranes. These extracts typically include ribosomes, transfer RNAs (tRNAs), messenger RNAs (mRNAs), aminoacyl-tRNA synthetases, initiation, elongation, and termination factors, as well as enzymes for transcription and energy metabolism. The natural complexity of these systems retains regulatory elements such as chaperones and folding factors, but also introduces impurities like nucleases and proteases that can limit reaction duration and efficiency.^[14] Prokaryotic cell extract-based systems, particularly those from Escherichia coli, are among the most widely used due to their high productivity for recombinant protein expression. The S30 extract, obtained as the supernatant after centrifugation at 30,000 × g, exemplifies this approach and contains the full machinery for coupled transcription-translation when supplemented with plasmid DNA and nucleotides. Developed through optimizations in the 1990s, the E. coli S30 system enables high-yield production of prokaryotic and simple eukaryotic proteins, often achieving 100–500 μg/mL in batch reactions under optimized conditions.^[15] Eukaryotic extracts offer advantages for synthesizing proteins requiring post-translational modifications, such as glycosylation and disulfide bond formation. Wheat germ extracts, prepared from embryonic tissues, provide a plant-based eukaryotic environment that supports efficient translation of mRNAs or DNAs, with particular utility for glycosylated proteins due to endogenous glycosyltransferases.^[14] Similarly, rabbit reticulocyte lysates, derived from anemic rabbit blood, facilitate mammalian-like folding and modifications, including phosphorylation and limited glycosylation, making them suitable for studying eukaryotic protein function. The broad spectrum of endogenous factors in cell extract-based systems enables seamless coupled transcription-translation, reducing the need for separate steps and allowing rapid prototyping of genetic constructs.^[16] Preparation commonly involves initial cell lysis followed by centrifugation to remove cellular debris and dialysis to exchange buffers, ensuring compatibility with added substrates.^[15] To sustain prolonged reactions, these systems incorporate energy regeneration mechanisms, such as the creatine phosphate/creatine kinase couple, which efficiently recycles ATP and GTP from ADP and GDP, preventing energy depletion.^[17]

Purified component-based systems

Purified component-based cell-free systems are assembled from individually purified or recombinantly produced biomolecules, enabling precise control over the reaction environment without cellular debris or unintended activities. These systems typically include ribosomes, translation initiation factors (such as IF1, IF2, and IF3), elongation factors (EF-Tu, EF-Ts, and EF-G), release factors (RF1, RF3), ribosome recycling factor (RRF), and 20 aminoacyl-tRNA synthetases, among other essential elements, all reconstituted in defined buffers with energy sources like ATP and GTP. The seminal PURE (Protein synthesis Using Recombinant Elements) system, for instance, comprises 31 such purified protein factors derived from Escherichia coli, allowing for modular assembly that minimizes off-target reactions inherent in crude extracts.^[13] A prominent example is the PURExpress kit, a commercial E. coli-based system that incorporates these purified components for in vitro transcription-translation, facilitating high-throughput protein synthesis with yields typically reaching 100–300 μg/mL in batch reactions. Another application involves synthetic minimal systems tailored for specific metabolic pathways, such as the reconstitution of glycolysis integrated with PURE components to drive energy production from sugars, demonstrating the versatility of these purified setups for pathway engineering. These systems are produced by overexpressing components in heterologous hosts like E. coli, followed by affinity purification to ensure homogeneity, which supports scalability and reproducibility.^[18]^[19] Key advantages of purified component-based systems include their high purity, which reduces nonspecific interactions and enables customization, such as the incorporation of orthogonal tRNAs for unnatural amino acid labeling without competing cellular elements. Optimized assembly ratios, for example, balance ribosomes at lower concentrations relative to amino acids and energy sources (often in the range of 1:10:100 by molar equivalents), to maximize efficiency while avoiding resource waste. Although yields are generally lower than those of extract-based systems—averaging around 160 μg/mL per hour in standard batch mode—the precision allows for targeted modifications, such as omitting specific release factors to enhance read-through for noncanonical translations.^[13]^[20]

Preparation

Extract preparation techniques

Cell-free systems rely on crude extracts derived from prokaryotic or eukaryotic sources to provide the necessary transcriptional and translational machinery. Preparation of these extracts involves cell disruption, clarification, and optimization to ensure high activity and low contamination.

Prokaryotic Extracts

Prokaryotic cell-free extracts are most commonly prepared from Escherichia coli due to its rapid growth, well-characterized genetics, and high yields of translationally active components. Cells are typically grown to mid-log phase (OD₆₀₀ ≈ 2–4) in rich media like 2× YT or LB, harvested by centrifugation at 5,000 × g for 10 min at 4°C, and washed in lysis buffer to remove media contaminants.^[21] Lysis is achieved through mechanical disruption to release cytoplasmic contents while minimizing damage to sensitive enzymes. Common methods include high-pressure homogenization using a French press at 20,000 psi (passed 2–3 times) or alumina grinding, where cells are mixed with acid-washed alumina powder (1:1 w/v) and ground in a chilled mortar for 10–15 min. These techniques yield intact membrane vesicles and high protein recovery, with French press preferred for scalability up to 30 mL batches.^[21]^[22] The crude lysate is clarified by centrifugation to produce the S30 extract, named for the sedimentation step at 30,000 × g for 30 min at 4°C, which pellets cell debris and unbroken cells while retaining ribosomes and soluble factors in the supernatant. Buffers for lysis and centrifugation typically consist of 10 mM Tris-HCl (pH 7.7–8.2), 60 mM potassium glutamate or acetate, 14 mM magnesium acetate, and 1–10 mM dithiothreitol (DTT) to maintain reducing conditions and ionic balance essential for ribosomal stability.^[21]^[23] Optimization steps enhance extract performance by reducing endogenous activity and stabilizing components. Treatment with DNase I or RNase A (10–50 μg/mL) for 15–30 min at 37°C degrades nucleic acids, preventing competition from host transcripts; this is quenched with EDTA or EGTA. Dialysis against S30 buffer for 3–18 h removes small molecules and inhibitors, though it is often omitted in high-throughput protocols without yield loss. Extracts are aliquoted, supplemented with 10–20% glycerol, flash-frozen in liquid nitrogen, and stored at −80°C for up to 6–12 months. Contamination is controlled by using RNase inhibitors (e.g., RNasin at 40 U/mL) during handling and endonuclease-deficient strains like BL21 to minimize DNA degradation.^[21]^[24] Yields from optimized E. coli extracts typically provide 20–30 mg/mL total protein, equivalent to approximately 20–30 mg protein per gram of wet cell weight, assuming 1–2 mL extract per gram. This supports protein synthesis yields up to 1–2 mg/mL in batch reactions.^[23]^[15]

Eukaryotic Extracts

Eukaryotic extracts, such as those from wheat germ or rabbit reticulocytes, incorporate post-translational modifications like glycosylation, making them suitable for complex eukaryotic proteins. Preparation emphasizes gentle lysis to preserve folding chaperones and initiation factors. For wheat germ extracts, embryos are first isolated from commercial wheat germ by flotation in a cyclohexane-carbon tetrachloride mixture (density 1.4 g/mL), yielding 30–40% viable material by weight. The dried embryos (5–10 g) are ground mechanically in a chilled mortar with an equal weight of acid-washed quartz sand or under liquid nitrogen to a fine powder, then homogenized in extraction buffer (e.g., 40 mM HEPES-KOH pH 7.6, 100 mM KOAc, 5 mM Mg(OAc)₂, 2 mM CaCl₂, 4 mM DTT). This disrupts plant cell walls without excessive heat. The homogenate is centrifuged at 23,000 × g for 10 min at 4°C (repeated), and the supernatant is desalted via gel filtration (e.g., Sephadex G-25) in column buffer to remove pigments and inhibitors.^[25] Rabbit reticulocyte lysates are prepared from anemia-induced rabbits bled via cardiac puncture, yielding reticulocytes (>90% purity). Cells are washed in buffered saline (e.g., 140 mM NaCl, 5 mM KCl, 5 mM glucose) and lysed hypotonically by resuspension in 1.5 volumes of ice-cold distilled water or low-ionic-strength buffer (e.g., 10 mM Tris-HCl pH 7.6, 10 mM KCl, 1.5 mM MgCl₂), followed by gentle mixing for 20–30 min at 0–4°C to induce osmotic bursting. In some protocols, 0.5% NP-40 detergent is added to aid membrane solubilization without denaturing factors. The lysate is clarified by centrifugation at 15,000 × g for 20 min at 2°C, optionally followed by microfiltration through 0.45 μm filters to remove particulates.^[24]^[26] Recent advances include the preparation of extracts from human cell lines such as HeLa or HEK293, which offer improved compatibility for human protein folding and modifications. These are typically prepared by harvesting cells, followed by hypotonic lysis or dual centrifugation at low temperatures (e.g., 10,000–20,000 × g for 10–20 min at 4°C) to obtain translation-competent supernatants, with total protein yields of 20–50 mg/mL and synthesis efficiencies up to 100–200 μg/mL. Extracts are optimized similarly with nuclease treatment and stored at −80°C.^[27] Shared optimization for eukaryotic extracts includes micrococcal nuclease treatment (10–40 U/mL, 15 min at 20°C) to eliminate endogenous mRNAs, quenched with 2 mM EGTA, ensuring exogenously added templates dominate. Dialysis against reaction buffer (e.g., 20 mM HEPES pH 7.6, 100 mM KOAc, 2 mM Mg(OAc)₂) refines ionic conditions, and extracts are stabilized with 10% glycerol, frozen in liquid N₂, and stored at −80°C. RNase inhibitors are routinely added during processing to protect added mRNAs. Wheat germ yields 10–20 mg/mL total protein, while reticulocyte lysates provide 50–100 mg/mL but with lower synthesis efficiency (up to 200 μg/mL protein).^[24]^[28]

Component purification methods

Component purification methods in cell-free systems involve isolating individual biomolecules such as ribosomes, translation factors, and aminoacyl-tRNA synthetases from overexpression hosts, typically Escherichia coli or yeast, to enable scalable production of recombinant components. Overexpression in E. coli is commonly employed due to its rapid growth and genetic tractability, allowing high-yield production of tagged proteins for subsequent purification. Affinity chromatography, often using His-tags on recombinant enzymes and factors, facilitates rapid isolation by binding to nickel or cobalt resins under denaturing or native conditions. Size-exclusion chromatography is applied for ribosomes to separate them based on molecular weight, ensuring removal of contaminants like free ribosomal subunits.^[13]^[29] Key protocols for isolating core components include sucrose gradient ultracentrifugation for ribosomes, where E. coli lysates are layered on a 10-40% sucrose gradient and centrifuged at 100,000g for several hours to pellet intact 70S ribosomes while separating 30S and 50S subunits. This method yields highly active ribosomes suitable for reconstitution, with buffers containing magnesium and ammonium ions to maintain assembly. For tRNA synthetases, ion-exchange chromatography using DEAE-Sepharose or DEAE-cellulose columns is standard; enzymes are eluted with increasing NaCl gradients (e.g., 0.3-1.0 M) after ammonium sulfate precipitation and gel filtration, achieving partial to high purity for specific amino acids like leucine or isoleucine. These protocols, often combined with overexpression of His-tagged variants in E. coli, support the modular assembly of defined systems like PURE.^[29]^[13] Assembly of purified components into functional cell-free systems requires mixing in precise ratios to mimic cellular concentrations and optimize translation efficiency. Typical formulations include approximately 1–1.5 μM ribosomes, 0.3 mM each amino acid, 1–2 mM each NTP, and 0.1–3 μM for individual translation factors, with energy regeneration components like creatine phosphate added at 10-20 mM.^[30] Quality control involves activity assays, such as in vitro translation of luciferase mRNA followed by luminescence measurement, to verify protein synthesis rates exceeding 100 μg/mL under optimized conditions. In the PURE system, components are purified to homogeneity (>95% purity via SDS-PAGE analysis), enabling precise control but highlighting challenges like factor instability during storage.^[13]^[31]^[13] To address instability of sensitive factors in reconstituted systems, strategies such as chemical modifications (e.g., PEGylation) have been explored to enhance solubility and thermal stability without compromising activity.^[32]

Advantages and limitations

Benefits over cell-based systems

Cell-free systems offer enhanced control over biochemical reactions compared to cell-based systems, allowing direct addition or removal of components such as enzymes, cofactors, or substrates without the constraints of cellular membranes or metabolic burdens.^[33] This enables the handling of toxic intermediates or products that would otherwise inhibit or kill living cells, facilitating the study and optimization of complex pathways.^[34] Additionally, the open reaction environment supports real-time monitoring of reaction dynamics using techniques like high-resolution mass spectrometry or fluorescence spectroscopy, providing immediate insights into metabolite concentrations and pathway bottlenecks.^[33] In terms of scalability and speed, cell-free systems enable rapid prototyping and high-throughput screening, with reactions completing in hours rather than days required for cell growth and expression.^[35] They are compatible with standard formats like 96-well plates, allowing parallel testing of variants for protein engineering or pathway optimization.^[36] Furthermore, these systems tolerate a broad range of non-physiological conditions, including pH values from below 5 to 9 and high salt concentrations, which would disrupt cellular integrity.^[37] Cell-free systems often achieve higher yields and purity than cell-based methods, with optimized extracts producing up to 2.3 mg/mL of protein in batch reactions while avoiding issues like inclusion body formation or proteolytic degradation.^[38] The absence of cellular machinery reduces off-target interactions, yielding cleaner products that require minimal purification.^[37] Specific examples highlight these advantages, such as their use in extreme environments like organic solvents for biocatalysis, where cell-free setups enable cascade reactions with nonpolar substrates that are incompatible with living cells.^[39] They also prove cost-effective for prototyping synthetic pathways, eliminating the need for cell culture media, cloning, and maintenance, thus reducing overall expenses and timelines.^[40]

Challenges and drawbacks

Cell-free systems, while offering precise control over biochemical reactions, suffer from inherent stability issues that limit their operational duration to typically a few hours due to the degradation of essential factors such as enzymes and energy sources.^[5] Protease contamination in cell extracts can further exacerbate this by degrading synthesized proteins.^[5] To mitigate these challenges, strategies like energy regeneration systems employing phosphoenolpyruvate help sustain ATP levels and prolong reaction times.^[17] Additionally, encapsulation within liposomes can protect components from degradation and mimic cellular environments, enhancing overall stability.^[35] The high cost and complexity of purified component-based systems represent another significant drawback, with commercial kits costing tens to hundreds of dollars per reaction depending on scale and provider.^[41] In contrast, extract-based systems are more affordable but prone to impurities, such as nucleases and variable enzyme activities, which introduce batch-to-batch inconsistencies and reduce reproducibility.^[40] These impurities can lead to unpredictable protein folding or activity, complicating reliable outcomes in repeated experiments.^[40] A core limitation of cell-free systems is their lack of native membranes and compartmentalization, which hinders the recapitulation of multi-organelle processes like those involving the endoplasmic reticulum for glycosylation or secretion.^[35] This absence restricts applications to simpler pathways and poses scalability bottlenecks for industrial use, as open reactions struggle with maintaining efficiency at larger volumes without cellular barriers to control diffusion and protect sensitive intermediates.^[5] Recent advancements, such as freeze-drying of extracts, address storage challenges by enabling stability for several months at room temperature, facilitating easier distribution and long-term preservation without refrigeration.^[42] As of 2025, innovations like AI-optimized extract compositions and nanomaterial integrations have further improved reaction yields and stability, mitigating some operational limitations.^[1]

Applications

Protein expression and synthesis

Cell-free protein synthesis primarily relies on coupled transcription-translation mechanisms, where a DNA template is simultaneously transcribed into mRNA and translated into protein by ribosomes. In prokaryotic systems, such as those derived from Escherichia coli, the bacteriophage T7 RNA polymerase drives efficient transcription from a T7 promoter, enabling rapid mRNA production that is immediately utilized by endogenous ribosomes for translation.^[3] This process supports high-fidelity protein production without cellular constraints, often using linear DNA templates that can achieve yields up to 1.5 mg/mL for specific proteins like trimeric outer membrane proteins.^[43] Optimizations in template design enhance expression efficiency, such as incorporating a 5' hammerhead ribozyme to precisely process the untranslated region (UTR) of the mRNA, removing extraneous sequences and improving translation initiation.^[44] Commercial systems, like Promega's TNT coupled transcription-translation kits based on rabbit reticulocyte lysates or wheat germ extracts, facilitate eukaryotic protein expression, while hybrid systems combining E. coli S30 and wheat germ extracts have been developed to boost yields of fluorescent proteins up to several-fold higher than individual extracts.^[45] In applications for vaccine production, cell-free systems enable the rapid synthesis of B-cell lymphoma antigens, such as single-chain variable fragment (scFv) fusion proteins, which elicit potent antilymphoma immune responses when administered to mice.^[46] For therapeutic proteins, these systems support the production of insulin precursors, allowing on-demand manufacturing of mature desB30-insulin in under 24 hours with yields suitable for preclinical evaluation.^[47] Eukaryotic cell-free extracts, such as rabbit reticulocyte lysates, enable certain post-translational modifications, including phosphorylation, which occurs post-translationally on synthesized polypeptides to regulate activity and folding.^[48] ATP depletion limits overall productivity in these systems.

Metabolic pathway reconstruction

Cell-free systems enable the reconstruction of metabolic pathways by assembling enzyme cascades outside living cells, allowing researchers to study and optimize biochemical networks in a controlled environment. These systems typically utilize crude cell extracts or purified enzyme sets to mimic natural metabolic routes, such as the 10-enzyme glycolysis pathway that converts glucose to lactate, providing insights into pathway efficiency without cellular interference. This approach facilitates the dissection of complex metabolisms, including non-native or engineered pathways, by enabling precise control over reaction conditions like pH, temperature, and cofactor availability.^[49] Key examples demonstrate the versatility of cell-free metabolic reconstruction. In one application, a cell-free system derived from Escherichia coli extracts produces dihydroxyacetone phosphate (DHAP) from glycerol using a four-enzyme cascade (glycerol kinase, acetate kinase, glycerol-3-phosphate dehydrogenase, and acylphosphatase), achieving up to 88% conversion yield under optimized conditions.^[50] Another notable reconstruction involves hydrogen production via the oxidative pentose phosphate pathway, where a 12-enzyme system from E. coli lysate converts glucose to 12 moles of H₂ per mole of glucose, highlighting the potential for biofuel synthesis with high theoretical yields.^[49] These cascades often incorporate redox balancing through added cofactors like NAD⁺/NADH, ensuring sustained activity over hours. Analysis of these reconstructed pathways relies on techniques to monitor and manipulate flux. Metabolite addition or depletion allows flux control, revealing bottlenecks; for instance, supplementing intermediates in cell-free glycolysis can shift carbon flux toward desired products like isobutanol. Kinetic modeling, such as the Michaelis-Menten equation v = \frac{V_{\max} [S]}{K_m + [S]}, quantifies enzyme behavior in these systems, where V_{\max} represents maximum velocity, [S] is substrate concentration, and K_m is the Michaelis constant, aiding predictions of pathway performance. Recent advances in the 2020s have enhanced pathway efficiency through compartmentalization. Multi-enzyme cascades assembled on DNA origami scaffolds spatially organize enzymes, reducing intermediate diffusion losses and boosting yields by mimicking cellular proximity.^[51] These innovations, often using purified components, underscore cell-free systems' role in scalable biomanufacturing.

Incorporation of unnatural amino acids

Cell-free systems enable the site-specific incorporation of unnatural amino acids (UAAs) into proteins through genetic code expansion, primarily by leveraging orthogonal tRNA/aminoacyl-tRNA synthetase (aaRS) pairs that do not cross-react with endogenous components. These pairs, often derived from archaeal or bacterial sources such as Methanocaldococcus jannaschii TyrRS/tRNA^CUA or Methanosarcina mazei PylRS/tRNA^CUA, charge the orthogonal tRNA with a specific UAA, allowing its insertion at a designated codon during translation.^[52] The process begins with the aminoacylation reaction, where the UAA binds to the tRNA catalyzed by the engineered aaRS, forming the charged aa-tRNA that is then recognized by the ribosome for incorporation.^[53] A common technique is amber suppression, utilizing the TAG stop codon reassigned to encode the UAA via an amber suppressor tRNA. In cell-free platforms, this is facilitated by supplementing the reaction with the orthogonal pair and the UAA, often in Escherichia coli-based extracts or reconstituted systems. Seminal work demonstrated this approach using a modified tyrosyl-tRNA synthetase to incorporate a keto-containing tyrosine analog at amber sites in cell-free translation.^[54] To enhance efficiency, extracts from recoded E. coli strains lacking release factor 1 (RF1) are used, as RF1 normally terminates translation at amber codons; such strains enable up to 2.5-fold higher yields of full-length proteins with UAAs.^[55] The PURE system, a reconstituted cell-free platform with purified ribosomal components, tRNAs, and translation factors from E. coli, achieves near-100% incorporation efficiency for certain UAAs due to its defined composition, which minimizes competition from natural amino acids.^[56]^[52] Optimizations involve supplementing the reaction mixture with the UAA, such as p-acetyl-L-phenylalanine, and engineering the aaRS for specificity; this has yielded 50-80% labeled protein in crude extract systems.^[52] Representative applications include the incorporation of fluorinated amino acids, like 4-fluorotryptophan, into proteins for ¹⁹F NMR spectroscopy, providing high-resolution structural insights without background interference from natural residues. Another example is the use of photocaged amino acids, such as O-nitrobenzyl-protected tyrosine, which allow light-controlled protein folding and activation in cell-free reactions, enabling temporal studies of protein dynamics. These methods expand the proteome's chemical diversity for biophysical and therapeutic studies, with ongoing refinements focusing on multi-site incorporation via RF1-depleted systems.

Emerging uses in synthetic biology and drug discovery

In synthetic biology, cell-free systems have enabled the prototyping of complex gene circuits, including those leveraging CRISPR-Cas mechanisms for in vitro DNA assembly and editing. For instance, cell-free protein synthesis (CFPS) platforms integrated with CRISPR-Cas9 have facilitated the development of portable diagnostic tools, such as freeze-dried circuits capable of detecting viral RNA like that of Ebola with high sensitivity in resource-limited settings.^[57] These circuits operate without cellular constraints, allowing rapid iteration and optimization of genetic logic gates, such as AND/OR operations, directly in crude extracts.^[57] Vesicle-encapsulated cell-free systems, known as cell-free vesicles (CFVs), serve as protocell mimics by combining CFPS with lipid or polymer membranes to emulate compartmentalized biology. Recent advances have demonstrated CFVs for synthesizing functional membrane proteins, like GPCRs, within liposomes or polymersomes, enabling studies of protein-ligand interactions in a controlled environment.^[58] In 2025, reviews highlighted CFVs' potential for bottom-up assembly of synthetic cells, with examples including RNA replication and glycosylation-enhanced protein production inside vesicles.^[58] Additionally, autotrophy engineering has progressed through cell-free CO2 fixation cascades, such as the synthetic CETCH cycle, which converts CO2 to malate and glycolate using over 15 purified enzymes, achieving efficiencies comparable to natural pathways while integrating light or electrical energy inputs.^[59] These cascades support sustainable biomanufacturing by decoupling fixation from cellular growth limitations.^[59] The PURE system, a reconstituted cell-free translation platform, has advanced synthetic biology by enabling the incorporation of unnatural nucleotides and amino acids, expanding the genetic code for novel biomolecular designs like modified aptamers. This system allows precise control over translation components, facilitating the synthesis of proteins with non-canonical building blocks for enhanced stability and function in aptamer-based sensors.^[60] In drug discovery, cell-free systems integrated with artificial intelligence and machine learning have accelerated antimicrobial peptide (AMP) screening by combining deep learning models to predict sequences with CFPS for rapid validation. A 2023 study utilized this pipeline to produce and test hundreds of de novo AMP variants from DNA templates in E. coli extracts, identifying broad-spectrum candidates against Gram-positive and Gram-negative bacteria with minimal toxicity.^[61] Machine learning optimization of CFPS buffers has further boosted yields, achieving up to 10-fold increases in protein production through active learning algorithms that explore combinatorial spaces of components like NTPs and amino acids.^[62] High-throughput GPCR production via CFPS supports binding assays for drug screening, with eukaryotic extracts yielding functional, membrane-embedded receptors like GLP-1R at 16 µg/mL for direct immobilization on beads.^[63] This enables radioligand competition assays to quantify affinity, bypassing cellular expression challenges for hard-to-produce targets.^[63] Emerging applications include scalable biomanufacturing with field-deployable CFPS kits for on-demand vaccine production, such as two-step conjugate vaccines against bacterial pathogens, optimized for decentralized manufacturing in 2025.^[64] Freeze-dried, programmable kits facilitate rapid deployment, producing immunogenic proteins without cold chain requirements.^[65] For drug delivery, a 2025 review on CFVs emphasized programmable release mechanisms, including pH-triggered disassembly in tumor microenvironments (pH ~6.5) for targeted therapeutic payloads like chemotherapeutics.^[58] These innovations underscore cell-free systems' role in bridging synthetic biology with therapeutic translation.

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ATP Regeneration from Pyruvate in the PURE System
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Cell-Free Protein Expression by a Reconstituted Transcription ...
Furthermore, SG-PURE synthesizes more protein than the basic commercial PURE system, which synthesizes less than 0.3 mg/mL of proteins (PUREfrex 1.0 or ...
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Chassis/Cell-Free Systems/Homemade E.coli S30/Preparation ...
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Place dried wheat germ in a pre-chilled 6 in. mortar, cover with liquid N2, and grind wheat germ to a fine powder with a pestle. The total grinding time ...
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One of the first CFPS systems was based on E. coli cell extracts, and developments of this system have aimed at enhancing the yields of de novo synthesized ...
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A critical comparison of cellular and cell-free bioproduction systems
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An all E. coli cell-free system was optimized to synthesize 2.3 mg/ml of protein using a maltose-based metabolism for ATP regeneration, lasting up to 10 hours.
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Sep 20, 2023 · Cell-free protein synthesis (CFPS) system is an innovative technology with a wide range of potential applications that could challenge current thinking.
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A Simple, Robust, and Low-Cost Method To Produce the PURE Cell ...
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Perspective: Solidifying the impact of cell-free synthetic biology ...
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Cell-Free Protein Synthesis: A Promising Option for Future Drug ...
By increasing the temperature from 27 to 30 °C, the protein yields were increased by almost 50%. ... Optimizing cell-free protein synthesis for increased yield ...
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Cell-free co-production of an orthogonal transfer RNA activates ...
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Development of high-yield autofluorescent protein microarrays using ...
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A vaccine directed to B cells and produced by cell-free protein ...
A vaccine directed to B cells and produced by cell-free protein synthesis generates potent antilymphoma immunity.
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On-demand insulin manufacturing using cell-free systems with an ...
Here we report on advancements in manufacturing insulin using cell-free protein synthesis (CFPS) systems to rapidly produce mature desB30-insulin in less than ...
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Differential post-translational modification of human type I keratins ...
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Tuned Protein Synthesis Machinery in Escherichia coli-Based Cell ...
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Advancing synthetic biology through cell-free protein synthesis - PMC
Devoid of living cells, CFPS system possess unique advantages for membrane proteins and has successfully overexpressed a large number of membrane proteins ...
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Cell-free protein synthesis and vesicle systems for programmable ...
Jun 5, 2025 · The open environment of CFPS offers precise control over protein synthesis by enabling the modulation of synthetic conditions. Additionally, ...
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Cell-Free Systems to Mimic and Expand Metabolism
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Cell-Free Synthesis of Proteins with Unnatural Amino Acids. The ...
In this chapter, the advantage of the reconstituted translation system, named PURE system, for the efficient incorporation of an unnatural amino acid at amber ...
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Cell-free biosynthesis combined with deep learning accelerates de ...
Nov 8, 2023 · We established a cell-free protein synthesis (CFPS) pipeline for the rapid and inexpensive production of antimicrobial peptides (AMPs) directly from DNA ...Missing: AI | Show results with:AI
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Large scale active-learning-guided exploration for in vitro protein ...
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A scalable cell-free manufacturing platform for two-step bioproduction of immunogenic conjugate vaccines
**Summary of Scalable Cell-Free Manufacturing for Vaccines and Field-Deployable Kits in 2025:**
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Automated and Programmable Cell-Free Systems for Scalable ...
Freeze-dried and remotely operable CFPS kits act as the “last-mile” of the synthetic biology pipeline, enabling programmable and scalable biological functions ...