Cell-Free Systems Intro: Advantages vs In Vivo TXTL

This video presents an integrated approach combining CRISPR-based in vivo metabolic engineering with cell-free biosynthesis for rapid prototyping of genes and pathways. The method uses tiled guide RNA libraries synthesized at JGI to systematically tune gene expression levels across entire metabolic networks, identifying optimal expression levels (not just maximum overexpression or knockout) that improve pathway performance. Rewired cells are then lysed to create enhanced cell-free extracts that enable faster reaction kinetics, higher productivities, and greater flexibility in reaction conditions compared to living cells. This bidirectional workflow allows researchers to discover beneficial genetic modifications in vivo and immediately test them in vitro, accelerating the design-build-test cycle for metabolic engineering.

Self-expression systems enable construction of biochemical systems outside living organisms using DNA programs. TXTL (Transcription-Translation) combines transcription and translation in vitro, also known as cell-free expression or IVTT. The All-E. coli TXTL system uses E. coli RNA polymerase core with sigma factor 70, providing hundreds of regulatory elements unlike T7-based systems. Performance has improved dramatically: Toolbox 1 (2012) achieved ~1 mg/mL GFP, Toolbox 3 ~4 mg/mL. For T7 phage expression, improvement from 10^8 to 10^14 PFU represents a 1000-fold increase. TXTL systems can be developed from various bacteria including Gram-positive and Gram-negative species. Using fluorescent aptamers and next-generation sequencing, researchers can characterize hundreds to 1000 different regulatory elements in a single tube. CRISPR technology works effectively in TXTL for prototyping. TXTL can express very large gene sets including entire phage genomes: MS2 (small RNA phage), 5x174 (11 kb), T7 phage (40 kb, 60 genes), and T4 phage (169 kb, almost 300 genes). Synthetic cells are created by encapsulating TXTL reactions in liposomes (phospholipid vesicles, 1-50 micrometers). Bottom-up synthetic cells are categorized into: protocells (origin of life, prebiotic molecules), artificial cells (non-natural molecules), and minimal cells (natural molecular machineries). Minimal cells require three essential components: compartment (physical boundary), metabolism (energy management), and information (genetic material). A signaling system was developed using the sigma-28 cascade to express MscL (mechanosensitive channel). A relay signaling system was constructed where external IPTG triggers MscL expression, allowing IPTG to leak inside, activating a switch to express MreB (cytoskeletal protein).

Acoustic liquid handling systems like the Labcyte Echo enable rapid, high-throughput prototyping of cell-free transcription-translation (TXTL) reactions by reducing reaction volumes to 2 microliters and accelerating setup times 5-10 times compared to manual pipetting, making it ideal for characterizing genetic circuits and CRISPR technologies in synthetic biology research.

Cell-free expression systems like myTXTL combined with high-fidelity gene fragments enable rapid protein production from DNA within hours rather than days, eliminating traditional cloning and cell culture steps; this approach allows researchers to screen protein libraries, analyze metabolic pathways, and test gene circuits efficiently by directly synthesizing proteins in an open reaction environment without requiring living cells.

TXTL (cell-free transcription-translation) systems enable rapid protein and RNA production without live cells. Researchers simply add DNA, and functional proteins appear within hours. This eliminates purification steps, avoids cytotoxicity, and enables parallel experimentation. Building on this capability, the IMPRINT method was developed to recreate bacterial methylation patterns. By expressing methyltransferases in TXTL reactions with SAM methyl donor, researchers produce plasmids with native methylation patterns. This approach can scale from 1-2 to many methyltransferases, accommodating the diverse methylation systems across bacterial species.

The cell-free hybrid system has multiple applications: scanning binding interfaces, screening protein binders, small molecule modulation assays, and biosensing. Limitations include detection restricted to high-affinity interactions (below 100 nM), difficulty with glycosylated proteins, and DNA synthesis costs. The system is most valuable as a pre-screening tool for large libraries, enabling rapid identification of top candidates before detailed characterization. The ability to integrate with computational design pipelines creates closed-loop design-build-test-learn cycles, democratizing protein engineering for biomedical applications including cancer research and therapeutic binder discovery.

The industrial perception of cell-free systems is undergoing a fundamental transformation driven by technological advances and demonstrated applications. Over recent years, costs have decreased while capabilities have expanded, leading to growing interest from pharmaceutical partners and manufacturers. Key applications include large-scale production of protein medicines and small molecules at scales ranging from 100 liters to thousands of liters. Beyond manufacturing, cell-free systems serve as powerful platforms for accelerating biological design through machine learning integration. The absence of cellular growth requirements enables dramatically faster DNA-to-protein timelines, eliminating cloning and transformation steps. Combined with automated liquid handling technologies, these systems support high-throughput screening essential for training machine learning models in protein engineering and synthetic biology design, positioning cell-free technology as a cornerstone of next-generation biomanufacturing and bioengineering workflows.

Cell-free expression systems like myTXTL combined with high-fidelity gene fragments enable rapid protein production from DNA within hours rather than days, eliminating traditional cloning and cell culture steps; this approach allows researchers to screen protein libraries, analyze metabolic pathways, and test gene circuits efficiently by directly synthesizing proteins in an open reaction environment without requiring living cells.

Cell-free systems enable three major applications: (1) Cell-free metabolism produces biofuels and chemicals by combining native E. coli enzymes with custom DNA-encoded enzymes; (2) Biosensors use transcription factors to detect environmental contaminants (antibiotics, toxins, heavy metals) with concentration-dependent signals; (3) Glycoengineering adds sugar groups to proteins (antibodies, vaccines) to enhance stability and immune recognition. Each application leverages the same core technology—mixing cell extract with DNA instructions to produce functional biological systems outside living cells.

Cell-free systems enable metabolic engineering outside living cells, allowing researchers to study and manipulate metabolic pathways without cellular constraints. These systems power transcription and translation in vitro using E. coli extracts supplemented with amino acids, NTPs, salts, co-factors, energy molecules, and template DNA. The historical development began with Edward Buchner's 1897 discovery that yeast extract could ferment sugar to alcohol without living cells, challenging the vitalism theory that organic chemicals could only be made by living organisms. This earned him the Nobel Prize in 1907. The first cell-free protein synthesis system was developed by Joan Folkers and Ernest Gil in 1955 using Staphylococcus aureus, followed by Lambert and Zamanik's E. coli-based system. Nirenberg and Matthaei's 1961 discovery of the first codon built upon these foundations. Jeffrey Zubet's foundational protocol around 1970-1980 introduced the S30 extract, which remains the standard reference for modern cell-free protein synthesis. In the early 2000s, Jim Schwarz and colleagues investigated glycolysis and extract metabolism optimization, while Vincent Noiret's group explored using maltodextrin as a fuel source for ATP synthesis. The field expanded to include cell-free synthetic biology, defined as understanding, utilizing, and expanding functions of natural biological systems through repurposed design, transformation, and resynthesis. Cell-free protein synthesis serves as the core technique known as in vitro protein transcription and translation technology. Cell-free systems enable two approaches to enzyme addition: pre-lysis enrichment (transforming cells with DNA expressing enzymes of interest) and post-lysis enrichment (synthesizing proteins directly in lysate using transcription-translation machinery). Researchers can mix and match lysates enriched with different enzymes to assemble enzymatic pathways, analyzing metabolites rather than protein concentration. Glycolysis and TCA cycle utilize PEP and glucose as starting molecules. Cell-free systems can produce 2,3-butanediol from glucose at titers greater than 80 grams per liter with peak productivities of 11 grams per liter per hour, much higher than cellular systems. Reverse beta-oxidation reverses the fatty acid cycle to make alcohols and acids, serving as a modular system that can produce many medium and long-chain fatty acids and alcohols by stitching acetyl-CoA molecules together and terminating the cycle at different points. Cell-free systems enable study of enzyme synergies in enzymatic pathways by screening homologues of enzymes at different positions and testing pair-wise combinations for better performance. SAMD-CoA mass spectrometry allows observation of different CoA constituents over 20-hour reactions, revealing bottlenecks and how enzyme concentrations affect pathway flux. Research has shown good correlation between cell-free system results and in vivo expression in organisms like Clostridium autoethanogenum, making cell-free systems effective for initial pathway screening. Cell-free systems have been extended to produce complex metabolic products including isoprenoids (bisabolene, pinene, limonene) and convert amino acids to polymer monomers like styrene from phenylalanine. For carbon dioxide fixation, researchers are building reform pathways that are thermodynamically favorable, tolerant to aerobic environments, composed of minimal enzymes, and bypass the rate-limiting CO2 step. Formate serves as an interesting intermediate because it is a reduction product of CO2. Since core reactions are not known to occur in nature, researchers use cell-free systems to synthesize DNA variants and produce proteins from them, testing enzyme homologues from nature and identifying key mutations through site-saturated mutagenesis to create enzymes with new substrate specificities.

Cell-free systems enable expansion beyond the 20 canonical amino acids by incorporating non-standard amino acids through new tRNAs, codons, and ribosomal modifications. However, amber stop codon suppression faces limitations including poor synthetase efficiency, transport protein binding challenges, and competition with endogenous release factors causing premature termination. Genomically recoded organisms with release factor one deletions solve these problems by eliminating competition, enabling production of proteins with multiple site-specific non-standard amino acids at yields up to 800 micrograms per milliliter. The bacterial ribosome comprises approximately 54 proteins (21 small subunit, 33 large subunit) plus rRNAs, with natural ribosomes constrained by the need to decode all cellular RNAs, limiting engineering possibilities. The Integrated Synthesis Assembly and Translation (ISAT) platform enables co-activation of ribosome RNA transcription, assembly, and protein translation within single reactions using cell-free crude extracts. Over four years, activity increased approximately 10,000-fold, approaching natural E. coli ribosome activity, enabling selection of modified ribosomes through antibiotic resistance screening.

The site-specific incorporation of unnatural amino acids into proteins in live cells was demonstrated by Schultz and coworkers in the early 2000s, achieved by identifying orthogonal tRNA-synthetase pairs from different organisms (particularly archaea) that can be engineered to incorporate non-natural amino acids without interfering with the native genetic code; this methodology involves selecting tRNAs that decode stop codons and synthetases that specifically recognize unnatural amino acids, followed by iterative protein engineering using combinatorial saturation mutagenesis and genetic selection to optimize orthogonality, enabling the incorporation of over 30 different unnatural amino acids as of 2006.

The Schultz method enables incorporation of unnatural amino acids into proteins by reassigning the amber stop codon (UAG) to encode an orthogonal tRNA-aminoacyl tRNA synthetase pair from Methanococcus jannaschii, which was engineered through directed evolution to avoid interactions with E. coli's native machinery; however, this method suffers from low efficiency (20-30% for single amino acids, dropping to less than 1% for multiple incorporations) due to competition between the orthogonal tRNA and release factor 1 at the ribosome's A-site, which can be partially addressed by using an orthogonal ribosome with specific 16S rRNA mutations.

Amber suppression enables incorporation of unnatural amino acids into proteins by exploiting the amber stop codon (UAG). The technique requires three components: a plasmid encoding the protein of interest with a UAG substitution at the desired position, a plasmid encoding orthogonal tRNA and aminoacyl-tRNA synthetase pairs that recognize only the unnatural amino acid, and the unnatural amino acid itself supplied during expression. This approach works in living cells, is bio-orthogonal to native cellular chemistry, and functions effectively even in the presence of cysteines. Applications include installing post-translational modifications (phosphorylated serine, acetylated lysine), fluorescent probes, EPR probes, and bio-orthogonal handles (norbornene, cyclooctynes) for targeted conjugation. Practical considerations include optimizing expression conditions, managing truncation products, avoiding isotope scrambling during C-13 labeling, and confirming incorporation via mass spectrometry.

Living systems use 20 standard amino acids as the basis for all proteins, but non-proteogenic amino acids can be successfully incorporated into growing polypeptide chains and proteins. This incorporation provides advantages including increased protein stability, resistance to proteasomal degradation, and extended protein activity. These non-natural building blocks also provide novel functionalities and enhance protein function. Applications include therapeutics, bioplastics, antimicrobial drugs, and precise biomolecule localization using fluorescent labels. The historical development began in 1966 with chemical modification approaches, progressed through recombinant DNA technology enabling site-directed mutagenesis, and culminated in the 1989 in vitro incorporation using tRNA ligase. In 2001, researchers demonstrated in vivo incorporation using orthogonal systems—enzymes that exist in normal cellular environments without cross-talk with host machinery. Multiple aminoacyl-tRNA synthetase and tRNA pairs were generated, enabling charging of multiple non-cognate amino acids. A critical bottleneck was the need for orthogonal ribosomes, which require orthogonal mRNA. Researchers mutated Shine-Dalgarno and anti-Shine-Dalgarno sequences through random mutation and selection to generate orthogonal ribosome and mRNA pairs.

Freeze-dried cell-free systems enable revolutionary approaches to diagnostics and manufacturing. Paper-based sensors spotted with wax-imprinted patterns allow distributed pathogen detection, with Ebola and Zika sensors developed in just five days. Distributed biomanufacturing delivers components and DNA separately for on-site production: 15 functional antibodies, violacein biosynthesis pathways, 10 antimicrobial peptides, and protein-based vaccines (diphtheria) that induce proper immune responses. This paradigm shifts from centralized factory production to localized, rapid manufacturing at the point of care.

Point-of-care diagnostics evolved from animal-based tests (rabbit pregnancy tests) to sophisticated paper-based sensors. The Clear Blue pregnancy test demonstrated commercial viability, selling 10 million units in its first year. Whitesides pioneered paper microfluidics using photolithography to create hydrophilic/hydrophobic patterns, but faced challenges including slow fluid flow and difficulty implementing wash steps. Hong Lou introduced origami folding to create three-dimensional devices, while slip chip technology enabled controlled reaction timing. Electrochemical sensing translates molecular recognition into measurable electrical signals: aptamers change conformation upon target binding, altering redox tag distances from electrodes. Cocaine detection demonstrated 50% signal increases. Hollow channel paper microfluidics replaced cellulose-filled channels with empty channels surrounded by wax barriers, achieving seven-fold speed improvements while producing textbook-quality cyclic voltammograms comparable to conventional laboratory equipment.

Cell-free biosensing uses lysed bacterial extracts containing transcription and translation machinery to detect biomarkers. This platform addresses limitations of traditional clinical testing by enabling finger-stick blood collection, low-cost assays (pennies per test), and deployment in resource-limited settings without electricity or specialized equipment. Polymer-based two-phase systems (dextran/PEG) enable compartmentalization of reactions into separate droplets, allowing simultaneous detection of nucleic acids, proteins, and small molecules from the same sample. Freeze-dried colorimetric reporters (X-gal/lacZ) provide visually interpretable readouts suitable for point-of-care applications.

Keith Pardee developed cell-free synthetic biology by extracting cellular machinery (proteins and DNA) and running experiments outside living cells. These extracts can be spotted on paper, freeze-dried, and stored at room temperature for extended periods without losing activity. This addresses the refrigeration requirements that limit synthetic biology in low-resource areas. Using regular computer printers, researchers can print arrays of paper strips containing diagnostic components. Molecular switches can be engineered to respond to specific molecules (like pathogen RNA) and produce visible outputs such as color changes. This enables rapid detection of antibiotic resistance genes in about 30 minutes, addressing the growing global problem of antibiotic resistance that now affects not just hospitals but also schools, campuses, and childcare centers.

Cell-free synthetic biology enables the creation of stable, portable diagnostic platforms by freeze-drying biological components onto paper, allowing rapid detection of pathogens (such as Ebola, Zika, antibiotic-resistant bacteria, HPV, and cancer-related biomarkers) without requiring living cells, specialized equipment, or refrigeration, with detection limits reaching femtomolar concentrations and costs as low as two cents per test.