Week 9 HW: Cell Free Systems

Part 1: Cell-Free Protein Synthesis

Explain the main advantages of cell-free protein synthesis over traditional in vivo methods, specifically in terms of flexibility and control over experimental variables. Name at least two cases where cell-free expression is more beneficial than cell production.

Cell-free protein synthesis (CFPS) removes the constraint of keeping a living cell alive. In a normal in vivo expression experiment, every design choice has to be compatible with growth, metabolism, membrane integrity, and host viability. In CFPS, the transcription and translation machinery is retained, but the cell itself is gone, so the reaction becomes an open biochemical system that can be directly tuned. DNA concentration, magnesium and potassium levels, redox state, chaperones, cofactors, detergents, lipids, noncanonical amino acids, and energy substrates can all be adjusted without worrying about whether the host will survive.

That open format gives two major advantages. First, CFPS is much more flexible for rapid prototyping: I can test many DNA templates, promoter/RBS designs, or reaction conditions in parallel in a few hours instead of building and transforming strains. Second, it gives tighter experimental control because every important variable is directly set by the user rather than indirectly filtered through cellular regulation. If translation drops, I can alter Mg2+ or template concentration immediately; if a membrane protein aggregates, I can add nanodiscs or detergent directly to the reaction.

Cell-free expression is especially beneficial in at least two cases:

• Toxic proteins: pore-forming toxins, nucleases, or strong metabolic enzymes often kill or stress living hosts, but can still be expressed in CFPS because there is no cell viability to protect.
• Membrane proteins: these are difficult to express in vivo because they misfold, aggregate, or overload the membrane insertion machinery; in CFPS, membrane mimics such as liposomes, nanodiscs, or mild detergents can be added directly.
• Rapid circuit prototyping: gene circuits, biosensors, and promoter libraries can be screened much faster without cloning into cells and waiting for growth.
• Proteins with noncanonical chemistry: CFPS is well suited for adding isotope labels, unnatural amino acids, or unusual cofactors that may be hard for living cells to tolerate or import.

Describe the main components of a cell-free expression system and explain the role of each component.

A typical cell-free expression reaction has several core parts:

• Cell extract or purified Tx/Tl machinery: This is the engine of the system. Crude lysates from E. coli, wheat germ, insect, or mammalian cells contain ribosomes, tRNAs, aminoacyl-tRNA synthetases, translation factors, and many metabolic enzymes. In PURE systems, these are supplied as purified components rather than as a crude extract.
• DNA or mRNA template: This encodes the protein of interest. If DNA is used, it must include promoter, ribosome binding site or Kozak sequence, coding sequence, and terminator/polyadenylation features appropriate to the system.
• Amino acids: These are the building blocks used by ribosomes to make the protein.
• Nucleotides (ATP, GTP, CTP, UTP): These are required for transcription and for many steps of translation and energy transfer.
• Energy source and regeneration system: Protein synthesis consumes large amounts of ATP and GTP, so the reaction needs both an initial energy pool and a way to recycle it.
• Salts and cofactors: Magnesium and potassium are especially important because they control ribosome function, RNA folding, and enzyme activity. Other cofactors such as spermidine, folate derivatives, or reducing agents may also be needed.
• Buffer system: This maintains the pH and ionic environment so the enzymes remain active throughout the reaction.
• Accessory additives: Chaperones, disulfide bond isomerases, detergents, nanodiscs, liposomes, RNase inhibitors, protease inhibitors, or crowding agents can be added depending on the target protein.

In short, CFPS works by reconstituting the minimum biochemical environment needed for transcription and translation, then tuning that environment for the protein or circuit of interest.

Why is energy regeneration critical in cell-free systems? Describe a method you could use to ensure continuous ATP supply in your cell-free experiment.

Energy regeneration is critical because protein synthesis is extremely energy-intensive. ATP is required for tRNA charging and many upstream metabolic steps, while GTP is consumed during translation initiation, elongation, and translocation. In a closed reaction, the energy pool is depleted quickly, and inhibitory byproducts such as inorganic phosphate can accumulate. If ATP collapses, transcription slows, translation stalls, and yield drops sharply even if all other components are present.

One practical way to maintain ATP is to use a phosphoenolpyruvate (PEP) plus pyruvate kinase regeneration system. In this setup, ATP is consumed during the reaction and converted to ADP. Pyruvate kinase then transfers the high-energy phosphate from PEP back onto ADP, regenerating ATP continuously. This method is common because it is simple, fast, and effective for short to medium CFPS reactions.

In my experiment, I would pair PEP regeneration with optimization of magnesium and phosphate balance, because even a good energy donor can fail if phosphate buildup poisons the reaction. For longer reactions, I would also consider slower-burning substrates such as 3-phosphoglycerate, glucose, or maltodextrin, which often improve reaction longevity by releasing energy more gradually.

Compare prokaryotic versus eukaryotic cell-free expression systems. Choose a protein to produce in each system and explain why.

A prokaryotic system is best when the goal is speed, low cost, and high yield for proteins that do not require elaborate post-translational processing. If I wanted to produce sfGFP, I would choose an E. coli CFPS system because sfGFP folds well in bacterial conditions, does not need glycosylation, and can be produced quickly at high yield. It is also an ideal reporter for reaction optimization because fluorescence gives a direct readout of productive expression.

A eukaryotic system is preferable when the protein requires a eukaryotic folding environment, disulfide bond formation, microsomal insertion, or other processing steps. If I wanted to produce human erythropoietin (EPO), I would choose a mammalian or insect-derived cell-free system because EPO is a secreted human glycoprotein whose activity and stability depend strongly on proper eukaryotic folding and post-translational processing. An E. coli lysate could make the polypeptide, but it would be much less likely to produce a properly folded, functional therapeutic-like product.

How would you design a cell-free experiment to optimize the expression of a membrane protein? Discuss the challenges and how you would address them in your setup.

To optimize a membrane protein, I would design the experiment around co-translational insertion into a membrane mimic rather than expressing the protein into free solution and hoping it folds afterward. As a concrete example, I would use an E. coli CFPS system to express the bacterial potassium channel KcsA with a C-terminal GFP tag for rapid screening. The reaction would include preassembled nanodiscs made from MSP1D1 scaffold protein and a POPC:POPG lipid mixture, because KcsA is far more likely to remain soluble and native-like if it inserts into a bilayer during translation.

The main challenges are:

• Aggregation: hydrophobic transmembrane segments tend to precipitate in aqueous solution.
• Misfolding: even if the protein is made, it may not adopt the correct conformation or oligomeric state.
• Poor membrane insertion: the reaction may produce full-length protein that never enters a lipid environment.
• Reaction inhibition: detergents, excess DNA, or incorrect salt balance can reduce overall translation efficiency.

To address these issues, I would screen a matrix of conditions:

• nanodiscs versus small liposomes versus mild detergents such as DDM or LMNG
• low versus moderate DNA concentration
• 25, 30, and 37 degrees C reaction temperatures
• magnesium concentration and potassium glutamate concentration
• optional chaperone supplementation such as DnaK/DnaJ/GrpE

I would measure three outputs: total protein made, soluble or membrane-associated fraction, and functional activity after reconstitution. Total yield could be checked by SDS-PAGE or in-gel GFP fluorescence. Membrane insertion could be assessed by co-migration with nanodisc fractions or flotation assays. Function could be tested with a potassium flux assay after purification or direct reconstitution. The best condition would not simply be the one with the most protein, but the one that gives the highest amount of correctly inserted and functional channel.

Imagine you observe a low yield of your target protein in a cell-free system. Describe three possible reasons for this and suggest a troubleshooting strategy for each.

Three common causes of low CFPS yield are:

1. Poor template design or poor template quality.
   If the promoter is weak, the RBS is poorly matched, the DNA is degraded, or the coding sequence has problematic secondary structure, transcription and translation can both suffer. I would troubleshoot by checking DNA quality, comparing plasmid versus linear template, redesigning the 5’ untranslated region, and testing a stronger promoter or codon-optimized construct.

2. Incorrect reaction chemistry.
   CFPS depends sensitively on magnesium, potassium, pH, and energy balance. A reaction that is slightly off can collapse even when all components are present. I would run a small design-of-experiments screen varying Mg2+, K+, DNA concentration, and energy substrate, while using a known positive-control template such as sfGFP to determine whether the problem is the reaction mixture or the target itself.

3. Protein instability, aggregation, or degradation.
   Some proteins fold poorly, are protease-sensitive, or precipitate as they are made. I would troubleshoot by lowering reaction temperature, shortening reaction time, adding chaperones, adding protease inhibitors, or including membrane mimics or redox helpers if the target is a membrane protein or disulfide-rich protein.

Low yield is usually not caused by one single factor. In practice, I would troubleshoot in the order of template quality, reaction chemistry, and protein-specific folding issues, because that sequence separates general reaction failure from target-specific failure.

Part 2: Design of a Useful Synthetic Minimal Cell

1. Pick a function and describe it.

I would design a synthetic minimal cell (SMC) that senses theophylline and, in response, activates a nearby engineered probiotic bacterium. The idea is to convert a small molecule that the bacterium does not naturally monitor into a standard bacterial induction signal.

• Function: user-controlled activation of a probiotic gene program
• Input: theophylline
• Output of the synthetic minimal cell: IPTG release
• Output of the whole hybrid system: sfGFP in a proof-of-principle strain of E. coli Nissle 1917, or a therapeutic payload in a future version

This function could not be realized by cell-free Tx/Tl alone without encapsulation. If IPTG were simply mixed into a bulk cell-free reaction, it would diffuse directly to the bacteria and there would be no gated actuator step. The membrane compartment is what lets the SMC store the output signal until the input molecule triggers pore formation.

This function could be realized by a genetically modified natural cell, but that would require engineering a living probiotic to directly sense theophylline and carry the entire logic internally. The synthetic-cell version is more modular: the same probiotic responder could be paired with many different SMC sensors just by swapping the sensing module.

The desired outcome is that the probiotic turns on only when theophylline is present, giving an external chemical control knob over bacterial behavior without permanently hard-wiring the sensing logic into the living cell.

2. Design all components that would need to be part of the synthetic cell.

I would use a bacterial Tx/Tl system, not a mammalian one, because the key regulatory element here is a small-molecule riboswitch and the output is just pore formation and inducer release. No mammalian glycosylation or nuclear machinery is needed.

The SMC would communicate with the environment in two steps. First, theophylline diffuses across the vesicle membrane and binds the riboswitch, turning on pore synthesis. Second, alpha-hemolysin inserts into the vesicle membrane and releases encapsulated IPTG, which then diffuses to the surrounding probiotic cells and activates their lac-regulated gene circuit.

3. Experimental details

Lipids and genes

• Lipids: POPC, cholesterol, DSPE-PEG2000
• Tx/Tl system: E. coli S30 extract or PURE system
• Energy system: 3-phosphoglycerate or PEP-based ATP regeneration
• Synthetic-cell gene: Staphylococcus aureus hla encoding alpha-hemolysin, controlled by a theophylline riboswitch
• Encapsulated small-molecule cargo: IPTG
• Responder-cell genes: constitutive lacI plus sfGFP under PlacUV5 or Ptac in E. coli Nissle 1917

Measurement strategy

I would measure function primarily through the GFP output of the responder bacteria. In the presence of theophylline, the SMC should synthesize alpha-hemolysin, release IPTG, and induce bacterial GFP. The cleanest readout would be flow cytometry or plate-reader fluorescence of the E. coli Nissle reporter strain.

Key controls would include:

• no theophylline
• no hla DNA
• SMCs without encapsulated IPTG
• responder bacteria without the lac-regulated reporter

If needed, IPTG release could also be confirmed indirectly by comparing fluorescence kinetics or directly by chemical assay of the supernatant.

Part 3: Freeze-Dried Cell-Free Systems in Materials

One-sentence pitch

I propose a soft-robotic skin with embedded freeze-dried cell-free microcapsules that detect damage, generate a visible warning signal, and locally produce a crosslinking enzyme to help seal small tears.

How will the idea work?

The robotic skin would contain patterned microcapsules loaded with freeze-dried cell-free reactions, a DNA template for a visible chromoprotein, and a DNA template for microbial transglutaminase. These capsules would be embedded inside an elastomer layer that also contains a thin repair hydrogel rich in crosslinkable residues. When the skin is punctured or torn, a built-in water reservoir or ambient moisture would rehydrate the damaged region and activate the local cell-free reactions. The chromoprotein would mark the damaged area for easy inspection, while transglutaminase would crosslink the repair layer and help slow crack growth or fluid leakage long enough for replacement.

What societal challenge or market need will this address?

Soft robots are increasingly used in medical devices, warehouse automation, and search-and-rescue environments, but their compliant materials are vulnerable to small tears, abrasion, and puncture. Today, many failures are only discovered after performance drops or a leak becomes severe. A self-reporting, partially self-sealing skin would reduce downtime, improve safety, and make soft robots more practical in environments where immediate maintenance is difficult.

How do you envision addressing the limitation of cell-free reactions?

I would address activation and stability by storing the reactions in trehalose-stabilized, vacuum-sealed microcapsules laminated inside the material until damage occurs. Water-triggering is actually useful here, because damage can be coupled to capsule rupture or exposure to a local hydration layer. The one-time-use limitation can be handled by making the sensing-and-repair elements modular and replaceable, like sacrificial patches in high-strain regions. For long shelf life, the material would use oxygen and moisture barrier films so the cell-free modules stay dormant until needed.

Part 4: Mock Genes in Space Proposal

1. Background information

Long-duration missions may depend on dried DNA templates for on-demand production of medicines, enzymes, and diagnostics. Space radiation and temperature cycling could damage these templates and reduce the reliability of cell-free manufacturing. I want to test how well lightweight shielding preserves the functional expression capacity of stored DNA. This matters for humanity because future crews will need compact, stable biotechnology systems far from Earth, and it is scientifically interesting because it connects the space environment directly to the survival of usable genetic information.

2. Molecular or genetic target

Plasmid DNA encoding sfGFP under a T7 promoter, plus the T7-promoter-to-sfGFP junction as a PCR integrity marker.

3. How the target relates to the challenge

If spaceflight damages the promoter or coding sequence, BioBits should produce less GFP even when the same amount of DNA is added. Measuring fluorescence therefore converts DNA integrity into a simple functional readout. By comparing shielded and unshielded templates, I can test whether stored genetic instructions remain usable for future in-space biomanufacturing and biosensing.

4. Hypothesis or research goal

My hypothesis is that DNA stored behind lightweight, hydrogen-rich shielding will retain higher functional expression capacity than unshielded DNA after space exposure. The goal is to compare practical storage strategies for preserving genetic templates that could later be used in cell-free systems aboard spacecraft. This hypothesis is based on the fact that ionizing radiation causes strand breaks and base damage, while hydrogen-rich materials can reduce secondary particle damage more effectively than many denser materials. A functional BioBits readout is especially useful because a template may still be amplifiable by PCR yet perform poorly in transcription or translation.

5. Experimental plan

I would test freeze-dried plasmid aliquots stored in three conditions: unshielded, polyethylene-shielded, and aluminum-shielded, with matched ground controls. After exposure, each sample would be rehydrated in BioBits and GFP output would be measured with the P51 Molecular Fluorescence Viewer at fixed time points. The miniPCR would amplify the T7-sfGFP region from the same samples as an integrity control. Fresh plasmid would serve as a positive control, and no-DNA reactions would serve as negative controls.