Week 9 HW: Cell Free Systems

General Questions

1. 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 gives you a level of control over the reaction environment that you simply cannot get when working inside a living cell. Because there’s no cell membrane, you can directly add or remove components, adjust concentrations in real time, and introduce molecules that would be toxic to a living cell without worrying about killing your chassis. You also get direct access to the product without needing to lyse cells or purify through layers of cellular debris.

Two cases where cell-free is better than cell-based production is

MS2 L-protein punches holes in membranes and kills bacteria, you can’t reasonably produce it inside a living E. coli because it would lyse its own host before you getting meaningful yield. Cell-free lets you synthesize toxic protein in a controlled environment without that problem. It also lets you iterate and test on dozens of variants quickly.

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

A cell-free expression system is essentially the inside of a cell, extracted and reconstituted in a tube. It conssits of:

  • Cell extract: This is the ‘machinery’ containing ribosomes, translation factors, chaperones, and all the machinery needed to read an mRNA and assemble a protein.

  • DNA template or mRNA: This is what you want expressed. You can add a plasmid, linear PCR product, or pre-transcribed mRNA depending on whether you want transcription to happen in the reaction or not.

  • RNA polymerase: Needed if you’re starting from DNA typically T7 RNAP is added for prokaryotic systems since it’s fast and highly processive.

  • Amino acids: The building blocks. You supply all 20 at defined concentrations so the ribosomes have raw material.

  • Energy regeneration system: ATP is consumed rapidly during translation. You need a system to regenerate it typically phosphocreatine + creatine kinase, or PEP (phosphoenolpyruvate).

3. Why is energy provision 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 translation is ATP- intensive. The cell-free reaction has a finite supply, and without regeneration the reaction stalls within minutes.

The most common approach is the phosphocreatine/creatine kinase system that catalyzes the transfer of a phosphate group from phosphocreatine to ADP, regenerating ATP. This is simple to add and works well for reactions up to a few hours.

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

Prokaryotic cell-free systems (E. coli-based) are faster to prepare, cheaper, and give higher yields for most simple proteins. The extract is easy to make in bulk and the system is well characterized. I’d use it to produce the MS2 L-protein, its natural context is E. coli, all the relevant chaperones are present in the E. coli extract, and I need high yield quickly for membrane insertion assays.

Eukaryotic systems are needed when your protein requires post-translational modifications like glycosylation, disulfide bond formation in the ER, or mammalian-specific folding chaperones. I’d use a mammalian cell-free system to produce human SOD1 it’s a cytosolic metalloenzyme that requires proper copper and zinc cofactor loading, and its folding energetics in the A4V mutant form are already perturbed, so having the right chaperone environment matters.

5. 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.

Membrane proteins are the hardest class to express in cell-free systems because they’re hydrophobic and aggregate instantly in aqueous solution without a membrane to insert into. The key is to provide a hydrophobic environment during synthesis.

I would design the experiment as follows: use an E. coli-based cell-free system supplemented with nanodiscs or liposomes added directly to the reaction so the protein co-translationally inserts into a lipid bilayer as it comes off the ribosome. For the L-protein specifically, I’d prepare nanodiscs made from POPC and MSP1D1 scaffold protein, add them at ~0.2 mg/mL to the cell-free reaction, and run the reaction to slow translation slightly and give the protein more time to fold beforethe next ribosome catches up.

The main challenges are: (1) aggregation before membrane insertion addressed by pre-adding nanodiscs before starting transcription; (2) low yield because hydrophobic proteins titrate out ribosomes, addressed by using a PURE system where you have more control over ribosome concentration; (3) confirming proper insertion addressed by running a protease protection assay where correctly inserted protein is shielded from externally added proteinase K.

6. 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.

Reason 1: The genetic template isn’t intact or there isn’t enough of it. The machinery can only build what it can read. If the DNA or RNA blueprint has degraded, or if there simply isn’t enough of it in the reaction, the output will be low no matter how healthy everything else is. To fix this, I’d first verify the quality and quantity of my template before adding it to the reaction. If the instructions are broken, no amount of tweaking elsewhere will help. I’d also protect the template from being destroyed mid-reaction by adding agents that block the enzymes responsible for degrading nucleic acids.

Reason 2: The energy or building blocks ran out. Protein synthesis is energy-hungry, and a cell-free reaction has a fixed starting supply. Once it is exhausted, the machinery stops, even if everything else is fine. Similarly, if the amino acid pool gets depleted partway through, the ribosomes stall. To troubleshoot this, I’d make sure the reaction includes an energy regeneration system so the fuel gets continuously recycled rather than just consumed, and I’d check that all twenty amino acids are present and well-supplied throughout the reaction.

Reason 3: The reaction environment isn’t right for this particular protein. The chemical conditions inside the tube things like salt balance and pH affect how well the machinery functions and whether the protein folds correctly after being made. A protein that misfolds immediately gets flagged and broken down, so even if translation is happening, the yield of intact product stays low. I’d troubleshoot this by running a small set of test reactions where I vary the buffer conditions slightly and see which environment gives the best result for my specific protein, rather than assuming the default conditions work for everything.

Homework Question from Kate Adamala

1. Function

a. What would your synthetic cell do? What is the input and what is the output?

My synthetic cell would act as a targeted antibiotic delivery vesicle for treating antibiotic-resistant bacterial infections. The input is a specific lipopolysaccharide (LPS) signature from a pathogenic gram-negative bacterium (e.g. K. pneumoniae). The output is localized release of a pore-forming peptide payload directly at the bacterial surface, lysing the pathogen without systemic antibiotic exposure.

b. Could this function be realized by cell-free Tx/Tl alone, without encapsulation?

No. Without encapsulation, there is no spatial specificity. The pore-forming peptide would be released everywhere and would be toxic to host cells as well. Encapsulation is what makes the delivery targeted: the synthetic cell only releases its payload when it docks onto a pathogen-specific surface signal.

c. Could this function be realized by a genetically modified natural cell?

Not easily. A living cell programmed to lyse bacteria would face serious immune clearance, regulatory hurdles, and the risk of horizontal gene transfer to other organisms. A synthetic minimal cell is non-replicating, non-living, and therefore much safer and more controllable.

d. Describe the desired outcome of your synthetic cell operation.

When the synthetic cell encounters a K. pneumoniae surface, a LPS-sensing aptamer on the membrane surface triggers expression of a pore-forming peptide (colistin mimetic) from the encapsulated Tx/Tl system. The peptide inserts into the bacterial membrane, causing lysis specifically at the site of infection, while host mammalian cells which lack LPS are untouched.

2. Component Design

a. What would the membrane be made of?

POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine) as the main structural lipid, supplemented with 30% cholesterol for membrane stability, and 5% DSPE-PEG2000 for steric stabilization and extended circulation time in biological fluids. The LPS-sensing aptamer would be conjugated to DSPE-PEG-maleimide on the outer leaflet.

b. What would you encapsulate inside?

  • Bacterial cell-free Tx/Tl system (E. coli S30 extract)
  • Linear DNA template encoding the pore-forming peptide under a T7 promoter with an aptazyme riboswitch responsive to LPS
  • ATP regeneration mix (phosphocreatine + creatine kinase)
  • All 20 amino acids at standard PURE system concentrations
  • Mg²⁺ optimized to 8 mM

c. Which organism will your Tx/Tl system come from?

Bacterial (E. coli S30 extract) — this is sufficient because the trigger is an aptazyme riboswitch, which works in bacterial Tx/Tl. No mammalian promoter system is needed since I’m not using Tet-ON or similar mammalian-specific inducible systems.

d. How will your synthetic cell communicate with the environment?

The LPS signal is detected by a surface-conjugated aptamer that, upon binding, triggers local membrane destabilization — releasing the Tx/Tl system contents or initiating fusion with the bacterial outer membrane. The pore-forming peptide produced inside the synthetic cell is hydrophobic enough to insert directly into the adjacent bacterial membrane upon release, without needing a dedicated membrane channel for export.

3. Experimental Details

a. List all lipids and genes:

Lipids:

  • POPC (main bilayer)
  • Cholesterol (30 mol%)
  • DSPE-PEG2000-maleimide (5 mol%, for aptamer conjugation)

Genes:

  • Pore-forming peptide gene: synthetic codon-optimized gene encoding Magainin-2 (a well-characterized antimicrobial peptide) under T7 promoter, with an LPS-responsive aptazyme (based on the OxyS aptazyme scaffold) in the 5’ UTR
  • T7 RNA polymerase gene: for transcription of the peptide gene inside the vesicle

Aptamer: LPS-binding aptamer sequence (Johnson et al., 2008, derived from SELEX against LPS from E. coli O111:B4) conjugated to DSPE-PEG-maleimide via thiol chemistry.

b. How will you measure the function of your system?

Primary readout: mix synthetic cells with K. pneumoniae in liquid culture and measure optical density at 600 nm over 6 hours a drop in OD600 indicates bacterial lysis. Secondary readout: add SYTOX Green (a membrane- impermeant DNA dye) to the co-culture. If bacteria are lysed, SYTOX enters and fluorescence increases, which can be quantified by plate reader or flow cytometry.

Homework Question from Peter Nguyen

Field chosen: Architecture

One-sentence pitch: A building facade material embedded with dormant slime mould networks and freeze-dried cell-free reporters that together map and visually display real-time moisture stress, structural load distribution, and ventilation dead zones across a building’s surface thereby turning the wall itself into a living diagnostic instrument.

How it works: Slime mould (Physarum polycephalum) is a remarkable organism that naturally grows its network along paths of least resistance, optimises for efficient transport between nodes, and retreats from dry or chemically hostile zones. These are exactly the same problems a building faces: where is moisture accumulating behind cladding? Where are thermal bridges concentrating stress? Where is air circulation failing?

The material would work in two layers. The first is a slime mould network layer a thin hydrogel matrix embedded in the interior face of a facade panel, seeded with dormant freeze-dried Physarum. When humidity inside the wall cavity rises above a threshold (indicating moisture ingress, condensation, or a failing vapour barrier), the slime mould rehydrates and begins growing. Because Physarum preferentially colonises humid corridors and avoids dry zones, its network topology after 24–48 hours of growth literally traces the moisture distribution map of that wall section — the densest growth appears where the problem is worst.

The second layer is a freeze-dried cell-free biosensor layer sitting just inside the visible surface of the panel. As the slime mould network grows, it releases metabolic byproducts specifically extracellular ATP and changes in local pH that diffuse into the cell-free layer. These chemical signals activate a riboswitch in the encapsulated Tx/Tl system, driving expression of a pigment or structural protein that causes a visible color shift on the panel’s surface. The wall literally marks its own problem zones in a colour visible from outside, without any wiring, sensors, or power supply.

When the moisture problem is resolved and the wall dries out, Physarum desiccates back into its dormant spore state, the cell-free reaction stops (no more trigger signal), and the panel resets, ready to respond again if the problem returns. Multiple panels across a facade create a distributed, self-reporting moisture map of the entire building skin.

Societal challenge addressed: Hidden moisture damage is one of the most expensive and dangerous failure modes in construction. It causes structural rot, mould growth, and insulation failure, and it is almost always detected too late because it is invisible until the damage is severe. Current monitoring requires either invasive physical inspection or expensive embedded electronic sensor networks that need power, maintenance, and replacement. A passive biological system that self-activates, self-maps, and self-resets would give architects and building managers a continuous, maintenance-free diagnostic layer in the fabric of the building itself is particularly valuable in social housing, schools, and infrastructure in lower-resource settings where sensor networks are not economically viable.

Addressing cell-free limitations: The one-time-use limitation is turned into a feature here. Each activation event corresponds to a real moisture event, and the system resetting when conditions improve means the panel is always ready for the next event rather than giving a permanent false positive. Stability is handled by the slime mould’s own biology by naturally encysting into desiccation-resistant sclerotia when dry, which can survive years without nutrients, and the freeze-dried cell-free layer sits dormant in the same conditions. Activation is not by externally added water but by the building’s own pathological moisture. The system only triggers when there is a genuine problem, not from rain on the outer surface or ambient humidity fluctuations. The spatial resolution of the diagnostic comes for free from Physarum’s network growth dynamics.

Homework Question from Ally Huang

Using BioBits® Cell-Free Protein Expression System

1. Background

Astronauts on long-duration missions experience significant immune dysregulation, including reduced lymphocyte function and increased susceptibility to latent viral reactivation. In space, standard laboratory-based immune monitoring is completely out of reach. Early detection of immune stress markers is critical for crew health, especially on future Mars missions where communication delays make real-time Earth-based medical support impossible. A lightweight, freeze-dried diagnostic system that can be activated on demand would directly address this gap.

2. Molecular Target

Interleukin-6 (IL-6) mRNA — an early biomarker of systemic immune activation, inflammation, and viral reactivation in astronauts.

3. How the target relates to the challenge

IL-6 spikes within hours of infection or physiological stress and has been documented at elevated levels in astronaut blood samples linked to latent herpesvirus reactivation during ISS missions. Detecting IL-6 mRNA using a cell-free toehold switch biosensor gives real-time immune status information without cold-chain reagents, trained personnel, or centrifuges.

4. Hypothesis

I hypothesize that a freeze-dried BioBits cell-free expression system programmed with an IL-6 mRNA-responsive toehold switch will reliably detect elevated IL-6 transcript levels aboard the ISS, producing a fluorescent output measurable by the P51 Molecular Fluorescence Viewer. The toehold switch keeps the ribosome binding site sequestered in a hairpin until the target IL-6 mRNA binds and unfolds it, triggering translation of sfGFP. A visible fluorescence signal indicates immune activation. The system will be validated against known IL-6 concentration standards before flight.

5. Experimental Plan

Freeze-dried BioBits pellets will be rehydrated with a small whole blood lysate sample from crew members at pre-flight, mid-mission, and post-flight timepoints. The miniPCR thermal cycler will maintain isothermal incubation conditions, and fluorescence will be read on the P51 viewer. Controls include a synthetic IL-6 mRNA positive control and a buffer-only negative control. Fluorescence presence or absence relative to a set threshold identifies immune activation events across mission timepoints.