Week 9 HW: Cell-Free Systems

Part A: General Homework Questions

Q1. 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.

The biggest appeal of cell-free systems is that you’re not fighting the cell anymore. In a living cell, you’re constantly competing with its own agenda it wants to grow, divide, manage stress. None of that is helpful when you just want to make a protein. In a cell-free system, you crack the cells open, take the machinery you actually need, and run the reaction completely on your own terms. You can tweak pH, redox state, ion concentrations mid-experiment, and add things that would kill a living cell outright.

Two situations where this really shines: first, expressing toxic proteins like pore-forming toxins or viral proteins that would kill a host cell before you ever got a decent yield. Second, incorporating non-natural amino acids — you just add the engineered tRNA and synthetase directly to the reaction without competing cellular pathways getting in the way.

Q2. Describe the main components of a cell-free expression system and explain the role of each component.

There are four main things you need. The cell extract is the heart of it basically the inside of a cell without the membrane, containing ribosomes, RNA polymerase, translation factors, and chaperones. Then you need a DNA template — the gene you want expressed, usually on a plasmid or linear PCR product with a promoter the system recognizes (T7 is standard for E. coli extracts). You also need an energy regeneration system because transcription and translation burn through ATP incredibly fast without replenishing it, your reaction dies in minutes. Finally, a reaction buffer supplies amino acids, magnesium, potassium, and any cofactors your specific protein needs.

Q3. 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.

Making protein is energetically expensive every peptide bond costs roughly 4 ATP equivalents, and that adds up fast. Without replenishment, the reaction stalls within 10–20 minutes, ribosomes stop, and you end up with truncated, useless fragments.

The classic fix is the phosphocreatine/creatine kinase (PCK) system. Creatine kinase continuously converts ADP back to ATP using phosphocreatine as the phosphate donor. It’s simple and well-characterized. Another solid option is phosphoenolpyruvate (PEP) with pyruvate kinase, which works on the same principle. For longer reactions, glucose-based systems tapping into glycolytic enzymes in the extract can sustain output for hours, though you have to watch out for acidification from acetate accumulation.

Q4. Compare prokaryotic versus eukaryotic cell-free expression systems. Choose a protein to produce in each system and explain why.

Prokaryotic systems, usually E. coli-based, are fast, cheap, high-yield, and easy to work with. The downside is they can’t do complex post-translational modifications like glycosylation. Eukaryotic systems like wheat germ, rabbit reticulocyte, or HeLa-based are slower and pricier, but they support glycosylation, complex disulfide bonds, and have proper chaperones for folding tricky proteins.

For a prokaryotic system, I’d produce T7 RNA polymerase it’s a bacterial protein, needs no glycosylation, and you want as much of it as possible as cheaply as possible. For a eukaryotic system, I’d go with erythropoietin (EPO) it’s heavily N-glycosylated, and that glycosylation is what determines its activity and serum half-life in vivo, so you really need a eukaryotic system to get a functional product.

Q5. 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 notoriously annoying because their hydrophobic transmembrane domains aggregate the moment they hit aqueous solution. In a cell-free system though, you have more tools than you would in vivo.

The main strategy is to supply a lipid environment directly in the reaction so the protein can fold as it’s being made. The cleanest way to do this is with nanodiscs ,a small, monodisperse lipid bilayer discs held together by membrane scaffold proteins. They’re soluble, well-defined, and the newly synthesized protein can insert cotranslationally. You can also add detergents at sub-CMC concentrations to stabilize hydrophobic regions without disrupting the ribosomes. On top of that, supplementing with chaperones like Skp or SurA helps prevent aggregation. Finally, you’d need to carefully titrate Mg²⁺ and K⁺ concentrations since membrane protein translation is particularly sensitive to ionic conditions. To check if it worked, you’d measure yield by a GFP fusion and verify function with a ligand-binding or transport assay.

Q6. 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 — Template degradation: If you’re using a linear PCR product, exonucleases in the extract will chew it up fast. Fix: switch to a circular plasmid, or add GamS protein (a RecBCD inhibitor from lambda phage) to protect linear DNA.

Reason 2 — Energy depletion: Your ATP regeneration system might not be keeping up, especially for longer reactions. The reaction acidifies as pyruvate accumulates, which tanks ribosome activity. Fix: titrate your phosphocreatine and creatine kinase concentrations, monitor pH during the reaction, and consider switching to a more sustained energy system like maltose/maltose-binding protein.

Reason 3 — Codon bias: Your gene might contain rare codons that deplete specific tRNA pools in the extract, causing ribosomes to stall mid-translation. Fix: codon-optimize your gene for the expression host, or directly supplement the reaction with total tRNA from the appropriate organism.

Homework Question from Kate Adamala

Design an example of a useful synthetic minimal cell.

What does it do, and what are the inputs/outputs? I’d design a synthetic cell that detects miRNA-21 , a microRNA that’s significantly overexpressed in many cancer types — and responds by releasing a fluorescent signal or a small therapeutic payload. The input is miRNA-21 diffusing into the synthetic cell from the surrounding environment. The output is release of an encapsulated cargo (fluorescent dye or drug) triggered by miRNA-21-driven gene expression inside the cell.

Could this work without encapsulation? No. The whole point is spatial separation between sensing and response. If you just had a cell-free reaction floating around without a membrane, the cargo would diffuse everywhere regardless of whether miRNA-21 was present. The encapsulation is what makes the release conditional on the input signal.

Could a GMO do this? Technically yes , you could engineer a cell with a miRNA-21-responsive circuit. But living cells are harder to control, replicate uncontrollably, and carry significant regulatory and safety concerns for therapeutic use. A synthetic cell is non-replicating, can’t transfer genes horizontally, and is much safer to deploy near human tissue.

Desired outcome: In the presence of tumor-associated miRNA-21, the synthetic cell detects the signal and releases its payload specifically at the tumor site , a clean, autonomous, targeted response.

Membrane: DOPC + cholesterol (roughly 7:3 ratio) , a stable, well-characterized lipid vesicle around 100–200 nm.

Encapsulated contents: PURE system (bacterial cell-free Tx/Tl), fluorescent cargo (calcein or FITC-dextran), and a DNA construct encoding alpha-hemolysin (aHL) under the control of a synthetic toehold switch riboswitch responsive to miRNA-21.

Which Tx/Tl system? Bacterial PURE system is fine here. The toehold switch is a synthetic RNA element that functions in a bacterial translation context, so there’s no need for mammalian machinery.

How does it communicate with the environment? miRNA-21 is small enough to cross or leak through the lipid bilayer passively. Once inside, it triggers the toehold switch, derepressing translation of aHL. The aHL protein inserts into the membrane as a pore, and the encapsulated cargo exits through it.

Lipids: DOPC, cholesterol (7:3)

Genes: hla (alpha-hemolysin from S. aureus) under a miRNA-21-responsive toehold switch (designed using the Green et al. 2014 framework)

How to measure it: A calcein dequenching assay works well — calcein is self-quenching at high concentrations inside the vesicle, and fluorescence increases sharply when pores form and it dilutes into the surrounding solution. In a more applied setting, you’d co-culture with MCF-7 breast cancer cells (high miRNA-21) versus MCF-10A normal cells (low miRNA-21) and image with confocal microscopy.

Homework Question from Peter Nguyen

Field: Textiles/Fashion

One-sentence pitch: A smart wound dressing that autonomously detects bacterial infection and produces an antimicrobial peptide on-site using freeze-dried cell-free reactions reactivated by wound fluid.

How it works: The fabric is impregnated with freeze-dried CF reactions containing a DNA construct encoding an antimicrobial peptide (like nisin or a defensin) under the control of a promoter responsive to bacterial quorum-sensing molecules specifically 3-oxo-C6-HSL, which E. coli and many other pathogens secrete as their population grows. When a wound gets infected, bacteria start producing these signaling molecules. They diffuse into the dressing, the wound exudate provides the water needed to rehydrate the CF system, and the reaction kicks off — producing the antimicrobial peptide directly within the fabric, right where it’s needed. No nurse intervention, no systemic antibiotics.

Societal challenge: Antibiotic-resistant wound infections, particularly MRSA and Pseudomonas aeruginosa are a major cause of mortality in hospital and battlefield settings. Current dressings either release antibiotics constantly (driving resistance) or require a clinical decision to escalate treatment. An autonomous-response dressing could reduce resistance pressure and be hugely valuable in low-resource or remote settings where clinical oversight isn’t always available.

Addressing CF limitations: For activation with water, wound exudate naturally provides the moisture, this is actually a feature, since the reaction only turns on when there’s active wound fluid, which correlates with infection. For stability, freeze-drying with trehalose and PVA stabilizers can preserve CF activity for over a year at room temperature, and the textile can be sealed with a moisture barrier during storage. For the one-time use limitation, wound dressings are already single-use medical devices, so this isn’t really a drawback here. One reaction window of 6–16 hours aligns perfectly with standard dressing-change intervals.

Homework Question from Ally Huang

Background: Long-duration spaceflight exposes astronauts to ionizing radiation from cosmic rays and solar particle events at rates far exceeding anything on Earth. This radiation causes DNA double-strand breaks and oxidative damage that accumulate over months-long missions. Right now, radiation damage in astronauts is mostly assessed after the mission, meaning crews have no real-time health data while they’re actually at risk. For missions to Mars, this becomes a serious problem. Astronauts could be accumulating dangerous levels of genomic damage with no way of knowing. A compact, resource-minimal diagnostic that works in real time aboard a spacecraft would be a genuine step forward for crew safety.

Molecular target: γH2AX — histone H2AX phosphorylated at serine 139, a direct, quantitative biomarker of DNA double-strand breaks in peripheral blood leukocytes.

How the target relates to the challenge: Every time ionizing radiation creates a double-strand break in DNA, H2AX gets phosphorylated at serine 139 within minutes, forming γH2AX foci that recruit repair machinery. The number of foci per cell is directly proportional to the number of breaks, which makes it one of the most sensitive and well-validated radiation damage markers we have. Measuring γH2AX in blood leukocytes requires only a small blood draw, minimal processing, and gives you a real-time snapshot of cumulative radiation dose — making it ideal for a spaceflight-compatible biosensor.

Hypothesis: We hypothesize that a freeze-dried BioBits cell-free expression system can be engineered as a quantitative biosensor for γH2AX, enabling real-time radiation dose monitoring aboard spacecraft. We’ll design a CF reaction where a synthetic antibody fragment (nanobody) targeting the phospho-S139 epitope of H2AX is fused to one half of a split-sfGFP reporter. When a blood lysate from an irradiated astronaut is added to the rehydrated BioBits reaction, γH2AX binds the nanobody fragment, bringing the split-GFP halves together and reconstituting fluorescence. Signal intensity will scale with γH2AX concentration, giving a quantitative readout of radiation dose. The entire workflow uses only the P51 fluorescence viewer already in the Genes in Space toolkit, requires no refrigeration, and produces results in under 4 hours.

Experimental plan: Samples will be lysates from TK6 human lymphoblastoid cells irradiated at 0, 0.5, 1, 2, and 4 Gy. Controls include unirradiated lysate (negative control), a recombinant γH2AX protein spike (positive control), and non-phosphorylated H2AX protein (specificity control). Each lysate is added to a rehydrated BioBits reaction, incubated for 4 hours at 29°C, then imaged with the P51 fluorescence viewer. We’ll measure fluorescence intensity per well and build a dose-response curve across three biological replicates to validate sensitivity and specificity.