Week 9 HW: Cell free systems

General homework 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 is more flexible than in vivo expression because we can directly control the reaction conditions, such as DNA concentration, salts, cofactors, temperature, and additives. The in vivo model limits out experimet by time since we have atcually grow cells and wait for results, in cell free systems the speed of these procedures is much faster. It is more beneficial to use cell free systems for toxic proteins, membrane proteins, and rapid prototyping or diagnostics, because we do not need to keep a living cell alive while producing the protein.

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

There are several components of a cell-free expression systems:

Cell extract:

This is the liquid fraction from broken cells that supplies the machinery needed to make proteins.

DNA template:

This is the gene blueprint that tells the system which protein to produce.

Amino acids:

These are the raw materials used to build the protein chain.

NTPs:

These molecules are used to make RNA and also help power the reaction.

Energy source:

This keeps the system active by replacing the energy used during protein synthesis.

Cofactors:

These keep the reaction environment suitable so the enzymes can work efficiently.

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 the cell free system does not provide naturally energy for wokring conditions. Transcription and translation consume ATP and GTP very quickly, so the reaction stops if energy runs out. A common method is to add an energy regeneration system such as phosphoenolpyruvate or creatine phosphate so ATP can be continuously recycled during the reaction.

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

Prokaryotic and eukaryotic cell-free systems are better for different kinds of proteins. A prokaryotic system, usually based on E. coli extract, is fast, inexpensive, and very efficient for making proteins that do not require complicated folding or post-translational modifications. Because of that, I would choose a simple bacterial protein, such as GFP or a bacterial enzyme, to produce in this system. These proteins are usually easier to express because they fold well in bacterial conditions and do not depend on glycosylation or other eukaryotic modifications.

A eukaryotic cell-free system, such as one based on wheat germ, insect, or mammalian extract, is better for proteins that are more complex and need additional folding help or cellular processing. I would choose a human membrane receptor or a secreted human protein for this system, because these proteins often need more than just translation to become functional. Eukaryotic systems are more suitable when the target protein needs correct folding, disulfide bond formation, or a more native-like environment to stay stable and active.

The main difference between the two systems is the source of the extract and also the type of protein they are best able to produce. Prokaryotic systems are usually preferred when speed, low cost, and high yield are most important. Eukaryotic systems are preferred when protein quality, folding, and biological realism are more important than maximum yield.

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.

To optimize the expression of a membrane protein in a cell-free system, I would choose a system that better supports membrane insertion and protein folding, such as a eukaryotic extract or an extract supplemented with membrane-mimicking components. Membrane proteins are difficult to express because their hydrophobic regions tend to aggregate in aqueous solution, so I would include liposomes, nanodiscs, or microsomal vesicles to provide a more natural lipid environment for folding and insertion.

I would also design the construct with a small fluorescent tag, such as GFP, so I can monitor whether the protein is being produced successfully and whether the expression level changes under different conditions. This would allow me to compare different reaction setups, such as varying temperature, magnesium concentration, extract type, and DNA template amount, to find the best conditions for expression. If the protein is especially sensitive, I would also test slower reaction temperatures because lower temperatures can sometimes improve folding and reduce aggregation.

Another important part of the design would be energy management. Since membrane protein synthesis can take a long time, I would use an energy regeneration system, such as creatine phosphate and creatine kinase, or a continuous exchange setup to keep ATP levels stable during the reaction. This would help extend the reaction and increase the chance of obtaining a properly folded product.

The main challenge is that membrane proteins are not only hard to synthesize, but also hard to keep soluble and functional after synthesis. To address this, I would compare several conditions side by side, including reactions with and without membrane mimics, and measure both yield and activity. In the end, the best setup would be the one that gives the highest amount of correctly folded membrane protein, not just the highest total protein amount.

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.

Some possible low yield target protein cases:

Poor DNA template quality or low translation efficiency:

One possible reason for low protein yield is that the DNA template is not clean, damaged, or not optimized for expression in the cell-free system. This can be improved by purifying the plasmid or PCR product more carefully, and in some cases by using codon optimization or adding an RNA inhibitor to reduce template degradation and improve expression.

Insufficient energy supply:

Another reason is that the reaction may run out of ATP and GTP too quickly, so the protein synthesis machinery cannot keep working. To fix this, you can improve the ATP regeneration system by adding a stronger energy source or a better recycling strategy so the reaction stays active for longer.

Unfavorable reaction conditions:

A third reason is that the magnesium level, salt concentration, temperature, or extract quality may not be ideal for the protein you are trying to make. You can troubleshoot this by testing several reaction conditions one by one, such as different temperatures or ion concentrations, until you find the setup that gives the best yield.

Homework question from Kate Adamala

Pick a function and describe it. What would your synthetic cell do? What is the input and what is the output?

The synthetic minimal cell (SMC) I want to design would detect TDP-43 protein aggregates (the pathological hallmark of Frontotemporal Dementia (FTD)) and respond by producing and releasing a therapeutic anti-aggregation peptide. TDP-43 is an RNA-binding protein that under disease conditions mislocalizes from the nucleus to the cytoplasm, where it forms toxic aggregates that drive neurodegeneration.

  • Input: Extracellular TDP-43 aggregates in the cerebrospinal fluid or interstitial brain environment
  • Output: A designed TDP-43 aggregation-inhibiting peptide released into the local environment to disrupt fibril formation and reduce proteotoxic stress

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

No. Without encapsulation, the cell-free Tx/Tl system would constitutively produce the anti-aggregation peptide regardless of whether TDP-43 aggregates are present. Encapsulation is essential to couple detection (input sensing via an aptamer) to production (output peptide expression), creating a conditional, signal-responsive system rather than a constitutive one.

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

Yes, in theory. Natural cell could be engineered with a TDP-43-responsive promoter driving anti-aggregation peptide expression. However, introducing living genetically modified cells into the brain raises several biosafety, immune rejection, and ethical concerns. A synthetic minimal cell offers a safer, non-replicating, and fully controllable alternative that degrades naturally once its payload is delivered.

Describe the desired outcome of your synthetic cell operation.

In the presence of TDP-43 aggregates, the SMC detects them via a surface-anchored RNA aptamer, triggers internal gene expression of the anti-aggregation peptide, and releases it through a membrane pore into the surrounding tissue. The result is localized, on-demand therapeutic peptide delivery specifically where and when TDP-43 aggregation is occurring, slowing the progression of FTD neurodegeneration.

Design All Components

What would the membrane be made of?

Phospholipids (POPC + POPE) and cholesterol to mimic a stable bilayer. Some biotinylated lipids would be incorporated to anchor the TDP-43-sensing aptamer on the outer membrane surface via streptavidin linkage.

What would you encapsulate inside?

  • Bacterial cell-free Tx/Tl system (PURE system)
  • DNA construct: gene encoding the anti-aggregation peptide under a T7 promoter coupled to a TDP-43 aptamer-responsive riboswitch
  • Gene encoding α-hemolysin (aHL) pore-forming protein, also under aptamer control, to enable peptide release upon activation
  • Small molecule cofactors for Tx/Tl (ATP, amino acids, NTPs)

Which organism will your Tx/Tl system come from?

Bacterial (PURE system from E. coli) is sufficient here, as the riboswitch used for TDP-43 detection is compatible with bacterial transcription/translation machinery. A mammalian system is not required since no mammalian-specific promoters (e.g., Tet-ON) are needed.

How will your synthetic cell communicate with the environment?

The outer membrane surface displays a TDP-43 RNA aptamer. When TDP-43 aggregates bind this aptamer, a conformational signal is transduced intracellularly, activating the riboswitch and initiating Tx/Tl of both the anti-aggregation peptide and α-hemolysin. The expressed aHL inserts into the membrane and forms a pore through which the therapeutic peptide is released into the extracellular environment.

Experimental Details

Lipids and genes:

  • Lipids: POPC, POPE, cholesterol, biotinylated-DPPE (for aptamer anchoring)
  • Genes:
    • ahlA (α-hemolysin from Staphylococcus aureus) — membrane pore for peptide release
    • Synthetic gene encoding TDP-43 aggregation-inhibiting peptide (e.g., based on the YQ-rich domain inhibitor design) — under T7 promoter + TDP-43 aptamer riboswitch
  • Aptamer: Anti-TDP-43 aggregate RNA aptamer anchored to outer membrane via streptavidin-biotin linkage

How will you measure the function of your system?

  1. Incubate SMCs with recombinant TDP-43 aggregates in vitro and measure peptide release via ELISA or fluorescently tagged peptide fluorescence
  2. Use a ThT (Thioflavin T) aggregation assay to confirm that released peptide reduces TDP-43 fibril formation compared to controls without SMC
  3. Confirm pore formation by aHL via dye leakage assay (encapsulate fluorescent dye, measure release upon TDP-43 addition)

Homework question from Peter Nguyen

One-sentence pitch: A living wall coating embedded with freeze-dried cell-free biosensors that detects black mold (Stachybotrys chartarum) VOC emissions and produces a visible color change before mold becomes visible to the naked eye.

How will the idea work?

Black mold (Stachybotrys chartarum) releases characteristic volatile organic compounds (VOCs) during early colonization — most notably 1-octen-3-ol — before any visible growth appears. The proposed system embeds freeze-dried cell-free Tx/Tl reactions into a breathable polymer wall coating (e.g., a porous silicone or hydrogel matrix). When ambient humidity reactivates the freeze-dried system and 1-octen-3-ol diffuses into the coating, it binds to an engineered transcription factor (based on a modified OBP — odorant binding protein) that activates a chromoprotein reporter gene. The wall visibly changes color (e.g., from clear to deep violet using the chromoprotein amilCP) in the region of mold colonization, providing a spatially precise early warning signal. No electronics, power, or human monitoring are required — the building itself becomes the sensor.

What societal challenge or market need does this address?

Black mold is a major public health hazard linked to respiratory illness, neurological symptoms, and immune disorders, particularly in children and immunocompromised individuals. Current detection relies on visible inspection or expensive air quality testing, by which point mold is already well-established and remediation is costly. An estimated 50% of buildings in developed countries have some form of problematic moisture/mold. An early, passive, low-cost detection system embedded directly into building materials would allow intervention before health impacts occur, reducing both healthcare costs and remediation expenses.

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

  • Activation with water: The freeze-dried system is formulated in a hygroscopic hydrogel matrix that reactivates only when local humidity exceeds the threshold typical of mold-favorable conditions (>70% RH), creating a built-in environmental trigger
  • Stability: Freeze-drying with trehalose as a cryoprotectant extends shelf life to 1–2 years at room temperature; the wall coating can be replaced as a panel every 2 years as part of standard building maintenance
  • One-time use: This is reframed as a feature — once the color change occurs, it serves as a permanent record of mold detection in that location, and the panel is replaced. Multiple overlapping panels can be layered to provide repeated sensing capacity over the building lifetime

Homework question from Ally Huang

Background (≤100 words)

Long-duration spaceflight profoundly suppresses astronaut immune function. A well-documented consequence is the reactivation of latent herpesviruses — including Epstein-Barr virus (EBV) and Varicella-Zoster virus (VZV) — which remain dormant in healthy individuals but reactivate under the immune dysregulation caused by microgravity, radiation, and psychological stress. Herpesvirus reactivation has been detected in over 50% of astronauts on ISS missions and poses risks ranging from mild illness to serious neurological complications on long-duration missions to the Moon or Mars, where return to Earth is not possible.

Molecular/Genetic Target (≤30 words)

EBV immediate-early gene BZLF1 (also called Zta/ZEBRA) — its expression is the molecular switch that triggers EBV reactivation from latency and is detectable in saliva.

How does the target relate to the challenge? (≤100 words)

BZLF1 mRNA expression is the earliest detectable signal of EBV reactivation — preceding viral shedding and any clinical symptoms by days. Detecting BZLF1 transcripts in astronaut saliva samples using a cell-free toehold switch biosensor would provide real-time, equipment-minimal immune status monitoring. Since EBV reactivation is directly driven by the cortisol-mediated immune suppression characteristic of spaceflight stress, BZLF1 acts as a functional readout of overall immune dysregulation, not just viral status — making it a highly informative single-target proxy for astronaut immune health.

Hypothesis/Research Goal (≤150 words)

Hypothesis: BZLF1 mRNA will be detectable in astronaut saliva samples collected during ISS missions using a freeze-dried BioBits® toehold switch biosensor, and its expression will correlate with mission duration and radiation exposure levels, providing a quantitative real-time index of immune suppression severity.

Rationale: Toehold switches are synthetic RNA regulators that activate cell-free reporter gene expression only when a specific target RNA sequence is present. By designing a toehold switch complementary to the BZLF1 mRNA sequence and freeze-drying it with a cell-free GFP reporter into BioBits® pellets, we can create a single-use, room-temperature-stable diagnostic that requires only saliva addition and the P51 fluorescence viewer to produce a quantitative readout. This requires no specialized laboratory equipment, making it fully compatible with ISS constraints and scalable to future deep space missions.

Experimental Plan (≤100 words)

Samples: Astronaut saliva collected at mission days 0, 30, 60, 90, and 180. Controls: BZLF1-positive saliva (EBV-reactivating donor, ground), BZLF1-negative saliva (healthy seronegative donor, ground), and a no-template BioBits® pellet.

Procedure: Add 2 µL saliva to rehydrated BioBits® toehold switch pellet. Incubate 37°C for 2 hours. Read GFP fluorescence using the P51 Molecular Fluorescence Viewer.

Data collected: GFP intensity (proxy for BZLF1 mRNA concentration), correlated with mission day, cumulative radiation dose (from personal dosimeters), and cortisol levels (parallel measurement). Statistical correlation analysis performed on ground post-mission.