Week 9 HW: Cell-Free Systems

Homework Part A: General and Lecturer-Specific Questions

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 provides multiple advantages over traditional methods due to its lack of membrane. You don’t have to cross the membrane, and the cell’s homeostasis controls don’t resist any experimentations. Furthermore, since everything is now a controllable variable, you have better control over composition (such as concentrations) of amino acids, substrates, etc. You also maintain better folding control and template flexibility (since you don’t have to clone into plasmids). Cases where this is beneficial include space travel (since cell-free synthesis allows for freeze-drying, which is more shelf-stable in storage), and anywhere else resources may be limited to run a fermenter (war zones, rural clinics, etc.) Another case is the expression of “unhostable” proteins, which would in other cases, kill its living cell. The cell-free environment means you bypass the actual cell host.

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

A cell-free system is essentially a cell with no membrane, and no homeostasis-related regulation. Components include:

DNA Template (you still need parts the system can read such as promoters, RBS, CDS, terminator)
RNA polymerase (for synthesizing mRNA in the cell-free system)
Ribosomes (for translation, added back individually and generally most expensive component of cell-free systems)
tRNAs (for reading the codons and delivering the amino acids)
Amino acids (all canonical ones supplied so proteins are synthesized)
NTPs (required monomers for transcription)
Energy regeneration system (translation burns a lot of ATP, some sort of system required to keep system running)
Salts and cofactors that are critical for ribosomes and polymerase activity
Cell extract (lysed E.Coli containing most of the listed above that end up in the cell-free system)
Buffer (“background” holding everything together)

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.

Translation is very energy “expensive”, requiring multiple high-energy phosphate bonds for one peptide bond. Without some sort of energy regeneration, the cell-free system would function very slowly and inefficiently (as ATP → ADP + Pi, eventually most of the system is ADP instead of what the cell needs). A way you could keep ATP flowing is a high-energy phosphate donor + its kinase, where a phosphate is transferred back onto an ADP to make ATP again. This phosphate can be donated from other molecules, (called a high-energy phosphate donor). Common donors include phosphoenolpyruvate, creatine phosphate, and glucose.

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

Prokaryotic cells are generally the fastest and cheapest (generally in synthetic biology), allowing for greater efficiency and affordability. Furthermore, because prokaryotic cells are generally simpler, it’s easier to prototype and use. However, that simplicity means no post-translational modifications, and certain proteins cannot form properly due to lack of disulfide bonds. Also, if expressing a eukaryotic gene, it lacks basic infrastructure needed.

Protein example: Luc2 (used in my final project), a cytosolic enzyme from Photinus pyralis that folds well in E. coli. If you add D-luciferin and ATP to it, you get bioluminescence proportional to its translation rate. This would be ideal to prototype in a tube before agroinfiltration (rehydrate a BioBits-style pellet with a linear PCR product of 35S+Ω → luc2+SKL → NOS, add luciferin, light appears)

Eukaryotic cell systems are generally more expensive, and often output lower yield with slower reactions, but do offer more variations and resources. For plants, a popular choice is wheat germ extract (WGE), as it handles eukaryotic protein folding well due to its eukaryotic translation factors and plant-derived folding machinery.

Protein example: The expression of a novel candidate (such as the one I’m using in my second translational unit), in a wheat germ extract. Since I am expressing cytochrome P450s, I know they are difficult to express in prokaryotic cells. Therefore, it is better to express in a eukaryotic system, despite the higher costs.

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.

Since CFPS have no membrane, you’d design a system environment specifically suited for that specific type of membrane protein. Challenges include the obvious lack of membrane (therefore, nowhere for the protein to insert), translation issues (since many membrane proteins translate as they are being inserted into the membrane), orientation of the membranes (since membrane proteins have a defined position in the membrane), and having the correct lipids and confactors. I think the most practical solution that should theoretically solve all these problems is a fake membrane added in the system. These “fake”, lipid-rich membranes would not surround anything, but float freely allowing for a location for the membrane protein to insert into. They would be added into the reaction while the membrane proteins are being translated.

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.

One: Not enough energy (ATP)/Energy is running out too early

If translation is burning ATP faster than it’s being created, even with an energy regenerative system, reactions would plateau or slow down. To troubleshoot you’d adjust concentrations of PEP, and/or add sodium oxalate to suppress the phosphatases that are destroying the PEPs. You can also upgrade to a continuous-exchange CFPS (CECF), where you have two compartments separated by a dialysis membrane, so that there is continuous exchange between the “fuel” energy supply and waste supply.

Two: Low magnesium ion concentrations

If the concentration of Mg²⁺ is too low or high, ribosomes and polymerase don’t function correctly. Troubleshooting involves running a Mg²⁺ titration to to find the optimal concentration.

Three: Bad lysate batch

A problem with CFPS is that freeze-dried batches are not consistent. Extracts can vary even with the same protocol, and ribosome activity is sensitive to freeze/thaw cycles. Troubleshooting would involve never re-freezing, strict aliquoting, or a new batch.

How could you keep energy flowing for continuous ATP supply in a cell-free system?

Prokaryotic vs eukaryotic cell free expression systems?

How can you add more energy to cell-free systems if your energy regenerative system is still sluggish?

What factors are cell-free protein synthesis systems sensitive to?

Homework question from Kate Adamala

Design an example of a useful synthetic minimal cell as follows: What would your synthetic cell do? What is the input and what is the output?

Detect luciferin precursors and produce visible light (photo emission) in response (for example, in a plant if it detects all the metabolic precursors needed, it lights up). This is useful because we don’t know all the precursors for firefly bioluminescence. This is a helpful debugging tool in case my novel candidates are correct, but don’t glow due to folding errors, expression errors, etc. The input would be p-coumaric acid (at least for the example of my cytochrome P450s candidate), and output would be visible light at 560nm.

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

Yes but with losses from the lack of encapsulation. The chemistry wouldn’t suffer much, but the application (deploying the genes into living plant tissue) could be problematic as you need encapsulation to deploy. Also, encapsulation protects the luc2 genes from nucleases, which may reduce light output.

Could this function be realized by genetically modified natural cell?

You could express luc2 in E.coli or directly in the plant, but synthetic clinical cells may be more efficient at converting higher light output, since you wouldn’t be losing ATP for cell housekeeping and resources due to other pathways. In theory, this would give a clearer, defined system where the only chemistry happening is luciferin-related.

Describe the desired outcome of your synthetic cell operation.

When the minimal cell encounters a p-coumaric acid or other precursor, light output rises depending on the concentration detected. This would validate the enzyme I am testing, to see if it actually generates luciferin precursors before committing to an agroinfiltration pipeline.

Design all components that would need to be part of your synthetic cell. What would be the membrane made of?

Phosphatidylcholine (POPC) to mimic the cell membrane and cholesterol to stiffen the bilayer enough to hold a pore stably without leakage.

What would you encapsulate inside? Enzymes, small molecules.

I would encapsulate a bacterial cell-free system (E.Coli lysate), plasmid encoding luc2, plasmid encoding α-hemolysin, plasmid encoding my novel candidate (PPYR_02911), ATP starting pool, PEP for energy regeneration, and Mg²⁺.

Which organism your Tx/Tl system will come from? Is bacterial OK, or do you need a mammalian system for some reason? (hint: for example, if you want to use small molecule modulated promotors, like Tet-ON, you need mammalian)

It would come from bacterial organisms (E.Coli) lysate since the luc2 (firefly luciferase) I’m using from iGEM folds well into E.Coli and does not need eukaryotic post-translational modifications. I do not need a mammalian system since I’m not using any mammalian parts.

How will your synthetic cell communicate with the environment? (hint: are substrates permeable? or do you need to express the membrane channel?)

P-coumaric acid (what I’m testing for) diffuses into the membrane through α-hemolysin pores (which are also narrow enough to retain luc2 and lysate machinery). Light would exit passively since photons are not affected by lipid bilayers. Therefore, no export channel is needed for output (since it’s photons), allowing for a simpler design.

Experimental details. List all lipids and genes. (bonus: find the specific genes; for example, instead of just saying “small molecule membrane channel” pick the actual gene.)

Lipids: POPC, cholesterol (at a 70:30 molar ratio)

Genes:

Luc2 (BBa_K389004, firefly luciferase)
Hla (α-hemolysin, membrane pore)
PPYR_02911 (candidate CYP4C cytochrome)
How will you measure the function of your system?
The function of my system will be measured by a luminometer (measured over time after adding higher concentrations of p-coumaric - to a suspension synthetic minimal cell. Another measurement can be captured with a camera to image the cells as bright spots.

Ai I am designing a useful synthetic minimal cell for my firefly plant project that detects luciferin precursors and produces visible light in response (to test my novel candidates. Lets use PPYR_02911). What should the membrane be made of? Should we use bacterial since we’re using luc2?

Why a 70:30 ratio for POPC and cholesterol?

Homework question from Peter Nguyen

Write a one-sentence summary pitch sentence describing your concept.

Freeze-dried cell-free coating embedded in clothing fabric that produces bioluminescence when activated by sweat, transforming clubwear, entertainment, and performance garments into living light at clubs/concerts/nightlife.

How will the idea work, in more detail? Write 3-4 sentences or more.

The fabric is coated (you can shade them onto the fabric with each use) with the freeze-dried cell-free reaction pellets containing the luciferase expression system discussed earlier. These are all lyophilized into a stable powder bound into the textile amtrak with water-soluble polymers. When the wearer sweats (or is misted with water), the pellets are rehydrated, triggering the luciferase-driven light emission. The pattern follows the pattern of moisture, where the back may glow lighter, following the dancer’s movements. The garment’s lifecycle is a single night out, growing brightest when the person wearing the garment is most active, and dimming with the night winding down.

What societal challenge or market need will this address?

Clubwear is already saturated with glowing garments, but with expensive LED-embedded garments that are difficult to wash, are heavy, require batteries, and fail often. A bioluminescent textile is light, soft, batteryless, washable, and biodegradable, being kinder for the environment. This taps into broader movements against single-use plastics/batteries, and biofabricated materials, where the garment is part of the larger story of how it’s made (“my garment is alive and reacts to me”). It could also be used with runners for safety wear (being visible in the dark), and other sport events where you need to track athletes.

How do you envision addressing the limitation of cell-free reactions (e.g., activation with water, stability, one-time use)?

This idea has many limitations, but all can be addressed:

Activation (If the wearer does not sweat, water can be applied externally with sprays or other means. Water is provided everywhere in nightlife due to safety.)
Shelf life (The garment could be on the shelf for many months or years before first activation. This is well within the shelf-stability of lyophilized cell-free systems)
One-time use (Design the garments with refillable pockets for freeze-fried pellets cartridges where the wearer refills them between uses.

Homework question from Ally Huang

Provide background information that describes the space biology question or challenge you propose to address. Explain why this topic is significant for humanity, relevant for space exploration, and scientifically interesting.

Long-duration spaceflights of space colonies require the crew to grow their own food. However, plants often show stress under microgravity, with impaired growth, weakened cell walls, and other unpredictable responses. Although we have successfully grown certain vegetables in space, our monitoring of plant stress is still heavily reliant on freezing samples, and sending them back to earth. I propose a field-deployable sensor that reports plant stress directly on orbit, allowing for early intervention, and protecting food security on long missions.

Name the molecular or genetic target that you propose to study. Examples of molecular targets include individual genes and proteins, DNA and RNA sequences, or broader -omics approaches.

I propose to study the Arabidopsis jasmonic acid responsive transcripts (Jaz1), which is an early mRNA marker of plant dress and wounds. This would be detected via biosensor.

Describe how your molecular or genetic target relates to the space biology question or challenge your proposal addresses.

Jasmonic acid signaling is one of the fastest plant stress responses, being activated within minutes of abiotic stress or wounds. JAZ1 transcripts spike sharply during this response making it an almost real-time readout of plant stress. Detecting JAZ1 through a tiny leaf punch (adjusted for the leaf punch response) would tell astronauts right away if the plant is healthy, or if the stress response is currently active. This could also explore if microgravity inherently affects this pathway in plants.

Clearly state your hypothesis or research goal and explain the reasoning behind it.

Hypothesis: Arabidopsis plants that are grown in microgravity exhibit chronically elevated JAZ1 transcripts, reflecting the chronic low-grade stress plants inhibit in response to space flight. This can be detected using freeze-fried BioBits switch biosensors.

Reasoning: Previous studies of spaceflight grown plants have implied chronic stress and cell-wall remodeling. However, these samples are freeze-dried and return to earth, where samples may be getting affected by storage and re-entry. A direct measurement during orbit of JAZ1 in fresh tissue can inform us how much is spaceflight stress, and how much is sampling-related. The number of JAZ1 is proportional to transcript abundance. Therefore, it can also allow astronauts real-time, same-day data on the stress of their crops throughout the course of their mission.

Outline your experimental plan - identify the sample(s) you will test in your experiment, including any necessary controls, the type of data or measurements that will be collected, etc.

The JAZ1 targeting switch plasmid would be designed on earth, and then freeze fried into BioBits pellets. During the orbit, astronauts collect small leaf punches from the Arabidopsis, which scientists on earth collect in parallel with ground-control plants. The total RNA is then extracted using a simple column kit, and then rehydrated with BioBits reactions. Fluorescence is measured over a few hours and imaged with an iPad. Controls include: 1) BioBits + water 2) BioBits + synthetic JAZ1 RNA 3) BioBits + mismatched trigger RNA 4) wounded ground control leaves

I am designing a proposal that incorporates BioBits cell-free protein expression system. My idea is a biosensor that detects stress signals in plants in space. What is a reliable gene I can target with my biosensor?

How does JAZ1 work?

Homework Part B: Individual Final Project

Put your chosen final project slide in the appropriate slide deck following the instructions on slide 1 (DONE)

Submit this Final Project selection form if you have not already.(DONE)

Prepare your first DNA order and put it in the “Twist (MIT)” or “Twist (Nodes)” tab of the 2026 HTGAA Ordering: DNA, Reagents, Consumables spreadsheet, as appropriate.