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 (CFPS) skips the cell membrane barrier, so the user has direct access to the reaction mixture. This means parameters like pH, redox state, magnesium and potassium concentrations, temperature, and amino acid pools can all be tuned freely without worrying about cell viability. There is also no need for cloning, transformation, or growing cultures, so the time from DNA template to protein is hours instead of days.

Two cases where CFPS is clearly better than in vivo production:

  • Toxic proteins: Antimicrobial peptides or cytotoxic enzymes would kill an E. coli host, but in a cell-free reaction there is no host to harm.
  • Non-canonical amino acids: Incorporating unnatural amino acids or isotopic labels is straightforward because the amino acid pool can be depleted and replaced directly in the reaction tube.
2. Describe the main components of a cell-free expression system and explain the role of each component.
  • Cell extract (lysate): Crude cytoplasm from E. coli, wheat germ, rabbit reticulocyte, or CHO cells. Provides ribosomes, tRNAs, aminoacyl-tRNA synthetases, and translation factors.
  • DNA template: Plasmid or linear PCR product encoding the target gene under a strong promoter (commonly T7). Carries the genetic information for the protein.
  • Energy mix and substrates: NTPs, dNTPs, amino acids, and salts (Mg²⁺, K⁺) that fuel transcription and translation.
  • Energy regeneration system: A secondary high-energy phosphate donor (e.g., phosphoenolpyruvate, creatine phosphate) plus its kinase, which keeps ATP and GTP replenished throughout the reaction.
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.

Protein synthesis consumes multiple high-energy phosphate bonds per peptide bond formed. Without a living metabolism in the tube, free ATP runs out within minutes and inorganic phosphate accumulates, inhibiting further translation. Regeneration is what determines how long the reaction lasts and how much protein it produces.

A common method is the PEP / pyruvate kinase system: phosphoenolpyruvate acts as a high-energy phosphate donor, and pyruvate kinase transfers that phosphate onto ADP to regenerate ATP. Creatine phosphate with creatine kinase works the same way. For longer reactions, a continuous-exchange setup (CECF) with a semipermeable membrane can also be used to feed in fresh substrates and remove inhibitory byproducts.

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

Prokaryotic systems (typically E. coli-based) are fast, cheap, and high-yield, often reaching mg/mL concentrations within a few hours. They lack post-translational modifications such as glycosylation and proper disulfide bond formation. Eukaryotic systems (wheat germ, rabbit reticulocyte, HeLa, CHO) are slower and produce less protein, but they carry the chaperones, ER/microsomal membranes, and modification enzymes needed for complex folding.

  • Prokaryotic choice — GFP: GFP folds correctly on its own and needs no glycosylation. E. coli lysate produces large amounts cheaply, which is ideal for reporters and screening.
  • Eukaryotic choice — Erythropoietin (EPO): EPO requires sialic acid glycosylation to be biologically active. A mammalian lysate (CHO or HEK293) provides the modification machinery prokaryotes lack.
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.

The main challenge is hydrophobicity. Membrane proteins have transmembrane domains that misfold and aggregate when translated in aqueous extract without a lipid environment.

The setup would add a hydrophobic phase to the reaction so the protein has somewhere to insert as it leaves the ribosome:

  • Nanodiscs: Pre-formed lipid bilayer discs stabilized by membrane scaffold proteins. The nascent protein inserts co-translationally and ends up in a native-like bilayer.
  • Detergent micelles: Mild detergents like DDM, Brij-35, or Triton X-100 can solubilize the protein during synthesis. Cheaper than nanodiscs but may interfere with downstream assays.
  • Liposomes or SUVs: Synthetic vesicles can also serve as insertion targets.

Additional optimizations: tune Mg²⁺ concentration, add chaperones (DnaK/DnaJ/GrpE), and use a slower expression strain or lower temperature to avoid aggregation. Activity can be confirmed with a functional assay (binding, transport, or ligand response) rather than just SDS-PAGE.

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.
  1. Energy depletion: ATP/GTP runs out before translation finishes. Fix: Add a stronger regeneration system (creatine phosphate + creatine kinase) or switch to a continuous-exchange format.
  2. Template degradation by nucleases or protein degradation by proteases: Endogenous RNase E, Lon, or OmpT can chew up mRNA or the product. Fix: Add RNase inhibitors, use a circular plasmid instead of linear template, or switch to extracts from knockout strains (Δlon, ΔompT).
  3. Inefficient codon usage: Rare codons stall ribosomes. Fix: Codon-optimize the gene for the extract organism, or supplement with rare-codon tRNAs.

Homework Question from Kate Adamala

Design a synthetic minimal cell: a pesticide-sensing SMC that signals to reporter bacteria

1. Function and system logic

a. What does the synthetic cell do? Input and output? The SMC detects the agricultural pollutant atrazine in the environment and converts that signal into release of IPTG, which then activates GFP expression in nearby reporter E. coli. Input: atrazine (diffuses across the membrane). Output: IPTG released into the surroundings.

b. Could this work as cell-free TX/TL alone, without encapsulation? No. Without a membrane, the IPTG would immediately reach the reporter bacteria regardless of atrazine, so there would be no conditional sensing. Encapsulation is what makes the sensor gating possible.

c. Could this work in a genetically modified natural cell? In principle yes — an atrazine-responsive riboswitch could be engineered into live bacteria. But using an SMC avoids genetic drift, biocontainment issues, and host fitness costs. The same SMC architecture can also be re-targeted to other small molecules by swapping the riboswitch, which is harder to do in living cells.

d. Desired outcome: In the presence of atrazine, the SMC synthesizes a membrane pore, IPTG escapes, and reporter E. coli glow green. Without atrazine, no pore forms and the bacteria stay dark.

2. Component design

a. Membrane: POPC (palmitoyl-oleoyl-phosphatidylcholine) with cholesterol to control fluidity and reduce leakage.

b. Encapsulated inside: E. coli TX/TL extract, a pool of free IPTG, and a plasmid encoding α-hemolysin (aHL) under control of an atrazine-binding RNA riboswitch.

c. TX/TL origin: Bacterial (E. coli), because the riboswitch is bacterial and integrates cleanly with prokaryotic translation.

d. Communication with the environment: Atrazine is small and lipophilic enough to cross the bilayer passively. IPTG is membrane-impermeable and stays inside until aHL is expressed and forms a pore.

3. Experimental details

a. Lipids and genes:

  • Lipids: POPC, cholesterol.
  • Gene: α-hemolysin (aHL) downstream of an engineered atrazine aptamer/riboswitch.
  • Reporter cells: E. coli transformed with GFP under a T7 promoter with a lac operator.

b. Measurement: Co-incubate SMCs with reporter E. coli and titrate atrazine across a concentration range. Read bulk GFP fluorescence over time on a plate reader (or use flow cytometry for single-cell distributions). A no-atrazine control and a free-IPTG positive control define the dynamic range.

Homework Question from Peter Nguyen

Application field: Textiles / Fashion

a. One-sentence pitch

A field-worker jacket woven with freeze-dried cell-free biosensor patches that change color when exposed to airborne organophosphate pesticides.

b. How it works (more detail)

Disposable paper-matrix patches embedded into the jacket’s sleeves and shoulders contain freeze-dried E. coli extract, a constitutive expression circuit, and a butyrylcholinesterase-based reporter. When sweat or ambient humidity rehydrates the patch, the cell-free reaction starts producing a colored enzymatic output as a baseline. Organophosphate pesticides in the air inhibit the cholinesterase, stopping the reaction and triggering a visible color change on the fabric. The worker sees the warning directly on their sleeve without any electronics.

c. Societal challenge / market need

Agricultural laborers are exposed to pesticide overspray daily, and acute and chronic organophosphate poisoning are major causes of occupational illness, especially in low-income farming regions. Existing electronic monitors are expensive and rare in the field. A passive textile sensor is cheap, requires no power, and gives immediate visual feedback.

d. Addressing cell-free limitations
  • Activation by water: The patch stays dry-stable and is activated by sweat or humidity during use. To prevent unwanted activation, patches can sit behind a thin water-permeable membrane.
  • Stability: Freeze-drying preserves the lysate for months at ambient temperature; patches are sealed in foil until first use.
  • One-time use: Patches are modular and clipped/stitched into the jacket so they can be unclipped and swapped after a positive event or after the day’s shift, without replacing the whole garment.

Homework Question from Ally Huang

Mock Genes in Space proposal — Monitoring radiation-induced DNA damage in astronauts using BioBits®

1. Background

Astronauts on long-duration missions face chronic cosmic radiation that causes DNA double-strand breaks and oxidative stress, accumulating cancer and tissue-damage risk over time. Current diagnostic tools for tracking this damage rely on bulky lab equipment that is impractical aboard a spacecraft. A lightweight, power-efficient, room-temperature-stable biological assay would let crews monitor their own radiation exposure in real time and test the effectiveness of shielding materials. Freeze-dried cell-free systems are an ideal fit because they require no live cells, no cold storage, and minimal hardware — exactly the constraints of deep-space missions.

2. Molecular target

The human p53 transcript and protein, a master regulator of the DNA damage response that is upregulated in proportion to genomic stress.

3. Relation of the target to the challenge

p53 is the central node of the DNA damage response: when double-strand breaks occur, ATM/ATR signaling stabilizes and activates p53, which then drives transcription of repair and apoptosis genes. The level of p53 expression therefore tracks the intensity of recent radiation exposure. By using p53-responsive reporters in a freeze-dried cell-free system, we can quantify radiation-induced damage in real time aboard the ISS without complex equipment, and compare exposures across different shielding configurations or mission phases.

4. Hypothesis / research goal

Hypothesis: A freeze-dried BioBits® cell-free reaction containing a p53-responsive fluorescent aptamer reporter can be rehydrated aboard the ISS and used to quantify DNA damage in astronaut-derived samples, producing fluorescence proportional to radiation dose.

The reasoning rests on three points. First, cell-free reactions remain stable in freeze-dried form for long periods and do not need live cell culture, which fits the operational constraints of a spaceflight environment. Second, p53 activation is a well-validated, dose-dependent biomarker of DNA damage, with extensive ground-based calibration data already available. Third, fluorescent aptamer reporters provide a directly visible readout that can be quantified with the P51 Molecular Fluorescence Viewer, requiring no benchtop fluorimeter. If validated, this platform becomes a general template for in-flight molecular diagnostics.

5. Experimental plan

Astronauts collect small blood or cheek-swab samples at scheduled intervals. Each sample is mixed with rehydrated BioBits® extract carrying a p53-responsive fluorescent aptamer plasmid. Reactions are incubated in the miniPCR® thermal cycler at 37 °C, and fluorescence is read with the P51 viewer. Controls include: (i) a ground-prepared positive control with known radiation-damaged DNA, (ii) a shielded ground sample to establish baseline, and (iii) a no-template negative control. Data collected: fluorescence intensity over time and across sampling sessions, correlated with onboard dosimeter readings to validate the assay against physical radiation measurements.

References

  • Lentini, R. et al. (2014). Integrating artificial with natural cells to translate chemical messages that direct E. coli behaviour. Nature Communications, 5, 4012.
  • Martini, L. & Mansy, S. S. (2011). Cell-like systems with riboswitch controlled gene expression. Chemical Communications, 47(38), 10734.
  • Pardee, K. et al. (2016). Portable, on-demand biomolecular manufacturing. Cell, 167(1), 248–259.
  • Stark, J. C. et al. (2018). BioBits™ Bright: A fluorescent synthetic biology education kit. Science Advances, 4(8), eaat5107.