Week 9 HW: hw-cell-free-systems

Week 9 — 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.

Benefits in flexibility and control

Cell‑free systems are “open‑top” – you can add whatever you want: salts, energy sources, DNA templates, detergents, even toxic things that cells can’t handle. No worries about killing cells.

The reaction is fast – you get results in a few hours, not one or two days like fermentation. Want to test conditions? Set up 10 different pH or magnesium concentrations in the morning, and you’ll have the answer by the afternoon.

Transcription and translation can be decoupled – separately add T7 polymerase, inhibitors, modify the mRNA structure… go wild.

You can sample anytime without disrupting cells.

Two cases where cell‑free works better than living cells

• Making toxic or membrane proteins – Some proteins (like pore‑forming toxins) kill the cells as soon as they’re expressed, so the culture never grows. In a cell‑free system, there are no live cells – just add detergents or lipids, and the protein stays dissolved. No problem.
• Incorporating unnatural amino acids – In live cells, getting them to take up and incorporate unnatural amino acids is a huge pain: low efficiency, often toxic. In a cell‑free system, you just dump the unnatural amino acid into the reaction tube. Want to attach a fluorescent label or a crosslinker? Super easy.

Main components of a cell‑free expression system and their roles

Cell lysate – That’s the “soup” you get after breaking open cells (like E. coli or wheat germ). It contains ribosomes, tRNAs, various enzymes, and translation factors. This is the core machine.

Energy regeneration system – Simply adding ATP isn’t enough; it gets used up quickly. So you need a system that continuously converts ADP back into ATP. The most common one is creatine phosphate + creatine kinase.

Amino acids – The building blocks. The 20 standard amino acids. If you want isotopic labelling or unnatural amino acids, just swap them.

Nucleotides (NTPs) – For transcription: ATP, GTP, CTP, UTP.

DNA template – The gene you want to express, with a promoter in front (e.g., T7).

Salts and buffer – Maintain pH and ionic strength. Magnesium ions are especially important – ribosomes can’t work without them.

RNA polymerase – If the lysate doesn’t have enough, add extra T7 RNA polymerase.

Optional additives – Chaperones, detergents, DTT (to prevent oxidation), PEG (to mimic the crowded intracellular environment).

Why is energy regeneration so important? How do you ensure a continuous ATP supply?

The reason is simple: protein translation is a huge consumer of ATP and GTP. The small amount of ATP you add at the beginning turns into AMP and phosphate within minutes. Without energy regeneration, the reaction quickly stops, and the yield is pitiful. So you need a mechanism to convert ADP back into ATP continuously.

Common method: the creatine phosphate / creatine kinase system.

You add creatine phosphate (around 10–50 mM) and creatine kinase (1–2 U/μL) to the reaction tube. Creatine kinase transfers a high‑energy phosphate group from creatine phosphate to ADP, regenerating ATP. This method is very stable, has few side effects, and is used in most cell‑free experiments.

Prokaryotic vs. eukaryotic cell‑free systems – pick one protein for each and explain why

Prokaryotic system: I would make GFP (green fluorescent protein). This guy is simple, needs no modifications, folds beautifully in E. coli lysate, gives high yield, and you can see it with your naked eye. It’s super convenient for testing and tuning system parameters.

Eukaryotic system: I would make the human EGFR kinase domain (the intracellular part of the epidermal growth factor receptor). This protein needs to form disulfide bonds correctly and tends to aggregate. In E. coli it comes out mostly inactive. The wheat germ system has eukaryotic chaperones that help it fold slowly, producing functional protein for enzyme activity assays.

How to design a cell‑free experiment to optimise membrane protein expression? Challenges and how to tackle them

The pitfalls of membrane proteins:

• Hydrophobic transmembrane domains aggregate in water, forming precipitates.
• Hard to fold correctly and insert into a lipid bilayer.
• Yield is usually very low.
• Adding detergents or lipids might inhibit the reaction.

My experimental design ：

• Use E. coli lysate – cheap and high‑yielding. Add some extra chaperones (e.g., GroEL/GroES) to help with folding.
• Try different membrane environments:
  • Detergents (DDM, LDAO, etc.) – start with low concentrations (0.01‑0.1%); don’t go too high, or the reaction will be inhibited.
  • Liposomes – prepare small liposomes in advance (e.g., from E. coli total lipids or synthetic lipids) and add them during the reaction so the protein inserts as it is being synthesised.
  • Nanodiscs – use membrane scaffold protein (MSP) plus lipids to make nanodiscs that mimic a real membrane environment.
• Use a continuous exchange cell‑free (CECF) system – a semipermeable membrane separates the reaction mixture from a feeding buffer, which continuously supplies energy and amino acids while removing inhibitors (e.g., phosphate). Especially useful for membrane proteins.
• Add a fusion tag – for example, attach GFP or MBP to the N‑terminus. This helps folding and allows real‑time monitoring of expression levels.
• Lower the temperature – for example, 20‑25 °C. Membrane proteins usually prefer cooler temperatures and are less prone to aggregation.
• Add stabilisers – glycerol (5‑10%), trehalose, or even the ligand/substrate that the protein binds to, to stabilise the native conformation.

Detection: Use the GFP fusion to follow fluorescence. At the end, separate the membrane fraction from the soluble fraction by ultracentrifugation, then run a gel (stain or Western blot).

Low yield in a cell‑free experiment: three possible reasons and how to fix them

Reason 1: Problem with the DNA template
For example, weak promoter, a very stable secondary structure at the 5′ end, or too much DNA that inhibits the reaction.
How to fix it: Switch to a strong T7 promoter, check the RBS sequence (for prokaryotic systems), remove hairpins at the 5′ end. Run a DNA concentration gradient (typically 5‑50 μg/mL) to find the optimal concentration. Also, resequence the template.

Reason 2: Energy runs out; the regeneration system isn’t working
Maybe the creatine phosphate has degraded, or there is a high ATPase activity in the reaction that consumes ATP too quickly.
How to fix it: Open a fresh vial of creatine phosphate, increase the amount of creatine kinase. Or switch to a different energy system (e.g., glucose + glycolysis). If that doesn’t work, use an ATP detection kit (luciferase‑based) to monitor ATP levels by taking samples during the reaction.

Reason 3: Accumulation of inhibitors
ATP hydrolysis produces a lot of phosphate, which chelates magnesium ions and shuts down the reaction. Also, aggregated protein precipitates can cause trouble.
How to fix it: Switch to a continuous exchange cell‑free (CECF) system to continuously remove small‑molecule inhibitors. Alternatively, add a phosphatase inhibitor, or use a phosphate binder (e.g., phosphatase substrate – but that’s a bit tricky). A simpler approach: dilute the reaction or exchange the buffer.

Homework question from Peter Nguyen

Design an example of a useful synthetic minimal cell as follows:

Pick a function and describe it

🤴Activatable bacterial sensor for inflammatory cytokine detection. The synthetic minimal cell (SMC) serves as a signal transduction capsule – it detects the presence of tumor necrosis factor alpha (TNF-α), a key pro-inflammatory cytokine elevated in conditions such as sepsis, rheumatoid arthritis, and inflammatory bowel disease, and then converts that protein signal into a small-molecule output that can be read by a standard reporter bacterium. This effectively “translates” a protein biomarker that bacteria cannot naturally sense into a format that a simple engineered bacterial reporter can process.

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

In short: TNF‑α (input) → SMC → IPTG (output) → E. coli reporter → purple colour (readout).

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

🐱‍🐉No. The synthetic cell membrane provides two essential functions that free Tx/Tl mixture cannot.

First, the membrane serves as a physical barrier that holds the IPTG inside until the α‑hemolysin pores are formed. If the Tx/Tl system were simply mixed in a test tube without encapsulation, IPTG would be free in solution from the start and would reach the reporter bacteria immediately — even without TNF‑α. Under those conditions, the bacterial reporter would always turn purple regardless of whether the cytokine was present. Thus, the SMC acts as an AND logic gate: output is only released when both (1) IPTG is present in the interior AND (2) TNF‑α triggers pore formation.

Second, the membrane allows the SMC to sense large macromolecular inputs (TNF‑α, ~17 kDa) using encapsulated cell‑free machinery. In an open mix, TNF‑α could potentially interfere directly with the reporter — but encapsulation compartmentalises the sensing event, preventing crosstalk.

Could this function be realized by genetically modified natural cell?

🐉Yes, but with significant limitations.

It is theoretically possible to engineer a living bacterium to detect TNF‑α by:

Expressing a mammalian TNF‑α receptor on its surface, and

Coupling receptor binding to an intracellular genetic circuit that produces a colourimetric output.

However, this approach faces several practical challenges that the SMC avoids:

Describe the desired outcome of your synthetic cell operation

In the presence of TNF-α above a clinically relevant threshold (e.g., >50 pM, within the human serum range for inflammatory conditions), the synthetic minimal cell should:

Sense TNF-α via the riboswitch embedded in the α‑hemolysin mRNA.

Translate and assemble α‑hemolysin pores in its own membrane within approximately 1–2 hours.

Release encapsulated IPTG diffusively through these pores into the surrounding medium.

The released IPTG should, upon contact with the E. coli reporter strain, induce sufficient LacZ expression to produce a detectable colour change (yellow to purple) using a substrate like CPRG or X‑gal.

The desired specificity is that no colour change occurs in the absence of TNF-α, even if other cytokines (e.g., IL‑6, IL‑1β) or unrelated proteins are present. In other words, the SMC should function as a specific, activatable signal transducer for TNF‑α.

Design all components that would need to be part of your synthetic cell.

Organism for Tx/Tl system

Bacterial (E. coli) is appropriate because:

• Riboswitches naturally function in prokaryotic systems and are well‑characterised in E. coli cell‑free extracts.
• There is no need for eukaryotic post‑translational modifications – α‑hemolysin is a bacterial toxin that folds and assembles correctly in an E. coli lysate environment.
• The output (IPTG) is specifically designed to activate an E. coli‑based reporter; using a prokaryotic Tx/Tl system keeps all components biologically compatible and minimises cross‑reaction risks.

How will your synthetic cell communicate with the environment?

Input (TNF‑α) is not permeable through the intact lipid bilayer (a 17 kDa protein cannot cross). However, the SMC does not need to import TNF‑α. The riboswitch is encoded on the DNA template inside the SMC, and the cell‑free Tx/Tl system produces the α‑hemolysin mRNA that contains the riboswitch in its 5′ UTR. For TNF‑α to trigger the riboswitch, TNF‑α must first cross the membrane into the interior – but large proteins are unable to do so without pores. How is this resolved? The input TNF‑α is actually added along with the Tx/Tl mix at the time of SMC assembly. The entire Tx/Tl system, including the riboswitch‑containing DNA, the energy mix, and TNF‑α itself, is co‑encapsulated during SMC formation. In other words, TNF‑α is present inside the SMC from the start, not imported later. The design takes advantage of the fact that riboswitches are co‑translational regulatory elements – the aptamer is part of the nascent mRNA, and TNF‑α binding occurs as the transcript is being produced.

Output (IPTG) is initially encapsulated. Upon TNF‑α‑triggered translation and assembly of α‑hemolysin pores, IPTG diffuses out through these channels. Pores are sufficiently large (≈1.4 nm diameter) to allow passage of small molecules like IPTG (MW ≈ 238 Da).

Experimental details

Lipids

• POPC (1‑palmitoyl‑2‑oleoyl‑sn‑glycero‑3‑phosphocholine) – the primary structural phospholipid.
• Cholesterol – stabilises the bilayer, reduces passive permeability, and enhances mechanical robustness.

Genes

• α‑hemolysin (αHL, hla) gene – from Staphylococcus aureus (293 amino acids; secretes as a water‑soluble monomer that assembles into a heptameric β‑barrel pore upon contact with lipid membranes). A TNF‑α‑responsive RNA aptamer is engineered into the 5′ UTR directly upstream of the αHL RBS to create a riboswitch that activates translation only upon TNF‑α binding [6†L8-L16].

• TNF‑α aptamer sequence – selected via SELEX or RNA‑compete against human TNF‑α. The aptamer is inserted into the 5′ UTR of the αHL gene, positioned such that ligand‑binding induces a structural rearrangement that exposes the Shine‑Dalgarno sequence and start codon, enabling translation initiation.

• Reporter gene (in E. coli reporter strain) – lacZ (encoding β‑galactosidase) under a T7‑lacO promoter (e.g., pET‑derived vector). The reporter strain also constitutively expresses T7 RNA polymerase (e.g., E. coli BL21(DE3)). When IPTG enters the cell, it binds LacI, causing dissociation from the lacO operator and allowing T7 RNA polymerase to transcribe lacZ.

Encapsulation method

Liposomes are formed by hydration of a dried lipid film (POPC:cholesterol) in a buffer containing:

• cell‑free Tx/Tl mixture
• IPTG (~2 mM)
• riboswitch‑αHL DNA template (10–50 nM)
• NTPs, amino acids, energy regeneration components
• TNF‑α (variable concentrations, from 0 to 100 nM)
• any other necessary cofactors

The lipid mixture is then subjected to multiple freeze‑thaw cycles and extrusion through polycarbonate membranes (e.g., 400 nm pore size) to produce unilamellar vesicles with encapsulated contents. Unencapsulated material is removed by gel filtration or centrifugal washing. Alternatively, water‑in‑oil emulsions can be used to generate monodisperse SMCs.

External reporter preparation

An E. coli BL21(DE3) strain is transformed with a plasmid carrying lacZ under a T7‑lacO promoter (e.g., pET‑lacZ). The strain is grown to early exponential phase (OD₆₀₀ ≈ 0.4), washed, and resuspended in fresh medium containing the colourimetric substrate (e.g., CPRG at 0.5 mg/mL).

Assay setup

Purified SMCs are mixed with the reporter E. coli suspension and incubated at 30 °C for 2–4 hours. Colour development is monitored spectrophotometrically (absorbance at 570 nm for CPRG cleavage product) or by direct visual inspection.

How will you measure the function of your system?

Cross‑reference to the Lentini et al. architecture

The following table compares the original system by Lentini et al. (2014) with the present design:

The Lentini paper inspired the architecture of the present design – a riboswitch‑controlled pore‑forming toxin inside an artificial cell acts as an “actuator” that gates the release of a small‑molecule output. However, this design swaps the input from a synthetic molecule (theophylline) to a clinically relevant human protein biomarker (TNF‑α) and changes the readout from fluorescence to a naked‑eye colourimetric signal, making it more suitable for point‑of‑care diagnostic applications.

Platform modularity and future directions

This synthetic minimal cell provides a proof‑of‑concept for a modular, reagent‑free cytokine detection platform. By swapping the aptamer in the riboswitch, the same SMC architecture can be reconfigured to detect virtually any protein biomarker for which a specific RNA aptamer can be selected – from other cytokines (IL‑6, IL‑1β, IFN‑γ) to viral proteins (e.g., SARS‑CoV‑2 spike protein) or cancer biomarkers (e.g., PSA, CA‑125).

Homework question from Peter Nguyen

💖Freeze-dried cell-free systems can be incorporated into all kinds of materials as biological sensors or as inducible enzymes to modify the material itself or the surrounding environment. Choose one application field — Architecture, Textiles/Fashion, or Robotics — and propose an application using cell-free systems that are functionally integrated into the material. Answer each of these key questions for your proposal pitch:

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

👩‍🦰We turn building walls into living, low‑cost air quality monitors by embedding freeze‑dried cell‑free sensors that detect indoor pollutants and produce a visible colour change upon rehydration.

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

🙌The freeze‑dried cell‑free system is incorporated directly into water‑based paint or plaster. It contains a toxin‑responsive riboswitch or transcription factor controlling the expression of a chromoprotein (e.g., a purple pigment) or an enzyme that produces a visible dye. When indoor humidity or a small water leak activates the system, it becomes functional. If a target pollutant (e.g., formaldehyde, benzene, or carbon monoxide) diffuses into the material, it binds to a sensor protein, triggering transcription‑translation of the reporter. Within a few hours, a clear colour patch appears on the wall, alerting occupants. Multiple sensors can be patterned as stripes or QR codes for semi‑quantitative detection.

What societal challenge or market need will this address?

✨Poor indoor air quality (IAQ) causes “sick building syndrome”, asthma, and long‑term health issues, yet conventional electronic monitors are expensive, require power, and provide no spatial resolution. This technology offers a passive, disposable, and ultra‑low‑cost sensor that works without batteries or maintenance. It is especially useful for schools, hospitals, and low‑income housing where continuous monitoring is needed but budgets are tight.

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

👩‍🦳Activation with water – The freeze‑dried system is stored in a desiccated state. We co‑encapsulate trehalose to protect during drying and storage. Activation occurs when ambient humidity exceeds a threshold (e.g., >60 % RH) or by a small water reservoir integrated behind the coating. For applications with very dry air, a micro‑encapsulated water bead can be crushed by hand or by a simple pH‑triggered release.

Stability – Freeze‑dried cell‑free extracts remain active for months at room temperature when protected from oxygen and moisture. We package the reactive powder in a bilayer: a permeable outer membrane for gas exchange and an inner water‑soluble layer that dissolves upon activation, exposing the reaction mix to the environment.

One‑time use – This is actually an advantage for disposable sensors: the wall patch gives a permanent colour change, serving as a “record” of past contamination. For reusability, we can design a biphasic system where the reporter is deposited on a removable test strip that can be swapped out, while the enzyme‑producing layer remains.

By combining these strategies, the system becomes a practical “canary on the wall” that requires no electronics, no skilled operation, and no external power – only a small amount of water or humidity to wake it up.

BioBits in Space: Detecting DNA damage caused by space radiation using a cell‑free repair assay

Background

Space radiation causes DNA double‑strand breaks (DSBs), which increase cancer risk and threaten crew health. Current dosimeters measure physical dose but not biological effect. A simple, cell‑free assay that reports on DSB frequency would provide direct biological impact data, enabling better risk assessment for long missions.

Genetic target

Linear DNA template encoding a split‑GFP – two halves of GFP are expressed separately only after ligation of a radiation‑induced DSB.

👩‍🦳When radiation breaks DNA, the linear template is fragmented. By measuring reconstituted GFP fluorescence after a cell‑free repair reaction, we quantify DSB frequency. This links physical radiation dose directly to a functional molecular outcome – DNA integrity – without living cells.

Hypothesis: Freeze‑dried cell‑free extracts containing DNA repair enzymes (e.g., E. coli ligase and polymerase) can ligate radiation‑fragmented DNA back into a full‑length template, restoring split‑GFP expression. Fluorescence intensity will correlate with radiation dose.

Rationale: Current space radiation monitoring relies on passive detectors that must be returned to Earth for analysis. We propose a real‑time, in‑flight assay: a BioBits pellet containing repair machinery and split‑GFP DNA. After in‑orbit radiation exposure, adding water activates the repair reaction; intact templates produce GFP. This transforms the pellet into a direct biological dosimeter that can be read within 2 hours using the P51 fluorescence viewer.

Experimental plan

Test three groups: flight (exposed to space radiation), ground (identical but on Earth), and shielded (wrapped in 2 mm Al, on‑orbit). Each group: 5 BioBits pellets. Post‑flight, add water + T4 DNA ligase buffer to all pellets simultaneously, incubate at 37 °C for 2 h. Measure GFP fluorescence (470 nm excitation, 520 nm emission) using P51. Controls: intact plasmid DNA (positive), no DNA (negative).