Week 11 HW: Cell-Free Protein Synthesis

HTGAA 2026 — Week 11 Homework

Cell-Free Protein Synthesis & Collaborative BioArt

What I Liked About the Project

The experiment beautifully demonstrated that biological creativity and collective computation can be bridged through a structured, yet open-ended collaborative format. What stood out most was the emergent complexity: each individual’s single-pixel decision was locally simple, but globally the artwork encoded recognizable biological imagery. This mirrors how distributed cellular systems encode complex phenotypes from individual gene expression events. The fact that the “canvas” was limited to 1,536 pixels (a deliberate constraint) also made each contribution weighty and meaningful — a great lesson in resource allocation at biological scale.

What Could Be Made Better

Future iterations could benefit from a live fluorescence visualization layer — rather than a static pixel grid, students could watch their pixel “express” as a simulated fluorescence signal over time, giving a tangible connection between the bioart assignment and the cell-free expression lab that follows. Additionally, providing each participant with a brief protocol preview of how fluorescent protein expression maps to pixel intensity would reinforce the cross-lab connection and motivate deeper engagement.

Part B: Cell-Free Protein Synthesis — Cell-Free Reagents

B1. Component Roles in the Cell-Free Reaction

E. coli Lysate — BL21 (DE3) Star Lysate (includes T7 RNA Polymerase)

The lysate is the core “cellular machinery” of the reaction: a clarified extract of disrupted E. coli BL21(DE3) cells that retains the ribosomes, translation factors, aminoacyl-tRNA synthetases, chaperones, and metabolic enzymes required for coupled transcription and translation. BL21(DE3) cells are specifically used because they harbor a chromosomally integrated T7 RNA polymerase gene under an IPTG-inducible promoter; the T7 RNAP is induced prior to lysis, so the extract natively drives transcription from T7-promoter-containing DNA templates without the need for exogenous polymerase supplementation (Pardee et al., 2016; Sun et al., 2013). The “Star” designation refers to a recA-minus, RNase E mutant background that improves RNA stability and protein yield in cell-free reactions.

Reference: Sun, Z. Z. et al. (2013). Protocols for Implementing an Escherichia coli Based TX-TL Cell-Free Expression System for Synthetic Biology. JoVE, 79, e50762.

Salts / Buffer

Potassium Glutamate

Potassium glutamate serves two functions simultaneously: it provides potassium ions (K⁺) at physiologically relevant concentrations, which are critical for ribosome function and RNA polymerase activity, and it provides the glutamate counter-ion, which can act as a secondary carbon/energy source via oxidative metabolism in the cell extract. Mimicking the intracellular ionic environment of E. coli (where K⁺ is the dominant cation) is essential for maximal translation efficiency (Jewett & Swartz, 2004; Cai et al., 2015).

Reference: Jewett, M. C. & Swartz, J. R. (2004). Substrate replenishment extends protein synthesis with an in vitro translation system designed to mimic the cytoplasm. Biotechnol. Bioeng., 87(4), 465–472.

HEPES-KOH pH 7.5

HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid) is a zwitterionic buffering agent that maintains the reaction pH near 7.5, mimicking the cytoplasmic pH of E. coli. Maintaining stable pH is critical because acidification (common when glucose or glycolytic intermediates are used as energy sources) can inhibit ribosome activity and denature enzymes. HEPES is preferred over phosphate buffers in certain formulations because it does not chelate magnesium ions, preserving Mg²⁺ availability for ribosomes (Calhoun & Swartz, 2005).

Reference: Calhoun, K. A. & Swartz, J. R. (2005). Energizing cell-free protein synthesis with glucose metabolism. Biotechnol. Bioeng., 90(5), 606–613.

Magnesium Glutamate

Magnesium is an indispensable cofactor for ribosome assembly and stability, for all nucleotide-utilizing enzymes (polymerases, kinases), and for numerous metalloenzymes involved in energy metabolism. The glutamate salt form is preferred over magnesium acetate because it mimics the ionic composition of the E. coli cytoplasm and avoids introduction of extraneous acetate ions. Mg²⁺ concentration must be carefully optimized — too low inhibits ribosomes, too high inhibits transcription — and is consistently identified as one of the most influential variables in CFPS performance (Caschera & Noireaux, 2015).

Reference: Caschera, F. & Noireaux, V. (2015). Synthesis of 2.3 mg/ml of protein with an all Escherichia coli cell-free transcription–translation system. Biochimie, 99, 162–168.

Potassium Phosphate Monobasic (KH₂PO₄) and Dibasic (K₂HPO₄)

These two phosphate salts together form a phosphate buffer pair that helps supplement pH buffering capacity alongside HEPES and also provides inorganic phosphate (Pᵢ) as a substrate for ATP regeneration through oxidative phosphorylation and substrate-level phosphorylation in the extract. Adequate Pᵢ availability is important for long-duration reactions because phosphate limitation is a known bottleneck for sustained ATP regeneration from glucose (Calhoun & Swartz, 2005).

Energy / Nucleotide System

Ribose

Ribose is the pentose sugar backbone of all ribonucleotides. In the NMP-Ribose-Glucose system, ribose can enter the pentose phosphate pathway in the extract, regenerating PRPP (5-phosphoribosyl-1-pyrophosphate) needed for nucleotide salvage synthesis, and also providing reducing equivalents (NADPH) that support energy metabolism.

Glucose

Glucose serves as the primary carbon and energy source for ATP regeneration through glycolysis and, in E. coli extract, through further oxidative metabolism via the TCA cycle. While early cell-free systems used phosphoenolpyruvate (PEP) as the energy substrate, Calhoun and Swartz demonstrated that glucose (with pH control) can sustain high-yield protein synthesis in a far more cost-effective manner, with the extract’s endogenous glycolytic enzymes converting glucose to pyruvate and then to acetate via the PANOx pathway, generating multiple ATP molecules per glucose (Calhoun & Swartz, 2005).

Reference: Calhoun, K. A. & Swartz, J. R. (2005). Energizing cell-free protein synthesis with glucose metabolism. Biotechnol. Bioeng., 90(5), 606–613.

AMP, CMP, GMP, UMP (Nucleoside Monophosphates)

These four nucleoside monophosphates are the building blocks for RNA synthesis. In the NMP-based system, the extract’s endogenous nucleoside monophosphate kinases and nucleoside diphosphate kinases phosphorylate NMPs → NDPs → NTPs, fueling both transcription (NTPs are the substrates for RNA polymerase) and providing GTP for translation (ribosome translocation, EF-Tu·GTP). Using NMPs instead of NTPs is substantially more economical and avoids the inhibitory phosphate accumulation that occurs when NTPs are directly hydrolyzed (Caschera & Noireaux, 2015).

Guanine

Although GMP is included as the primary guanosine source, free guanine (the nucleobase without ribose or phosphate) is added as a supplement for the purine salvage pathway. See the Bonus Question below for a detailed explanation.

Translation Mix — Amino Acids

17 Amino Acid Mix

This mix contains 17 of the 20 canonical amino acids that are chemically stable in standard stocks and can be dissolved together without degradation or precipitation. These are the direct building blocks for polypeptide synthesis during ribosomal translation.

Tyrosine

Tyrosine is provided separately because it has very low aqueous solubility at neutral pH and must be prepared and added as a dilute suspension or alkaline solution to avoid precipitation that would reduce its bioavailability to the ribosome.

Cysteine

Cysteine is added separately because it is highly reactive and prone to oxidation (to cystine or sulfinic acid) in the presence of air or metal ions; it must be kept in a reducing environment (or added fresh) to remain in its reduced, translatable form.

Additives

Nicotinamide (NAD⁺ precursor / NAD⁺ supplement)

Nicotinamide (vitamin B₃) is a precursor of NAD⁺, a critical cofactor for redox reactions central to energy metabolism (e.g., glycolysis, TCA cycle, oxidative phosphorylation) within the cell extract. Supplementing NAD⁺ or its precursor helps sustain the extract’s capacity to oxidize NADH back to NAD⁺, keeping energy regeneration active and extending reaction longevity (Jewett & Swartz, 2004).

Backfill — Nuclease-Free Water

Nuclease-free water is used to bring the reaction to its final target volume (the “backfill” volume). It is critical that this water is free of RNases and DNases, which would otherwise degrade the mRNA and DNA template, terminating the reaction prematurely.

B2. Main Differences Between the 1-Hour Optimized PEP-NTP Master Mix and the 20-Hour NMP-Ribose-Glucose Master Mix

The most fundamental difference is the energy and nucleotide supply strategy. The 1-hour PEP-NTP system relies on phosphoenolpyruvate as the energy source and pre-formed NTPs as direct substrates for transcription. PEP is a high-energy phosphate compound that rapidly regenerates ATP through pyruvate kinase, delivering fast but short-lived energy that is typically exhausted within 1–2 hours (Kim & Swartz, 2001). The 20-hour NMP-Ribose-Glucose system, by contrast, supplies nucleoside monophosphates (NMPs) and glucose as the primary substrates, relying on the extract’s endogenous kinase cascade (NMP → NDP → NTP) and glycolytic pathway to continuously regenerate NTPs and ATP over many hours at lower cost — extending productive reaction time but requiring a slower initial ramp-up phase (Caschera & Noireaux, 2015; Calhoun & Swartz, 2005).

A second key difference is the pH management requirement: glucose metabolism through glycolysis generates acidic by-products (lactate, acetate) that acidify the reaction over time, which is why the 20-hour formulation typically requires more robust buffering (e.g., higher HEPES or phosphate concentrations) and may include pH stabilizers such as K₂HPO₄/KH₂PO₄ in adjusted ratios. This pH drift problem is minimal in the short PEP-NTP system because the reaction ends before significant acidification occurs (Calhoun & Swartz, 2005).

Finally, the cost and complexity differ significantly: the NMP-Ribose-Glucose system is more cost-effective per reaction (NMPs and glucose are cheaper than PEP and NTPs), making it preferable for large-scale or long-duration experiments like the 36-hour global artwork incubation, whereas the PEP-NTP system is simpler to optimize quickly and delivers consistent burst expression, making it ideal for rapid prototyping within a 1-hour window.

References:
Kim, D. M. & Swartz, J. R. (2001). Regeneration of adenosine triphosphate from glycolytic intermediates for cell-free protein synthesis. Biotechnol. Bioeng., 74(4), 309–316.
Calhoun, K. A. & Swartz, J. R. (2005). Energizing cell-free protein synthesis with glucose metabolism. Biotechnol. Bioeng., 90(5), 606–613.
Caschera, F. & Noireaux, V. (2015). Synthesis of 2.3 mg/ml of protein with an all E. coli cell-free transcription–translation system. Biochimie, 99, 162–168.

B3. Bonus Question: How Can Transcription Occur If GMP Is Not Included But Guanine Is?

Transcription can occur because the E. coli extract retains active purine salvage pathway enzymes, specifically hypoxanthine-guanine phosphoribosyltransferase (HGPRT/Gpt), which catalyzes the reaction:

Guanine + PRPP → GMP + PPᵢ

In this salvage reaction, the free guanine base is combined with phosphoribosyl pyrophosphate (PRPP) — generated from ribose-5-phosphate and ATP — to yield GMP in a single, energetically efficient step (Fiveable, 2026; BOC Sciences, 2025). The resulting GMP is then phosphorylated sequentially by guanylate kinase (GMP → GDP) and nucleoside diphosphate kinase (GDP + ATP → GTP + ADP) to produce GTP, which is then available as a substrate for T7 RNA polymerase during transcription.

This is why the NMP-Ribose-Glucose formulation includes both GMP and free guanine: the guanine provides an additional flux route into the GTP pool via the salvage pathway, supplementing the kinase-driven NMP phosphorylation route and ensuring that GTP availability does not become limiting during prolonged transcription over the 20-hour reaction (Construction of a GTP Regeneration System…, ScienceDirect, 2024).

References:
Kang, S.H. et al. (2024). Construction of a GTP regeneration system by regulating gene expression in the pathway. Biochem. Eng. J., Elsevier.
Berg, J. M., Tymoczko, J. L., & Stryer, L. (2015). Biochemistry, 8th ed. W. H. Freeman. (Nucleotide metabolism chapter.)

Part C: Planning the Global Experiment — Cell-Free Master Mix Design

C1. Biophysical/Functional Properties of Each Fluorescent Protein Relevant to Cell-Free Expression

sfGFP (Superfolder Green Fluorescent Protein)

Key Property: Exceptionally robust folding / resistance to aggregation

sfGFP was engineered with six specific mutations relative to EGFP (including F99S, M153T, V163A, and others) that dramatically improve its folding kinetics and resistance to misfolding even when fused to poorly folding proteins or expressed under challenging conditions (Pédelacq et al., 2006). In a cell-free context, where the molecular chaperone network is diluted relative to an intact cell and protein concentration can accumulate rapidly, sfGFP’s superior folding robustness means a higher fraction of translated polypeptides successfully form a fluorescent chromophore rather than aggregating as non-fluorescent inclusion bodies. Chromophore formation still requires molecular oxygen (O₂) and proceeds via an autocatalytic cyclization and oxidation mechanism, making it oxygen-dependent in cell-free systems incubated under aerobic conditions.

Reference: Pédelacq, J. D. et al. (2006). Engineering and characterization of a superfolder green fluorescent protein. Nature Biotechnology, 24(1), 79–88.

mRFP1 (Monomeric Red Fluorescent Protein 1)

Key Property: Relatively slow and incomplete chromophore maturation

mRFP1 was the first true monomeric RFP derived from DsRed (Discosoma sp.) via 33 directed evolution mutations designed to break the obligate tetrameric interface while restoring fluorescence (Campbell et al., 2002). However, mRFP1’s maturation rate is significantly slower than its successor mCherry (~60 min half-time vs. ~15 min for mCherry) and it is known to mature incompletely — a significant fraction of translated mRFP1 polypeptides undergo the initial cyclization but fail to complete the second, oxidative step required to produce the fully red-shifted chromophore, yielding a “green-intermediate” species (Shaner et al., 2004). In a cell-free system, this incomplete maturation is a critical limitation: even after extensive translation, a substantial fraction of mRFP1 fluorescent signal may be delayed or lost, potentially underestimating expression relative to faster-maturing proteins.

References:
Campbell, R. E. et al. (2002). A monomeric red fluorescent protein. PNAS, 99(12), 7877–7882.
Shaner, N. C. et al. (2004). Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein. Nature Biotechnology, 22(12), 1567–1572.

mKO2 (Monomeric Kusabira-Orange 2)

Key Property: Moderate acid sensitivity and a multi-step maturation mechanism

mKO2 is a monomeric orange fluorescent protein (excitation ~551 nm, emission ~565 nm) derived from the coral Verrillofungia concinna (formerly Fungia concinna) (FPbase). In vivo studies have characterized mKO2’s maturation as following a three-state model: translated → folded (non-fluorescent) → matured (fluorescent), with the second transition having a maturation half-time of approximately 135 minutes in living cells (Chabry et al., 2016 — PLoS ONE). This long-lived non-fluorescent intermediate means that in a short or even moderate-length cell-free reaction, a large pool of mKO2 may remain in the non-fluorescent folded state, suppressing observed fluorescence despite successful translation. Additionally, mKO2 has moderate acid sensitivity (FPbase), meaning that if the pH drifts below ~7 during long incubations (as can occur with glucose metabolism), fluorescence yield will decrease.

References:
FPbase entry for mKO2: https://www.fpbase.org/protein/mko2/
Chabry, D. et al. (2016). Sensitive and Quantitative Three-Color Protein Imaging in Fission Yeast Using Spectrally Diverse, Recoded Fluorescent Proteins with Experimentally-Characterized In Vivo Maturation Kinetics. PLoS ONE, 11(8), e0159292.

mTurquoise2

Key Property: Oxygen-dependent chromophore maturation with slow kinetics

mTurquoise2 is currently the brightest cyan fluorescent protein derived from Aequorea victoria GFP (excitation ~434 nm, emission ~474 nm). Its chromophore maturation follows a complex, multi-step oxidative mechanism — the rate-determining step is the reaction of the pre-cyclized chromophore with molecular O₂ to generate the mature fluorescent species (Goedhart et al., 2012). Studies characterizing mTurquoise2 maturation report a half-time of approximately 36–45 minutes in bacterial systems (Bindels et al., 2017), which is substantially slower than sfGFP. In an aerobic cell-free system this is manageable over a 20–36 hour incubation, but if the reaction becomes oxygen-limited (which can occur as O₂ is consumed by both chromophore maturation chemistry and metabolic enzymes in the lysate), fluorescence readout will be substantially underestimated relative to protein output.

References:
Goedhart, J. et al. (2012). Structure-guided evolution of cyan fluorescent proteins towards a quantum yield of 93%. Nature Communications, 3, 751.
Bindels, D. S. et al. (2017). mScarlet: a bright monomeric red fluorescent protein for cellular imaging. Nature Methods, 14(1), 53–56.

mScarlet-I

Key Property: Fast maturation and high brightness — favorable for cell-free expression

mScarlet-I is a monomeric red fluorescent protein (excitation ~569 nm, emission ~593 nm) optimized for intracellular brightness and rapid maturation (Bindels et al., 2017). Compared to mRFP1, mScarlet-I matures substantially faster and achieves a much higher quantum yield (QY ~0.54), making it one of the brightest monomeric red FPs currently available. A notable biophysical property relevant to cell-free systems is that mScarlet-I contains no native cysteines in its Anthozoa-derived sequence, which means it does not form adventitious disulfide bonds in the oxidative microenvironment of the reaction mix — an important advantage since disulfide-linked aggregates are non-fluorescent (Bindels et al., 2017). Its rapid maturation means fluorescence signals will emerge quickly during incubation, providing an early and reliable readout.

Reference: Bindels, D. S. et al. (2017). mScarlet: a bright monomeric red fluorescent protein for cellular imaging. Nature Methods, 14(1), 53–56.

Electra2 (Blue Fluorescent Protein)

Key Property: High intracellular brightness relative to other BFPs, but Stokes-shifted chromophore maturation sensitive to β-barrel folding fidelity

Electra2 is a recently developed blue fluorescent protein (derived from Aequorea-class GFP) optimized using a dual bacterial/mammalian expression screening system (Papadaki et al., 2022). Its chromophore requires the Y66H substitution (relative to GFP Y66) that generates the blue-shifted tyrosine-derived chromophore, which undergoes the same O₂-dependent autocatalytic maturation as standard GFPs. In cell-free systems, Electra2’s high intracellular brightness in mammalian and bacterial contexts (reported as ~2.1-fold brighter than mTagBFP2 in bacteria; Papadaki et al., 2022) suggests good translation efficiency and folding fidelity across expression systems. A key limitation for cell-free readout is that the blue chromophore has a relatively low extinction coefficient compared to green/red FPs, meaning that even with efficient maturation, fluorescence intensity per molecule is lower, requiring sensitive plate reader settings for detection.

References:
Papadaki, G. F. et al. (2022). Dual-expression system for blue fluorescent protein optimization. Nature Communications, 13, 2887.
Hashimura, H. et al. (2025). Use of blue fluorescent protein Electra2 for live-cell imaging in Dictyostelium discoideum. microPublication Biology, 10.17912/micropub.biology.001774.

C2. Hypothesis for Reagent Optimization to Maximize Fluorescence Over 36-Hour Incubation

Target protein: mKO2

Identified limitation: mKO2’s three-state maturation pathway results in accumulation of a non-fluorescent folded intermediate with a ~135-minute half-time for the folded → matured transition. This transition is the rate-limiting oxidative step that requires molecular O₂.

Hypothesis:

Increasing the dissolved oxygen availability in the reaction (by periodic brief vortexing/agitation or by supplementing with an O₂-releasing compound such as hydrogen peroxide at sub-inhibitory concentrations ~0.01–0.05 mM), combined with increasing magnesium glutamate concentration by 2–4 mM above the baseline level, will accelerate mKO2 chromophore maturation and increase total fluorescence signal at 36 hours compared to the standard unadjusted master mix.

Rationale: Since the rate-determining step in mKO2 maturation is the O₂-dependent oxidation of the pre-chromophore (as modeled in the three-state kinetic study by Chabry et al., 2016), increasing effective O₂ availability should shift more of the folded intermediate pool toward the mature fluorescent state within the 36-hour window. Simultaneously, magnesium is the key cofactor for ribosome activity and for the NMP kinase cascade that sustains NTP supply; a modest Mg²⁺ increase (within the 10–20 mM optimal range established by Caschera & Noireaux, 2015) could sustain a higher translation rate, expanding the total translated mKO2 pool that feeds the maturation pipeline.

Expected effect: A 1.5–2× increase in total mKO2 fluorescence at the 36-hour endpoint, with fluorescence increase beginning earlier in the time course as O₂ supplementation accelerates the maturation bottleneck.

Reagent to modify (custom 2 µL supplement):

1 µL of 0.1 mM H₂O₂ (diluted fresh in nuclease-free water) → final ~0.01 mM in 20 µL reaction
1 µL of 40 mM MgGlu → adds +2 mM MgGlu to the reaction

References:
Chabry, D. et al. (2016). PLoS ONE, 11(8), e0159292.
Caschera, F. & Noireaux, V. (2015). Biochimie, 99, 162–168.
Goedhart, J. et al. (2012). Nature Communications, 3, 751. (O₂ dependence of GFP-class chromophore maturation)