Week 9 HW: Cell free systems

Homework Part A: General and Lecturer-Specific Questions

General homework 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 (CFPS) offers a fundamentally distinct paradigm compared to conventional in vivo expression, primarily because it decouples protein production from the constraints of cellular viability. In traditional cell-based systems, the researcher is obligated to work within the biological boundaries of a living organism, meaning that toxic proteins, membrane-disrupting compounds, or non-natural amino acids cannot be introduced without compromising cell survival. Cell-free systems eliminate this constraint entirely, granting direct and unobstructed access to the transcription-translation machinery.

The flexibility afforded by CFPS is unparalleled in several dimensions. The reaction conditions, including pH, ionic strength, redox potential, and temperature, can be modulated independently without concern for cellular homeostasis. Researchers can introduce unnatural amino acids via suppressor tRNA technology, adjust the concentration of individual components such as ribosomes or chaperones, and even pause or restart reactions at defined time points. This level of granular control over experimental variables is simply not achievable in a living cell, where compensatory regulatory mechanisms continuously buffer against perturbations.

Two cases where cell-free expression is demonstrably superior to cell-based production are as follows. First, the synthesis of cytotoxic proteins, such as bacteriocins or certain viral proteins, is practically impossible in living cells because the expressed product kills the host before sufficient accumulation occurs. In a cell-free system, there is no host to protect, and the protein can be synthesized to desired concentrations without any viability concern. Second, the incorporation of non-canonical amino acids for site-specific labeling or the production of proteins with novel chemical functionalities is far more tractable in CFPS, where the genetic code can be reprogrammed by supplementing the reaction with orthogonal aminoacyl-tRNA synthetase and suppressor tRNA pairs, bypassing the need for genome-wide engineering of a living organism.

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

A cell-free expression system is a reconstituted biochemical environment designed to support the complete process of gene expression outside of a living cell. Each component plays a specific and indispensable role in ensuring that the transcription and translation machinery functions with fidelity and efficiency.

The cell extract, typically derived from prokaryotic sources such as Escherichia coli or from eukaryotic sources such as wheat germ, rabbit reticulocyte lysate, or insect cell lysate, constitutes the foundational element of any CFPS reaction. This extract supplies the ribosomes, translation factors, aminoacyl-tRNA synthetases, chaperones, and other endogenous enzymes necessary for polypeptide synthesis. The quality and preparation method of the extract are critical determinants of overall system productivity.

The DNA or mRNA template provides the genetic information encoding the target protein. In systems that include an RNA polymerase, a plasmid or linear DNA template can be used directly, allowing transcription to occur in situ. In translation-only systems, a pre-synthesized mRNA bearing a suitable 5’ untranslated region and Kozak or Shine-Dalgarno sequence is supplied directly to the ribosomes.

Amino acids are the building blocks of the nascent polypeptide chain and must be supplied exogenously at concentrations sufficient to sustain the entire synthesis reaction without becoming rate-limiting. An energy regeneration system, typically composed of ATP, GTP, creatine phosphate, and creatine kinase or phosphoenolpyruvate and pyruvate kinase, provides the thermodynamic driving force for aminoacyl-tRNA charging, ribosome translocation, and other energy-dependent steps. Salts and buffer components, including magnesium acetate, potassium glutamate, and HEPES or Tris buffer, maintain the ionic environment required for ribosome integrity and enzymatic activity. Finally, an RNA polymerase such as T7 RNA polymerase is often added exogenously to drive robust transcription from a T7 promoter-containing template.

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 a critical bottleneck in cell-free protein synthesis because, unlike living cells, CFPS reactions lack the metabolic infrastructure to continuously generate ATP through oxidative phosphorylation or glycolysis. The translation of each peptide bond consumes multiple high-energy phosphate equivalents, and the rapid depletion of ATP leads to premature termination of the reaction, severely limiting protein yield. Without a mechanism to regenerate ATP from ADP and AMP, the reaction becomes thermodynamically unfavorable within minutes, making energy regeneration not merely important but absolutely essential for sustained productivity.

The most widely employed strategy for continuous ATP supply in cell-free experiments is the use of a secondary energy substrate coupled to a regenerating enzyme. The creatine phosphate and creatine kinase system is among the most established approaches. In this method, creatine phosphate serves as a phosphate donor, and creatine kinase catalyzes the transfer of the high-energy phosphate group to ADP, regenerating ATP in a rapid and reversible reaction. This system is particularly effective in short-duration reactions and is well-characterized in both prokaryotic and eukaryotic CFPS.

An alternative and increasingly preferred method is the use of phosphoenolpyruvate (PEP) as the energy substrate, coupled with pyruvate kinase present endogenously in the cell extract. PEP donates its phosphate group to ADP to regenerate ATP, with pyruvate as the byproduct. This system has the advantage of being self-contained when using crude cell extracts, as pyruvate kinase activity is naturally present. More recently, glucose-based systems that couple glycolysis to ATP regeneration have been developed, offering a cost-effective and longer-lasting energy supply by metabolizing glucose through the endogenous glycolytic enzymes retained in the extract, thereby sustaining protein synthesis for several hours.

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

Prokaryotic and eukaryotic cell-free expression systems differ fundamentally in their translational machinery, post-translational modification capacity, and the classes of proteins they are best suited to produce. Understanding these differences is essential for selecting the appropriate platform for a given target protein.

Prokaryotic CFPS systems, most commonly derived from E. coli, are characterized by high productivity, low cost, rapid preparation, and well-established protocols. The E. coli extract supports efficient translation from mRNAs bearing Shine-Dalgarno sequences and is compatible with T7 RNA polymerase-driven transcription. However, prokaryotic systems lack the endomembrane compartments, glycosylation machinery, and eukaryote-specific chaperones required for the proper folding and modification of complex eukaryotic proteins. An appropriate protein to produce in a prokaryotic CFPS system would be a bacteriophage structural protein such as the T4 major capsid protein gp23. This protein is of prokaryotic origin, does not require glycosylation or disulfide bond formation under reducing cytoplasmic conditions, and benefits from the high translational efficiency of the E. coli extract. The prokaryotic system is therefore ideally matched to the biochemical requirements of this target.

Eukaryotic CFPS systems, including those derived from wheat germ embryos, rabbit reticulocytes, insect cells (Sf21 or Sf9), or Chinese hamster ovary cells, contain the molecular chaperones, signal recognition particles, and in some cases microsomal membranes necessary for the synthesis of complex eukaryotic proteins with authentic post-translational modifications. A protein well-suited for eukaryotic CFPS would be human erythropoietin (EPO), a heavily glycosylated cytokine whose biological activity and serum half-life are critically dependent on N-linked glycosylation at specific asparagine residues. Producing EPO in a prokaryotic system would yield an aglycosylated, biologically suboptimal product, whereas a eukaryotic CFPS system, particularly one supplemented with microsomal membranes, can support signal peptide cleavage, translocation into the endoplasmic reticulum lumen, and glycan addition, producing a protein far closer to its native form.

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.

Membrane proteins represent one of the most challenging classes of targets in recombinant protein production, whether in vivo or in vitro, due to their inherent hydrophobicity, tendency to aggregate in aqueous environments, and requirement for a lipid bilayer or membrane-mimetic environment to achieve proper folding and function. A well-designed cell-free experiment for membrane protein expression must address these challenges systematically at every stage of the experimental setup.

The first and most fundamental consideration is the choice of hydrophobic environment into which the nascent membrane protein will be co-translationally or post-translationally inserted. Three principal strategies are employed in the field. The first is the addition of detergent micelles, such as digitonin, DDM (n-dodecyl-β-D-maltoside), or LMNG (lauryl maltose neopentyl glycol), directly to the cell-free reaction at concentrations above the critical micelle concentration. The detergent provides a hydrophobic core that accommodates the transmembrane domains and prevents aggregation. The second strategy involves the inclusion of preformed liposomes or nanodiscs, which are phospholipid bilayer discs stabilized by membrane scaffold proteins. Nanodiscs are particularly advantageous because they provide a defined, native-like lipid bilayer environment of controlled size and lipid composition, enabling the study of lipid-protein interactions. The third approach uses lipid-polymer hybrid systems or bicelles, which offer intermediate properties between micelles and bilayers.

The cell extract itself should be selected carefully. An E. coli-based extract is commonly used for prokaryotic membrane proteins, while insect cell or HeLa cell extracts are preferred for eukaryotic targets requiring specific lipid environments or post-translational modifications. The magnesium concentration must be optimized empirically, as membrane protein translation is particularly sensitive to ionic conditions that affect ribosome association with the membrane-mimetic surface. Chaperone supplementation, particularly with SecYEG translocon components or YidC insertase for bacterial inner membrane proteins, or with Sec61 and TRAP complexes for eukaryotic targets, can substantially improve folding efficiency. Functional validation of the expressed membrane protein should be performed using activity assays, ligand binding studies, or reconstitution into proteoliposomes, since yield alone is an insufficient measure of success for this protein class.

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. Observing low yield of a target protein in a cell-free system is a common but multifactorial problem that requires systematic diagnosis rather than arbitrary modification of reaction conditions. Three of the most frequent underlying causes, along with their respective troubleshooting strategies, are described below.

The first possible reason is suboptimal template quality or concentration. If the DNA or mRNA template is degraded, contaminated with inhibitory substances such as residual phenol or ethanol from nucleic acid purification, or present at a concentration outside the optimal range, transcription and translation efficiency will be severely compromised. To address this, the template should be re-purified using a column-based method and quantified spectrophotometrically, with the A260/A280 ratio confirmed to be between 1.8 and 2.0. A titration experiment spanning a range of template concentrations, typically from 1 to 50 nM for plasmid DNA or 10 to 200 nM for mRNA, should be conducted to identify the optimal input. Additionally, the integrity of the mRNA should be verified by denaturing gel electrophoresis before use in translation-only systems.

The second possible reason is rapid degradation of the synthesized protein by proteases present in the cell extract. Crude cell extracts inevitably contain endogenous proteases that, if not adequately suppressed, will degrade the target protein as fast as it is synthesized, resulting in low apparent yield despite active translation. The troubleshooting strategy here involves the addition of protease inhibitor cocktails to the reaction, the use of protease-deficient extract strains such as E. coli BL21 Star or dedicated CFPS strains with deletions in major protease genes, or the incorporation of a purification tag such as His6 or Strep-tag II that allows rapid affinity capture of the protein immediately after synthesis, minimizing exposure time to proteolytic activity.

The third possible reason is inefficient energy regeneration leading to premature ATP depletion. As discussed previously, the translation machinery is highly energy-dependent, and if the ATP regeneration system is inadequate, either due to insufficient concentration of the phosphate donor or inactivation of the regenerating enzyme, the reaction will stall early. To troubleshoot this, the concentration of creatine phosphate or PEP should be increased incrementally, and the activity of the regenerating enzyme should be verified independently. Switching from a creatine phosphate system to a more sustained glucose-based or maltose-based energy regeneration platform can extend the productive phase of the reaction significantly. Monitoring ATP concentration over the course of the reaction using a luciferase-based ATP assay provides direct evidence of whether energy depletion is the limiting factor, allowing targeted intervention rather than broad empirical optimization.

Homework question from Kate Adamala

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

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

The synthetic minimal cell is designed to expand the chemical sensing capacity of mammalian reporter cells that cannot directly respond to extracellular glucose fluctuations in a programmable, synthetic manner. The input to the SMC is glucose, which passively permeates the liposomal membrane via encapsulated glucose transporter activity or through membrane permeability at low concentrations. Inside the SMC, glucose activates a DNA aptamer-regulated gene circuit that drives the expression of adenylate cyclase (CyaA), which converts ATP to cyclic AMP (cAMP). The output of the SMC is cAMP, which is released into the extracellular environment through an expressed membrane pore and subsequently activates a cAMP-responsive element (CRE)-driven luciferase reporter in co-cultured mammalian HEK293 cells. The overall system output is therefore measurable luciferase bioluminescence proportional to extracellular glucose concentration.

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

No. Without encapsulation, the cAMP produced by the cell-free transcription-translation reaction would immediately diffuse into the bulk medium and enter the mammalian reporter cells regardless of whether glucose was present or absent. The entire logic of the system depends on glucose-triggered gene expression occurring inside the SMC, followed by controlled release of cAMP through a pore that is itself expressed only upon glucose sensing. Without the physical compartmentalization provided by the lipid membrane, the conditional gating mechanism is entirely lost and the system would constitutively activate the reporter, eliminating its utility as a biosensor.

c. Could this function be realized by genetically modified natural cell?

In principle, yes. A mammalian cell could be engineered to express a synthetic glucose-sensing riboswitch or aptazyme upstream of an adenylate cyclase transgene, thereby coupling extracellular glucose to intracellular cAMP production. However, this approach is far less generalizable than the SMC strategy. Engineering a new sensing modality into a living cell requires stable genomic integration, extensive optimization of gene regulation, and the risk of interference from endogenous metabolic pathways, particularly since mammalian cells already possess complex glucose sensing and cAMP signaling networks that would confound the synthetic circuit. The SMC approach is modular, meaning the glucose-sensing aptamer can in principle be swapped for any other aptamer without re-engineering the reporter cell, making a single reporter cell line compatible with detection of many different analytes.

d. Describe the desired outcome of your synthetic cell operation.

In the presence of the SMC population co-cultured with CRE-luciferase HEK293 reporter cells, the system as a whole gains the capacity to sense extracellular glucose and transduce that signal into a quantifiable bioluminescent output. In the absence of glucose, no cAMP is released and the reporter cells remain quiescent. Upon glucose addition, the SMC internal gene circuit is activated, cAMP is synthesized and released, and the reporter cells produce luciferase in a dose-dependent manner.

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

The membrane would be composed of POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine) and cholesterol in a molar ratio of approximately 7:3. Cholesterol is included to reduce membrane fluidity and improve vesicle stability at physiological temperature. This lipid composition produces a stable unilamellar liposome that is permeable to small uncharged molecules such as glucose at low concentrations, while retaining larger molecules such as cAMP and nucleic acids inside the lumen.

b. What would you encapsulate inside? Enzymes, small molecules.

The SMC lumen would contain a bacterial cell-free Tx/TI system (E. coli S30 extract supplemented with T7 RNA polymerase, amino acids, NTPs, and an ATP regeneration system based on phosphoenolpyruvate and pyruvate kinase), a plasmid encoding the pore-forming protein alpha-hemolysin (aHL) under the control of a theophylline-responsive riboswitch, and a second plasmid encoding a truncated, constitutively active adenylate cyclase domain (the catalytic domain of Bordetella pertussis CyaA, residues 1 to 364) under the control of a glucose-responsive aptazyme regulatory element. ATP would also be encapsulated as the substrate for cAMP synthesis.

c. 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)

A bacterial cell-free Tx/TI system derived from E. coli is appropriate for this design. The regulatory elements used, specifically the theophylline riboswitch and the aptazyme-based glucose sensor, are RNA-level regulatory elements that function within prokaryotic transcription-translation contexts and do not require the eukaryotic spliceosomal or cap-dependent translation machinery. A mammalian system would be unnecessary and would add cost and complexity without functional benefit for this particular gene circuit architecture.

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

The SMC membrane is passively permeable to glucose, which enters the lumen and activates the internal gene circuit. The output molecule, cAMP, is a small, charged molecule that does not readily cross an intact phospholipid bilayer on its own. Therefore, the SMC is designed to express alpha-hemolysin (aHL), a self-assembling heptameric pore-forming protein that inserts into the lipid bilayer and creates a non-selective aqueous channel with a diameter of approximately 1.4 nm. This pore is large enough to permit passive diffusion of cAMP (molecular weight 329 Da) from the SMC lumen into the extracellular medium, where it can then act on the co-cultured mammalian reporter cells.

3. Experimental details a. 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 and cholesterol (7:3 molar ratio), used to form unilamellar liposomes by extrusion through a 200 nm polycarbonate membrane.

Genes and constructs Alpha-hemolysin (aHL) from Staphylococcus aureus (gene: hla, UniProt P09616), cloned downstream of a T7 promoter and a theophylline-responsive riboswitch (aptamer sequence from Martini and Mansy, 2011) to allow translational activation upon glucose-triggered internal signaling. Note: in a simplified first-generation design, theophylline can be co-encapsulated to constitutively activate aHL expression after a defined lag period, ensuring pore formation occurs before cAMP accumulates.

Adenylate cyclase catalytic domain from Bordetella pertussis CyaA (gene: cyaA, residues encoding amino acids 1 to 364, which constitute the calmodulin-independent catalytic fragment), cloned downstream of a T7 promoter and a glucose-sensing aptazyme derived from the glucosamine-6-phosphate riboswitch (glmS ribozyme system), which undergoes self-cleavage in the absence of glucose-6-phosphate and is stabilized in its full-length, translationally competent form when glucose-6-phosphate is present intraluminally.

Reporter construct in biological cells: HEK293 cells stably transfected with a CRE (cAMP response element) driving firefly luciferase (pCRE-Luc, commercially available from Promega as the pCRE-Luc vector).

b. How will you measure the function of your system?

The primary readout is bioluminescence from the CRE-luciferase HEK293 reporter cells. After co-incubation of SMCs with reporter cells in the presence of varying concentrations of glucose, cells are lysed and luciferase activity is quantified using a luminometer with a luciferin substrate reagent such as the Promega Bright-Glo system. A dose-response curve of luciferase signal versus glucose concentration establishes the dynamic range and sensitivity of the biosensor. As a secondary validation, intracellular cAMP concentration in the reporter cells can be measured independently using a competitive ELISA-based cAMP assay kit to confirm that the luciferase signal is driven by cAMP and not by an off-target effect. Negative controls include SMCs lacking the CyaA gene, SMCs prepared without glucose, and free cell-free extract added to reporter cells without encapsulation, the last of which directly tests whether encapsulation is necessary for conditional function.

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.

A structural building material, specifically a load-bearing wall panel, embedded with freeze-dried cell-free biosensor systems that rehydrate upon water ingress to produce a colorimetric signal, enabling passive, real-time detection of moisture damage, mold-conducive conditions, and structural water infiltration without any electronic infrastructure.

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

The core concept involves embedding freeze-dried cell-free transcription-translation (Tx/TI) pellets into a porous polymer matrix or cement composite that forms the interior layer of a wall panel. Under normal dry conditions, the cell-free components remain completely inert and stable within the material. When water infiltrates the wall due to a leak, condensation buildup, or flood damage, the moisture rehydrates the freeze-dried pellets locally at the site of ingress, activating the encapsulated gene circuit.

The activated cell-free system expresses a reporter enzyme, specifically laccase or beta-galactosidase, which reacts with a co-embedded chromogenic substrate such as X-gal or a catechol derivative to produce a visible color change at the precise location of water infiltration. This colorimetric output diffuses to the surface of the panel or is visible through a thin translucent indicator strip laminated onto the interior wall surface, alerting building occupants or maintenance personnel to the exact location and approximate extent of moisture damage without requiring any sensors, wiring, or digital infrastructure.

A secondary circuit layer could be designed to detect the presence of volatile organic compounds associated with mold growth, such as 1-octen-3-ol, by incorporating an aptamer-regulated gene circuit responsive to fungal metabolites, thereby providing an early biological warning before visible mold colonization occurs. The entire system is entirely passive, requires no power source, and operates on the same principle as a one-time biological fuse triggered by the environmental condition it is designed to detect.

• What societal challenge or market need will this address?

Water damage is one of the most economically devastating and health-relevant problems in the built environment. According to insurance industry data, water infiltration and moisture-related structural damage account for billions of dollars in repair costs annually across residential and commercial buildings worldwide. More critically, hidden moisture accumulation within walls creates conditions favorable to toxic mold growth, particularly Stachybotrys chartarum and Aspergillus species, which are associated with severe respiratory illness, neurological symptoms, and long-term occupant health consequences. Current detection methods rely either on expensive electronic moisture sensors that require installation and maintenance, or on visual inspection after damage has already become extensive. There is a clear and unmet market need for a passive, low-cost, infrastructure-free early warning system that can be built directly into construction materials at the point of manufacture, particularly in affordable housing, schools, hospitals, and disaster-relief temporary structures where electronic monitoring is impractical or cost-prohibitive.

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

The three principal limitations of cell-free systems in this context are activation specificity, long-term stability, and the one-time use constraint, each of which can be addressed through deliberate material engineering.

Regarding activation specificity, the risk that ambient humidity rather than true water infiltration triggers the system prematurely is addressed by embedding the freeze-dried pellets within a hydrophobic polymer shell, such as a poly(lactic-co-glycolic acid) microsphere coating, that requires sustained liquid water contact rather than vapor-phase humidity to dissolve and release the rehydrating water to the cell-free core. This ensures that the system is activated only by genuine liquid ingress events and not by normal fluctuations in indoor relative humidity.

Regarding long-term stability, freeze-dried cell-free systems supplemented with trehalose and bovine serum albumin as lyoprotectants have been demonstrated to retain activity for over one year at room temperature and for multiple years under cool, dry storage conditions, as shown by Pardee et al. (2016) and subsequent work from the Jewett and Noireaux laboratories. Incorporating these stabilizers into the pellet formulation, combined with the protective hydrophobic shell described above, is expected to provide a shelf life compatible with the decades-long service life of building materials.

Regarding the one-time use constraint, this is reframed as a feature rather than a limitation in this application. A building wall panel that produces an irreversible color change upon first water contact provides a permanent, tamper-evident record of moisture infiltration history, which is directly valuable for insurance assessment, building inspection, and liability documentation. For applications where repeated sensing is desired, a modular panel design could allow the indicator strip layer to be replaced during routine maintenance while the structural panel itself remains in place, effectively resetting the sensing capacity at low cost.

Homework question from Ally Huang

Freeze-dried cell-free reactions have great potential in space, where resources are constrained. As described in my talk, the Genes in Space competition challenges students to consider how biotechnology, including cell-free reactions, can be used to solve biological problems encountered in space. While the competition is limited to only high school students, your assignment will be to develop your own mock Genes in Space proposal to practice thinking about biotech applications in space!

For this particular assignment, your proposal is required to incorporate the BioBits® cell-free protein expression system, but you may also use the other tools in the Genes in Space toolkit (the miniPCR® thermal cycler and the P51 Molecular Fluorescence Viewer). For more inspiration, check out https://www.genesinspace.org/ .

1. 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. (Maximum 100 words)

Astronauts on long-duration missions beyond low Earth orbit are exposed to chronic galactic cosmic radiation and solar particle events at doses far exceeding terrestrial limits. This ionizing radiation causes double-strand DNA breaks, oxidative stress, and genomic instability, increasing cancer risk and impairing immune function. Current onboard diagnostics cannot monitor molecular-level radiation damage in real time. Developing a rapid, resource-minimal biosensor capable of detecting radiation biomarkers directly aboard spacecraft is critical for crew health monitoring, mission safety decision-making, and advancing our understanding of how the human body responds to the space radiation environment.

2. 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. (Maximum 30 words)

The molecular targets are the mRNA transcripts of two radiation-responsive genes: CDKN1A (p21), a DNA damage checkpoint effector, and GADD45A, a stress-inducible DNA repair gene, measured in astronaut saliva-derived cell-free RNA.

3. Describe how your molecular or genetic target relates to the space biology question or challenge your proposal addresses. (Maximum 100 words)

CDKN1A and GADD45A are both transcriptionally upregulated within hours of ionizing radiation exposure as part of the p53-mediated DNA damage response pathway. Their mRNA levels in biofluids such as saliva have been validated as non-invasive, quantitative biomarkers of radiation dose in terrestrial clinical and occupational settings. In the context of spaceflight, monitoring the upregulation of these transcripts in real time would provide a direct molecular readout of cumulative radiation-induced cellular stress in the astronaut, enabling dose assessment without blood draws, laboratory equipment, or ground-based analysis, all of which are impractical during deep space missions.

4. Clearly state your hypothesis or research goal and explain the reasoning behind it. (Maximum 150 words)

The central hypothesis is that freeze-dried BioBits® cell-free transcription-translation reactions can be engineered to function as toehold switch-based biosensors that produce a detectable fluorescent signal specifically in the presence of CDKN1A or GADD45A mRNA extracted from astronaut saliva, and that the intensity of this fluorescent signal will correlate positively with the cumulative radiation dose received by the crew member.

The reasoning behind this goal is as follows. Toehold switches are synthetic RNA regulatory elements that remain translationally repressed in the absence of their cognate trigger RNA and are activated upon binding of the target mRNA, driving expression of a reporter such as sfGFP. By designing toehold switches specific to CDKN1A and GADD45A transcripts and freeze-drying them into BioBits® reactions, a stable, single-use diagnostic cartridge can be created that requires only saliva RNA input and water to operate, making it fully compatible with the resource and infrastructure constraints of spaceflight.

5. 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. (Maximum 100 words)

Saliva samples will be collected from astronauts at defined intervals: pre-flight baseline, weekly during mission, and post-flight. Cell-free RNA will be extracted using a simple silica column protocol and added directly to rehydrated BioBits® toehold switch reactions targeting CDKN1A and GADD45A mRNA. Fluorescence output will be measured using the P51 Molecular Fluorescence Viewer. Controls include a no-template negative control, a synthetic trigger RNA positive control at known concentrations, and a housekeeping gene toehold switch targeting ACTB mRNA to normalize expression. Signal intensity will be quantified as a ratio of target to housekeeping fluorescence and correlated with dosimeter-recorded radiation exposure data.