Methods for Embedding Cell-Free Protein Synthesis Reactions in Macro-Scale Hydrogels | Text Page (2024)

Synthetic biology integrates diverse engineering disciplines to design and engineer biologically based parts, devices, and systems that can perform functions that are not found in nature. Most synthetic biology approaches are still bound to living cells. By contrast, cell-free synthetic biology systems facilitate unprecedented levels of control and freedom in design, enabling increased flexibility and a shortened time for engineering biological systems while eliminating many of the constraints of traditional cell-based gene expression methods^1,2,3. CFPS is being used in an increasing number of applications across numerous disciplines, including constructing artificial cells, prototyping genetic circuits, developing biosensors, and producing metabolites^4,5,6. CFPS has also been particularly useful for producing recombinant proteins that cannot be readily expressed in living cells, such as aggregation-prone proteins, transmembrane proteins, and toxic proteins^6,7,8.

CFPS is typically performed in liquid reactions. This, however, may restrict their deployment in some situations, as any liquid cell-free device must be contained within a reaction vessel. The rationale for the development of the methods presented here was to provide robust protocols for embedding cell-free synthetic biology devices into hydrogels, not as a protein production platform per se but, instead, to allow the use of hydrogels as a physical chassis for the deployment of cell-free devices beyond the laboratory. The use of hydrogels as CFPS chassis has several advantages. Hydrogels are polymeric materials that, despite a high water content (sometimes in excess of 98%), possess solid properties^9,10,11. They have uses as pastes, lubricants, and adhesives and are present in products as diverse as contact lenses, wound dressings, marine adhesive tapes, soil improvers, and baby diapers^{9,11,12,13,14}. Hydrogels are also under active investigation as drug delivery vehicles^9,15,16,17. Hydrogels may also be biocompatible, biodegradable, and possess some stimuli responses of their own^9,18,19,20. Therefore, the goal here is to create a synergy between molecular biology-derived functionality and materials science. To this end, efforts have been made to integrate cell-free synthetic biology with a range of materials, including collagen, laponite, polyacrylamide, fibrin, PEG-peptide, and agarose^11,21,22, as well as to coat surfaces of glass, paper, and cloth^11,23,24 with CFPS devices. The protocols presented here demonstrate two methods for embedding CFPS reactions in macro-scale (i.e., >1 mm) hydrogel matrices, using agarose as the exemplar material. Agarose was chosen due to its high water absorption capacity, controlled self-gelling properties, and tuneable mechanical properties^11,24,25,26. Agarose also supports functional CFPS, is cheaper than many other hydrogel alternatives, and is biodegradable, making it an attractive choice as an experimental model system. These methods have, however, been previously demonstrated as appropriate for embedding CFPS in a range of alternative hydrogels¹¹. Considering the wide range of applications of hydrogels and the functionality of CFPS, the methods demonstrated here can provide a basis from which researchers are able to develop biologically enhanced hydrogel materials suited to their own ends.

In previous studies, microgel systems with a size range of 1 µm to 400 µm have been used to perform CFPS in hydrogels submerged in reaction buffer^{23,27,28,29,30,31}. The requirement to submerge hydrogels within CFPS reaction buffers, however, limits opportunities for their deployment as materials in their own right. The protocols presented here allow CFPS reactions to occur within hydrogels without the need to submerge the gels in reaction buffers. Second, the use of macroscale gels (between 2 mm and 10 mm in size) allows the study of the physical interaction between hydrogels and cell-free gene expression. For example, with this technique, it is possible to study how the hydrogel matrix affects CFPS reactions¹¹ and how CFPS reactions can impact the hydrogel matrix³¹. Larger sizes of hydrogels also allow for the development of novel, bio-programmable materials³². Finally, by embedding CFPS reactions into hydrogels, there is also a potential reduction in the requirement for plastic reaction vessels. For the deployment of cell-free sensors, this has clear advantages over devices dependent on plasticware. Taken together, embedding CFPS reactions in hydrogels provides several advantages for the deployment of cell-free devices beyond the laboratory.

The overall goal of the methods presented here is to allow the operation of CFPS reactions within hydrogel matrices. Two different methods are demonstrated for embedding cell-free protein production reactions in macro-scale hydrogel materials (Figure 1). In Method A, CFPS components are introduced to lyophilized agarose hydrogels to form an active system. In Method B, molten agarose is mixed with CFPS reaction components to form a complete CFPS hydrogel system, which is then lyophilized and stored until needed. These systems can be rehydrated with a volume of water or buffer and analyte to commence the reaction.

This study uses Escherichia coli cell lysate-based systems. These are some of the most popular experimental CFPS systems, as E. coli cell lysate preparation is simple, inexpensive, and achieves high protein yields. The cell lysate is complemented with the macromolecular components needed to perform transcription and translation, including ribosomes, tRNAs, aminoacyl-tRNA synthetases, and initiation, elongation, and termination factors. Specifically, this paper demonstrates the production of eGFP and mCherry in agarose hydrogels using E. coli cell lysates and monitors the appearance of fluorescence using a plate reader and confocal microscopy. Representative results for the microtiter plate reader can be seen in Whitfield et al.³¹, and the underlying data are publicly available³³. Further, the expression of fluorescent proteins throughout the gels is confirmed using confocal microscopy. The two protocols demonstrated in this paper allow for the assembly and storage of CFPS-based genetic devices in materia with the ultimate goal of creating a suitable physical environment for the distribution of cell-free gene circuitry in a manner supportive of field deployment.

1. Cell lysate buffer and media preparation

1. Preparation of 2x YT+P agar and medium
   1. Prepare 2x YT+P agar by measuring out 16 g/L tryptone, 10 g/L yeast extract, 5 g/L NaCl, 40 mL/L 1 M K₂HPO₄, 22 mL/L 1 M KH₂PO₄, and 15 g/L agar. For the 2x YT+P broth, follow the previous composition but omit the agar.
   2. Sterilize by autoclaving the 2x YT+P.
2. Preparation of the S30A buffer
   1. Prepare the S30A buffer with 5.88 g/L Mg-glutamate, 12.195 g/L K-glutamate, and 25 mL/L 2 M Tris, adjusted to pH 7.7 with acetic acid.
   2. Store the S30A buffer at 4 °C for up to 1 week.
3. Preparation of the S30B buffer
   1. Prepare the S30B buffer with 5.88 g/L Mg-glutamate, and 12.195 g/L K-glutamate, adjusted to pH 8.2 with 2 M Tris.
   2. Store the S30B buffer at 4 °C for up to 1 week.

2. Cell lysate preparation (4 day protocol)

1. Day 1
   1. Pour the 2x YT+P into agar plates supplemented with 100 µg/mL ampicillin.
   2. Streak E. coli from −80 °C glycerol stock, and incubate overnight at 37 °C.
2. Day 2
   1. Inoculate a single colony from the 2x YT+P plate into 400 mL of 2x YT+P broth (supplemented with ampicillin) in a 1 L baffled flask.
   2. Incubate for 24 h at 37 °C with 220 rpm shaking.
3. Day 3
   1. Measure and record the OD₆₀₀ of the 24 h cultures; growth is sufficient at an OD₆₀₀ of 3-4.
      NOTE: Take a 1 mL aliquot of bacterial culture, and perform a serial dilution with medium (2x YT+P broth) prior to measuring the OD₆₀₀ nm. Allow for the dilution effect when calculating the OD₆₀₀.
   2. Transfer the culture evenly between pre-chilled 500 mL centrifuge bottles.
   3. Centrifuge at 4,500 x g for 15 min at 4 °C, and then discard the supernatant.
   4. While centrifuging, complete the S30A buffer preparation by adding 2 mL/L of 1 M DTT to the previously prepared S30A buffer, mixing, and maintaining the buffer on ice.
   5. Resuspend the cell pellets in 200 mL of S30A buffer.
   6. Vortex the bottles vigorously until the whole pellet is completely solubilized, keeping the cells on ice as much as possible.
   7. Centrifuge at 4,500 x g for 15 min at 4 °C, and then discard the supernatant.
   8. Repeat steps 2.3.5-2.3.7 (inclusive).
   9. Add 40 mL of S30A buffer to each centrifuge bottle, resuspend the pellets, and transfer to pre-weighed 50 mL centrifuge tubes.
   10. Centrifuge at 4,000 x g for 15 min at 4 °C. Discard the supernatant by decanting, and remove the residual supernatant by pipette, keeping it on ice as much as possible.
   11. Re-weigh the centrifuge tubes, record the new mass, and calculate the mass of the pellets.
   12. Flash-freeze pellets in liquid nitrogen, and store them at −80 °C.
4. Day 4
   1. Thaw the pellets on ice.
   2. Per 1 g of pellet, add the following: 1.2 mL of S30A buffer (DTT added before use), 275 µL of protease inhibitor (8 mM stock), and 25 µL of lysozyme (10 mg/mL stock).
   3. Vortex vigorously until the pellets are completely solubilized with no remaining clumps, keeping them on ice as much as possible.
   4. In 50 mL centrifuge tubes, sonicate the cell suspensions at 120 W, 20 kHz, 30% amplitude, and 20 s/40 s on/off pulses for 5 min of sonication.
   5. Centrifuge at 4,000 x g for 1 h at 4 °C to pellet the cell debris.
   6. Transfer the supernatant to fresh 50 mL centrifuge tubes.
   7. Incubate tubes at 37 °C at 220 rpm for 80 min.
   8. Prepare dialysis materials by adding 1 mL/L of 1 M DTT to S30B buffer. Mix and add 900 mL to a 1 L sterile beaker. Add a magnetic stirrer, and keep it at 4 °C.
   9. Following the runoff reaction, transfer the cell extract from step 2.4.7 into 2 mL microcentrifuge tubes, and centrifuge at 20,000 x g for 10 min at 4 °C.
   10. Consolidate the pellet-free supernatant on ice in a 50 mL centrifuge tube.
   11. Determine the total volume of cell extract produced, and hydrate the necessary number of 10K MWCO dialysis cassettes by submerging them in S30B for 2 min.
   12. Load the cassettes via a syringe with up to 3 mL of extract. Dialyze up to three cassettes per 1 L beaker, stirring at 4 °C for 3 h.
   13. After the dialysis is complete, which clarifies the extract, transfer the extract to 2 mL micro-centrifuge tubes. Centrifuge at 20,000 x g for 10 min at 4 °C.
   14. Consolidate the clarified extract into a fresh centrifuge tube on ice, and vortex briefly to mix.
   15. Determine the protein concentration of the extract using a fluorometer.
   16. Dilute the extract to a concentration of 44.5 mg/mL using pre-chilled ddH₂O.
   17. Aliquot into 200 µL aliquots, flash-freeze in liquid nitrogen, and store at −80 °C.

3. Preparation of lyophilized hydrogels (Method A)

1. Weigh 0.75 g of agarose, add ddH₂O to a volume of 100 mL, and microwave in 30 s bursts at high temperatures to create molten agarose stock.
2. Using a pipette, transfer 50 µL of molten agarose to a 1.5 mL microcentrifuge tube, and allow the gel to polymerize for 20-30 min (polymerization occurs faster if the gels are transferred to a fridge).
3. Flash-freeze the microcentrifuge tubes, containing the polymerized gels, in liquid nitrogen.
4. Remove the lid of the centrifuge tubes, cover the centrifuge tube openings with wax film, and pierce the film several times with a needle or pipette tip.
5. Transfer the microcentrifuge tubes containing the polymerized gels to −80 °C storage for 1-2 h.
6. Recover the centrifuge tubes from −80 °C storage, and place in a freeze-drier, ensuring the tubes are completely dry.
7. Engage the freeze-drier with the following settings: temperature: −20 °C, pressure: 0.1 mbar.
8. Leave the gel to freeze-dry overnight.
9. Recover the gels from the freeze-drier, and place them into −80 °C storage until required.
   ​NOTE: Gels can be stored freeze-dried for up to 1 year.

4. Preparation of the 14x energy solution stock

1. Combine all the reaction components (Table 1) in a 15 mL centrifuge tube on ice to produce a 14x energy solution.
2. Aliquot the 14x energy solution into 1.5 mL microcentrifuge tubes, and store at −80 °C (smaller-volume aliquots can be used).
   ​NOTE: The 14x energy solution can be stored for several months at −80 °C if repeated freeze-thawing of the tubes is avoided.

5. Preparation of the 4x amino acid stock

1. Thaw, on ice, all 20 amino acid components of an RTS Amino Acid Sampler kit. Vortex the amino acids to ensure they are completely dissolved.
2. In a 50 mL centrifuge tube, combine all 20 amino acids, and add 12 mL of sterile ddH₂O. Vortex until the solution becomes clear, with only a slight haze of white coloration.
3. Aliquot the 4x amino acids into 1.5 mL microcentrifuge tubes (500 µL aliquots).
4. Flash-freeze the 4x amino acid solution aliquots in liquid nitrogen, and move the aliquots to −80 °C storage.

6. Cell-free buffer calibration

NOTE: The CFPS buffer used for the hydrogel reactions was calibrated for optimal DTT, Mg-glutamate, and K-glutamate concentrations following the protocol in Banks et al.³⁴ modified from Sun et al.³⁵. The calibration reaction compositions were selected using the design of experiments (DOE) method, with seven factor levels being selected for K-glutamate (200 mM, 400 mM, 600 mM, 1,000 mM, 1,200 mM, 1,400 mM), Mg-glutamate (0 mM, 10 mM, 20 mM, 30 mM, 40 mM, 50 mM, 60 mM), and DTT (0 mM, 5, 10 mM, 15 mM, 20 mM, 25 mM, 30 mM) stocks. JMP Pro 15 was used to generate a custom DOE design (Table 2) and conduct the analysis to determine the optimal factor concentrations.

1. Recover the cell-free extract from −80 °C storage, and thaw on ice.
2. Thaw the 4x amino acids, 14x energy solution, 40% PEG-8000, plasmid DNA, 3 M K-glutamate, 100 mM Mg-glutamate, and 100 mM DTT stocks on ice.
3. Create a calibration master mix by combining 12.5 µL of the 4x amino acids, 3.57 µL of the 14x energy solution, 2.5 µL of 40% PEG-8000, 3 µg of plasmid DNA, and 10 µL of cell-free extract (44.5 mg/mL), and make up to 35 µL per reaction with ddH₂O.
4. Distribute the 35 µL of master mix into the wells of a black 384-well microtiter plate.
5. Following the DOE design, add the appropriate volume of Mg-glutamate, K-glutamate, and DTT to each reaction, along with ddH₂O to make each reaction up to 50 µL.
6. Transfer the microtiter plate to a plate reader for fluorescence detection and analysis using the plate reader settings described (for mCherry): excitation: 587 nm, emission: 610 nm, wavelength bandwidth: 12 nm, optics: top, temperature: 37 °C, with fluorescence readings being collected every 5 min for 4 h of total read time. Incubate the plates with shaking on a pulsed setting at 60 rpm.
7. Upload the results into the JMP DOE design, and generate a prediction profile to determine the optimal concentration levels for K-glutamate, Mg-glutamate, and DTT based on desired response variables; in this case, the maximum fluorescence and reaction rate were the response variables.
8. Combine the reagents as shown in Table 3 to create the 2X CFPS buffer
9. Aliquot and store at −80 °C

7. CFPS in rehydrated lyophilized hydrogels (Method A)

NOTE: The plasmids used in the CFPS reactions were created using components from the EcoFlex modular toolkit for E. coli³⁶ and described in Whitfield et al.¹¹The mCherry and eGFP were under the control of the constitutive JM23100 promoter. These components are available from Addgene.

1. Recover the cell-free extract from −80 °C storage, and thaw on ice for approximately 20 min.
2. Collect the 2x CFPS buffer and plasmid DNA, and thaw on ice.
3. Collect the freeze-dried hydrogels, and allow them to reach room temperature for 15 min by leaving the gels to stand on the bench.
4. Transfer the gels to 1.5 mL microcentrifuge tubes, trimming the gels with a scalpel if necessary to make them fit in the wells.
5. In a 1.5 mL microcentrifuge tube, combine 10 µL of cell-free extract with 25 µL of 2x CFPS buffer and 4 µg of plasmid DNA; make up to a total volume of 50 µL with ddH₂O to create the CFPS solution.
6. Pipette the CFPS solution onto the freeze-dried hydrogels.
7. Allow the gels to soak in the CFPS system for 5-10 min at room temperature.
8. Transfer the gels to a black 384-well microtiter plate using a spatula.
9. Transfer the microtiter plate to a plate reader for fluorescence detection and analysis using the plate reader settings described in step 6.6. Representative results are available in previous studies^31,33.

8. Cell-free protein synthesis in deployable hydrogels (Method B)

1. Recover cell-free extract from −80 °C storage, and thaw on ice for approximately 20 min.
2. Collect the plasmid DNA, and thaw on ice.
3. In a 1.5 mL microcentrifuge tube on ice, combine 10 µL of cell-free extract with 3µg of plasmid DNA and 25 µL of 2xCFPS buffer,makingup to a total volume of 37.5 µL.
4. Measure 3g of agarose, and add to 100 mL of ddH₂O buffer to create 3% agarose.
5. Microwave the 3% agarose in 30 s bursts at high power.
6. Pipette 12.5µL of molten agarose into 1.5 mL microcentrifuge tubes containing CFPS mix to a total volume of 50 µL.
7. Allow the molten agarose to cool, but not polymerize, leaving the agarose in a heat block set to 50 °C.
8. Combine the molten agarose with the CFPS solution; mix via pipetting and stirring with the tip, and ensure to move quickly to avoid gel polymerization before the CFPS solution can be mixed in.
9. Allow the gels to cool at room temperature and polymerize for approximately 2 min.
10. Transfer the polymerized agarose to 1.5 mL microcentrifuge tubes with a spatula, and flash-freeze in liquid nitrogen.
11. Place the flash-frozen hydrogels into −80 °C storage for 1 h.
12. Recover the gels from storage; remove the microcentrifuge tube lids, cover the tubes with wax film, and pierce the film to allow the moisture to be dried off.
13. Engage the freeze-drier with the following settings: temperature: −20 °C, pressure: 0.1 mbar, freeze drying for 18 h (overnight).
14. Store the freeze-dried CFPS devices in −80 °C storage until use.
15. The freeze-dried CFPS devices, with an approximate wet volume of 50 µL, can be rehydrated with 50 µL of ddH₂O without excess liquid. Rehydration takes approximately 30 min.
16. Transfer the gels to a black 384-well microtiter plate using a spatula.
17. Transfer the microtiter plate to a plate reader for fluorescence detection and analysis using the plate reader settings described in step 6.6. Representative results are available in previous publications^31,33.

This protocol details two methods for embedding CFPS reactions into hydrogel matrices, with Figure 1 presenting a schematic overview of the two approaches. Both methods are amenable to freeze-drying and long-term storage. Method A is the most utilized methodology for two reasons. First, it has been shown to be the most applicable method for working with a range of hydrogel materials¹¹.Second, Method A allows for the parallel testing of genetic constructs. Method B is more appropriate for the fabrication of an optimized system and field deployment. Both protocols allow many samples to be prepared in one go to aid in experimental reproducibility. This feature is also useful for the long-term development of the technology, as freeze-dried devices may be shipped in a dry state and reconstituted on site when needed.

The approach outlined in the protocol and Figure 1 can be used for the expression of single gene constructs or for the co-expression of multiple genes. The data presented in Figure 2 show the expression of both eGFP and mCherry in a 0.75% agarose gel. Confocal microscopy was used to confirm that protein expression was hom*ogenous throughout the hydrogel, including within the internal planes. Specifically, protein synthesis was not confined to the outer edges of the hydrogel, and internal fluorescence was not the result of protein diffusion. To confirm this, by placing an eGFP-expressing hydrogel in physical contact with an mCherry-expressing hydrogel, it was possible to see protein diffusion from one hydrogel to another. The rate of diffusion between the two was insufficient to explain the extensive localization of either red or green fluorescence inside the material. This experiment also illustrates a key advantage of deploying cell-free devices in hydrogels-the device functionality can be spatially organized in a manner that is not possible in liquid cell-free reactions. In addition, for the creation of gene networks, the simultaneous synthesis of more than a single gene product is needed. The results shown in Figure 2 (bottom row) confirmed the co-expression of both mCherry and eGFP in agarose. In this work, both proteins were expressed, and there was no spatial competition between the proteins. Again, an overlay of the red and green wavelength range demonstrates the even spatial distribution of both proteins within the hydrogel.

Table 1: The 14x energy solution stocks. Please click here to download this Table.

Table 2: Design of an experimental array for the optimization of DTT, Mg-glutamate, and K-glutamate within the cell-free protein synthesis reactions. Please click here to download this Table.

Table 3: The 2x CFPS buffer components. Please click here to download this Table.

Figure 1: Schematic of the two protocols. In the first method (Method A, demonstrated in this paper) hydrogel materials are prepared first and then freeze-dried (step 1) without cell-free components. These dried hydrogels can be stored and reconstituted when required (step 2) with the correct volume of cell-free reaction prior to incubation for protein production (step 3) The variant method, Method B, incorporates all, or some, of the cell-free reaction components in the initial hydrogel fabrication. Following freeze-drying (step 1), the hydrogels may then be reconstituted in water alone or in buffer containing an analyte of interest (step 2). Protein production (step 3) continues as before. A third method, in which freeze-dried cell-free components are reconstituted with hydrogel polymers, is described in Whitfield et al.¹¹ but has found use with only a limited number of hydrogels to date. Please click here to view a larger version of this figure.

Figure 2: Cell-free protein synthesis of eGFP and mCherry in a hydrogel using E. coli cell lysates. Agarose gels (0.75%) were prepared without DNA template (top) with 4 µg of either eGFP or mCherry template (middle) or with 4 µg of both eGFP and mCherry template (bottom). The hydrogels were incubated for 4 h before confocal microscopy in the red and green channels. An overlay of the two channels is also shown, and the overlay includes the differential interference contrast (DIC) image. Hydrogels containing either eGFP or mCherry template were prepared separately but incubated in physical contact with each other. The gel diameter is 6 mm. Please click here to view a larger version of this figure.

Outlined here are two protocols for the incorporation of E. coli cell lysate-based CFPS reactions into agarose hydrogels. These methods allow for simultaneous gene expression throughout the material. The protocol can be adapted for other CFPS systems and has been successfully conducted with commercially available CFPS kits in addition to the laboratory-prepared cell lysates detailed here. Importantly, the protocol allows for gene expression in the absence of an external liquid phase. This means that the system is self-contained and does not require a cell-free reaction bath. Distinct from previous methods in which CFPS reactions occurred within hydrogels, the requirement for external buffers and reaction vessels is removed with this method. It is anticipated that these methods will allow researchers to explore the use of hydrogel materials as chassis for cell-free synthetic biology devices, ultimately providing a route for moving such devices out of the laboratory.

Critical to the success of both approaches is the use of high-quality cell lysates. A high-quality cell lysate should have a high protein concentration (>40 mg/mL), should be kept cold for the duration of preparation, and should not have undergone repeated freeze-thaw cycles. In addition, it is critical that the DNA templates are of high purity. To achieve this, plasmids should be extracted from cells using a high-purity, high-yield plasmid preparation kit for transcriptional reactions to proceed within the CFPS environment. These critical stages are common to all CFPS applications and are not unique to this method. Nonetheless, it is the preparation of CFPS components that has the biggest impact on the effectiveness of the protocols. In terms of the materials, effective, rapid freeze-drying of the hydrogels needs to be achieved. If the freeze-drier does not reach a sufficiently cold temperature (below −45 °C), the gel will be dried rather than freeze-dried, which can compromise the internal structure of the hydrogel matrix and may cause the degradation of the embedded CFPS systems. Method B also has a unique concern; specifically, when adding the CFPS mix to the molten agarose to form the gel, the molten agarose must be left to cool sufficiently before the CFPS mixture is added to prevent thermal degradation to the CFPS reactions. However, allowing the agarose to cool too much will result in the polymerization of the gel and prevent the addition of the CFPS system. Finally, hydrogels possess a degree of autofluorescence, which can conflict with the fluorescence signals generated during the CFPS reactions. It is important, therefore, to include appropriate negative controls and genetic reporters that do not have the same fluorescence characteristics as the material.

The protocols demonstrated here center on the use of agarose as a model hydrogel. Agarose is an excellent model system for this work as it is widely available, inexpensive, and demonstrates excellent protein production capabilities. Previous investigations have also demonstrated that agarose can be a substitute for the crowding agent PEG in CFPS reactions, indicating that the interactions between the polymer chassis and the CFPS reactions may enhance the protein production¹¹. These methods, however, are also amenable to modifications using a wide range of alternative hydrogel matrices with varying physical and chemical properties. The method has been applied to xanthan gum, alginate, acyl gellan gum, polyacrylamide, hyaluronic acid hydrogels, and the poloxamer gels F-108 and F-127⁹. In some scenarios, reduced protein production is observed compared to liquid controls or other hydrogels. Nonetheless, in these scenarios, a trade-off may be met, where the functionality of the material itself (e.g., its adhesive properties or biocompatibility) is of significant benefit such that a reduction in the protein production may be accepted. Modifications to the concentrations of the hydrogels can also be made. For agarose, the methods are amenable to polymer concentrations in the range of 0.5%-1.5% w/v, for example. A higher gel concentration typically results, in a material sense, in a more robust device that is more resilient to handling and better preserves the CFPS reaction within. The trade-off with increased gel concentration, however, is that the reaction may have a reduced production rate or protein yield^11,37. The relationship between the polymer concentration and protein production, however, is not linear and varies depending on the hydrogel used¹¹. As such, for any new hydrogel polymer, a degree of experimentation is required to balance material functionality against cell-free functionality.

Embedding CFPS reactions in hydrogel materials without the need for external buffers represents an interesting route for the deployment of cell-free devices. Cell-free devices carry the benefit of removing the need for the release of genetically engineered organisms outside a controlled laboratory environment. In addition, embedding reactions in hydrogels removes the need for reaction vessels (typically plasticware) following reconstitution. Given that many hydrogels are biodegradable, this offers a potential route for reducing plastic waste. Likewise, the protocol detailed here also demonstrates that both hydrogel and CFPS are amenable to freeze-drying. While repeated cycles of lyophilization are not recommended and will likely damage the CFPS and hydrogel function, for single-use devices. The ability to freeze-dry and reconstitute the components reduces the device weight and removes the requirement for a cold chain during transportation. These properties ultimately simplify the logistics and reduce the transportation costs for the devices. Further, hydrogels have many useful functional properties of their own; for example, they can act as pastes, lubricants, and adhesives. Hydrogels may also be biocompatible, biodegradable, and possess stimuli-responses in their own right. Taken together, a synergy can be achieved between biological functionality and materials science to create a new class of bioprogrammable materials.