The authors greatly acknowledge the support of the Biotechnology and Biological Sciences Research Council awards BB/V017551/1 (S.K., T.P.H.) and BB/W01095X/1 (A.L., T.P.H.), and the Engineering and Physical Sciences Research Council - Defence Science and Technology Laboratories award EP/N026683/1 (C.J.W., A.M.B., T.P.H.). Data supporting this publication are openly available at: 10.25405/data.ncl.22232452. For the purpose of open access, the author has applied a Creative Commons Attribution (CC BY) license to any Author Accepted Manuscript version arising.

1. Cell lysate buffer and media preparation
<ol>
	<li>Preparation of 2x YT+P agar and medium
	<ol>
		<li>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.</li>
		<li>Sterilize by autoclaving the 2x YT+P.</li>
	</ol>
	</li>
	<li>Preparation of the S30A buffer
	<ol>
		<li>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.</li>
		<li>Store the S30A buffer at 4 &#176;C for up to 1 week.</li>
	</ol>
	</li>
	<li>Preparation of the S30B buffer
	<ol>
		<li>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.</li>
		<li>Store the S30B buffer at 4 &#176;C for up to 1 week.</li>
	</ol>
	</li>
</ol>
2. Cell lysate preparation (4 day protocol)
<ol>
	<li>Day 1
	<ol>
		<li>Pour the 2x YT+P into agar plates supplemented with 100 &#181;g/mL ampicillin.</li>
		<li>Streak E. coli from &#8722;80 &#176;C glycerol stock, and incubate overnight at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Day 2
	<ol>
		<li>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.</li>
		<li>Incubate for 24 h at 37 &#176;C with 220 rpm shaking.</li>
	</ol>
	</li>
	<li>Day 3
	<ol>
		<li>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.</li>
		<li>Transfer the culture evenly between pre-chilled 500 mL centrifuge bottles.</li>
		<li>Centrifuge at 4,500 x g for 15 min at 4 &#176;C, and then discard the supernatant.</li>
		<li>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.</li>
		<li>Resuspend the cell pellets in 200 mL of S30A buffer.</li>
		<li>Vortex the bottles vigorously until the whole pellet is completely solubilized, keeping the cells on ice as much as possible.</li>
		<li>Centrifuge at 4,500 x g for 15 min at 4 &#176;C, and then discard the supernatant.</li>
		<li>Repeat steps 2.3.5-2.3.7 (inclusive).</li>
		<li>Add 40 mL of S30A buffer to each centrifuge bottle, resuspend the pellets, and transfer to pre-weighed 50 mL centrifuge tubes.</li>
		<li>Centrifuge at 4,000 x g for 15 min at 4 &#176;C. Discard the supernatant by decanting, and remove the residual supernatant by pipette, keeping it on ice as much as possible.</li>
		<li>Re-weigh the centrifuge tubes, record the new mass, and calculate the mass of the pellets.</li>
		<li>Flash-freeze pellets in liquid nitrogen, and store them at &#8722;80 &#176;C.</li>
	</ol>
	</li>
	<li>Day 4
	<ol>
		<li>Thaw the pellets on ice.</li>
		<li>Per 1 g of pellet, add the following: 1.2 mL of S30A buffer (DTT added before use), 275 &#181;L of protease inhibitor (8 mM stock), and 25 &#181;L of lysozyme (10 mg/mL stock).</li>
		<li>Vortex vigorously until the pellets are completely solubilized with no remaining clumps, keeping them on ice as much as possible.</li>
		<li>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.</li>
		<li>Centrifuge at 4,000 x g for 1 h at 4 &#176;C to pellet the cell debris.</li>
		<li>Transfer the supernatant to fresh 50 mL centrifuge tubes.</li>
		<li>Incubate tubes at 37 &#176;C at 220 rpm for 80 min.</li>
		<li>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 &#176;C.</li>
		<li>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 &#176;C.</li>
		<li>Consolidate the pellet-free supernatant on ice in a 50 mL centrifuge tube.</li>
		<li>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.</li>
		<li>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 &#176;C for 3 h.</li>
		<li>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 &#176;C.</li>
		<li>Consolidate the clarified extract into a fresh centrifuge tube on ice, and vortex briefly to mix.</li>
		<li>Determine the protein concentration of the extract using a fluorometer.</li>
		<li>Dilute the extract to a concentration of 44.5 mg/mL using pre-chilled ddH2O.</li>
		<li>Aliquot into 200 &#181;L aliquots, flash-freeze in liquid nitrogen, and store at &#8722;80 &#176;C.</li>
	</ol>
	</li>
</ol>
3. Preparation of lyophilized hydrogels (Method A)
<ol>
	<li>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.</li>
	<li>Using a pipette, transfer 50 &#181;L 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).</li>
	<li>Flash-freeze the microcentrifuge tubes, containing the polymerized gels, in liquid nitrogen.</li>
	<li>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.</li>
	<li>Transfer the microcentrifuge tubes containing the polymerized gels to &#8722;80 &#176;C storage for 1-2 h.</li>
	<li>Recover the centrifuge tubes from &#8722;80 &#176;C storage, and place in a freeze-drier, ensuring the tubes are completely dry.</li>
	<li>Engage the freeze-drier with the following settings: temperature: &#8722;20 &#176;C, pressure: 0.1 mbar.</li>
	<li>Leave the gel to freeze-dry overnight.</li>
	<li>Recover the gels from the freeze-drier, and place them into &#8722;80 &#176;C storage until required. 
	&#8203;NOTE: Gels can be stored freeze-dried for up to 1 year.</li>
</ol>
4. Preparation of the 14x energy solution stock
<ol>
	<li>Combine all the reaction components (Table 1) in a 15 mL centrifuge tube on ice to produce a 14x energy solution.</li>
	<li>Aliquot the 14x energy solution into 1.5 mL microcentrifuge tubes, and store at &#8722;80 &#176;C (smaller-volume aliquots can be used). 
	&#8203;NOTE: The 14x energy solution can be stored for several months at &#8722;80 &#176;C if repeated freeze-thawing of the tubes is avoided.</li>
</ol>
5. Preparation of the 4x amino acid stock
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Aliquot the 4x amino acids into 1.5 mL microcentrifuge tubes (500 &#181;L aliquots).</li>
	<li>Flash-freeze the 4x amino acid solution aliquots in liquid nitrogen, and move the aliquots to &#8722;80 &#176;C storage.</li>
</ol>
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.
<ol>
	<li>Recover the cell-free extract from &#8722;80 &#176;C storage, and thaw on ice.</li>
	<li>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.</li>
	<li>Create a calibration master mix by combining 12.5 &#181;L of the 4x amino acids, 3.57 &#181;L of the 14x energy solution, 2.5 &#181;L of 40% PEG-8000, 3 &#181;g of plasmid DNA, and 10 &#181;L of cell-free extract (44.5 mg/mL), and make up to 35 &#181;L per reaction with ddH2O.</li>
	<li>Distribute the 35 &#181;L of master mix into the wells of a black 384-well microtiter plate.</li>
	<li>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 &#181;L.</li>
	<li>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 &#176;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.</li>
	<li>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.</li>
	<li>Combine the reagents as shown in Table 3 to create the 2X CFPS buffer</li>
	<li>Aliquot and store at &#8722;80 &#176;C</li>
</ol>
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.11&#160;The mCherry and eGFP were under the control of the constitutive JM23100 promoter. These components are available from Addgene.
<ol>
	<li>Recover the cell-free extract from &#8722;80 &#176;C storage, and thaw on ice for approximately 20 min.</li>
	<li>Collect the 2x CFPS buffer and plasmid DNA, and thaw on ice.</li>
	<li>Collect the freeze-dried hydrogels, and allow them to reach room temperature for 15 min by leaving the gels to stand on the bench.</li>
	<li>Transfer the gels to 1.5 mL microcentrifuge tubes, trimming the gels with a scalpel if necessary to make them fit in the wells.</li>
	<li>In a 1.5 mL microcentrifuge tube, combine 10 &#181;L of cell-free extract with 25 &#181;L of 2x CFPS buffer and 4 &#181;g of plasmid DNA; make up to a total volume of 50 &#181;L with ddH2O to create the CFPS solution.</li>
	<li>Pipette the CFPS solution onto the freeze-dried hydrogels.</li>
	<li>Allow the gels to soak in the CFPS system for 5-10 min at room temperature.</li>
	<li>Transfer the gels to a black 384-well microtiter plate using a spatula.</li>
	<li>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 studies31,33.</li>
</ol>
8. Cell-free protein synthesis in deployable hydrogels (Method B)
<ol>
	<li>Recover cell-free extract from &#8722;80 &#176;C storage, and thaw on ice for approximately 20 min.</li>
	<li>Collect the plasmid DNA, and thaw on ice.</li>
	<li>In a 1.5 mL microcentrifuge tube on ice, combine 10 &#181;L of cell-free extract with 3&#160;&#181;g of plasmid DNA and 25 &#181;L of 2x&#160;CFPS buffer,&#160;making&#160;up to a total volume of 37.5 &#181;L.</li>
	<li>Measure 3&#160;g of agarose, and add to 100 mL of ddH2O buffer to create 3% agarose.</li>
	<li>Microwave the 3% agarose in 30 s bursts at high power.</li>
	<li>Pipette 12.5&#160;&#181;L of molten agarose into 1.5 mL microcentrifuge tubes containing CFPS mix to a total volume of 50 &#181;L.</li>
	<li>Allow the molten agarose to cool, but not polymerize, leaving the agarose in a heat block set to 50 &#176;C.</li>
	<li>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.</li>
	<li>Allow the gels to cool at room temperature and polymerize for approximately 2 min.</li>
	<li>Transfer the polymerized agarose to 1.5 mL microcentrifuge tubes with a spatula, and flash-freeze in liquid nitrogen.</li>
	<li>Place the flash-frozen hydrogels into &#8722;80 &#176;C storage for 1 h.</li>
	<li>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.</li>
	<li>Engage the freeze-drier with the following settings: temperature: &#8722;20 &#176;C, pressure: 0.1 mbar, freeze drying for 18 h (overnight).</li>
	<li>Store the freeze-dried CFPS devices in &#8722;80 &#176;C storage until use.</li>
	<li>The freeze-dried CFPS devices, with an approximate wet volume of 50 &#181;L, can be rehydrated with 50 &#181;L of ddH2O without excess liquid. Rehydration takes approximately 30 min.</li>
	<li>Transfer the gels to a black 384-well microtiter plate using a spatula.</li>
	<li>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 publications31,33.</li>
</ol>

Embedding CFPS Reactions in a Variety of Hydrogel Matrices

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<li>Salati, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Agarose-based+biomaterials%3A+Opportunities+and+challenges+in+cartilage+tissue+engineering%5BTitle%5D)+AND+%22Polymers%22%5BJournal%5D)">Agarose-based biomaterials: Opportunities and challenges in cartilage tissue engineering.</a> Polymers. 12 (5), 1150(2020).</li>
<li>Buddingh, B. C., Van Hest, J. C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Artificial+cells%3A+Synthetic+compartments+with+life-like+functionality+and+adaptivity%5BTitle%5D)+AND+%22Accounts+of+Chemical+Research%22%5BJournal%5D)">Artificial cells: Synthetic compartments with life-like functionality and adaptivity.</a> Accounts of Chemical Research. 50 (4), 769-777 (2017).</li>
<li>Kahn, J. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((DNA+microgels+as+a+platform+for+cell-free+protein+expression+and+display%5BTitle%5D)+AND+%22Biomacromolecules%22%5BJournal%5D)">DNA microgels as a platform for cell-free protein expression and display.</a> Biomacromolecules. 17 (6), 2019-2026 (2016).</li>
<li>Yang, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Enhanced+transcription+and+translation+in+clay+hydrogel+and+implications+for+early+life+evolution%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Enhanced transcription and translation in clay hydrogel and implications for early life evolution.</a> Scientific Reports. 3, 3165(2013).</li>
<li>Zhou, X., Wu, H., Cui, M., Lai, S. N., Zheng, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long-lived+protein+expression+in+hydrogel+particles%3A+Towards+artificial+cells%5BTitle%5D)+AND+%22Chemical+Science%22%5BJournal%5D)">Long-lived protein expression in hydrogel particles: Towards artificial cells.</a> Chemical Science. 9 (18), 4275-4279 (2018).</li>
<li>Whitfield, C. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cell-free+genetic+devices+confer+autonomic+and+adaptive+properties+to+hydrogels%5BTitle%5D)+AND+%22BioRxiv%22%5BJournal%5D)">Cell-free genetic devices confer autonomic and adaptive properties to hydrogels.</a> BioRxiv. , (2019).</li>
<li>Feng, L., Jianpu, T., Jinhui, G. D., Luo, D. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Polymeric+DNA+hydrogel%3A+Design%2C+synthesis+and+applications%5BTitle%5D)+AND+%22Progress+in+Polymer+Science%22%5BJournal%5D)">Polymeric DNA hydrogel: Design, synthesis and applications.</a> Progress in Polymer Science. 98, 101163(2019).</li>
<li>Howard, T., et al. Datasets for Whitfield et al. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aHoward%2C+T+%222020+Chemical+Communications%22">2020 Chemical Communications</a>. , Newcastle University. UK. (2020).</li>
<li>Banks, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Key+reaction+components+affect+the+kinetics+and+performance+robustness+of+cell-free+protein+synthesis+reactions%5BTitle%5D)+AND+%22Computational+and+Structural+Biotechnology+Journal%22%5BJournal%5D)">Key reaction components affect the kinetics and performance robustness of cell-free protein synthesis reactions.</a> Computational and Structural Biotechnology Journal. 20, 218-229 (2022).</li>
<li>Sun, Z. Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Protocols+for+implementing+an+Escherichia+coli-based+TX-TL+cell-free+expression+system+for+synthetic+biology%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%22%5BJournal%5D)">Protocols for implementing an Escherichia coli-based TX-TL cell-free expression system for synthetic biology.</a> Journal of Visualized Experiments. (79), e50762(2013).</li>
<li>Moore, S. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((EcoFlex%3A+A+multifunctional+MoClo+kit+for+E.+coli+synthetic+biology%5BTitle%5D)+AND+%22ACS+Synthetic+Biology%22%5BJournal%5D)">EcoFlex: A multifunctional MoClo kit for E. coli synthetic biology.</a> ACS Synthetic Biology. 5 (10), 1059-1069 (2016).</li>
<li>Benítez-Mateos, A. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Micro+compartmentalized+cell-free+protein+synthesis+in+hydrogel+%C2%B5-channels%5BTitle%5D)+AND+%22ACS+Synthetic+Biology%22%5BJournal%5D)">Micro compartmentalized cell-free protein synthesis in hydrogel µ-channels.</a> ACS Synthetic Biology. 9 (11), 2971-2978 (2020).</li>
</ol>

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 (&#62;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 &#8722;45 &#176;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 yield11,37. 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.

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 methods1,2,3. CFPS is being used in an increasing number of applications across numerous disciplines, including constructing artificial cells, prototyping genetic circuits, developing biosensors, and producing metabolites4,5,6. 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 proteins6,7,8.
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 properties9,10,11. 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 diapers9,11,12,13,14. Hydrogels are also under active investigation as drug delivery vehicles9,15,16,17. Hydrogels may also be biocompatible, biodegradable, and possess some stimuli responses of their own9,18,19,20. 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 agarose11,21,22, as well as to coat surfaces of glass, paper, and cloth11,23,24 with CFPS devices. The protocols presented here demonstrate two methods for embedding CFPS reactions in macro-scale (i.e., &#62;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 properties11,24,25,26. 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 &#181;m to 400 &#181;m have been used to perform CFPS in hydrogels submerged in reaction buffer23,27,28,29,30,31. 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.

<table><tbody><tr><td>&lt;strong&gt;Material&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>3-PGA</td><td>Santa Cruz Biotechnology</td><td>sc-214793B</td><td></td></tr><tr><td>Acetic Acid</td><td>Sigma-Aldrich</td><td>A6283</td><td></td></tr><tr><td>Agar</td><td>Thermo Fisher Scientific</td><td>A10752.22</td><td></td></tr><tr><td>Agarose</td><td>Severn Biotech</td><td>30-15-50</td><td></td></tr><tr><td>Amino Acid Sampler Kit</td><td>VWR</td><td>BTRABR1401801</td><td></td></tr><tr><td>ATP</td><td>Sigma-Aldrich</td><td>A8937-1G</td><td></td></tr><tr><td>cAMP</td><td>Sigma-Aldrich</td><td>A9501-1G</td><td></td></tr><tr><td>Coenzyme A (CoA)</td><td>Sigma-Aldrich</td><td>C4282-100MG</td><td></td></tr><tr><td>CTP</td><td>Alfa Aesar</td><td>J14121.MC</td><td></td></tr><tr><td>DTT</td><td>Thermo Fisher Scientific</td><td>R0862</td><td></td></tr><tr><td>Folinic Acid</td><td>Sigma-Aldrich</td><td>F7878-100MG</td><td></td></tr><tr><td>GTP</td><td>Carbosynth</td><td>NG01208</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034-25G</td><td></td></tr><tr><td>K-glutamate</td><td>Sigma-Aldrich</td><td>G1149-100G</td><td></td></tr><tr><td>Lysozyme</td><td>Sigma-Aldrich</td><td>L6876-1G</td><td></td></tr><tr><td>Mg-glutamate</td><td>Sigma-Aldrich</td><td>49605-250G</td><td></td></tr><tr><td>NAD</td><td>Sigma-Aldrich</td><td>N6522-250MG</td><td></td></tr><tr><td>PEG-8000</td><td>Promega</td><td>V3011</td><td></td></tr><tr><td>Potassium Hydroxide (KOH)</td><td>Sigma-Aldrich</td><td>757551-5G</td><td></td></tr><tr><td>Potassium Phosphate Dibasic (K2HPO4)</td><td>Sigma-Aldrich</td><td>P3786-500G</td><td></td></tr><tr><td>Potassium Phosphate Monobasic (KH2PO4)</td><td>Sigma-Aldrich</td><td>RDD037-500G</td><td></td></tr><tr><td>Protease Inhibitor cocktail</td><td>Sigma-Aldrich</td><td>P2714-1BTL</td><td></td></tr><tr><td>Qubit Protein concentration kit</td><td>Thermo Fisher Scientific</td><td>A50668</td><td></td></tr><tr><td>Rossetta 2 DE 3 E.coli</td><td>Sigma-Aldrich</td><td>71397-3</td><td></td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Sigma-Aldrich</td><td>S9888-500G</td><td></td></tr><tr><td>Spermidine</td><td>Sigma-Aldrich</td><td>85558-1G</td><td></td></tr><tr><td>Tryptone</td><td>Thermo Fisher Scientific</td><td>211705</td><td></td></tr><tr><td>Tris</td><td>Sigma-Aldrich</td><td>GE17-1321-01</td><td></td></tr><tr><td>tRNA</td><td>Sigma-Aldrich</td><td>10109541001</td><td></td></tr><tr><td>UTP</td><td>Alfa Aesar</td><td>J23160.MC</td><td></td></tr><tr><td>Yeast Extract</td><td>Sigma-Aldrich</td><td>Y1625-1KG</td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1.5 mL microcentrifuge tubes</td><td>Sigma-Aldrich</td><td>HS4323-500EA</td><td></td></tr><tr><td>10K MWCO dialysis cassettes</td><td>Thermo Fisher Scientific</td><td>66381</td><td></td></tr><tr><td>15 mL centrifuge tube</td><td>Sarstedt</td><td>62.554.502</td><td></td></tr><tr><td>50 mL centrifuge bottles</td><td>Sarstedt</td><td>62.547.254</td><td></td></tr><tr><td>500 mL centrifuge bottles</td><td>Thermo Fisher Scientific</td><td>3120-9500</td><td></td></tr><tr><td>Alpha 1-2 LD Plus freeze-dryer</td><td>Christ</td><td>part no. 101521, 101522, 101527</td><td></td></tr><tr><td>Benchtop Centrifuge</td><td>Thermo Fisher Scientific</td><td>H-X3R</td><td></td></tr><tr><td>Black 384 well microtitre plates</td><td>Fischer Scientific</td><td>66</td><td></td></tr><tr><td>Cuvettes</td><td>Thermo Fisher Scientific</td><td>222S</td><td></td></tr><tr><td>Elga Purelab Chorus</td><td>Elga</td><td>#####</td><td></td></tr><tr><td>Eppendorf Microcentrifuge 5425R</td><td>Eppendorf</td><td>EP00532</td><td></td></tr><tr><td>High Speed Centrifuge</td><td>Beckman Coulter</td><td>B34183</td><td></td></tr><tr><td>JMP license</td><td>SAS Institute</td><td>15</td><td></td></tr><tr><td>Magnetic Stirrer</td><td>Fischer Scientific</td><td>15353518</td><td></td></tr><tr><td>Parafilm</td><td>Amcor</td><td>PM-966</td><td></td></tr><tr><td>Photospectrometer (Biophotometer)</td><td>Eppendorf</td><td>16713</td><td></td></tr><tr><td>Pipettes and tips</td><td>Gilson</td><td>#####</td><td></td></tr><tr><td>Precision Balance</td><td>Sartorius</td><td>16384738</td><td></td></tr><tr><td>Qubit 2.0 Fluorometer</td><td>Thermo Fisher Scientific</td><td>Q32866</td><td></td></tr><tr><td>Shaking Incubator</td><td>Thermo Fisher Scientific</td><td>SHKE8000</td><td></td></tr><tr><td>Sonic Dismembrator (Sonicator)</td><td>Thermo Fisher Scientific</td><td>12893543</td><td></td></tr><tr><td>Static Incubator</td><td>Sanyo</td><td>MIR-162</td><td></td></tr><tr><td>Syringe and needles</td><td>Thermo Fisher Scientific</td><td>66490</td><td></td></tr><tr><td>Thermo max Q8000 (Shaking Incubator)</td><td>Thermo Fisher Scientific</td><td>SHKE8000</td><td></td></tr><tr><td>Varioskan Lux platereader</td><td>Thermo Fisher Scientific</td><td>VLBL00GD1</td><td></td></tr><tr><td>Vortex Genie 2</td><td>Cole-parmer</td><td>OU-04724-05</td><td></td></tr><tr><td>VWR PHenomenal pH 1100 L, ph/mv/&amp;deg;c meter</td><td>VWR</td><td>662-1657</td><td></td></tr></tbody></table>

methods for embedding cell-free protein synthesis reactions in macro-scale hydrogels

Synthetic gene networks provide a platform for scientists and engineers to design and build novel systems with functionality encoded at a genetic level. While the dominant paradigm for the deployment of gene networks is within a cellular chassis, synthetic gene networks may also be deployed in cell-free environments. Promising applications of cell-free gene networks include biosensors, as these devices have been demonstrated against biotic (Ebola, Zika, and SARS-CoV-2 viruses) and abiotic (heavy metals, sulfides, pesticides, and other organic contaminants) targets. Cell-free systems are typically deployed in liquid form within a reaction vessel. Being able to embed such reactions in a physical matrix, however, may facilitate their broader application in a wider set of environments. To this end, methods for embedding cell-free protein synthesis (CFPS) reactions in a variety of hydrogel matrices have been developed. One of the key properties of hydrogels conducive to this work is the high-water reconstitution capacity of hydrogel materials. Additionally, hydrogels possess physical and chemical characteristics that are functionally beneficial. Hydrogels can be freeze-dried for storage and rehydrated for use later. Two step-by-step protocols for the inclusion and assay of CFPS reactions in hydrogels are presented. First, a CFPS system can be incorporated into a hydrogel via rehydration with a cell lysate. The system within the hydrogel can then be induced or expressed constitutively for complete protein expression through the hydrogel. Second, cell lysate can be introduced to a hydrogel at the point of polymerization, and the entire system can be freeze-dried and rehydrated at a later point with an aqueous solution containing the inducer for the expression system encoded within the hydrogel. These methods have the potential to allow for cell-free gene networks that confer sensory capabilities to hydrogel materials, with the potential for deployment beyond the laboratory.

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.&#160;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 homogenous 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. <a href="https://www.jove.com/files/ftp_upload/65500/JOVE_KAVIL Table 1 14X energy solution stock solutions.xlsx" target="_blank">Please click here to download this Table.</a>
Table 2: Design of an experimental array for the optimization of DTT, Mg-glutamate, and K-glutamate within the cell-free protein synthesis reactions. <a href="https://www.jove.com/files/ftp_upload/65500/JOVE_KAVIL Table 2 Designed Experimental Array.xlsx" target="_blank">Please click here to download this Table.</a>
Table 3: The 2x CFPS buffer components. <a href="https://www.jove.com/files/ftp_upload/65500/JOVE_KAVIL Table 3 2X CFPS Buffer Components.xlsx" target="_blank">Please click here to download this Table.</a>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65500/65500fig01.jpg" /> 
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. <a href="https://www.jove.com/files/ftp_upload/65500/65500fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65500/65500fig02v2.jpg" /> 
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 &#181;g of either eGFP or mCherry template (middle) or with 4 &#181;g 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. <a href="https://www.jove.com/files/ftp_upload/65500/65500fig02largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Novel Approach for Embedding Cell-Free Protein Synthesis Reactions in Hydrogels at JoVE.com

Author Spotlight: A Novel Approach for Embedding Cell-Free Protein Synthesis Reactions in Hydrogels 

Here, we present two protocols for embedding cell-free protein synthesis reactions in macro-scale hydrogel matrices without the need for an external liquid phase.

Methods for Embedding Cell-Free Protein Synthesis Reactions in Macro-Scale Hydrogels

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Research

JoVE Journal

Biology

1.1K Views.  Newcastle University. Here, we present two protocols for embedding cell-free protein synthesis reactions in macro-scale hydrogel matrices without the need for an external liquid phase.

Video: Author Spotlight: A Novel Approach for Embedding Cell-Free Protein Synthesis Reactions in Hydrogels 

Microfabricated Platforms for Mechanically Dynamic Cell Culture

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or fundamental cell biology studies is limited by current bioreactor technologies, which cannot simultaneously apply a variety of mechanical stimuli to cultured cells. In order to address this issue, we have developed a series of microfabricated platforms designed to screen for the effects of mechanical stimuli in a high-throughput format. In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and further demonstrate how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

In this protocol, we demonstrate the fabrication of a microactuator array of vertically displaced posts on which the technology is based, and how this base technology can be modified to conduct high-throughput mechanically dynamic cell culture in both two-dimensional and three-dimensional culture paradigms.

The ability to systematically probe in vitro cellular response to combinations of mechanobiological stimuli for tissue engineering, drug discovery or ...

Bioengineering

Fabrication of Micropatterned Hydrogels for Neural Culture Systems using Dynamic Mask Projection Photolithography

Increasingly, patterned cell culture environments are becoming a relevant technique to study cellular characteristics, and many researchers believe in the need for 3D environments to represent in vitro experiments which better mimic in vivo qualities 1-3. Studies in fields such as cancer research 4, neural engineering 5, cardiac physiology 6, and cell-matrix interaction7,8have shown cell behavior differs substantially between traditional monolayer cultures and 3D constructs.
Hydrogels are used as 3D environments because of their variety, versatility and ability to tailor molecular composition through functionalization 9-12. Numerous techniques exist for creation of constructs as cell-supportive matrices, including electrospinning13, elastomer stamps14, inkjet printing15, additive photopatterning16, static photomask projection-lithography17, and dynamic mask microstereolithography18. Unfortunately, these methods involve multiple production steps and/or equipment not readily adaptable to conventional cell and tissue culture methods. The technique employed in this protocol adapts the latter two methods, using a digital micromirror device (DMD) to create dynamic photomasks for crosslinking geometrically specific poly-(ethylene glycol) (PEG) hydrogels, induced through UV initiated free radical polymerization. The resulting "2.5D" structures provide a constrained 3D environment for neural growth. We employ a dual-hydrogel approach, where PEG serves as a cell-restrictive region supplying structure to an otherwise shapeless but cell-permissive self-assembling gel made from either Puramatrix or agarose. The process is a quick simple one step fabrication which is highly reproducible and easily adapted for use with conventional cell culture methods and substrates.
Whole tissue explants, such as embryonic dorsal root ganglia (DRG), can be incorporated into the dual hydrogel constructs for experimental assays such as neurite outgrowth. Additionally, dissociated cells can be encapsulated in the photocrosslinkable or self polymerizing hydrogel, or selectively adhered to the permeable support membrane using cell-restrictive photopatterning. Using the DMD, we created hydrogel constructs up to ~1mm thick, but thin film (&lt;200 &mu;m) PEG structures were limited by oxygen quenching of the free radical polymerization reaction. We subsequently developed a technique utilizing a layer of oil above the polymerization liquid which allowed thin PEG structure polymerization.
In this protocol, we describe the expeditious creation of 3D hydrogel systems for production of microfabricated neural cell and tissue cultures. The dual hydrogel constructs demonstrated herein represent versatile in vitro models that may prove useful for studies in neuroscience involving cell survival, migration, and/or neurite growth and guidance. Moreover, as the protocol can work for many types of hydrogels and cells, the potential applications are both varied and vast.

Simple techniques are described for the rapid production of microfabricated neural culture systems using a digital micromirror device for dynamic mask projection lithography on regular cell culture substrates. These culture systems may be more representative of natural biological architecture, and the techniques described could be adapted for numerous applications.

Increasingly, patterned cell culture environments are becoming a relevant technique to study cellular characteristics, and many researchers believe in the ...

Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field of 3D templates is rising due to the potential resemblance of those materials to the natural extracellular matrix. Protein-based hydrogels are particularly promising because they can easily be functionalized and can achieve defined structures with adjustable physicochemical properties. However, the production of macroporous 3D templates for cell culture applications using natural materials is often limited by their weaker mechanical properties compared to those of synthetic materials. Here, different methods were evaluated to produce macroporous bovine serum albumin (BSA)-based hydrogel systems, with adjustable pore sizes in the range of 10 to 70 &#181;m in radius. Furthermore, a method to generate channels in this protein-based material that are several hundred microns long was established. The different methods to produce pores, as well as the influence of pore size on material properties such as swelling ratio, pH, temperature stability, and enzymatic degradation behavior, were analyzed. Pore sizes were investigated in the native, swollen state of the hydrogels using confocal laser scanning microscopy. The feasibility for cell culture applications was evaluated using a cell-adhesive RGD peptide modification of the protein system and two model cell lines: human breast cancer cells (A549) and adenocarcinomic human alveolar basal epithelial cells (MCF7).

Different methods to manipulate three-dimensional architecture in protein-based hydrogels are evaluated here with respect to material properties. The macroporous networks are functionalized with a cell-adhesive peptide, and their feasibility in cell culture is evaluated using two different model cell lines.

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field ...

Biochemistry

Synthesis of an Intein-mediated Artificial Protein Hydrogel

We present the synthesis of a highly stable protein hydrogel mediated by a split-intein-catalyzed protein trans-splicing reaction. The building blocks of this hydrogel are two protein block-copolymers each containing a subunit of a trimeric protein that serves as a crosslinker and one half of a split intein. A highly hydrophilic random coil is inserted into one of the block-copolymers for water retention. Mixing of the two protein block copolymers triggers an intein trans-splicing reaction, yielding a polypeptide unit with crosslinkers at either end that rapidly self-assembles into a hydrogel. This hydrogel is very stable under both acidic and basic conditions, at temperatures up to 50 &#176;C, and in organic solvents. The hydrogel rapidly reforms after shear-induced rupture. Incorporation of a &#34;docking station peptide&#34; into the hydrogel building block enables convenient incorporation of &#34;docking protein&#34;-tagged target proteins. The hydrogel is compatible with tissue culture growth media, supports the diffusion of 20 kDa molecules, and enables the immobilization of bioactive globular proteins. The application of the intein-mediated protein hydrogel as an organic-solvent-compatible biocatalyst was demonstrated by encapsulating the horseradish peroxidase enzyme and corroborating its activity.

We present the synthesis of a split-intein-mediated protein hydrogel. The building blocks of this hydrogel are two protein copolymers each containing a subunit of a trimeric protein that serves as a crosslinker and one half of a split intein. Mixing of the two protein copolymers triggers an intein trans-splicing reaction, yielding a polypeptide unit that self-assembles into a hydrogel. This hydrogel is highly pH- and temperature-stable, compatible with organic solvents, and easily incorporates functional globular proteins.

We present the synthesis of a highly stable protein hydrogel mediated by a split-intein-catalyzed protein trans-splicing reaction. The building blocks of ...

Cellular Encapsulation in 3D Hydrogels for Tissue Engineering 

The 3D encapsulation of cells within hydrogels represents an increasingly important and popular technique for culturing cells and towards the development of constructs for tissue engineering. This environment better mimics what cells observe in vivo, compared to standard tissue culture, due to the tissue-like properties and 3D environment. Synthetic polymeric hydrogels are water-swollen networks that can be designed to be stable or to degrade through hydrolysis or proteolysis as new tissue is deposited by encapsulated cells. A wide variety of polymers have been explored for these applications, such as poly(ethylene glycol) and hyaluronic acid. Most commonly, the polymer is functionalized with reactive groups such as methacrylates or acrylates capable of undergoing crosslinking through various mechanisms. In the past decade, much progress has been made in engineering these microenvironments - e.g., via the physical or pendant covalent incorporation of biochemical cues - to improve viability and direct cellular phenotype, including the differentiation of encapsulated stem cells (Burdick et al.).
The following methods for the 3D encapsulation of cells have been optimized in our and other laboratories to maximize cytocompatibility and minimize the number of hydrogel processing steps. In the following protocols (see Figure 1 for an illustration of the procedure), it is assumed that functionalized polymers capable of undergoing crosslinking are already in hand; excellent reviews of polymer chemistry as applied to the field of tissue engineering may be found elsewhere (Burdick et al.) and these methods are compatible with a range of polymer types. Further, the Michael-type addition (see Lutolf et al.) and light-initiated free radical (see Elisseeff et al.) mechanisms focused on here constitute only a small portion of the reported crosslinking techniques. Mixed mode crosslinking, in which a portion of reactive groups is first consumed by addition crosslinking and followed by a radical mechanism, is another commonly used and powerful paradigm for directing the phenotype of encapsulated cells (Khetan et al., Salinas et al.).

Cellular Encapsulation in 3D Hydrogels for Tissue Engineering

We present protocols for the 3-dimensional (3D) encapsulation of cells within synthetic hydrogels. The encapsulation procedure is outlined for two commonly used methods of crosslinking (michael-type addition and light-initiated free radical mechanisms), as well as a number of techniques for assessing encapsulated cell behavior.

The 3D encapsulation of cells within hydrogels represents an increasingly important and popular technique for culturing cells and towards the development ...

Light-mediated Formation and Patterning of Hydrogels for Cell Culture Applications

Click chemistries have been investigated for use in numerous biomaterials applications, including drug delivery, tissue engineering, and cell culture. In particular, light-mediated click reactions, such as photoinitiated thiol&#8722;ene and thiol&#8722;yne reactions, afford spatiotemporal control over material properties and allow the design of systems with a high degree of user-directed property control. Fabrication and modification of hydrogel-based biomaterials using the precision afforded by light and the versatility offered by these thiol&#8722;X photoclick chemistries are of growing interest, particularly for the culture of cells within well-defined, biomimetic microenvironments. Here, we describe methods for the photoencapsulation of cells and subsequent photopatterning of biochemical cues within hydrogel matrices using versatile and modular building blocks polymerized by a thiol&#8722;ene photoclick reaction. Specifically, an approach is presented for constructing hydrogels from allyloxycarbonyl (Alloc)-functionalized peptide crosslinks and pendant peptide moieties and thiol-functionalized poly(ethylene glycol) (PEG) that rapidly polymerize in the presence of lithium acylphosphinate photoinitiator and cytocompatible doses of long wavelength ultraviolet (UV) light. Facile techniques to visualize photopatterning and quantify the concentration of peptides added are described. Additionally, methods are established for encapsulating cells, specifically human mesenchymal stem cells, and determining their viability and activity. While the formation and initial patterning of thiol-alloc hydrogels are shown here, these techniques broadly may be applied to a number of other light and radical-initiated material systems (e.g., thiol-norbornene, thiol-acrylate) to generate patterned substrates.

We describe a sequential process for light-mediated formation and subsequent biochemical patterning of synthetic hydrogel matrices for three-dimensional cell culture applications. The construction and modification of hydrogels with cytocompatible photoclick chemistry is demonstrated. Additionally, facile techniques to quantify and observe patterns and determine cell viability within these hydrogels are presented.

Click chemistries have been investigated for use in numerous biomaterials applications, including drug delivery, tissue engineering, and cell culture. In ...

Fabricating Degradable Thermoresponsive Hydrogels on Multiple Length Scales via Reactive Extrusion, Microfluidics, Self-assembly, and Electrospinning

While various smart materials have been explored for a variety of biomedical applications (e.g., drug delivery, tissue engineering, bioimaging, etc.), their ultimate clinical use has been hampered by the lack of biologically-relevant degradation observed for most smart materials. This is particularly true for temperature-responsive hydrogels, which are almost uniformly based on polymers that are functionally non-degradable (e.g., poly(N-isopropylacrylamide) (PNIPAM) or poly(oligoethylene glycol methacrylate) (POEGMA)). As such, to effectively translate the potential of thermoresponsive hydrogels to the challenges of remote-controlled or metabolism-regulated drug delivery, cell scaffolds with tunable cell-material interactions, theranostic materials with the potential for both imaging and drug delivery, and other such applications, a method is required to render the hydrogels (if not fully degradable) at least capable of renal clearance following the required lifetime of the material. To that end, this protocol describes the preparation of hydrolytically-degradable hydrazone-crosslinked hydrogels on multiple length scales based on the reaction between hydrazide and aldehyde-functionalized PNIPAM or POEGMA oligomers with molecular weights below the renal filtration limit. Specifically, methods to fabricate degradable thermoresponsive bulk hydrogels (using a double barrel syringe technique), hydrogel particles (on both the microscale through the use of a microfluidics platform facilitating simultaneous mixing and emulsification of the precursor polymers and the nanoscale through the use of a thermally-driven self-assembly and cross-linking method), and hydrogel nanofibers (using a reactive electrospinning strategy) are described. In each case, hydrogels with temperature-responsive properties similar to those achieved via conventional free radical cross-linking processes can be achieved, but the hydrazone cross-linked network can be degraded over time to re-form the oligomeric precursor polymers and enable clearance. As such, we anticipate these methods (which may be generically applied to any synthetic water-soluble polymer, not just smart materials) will enable easier translation of synthetic smart materials to clinical applications.

Protocols are described for the fabrication of degradable thermoresponsive hydrogels based on hydrazone cross-linking of polymeric oligomers on the bulk scale, microscale, and nanoscale, the latter for preparation of both gel nanoparticles and nanofibers.

While various smart materials have been explored for a variety of biomedical applications (e.g., drug delivery, tissue engineering, bioimaging, etc.), ...

Patterning Bioactive Proteins or Peptides on Hydrogel Using Photochemistry for Biological Applications

There are many biological stimuli that can influence cell behavior and stem cell differentiation. General cell culture approaches rely on soluble factors within the medium to control cell behavior. However, soluble additions cannot mimic certain signaling motifs, such as matrix-bound growth factors, cell-cell signaling, and spatial biochemical cues, which are common influences on cells. Furthermore, biophysical properties of the matrix, such as substrate stiffness, play important roles in cell fate, which is not easily manipulated using conventional cell culturing practices. In this method, we describe a straightforward protocol to provide patterned bioactive proteins on synthetic polyethylene glycol (PEG) hydrogels using photochemistry. This platform allows for the independent control of substrate stiffness and spatial biochemical cues. These hydrogels can achieve a large range of physiologically relevant stiffness values. Additionally, the surfaces of these hydrogels can be photopatterned with bioactive peptides or proteins via thiol-ene click chemistry reactions. These methods have been optimized to retain protein function after surface immobilization. This is a versatile protocol that can be applied to any protein or peptide of interest to create a variety of patterns. Finally, cells seeded onto the surfaces of these bioactive hydrogels can be monitored over time as they respond to spatially specific signals.

In this method, we use photopolymerization and click chemistry techniques to create protein or peptide patterns on the surface of polyethylene glycol (PEG) hydrogels, providing immobilized bioactive signals for the study of cellular responses in vitro.

There are many biological stimuli that can influence cell behavior and stem cell differentiation. General cell culture approaches rely on soluble factors ...

Production of Elastin-like Protein Hydrogels for Encapsulation and Immunostaining of Cells in 3D

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue culture systems lack a three-dimensional (3D) matrix, resulting in a significant disconnect between results collected in vitro and in vivo. To address this limitation, researchers have engineered 3D hydrogel tissue culture platforms that can mimic the biochemical and biophysical properties of the in vivo cell microenvironment. This research has motivated the need to develop material platforms that support 3D cell encapsulation and downstream biochemical assays. Recombinant protein engineering offers a unique toolset for 3D hydrogel material design and development by allowing for the specific control of protein sequence and therefore, by extension, the potential mechanical and biochemical properties of the resultant matrix. Here, we present a protocol for the expression of recombinantly-derived elastin-like protein (ELP), which can be used to form hydrogels with independently tunable mechanical properties and cell-adhesive ligand concentration. We further present a methodology for cell encapsulation within ELP hydrogels and subsequent immunofluorescent staining of embedded cells for downstream analysis and quantification.

Recombinant protein-engineered hydrogels are advantageous for 3D cell culture as they allow for complete tunability of the polymer backbone and therefore, the cell microenvironment. Here, we describe the process of recombinant elastin-like protein purification and its application in 3D hydrogel cell encapsulation.

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue ...

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) culture systems. However, measurement of cell function in 3D culture is often more challenging. Many biological assays require retrieval of cellular material which can be difficult in 3D cultures. One way to address this challenge is to develop new materials that enable measurement of cell function within the material. Here, a method is presented for measurement of cellular matrix metalloproteinase (MMP) activity in 3D hydrogels in a 96-well format. In this system, a poly(ethylene glycol) (PEG) hydrogel is functionalized with a fluorogenic MMP cleavable sensor. Cellular MMP activity is proportional to fluorescence intensity and can be measured with a standard microplate reader. Miniaturization of this assay to a 96-well format reduced the time required for experimental set up by 50% and reagent usage by 80% per condition as compared to the previous 24-well version of the assay. This assay is also compatible with other measurements of cellular function. For example, a metabolic activity assay is demonstrated here, which can be conducted simultaneously with MMP activity measurements within the same hydrogel. The assay is demonstrated with human melanoma cells encapsulated across a range of cell seeding densities to determine the appropriate encapsulation density for the working range of the assay. After 24 h of cell encapsulation, MMP and metabolic activity readouts were proportional to cell seeding density. While the assay is demonstrated here with one fluorogenic degradable substrate, the assay and methodology could be adapted for a wide variety of hydrogel systems and other fluorescent sensors. Such an assay provides a practical, efficient and easily accessible 3D culturing platform for a wide variety of applications.

Here, a protocol is presented for encapsulating and culturing cells in poly(ethylene glycol) (PEG) hydrogels functionalized with a fluorogenic matrix metalloproteinase (MMP)-degradable peptide. Cellular MMP and metabolic activity are measured directly from the hydrogel cultures using a standard microplate reader.

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) ...

Methods for Embedding Cell-Free Protein Synthesis Reactions in Macro-Scale Hydrogels

Summary

Explore More Videos

Microfabricated Platforms for Mechanically Dynamic Cell Culture

Fabrication of Micropatterned Hydrogels for Neural Culture Systems using Dynamic Mask Projection Photolithography

Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Synthesis of an Intein-mediated Artificial Protein Hydrogel

Cellular Encapsulation in 3D Hydrogels for Tissue Engineering

Light-mediated Formation and Patterning of Hydrogels for Cell Culture Applications

Fabricating Degradable Thermoresponsive Hydrogels on Multiple Length Scales via Reactive Extrusion, Microfluidics, Self-assembly, and Electrospinning

Patterning Bioactive Proteins or Peptides on Hydrogel Using Photochemistry for Biological Applications

Production of Elastin-like Protein Hydrogels for Encapsulation and Immunostaining of Cells in 3D

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels