Authors would like to acknowledge Dr. Jennifer VanderKelen, Andrea Laubscher, and Tony Turretto for technical support, Wesley Kao, Layne Williams, and Christopher Hight for helpful discussions. Authors also acknowledge funding support from the Bill and Linda Frost Fund, Center for Applications in Biotechnology&#39;s Chevron Biotechnology Applied Research Endowment Grant, Cal Poly Research, Scholarly, and Creative Activities Grant Program (RSCA 2017), and the National Science Foundation (NSF-1708919). MZL acknowledges the California State University Graduate Grant. MCJ acknowledges the Army Research Office W911NF-16-1-0372, National Science Foundation grants MCB-1413563 and MCB-1716766, the Air Force Research Laboratory Center of Excellence Grant FA8650-15-2-5518, the Defense Threat Reduction Agency Grant HDTRA1-15-10052/P00001, the David and Lucile Packard Foundation, the Camille Dreyfus Teacher-Scholar Program, the Department of Energy BER Grant DE-SC0018249, the Human Frontiers Science Program (RGP0015/2017), the DOE Joint Genome Institute ETOP Grant, and the Chicago Biomedical Consortium with support from the Searle Funds at the Chicago Community Trust for support.

1. Media Preparation and Cell Growth
<ol>
	<li>Day 1
	<ol>
		<li>Streak E. coli BL21*(DE3) cells from a glycerol stock onto an LB agar plate and incubate for at least 18 h at 37 &#176;C.</li>
		<li>Prepare 50 mL of LB media and autoclave the solution on a liquid cycle for 30 min at 121 &#176;C. Store at room temperature.</li>
	</ol>
	</li>
	<li>Day 2
	<ol>
		<li>Prepare 750 mL of 2x YTP media and 250 mL of 0.4 M D-Glucose solution as described in the supplemental information.</li>
		<li>Pour the 2x YTP media into an autoclaved 2.5 L baffled flask and the D-Glucose solution into an autoclaved 500 mL glass bottle. Autoclave both solutions on a liquid cycle for 30 min at 121 &#176;C.</li>
		<li>Ensure that both sterile solutions are stored at 37 &#176;C if cell growth is being performed on the next day, to maximize growth rates upon inoculation. Do not combine solutions until inoculation. 
		NOTE: Solutions can be stored at 4 &#176;C for 1-2 d if needed, though the 2x YTP media is highly prone to contamination.</li>
		<li>Start an overnight culture of BL21(DE3) by inoculating 50 mL of LB media with a single colony of BL21(DE3) using a sterilized loop and sterile technique to avoid contamination.</li>
		<li>Place the 50 mL of BL21*(DE3) LB culture into a 37 &#176;C 250 rpm shaking incubator and grow overnight for 15-18 h.</li>
		<li>Prepare and sterilize all materials required for days 3 and 4, including: two 1 L centrifuge bottles, 4x cold 50 mL conical tubes (weigh and record masses of three), and many 1.5 mL microfuge tubes.</li>
	</ol>
	</li>
	<li>Day 3
	<ol>
		<li>Remove the 50 mL overnight culture of BL21*(DE3) in LB from the shaking incubator and measure the OD600 on a spectrophotometer using a 1:10 dilution with LB media. Calculate the volume of overnight culture necessary to add to 1 L of media for a starting OD600 of 0.1 (For example, if an OD600 of a 1:10 dilution is read as 0.4, inoculate 25 mL of the undiluted OD600 = 4.0 overnight culture into 1 L of 2x YTPG).</li>
		<li>Remove the warmed 2x YTP media and D-Glucose solutions from the 37 &#176;C incubator along with the 50 mL of LB culture. Using sterile technique, carefully pour the D-Glucose solution into the 2x YTP media (avoiding the sides of the baffled flask). 
		NOTE: Addition of D-Glucose completes the recipe for 1 L of 2x YTPG.</li>
		<li>Maintaining sterile technique, inoculate the 1 L of 2x YTPG solution with the appropriate amount of the 50 mL culture to begin the 1 L culture at a 0.1 OD600. Immediately place the inoculated 1 L culture into a 37 &#176;C shaking incubator at 200 rpm.</li>
		<li>Take the first OD600 reading after the first hour of growth (lag phase typical takes 1 h). Do not dilute the culture. Continue taking OD600 measurements approximately every 20-30 min until OD600 reaches 0.6.</li>
		<li>Upon reaching OD600 = 0.6, add 1 mL of 1M IPTG (final concentration in 1 L culture = 1 mM) to the 2x YTPG culture. 
		NOTE: Ideal induction OD600 is 0.6; however, a range of 0.6-0.8 is acceptable. Induction by IPTG is for endogenous production of T7 RNA Polymerase (T7RNAP).</li>
		<li>After induction, measure the OD600 approximately every 20-30 min until it reaches 3.0. 
		NOTE: Cool down the centrifuge to 4 &#176;C during this time. Prepare cold S30 buffer as detailed in the Supplementary Information. If the S30 buffer is prepared in advance, ensure that DTT is not added until the day of use.</li>
		<li>Once the OD600 reaches 3.0 (Figure 2A), pour the culture into a cold 1 L centrifuge bottle in an ice-water bath. Prepare a water-filled 1 L centrifuge bottle of equal weight to be used as a balance in the centrifuge. 
		NOTE: Absorbance values vary from instrument-to-instrument. While the OD600 of harvest of BL21(DE3) is not a sensitive variable, it is recommended that the user evaluate and optimize this variable as a troubleshooting measure. Larger spectrophotometers may result in relatively lower OD600 readings compared to smaller cuvette-based spectrophotometers.</li>
		<li>Centrifuge the 1 L bottles for 10 min at 5,000 x g and 10 &#176;C to pellet cells.</li>
		<li>Slowly pour off the supernatant and dispose of it according to the institution&#39;s biological waste procedures. Place the pellet on ice.</li>
		<li>Using a sterile spatula, scrape the cell pellet from the centrifuge bottle and transfer it to a cold 50 mL conical tube.</li>
		<li>Add 30 mL of cold S30 buffer to the conical tube and resuspend the cell pellet by vortexing with short bursts (20 - 30 s) and rest periods (1 min) on ice until fully resuspended with no chunks.</li>
		<li>Once the pellet is fully resuspended, use another 50 mL conical tube with water as a balance and centrifuge for 10 min at 5000 x g and 10 &#176;C (pre-cooled to 4 &#176;C). 
		NOTE: This completes the 1st of 3 washes required when harvesting the cells.</li>
		<li>Pour out the supernatant and dispose of it according to the institution&#39;s biological waste procedures. Resuspend the pellet with 20-25 mL of cold S30 buffer and centrifuge for 10 min at 5000 x g and 10 &#176;C (pre-cooled to 4 &#176;C). 
		NOTE: This completes the 2nd of 3 washes.</li>
		<li>Again, pour out the supernatant and dispose of it according to the institution&#39;s biological waste procedures. Add exactly 30 mL of S30 buffer and vortex again to resuspend the pellet.</li>
		<li>Using the 3 pre-weighed, cold 50 mL conical tubes and a serological pipette filler with a sterile pipette, aliquot 10 mL of resuspended pellet/S30 buffer mixture into each of the 3 conical tubes. 
		NOTE: Splitting the cells into 3 tubes is not required, but this step results in smaller cell pellets (~ 1 g) for increased convenience at later steps.</li>
		<li>Centrifuge all tubes, using appropriate balances as needed, for 10 min at 5000 x g and 10 &#176;C (pre-cooled to 4 &#176;C). 
		NOTE: This completes the final wash step.</li>
		<li>Pour out the supernatant and dispose of it according to the institution&#39;s biological waste procedures. Remove the excess S30 buffer by carefully wiping the inside of the conical tube and cap with a clean tissue; avoid touching the pellet.</li>
		<li>Reweigh the tubes on an analytical balance and record the final pellet weight on each tube. 
		NOTE: The protocol can be paused at this point. The pellets can be flash frozen in liquid nitrogen and stored at -80 &#176;C for up to a year until needed for extract preparation.</li>
	</ol>
	</li>
</ol>
2. Crude Cell Extract Preparation - Day 4
<ol>
	<li>For extract preparation, keep cells cold on ice during each step. Add 1 mL of cold S30 buffer per 1 g of cell mass of the pellet. Ensure that dithiothreitol (DTT) has been supplemented to the S30 buffer to a final concentration of 2 mM. 
	NOTE: Cool down the microcentrifuge to 4 &#176;C during this time.</li>
	<li>Resuspend the cell pellet by vortexing with short bursts (20 - 30 s) and rest periods (1 min) on ice until fully resuspended. If resuspension is difficult, leave the pellets on ice for 30 min to defrost.</li>
	<li>Transfer 1.4 mL of resuspended cells into a 1.5 mL microfuge tube.</li>
	<li>Place one 1.5 mL tube containing 1.4 mL of resuspended cells into an ice water bath in a beaker. Sonicate for 45 s on followed by 59 s off for 3 total cycles, with amplitude set at 50%. Close and invert the tubes to gently mix during the off periods. In total, deliver 800-900 J of energy to each 1.5 mL microfuge tube containing 1.4 mL of resuspended cells (Figure 3A &#38; 3B). 
	NOTE: This step is sensitive to the sonicator type and model used and should be optimized if equipment is different than listed for this procedure. Two complementary approaches can be used to scale-up the amount of extract prepared during this step: 1) multiple 1.5 mL microfuge tubes can be sonicated in parallel, and/or 2) larger volumes can be sonicated in conical tubes (up to 15 mL of cell resuspension per tube), scaling the amount of energy delivered as previously described 29,45.</li>
	<li>Immediately after sonication is complete, add 4.5&#160;&#181;L of 1 M DTT (supplementing an additional 2 mM DTT) into the 1.4 mL of lysate and invert several times to mix. Place the tube on ice. Repeat steps 2.4 and 2.5 for any additional tubes of resuspended cells before proceeding to centrifugation.</li>
	<li>Microcentrifuge samples at 18,000 x g and 4 &#176;C for 10 min (Figure 3C).</li>
	<li>Pipette the supernatant into a new 1.5 mL microfuge tube. Do not disturb the pellet; it is preferable to leave some supernatant behind to maintain purity than to disrupt the pellet in efforts to maximize yield.</li>
	<li>Incubate the supernatant from the previous step at 250 rpm and 37 &#176;C for 60 min by taping the tubes to the shaking platform of the incubator (this is the runoff reaction).</li>
	<li>Microcentrifuge samples at 10,000 x g and 4 &#176;C for 10 min.</li>
	<li>Remove the supernatant without disturbing the pellet and transfer it to a new tube. Create many 100 &#181;L aliquots of extract for storage. 
	NOTE: The protocol can be paused here, and the extract can be flash frozen in liquid nitrogen and stored at -80 &#176;C for up to a year until needed for CFPS reactions. At least 5 freeze-thaw cycles can be undergone without detriment to extract productivity (Figure 4).</li>
</ol>
3. Cell-Free Protein Synthesis Batch Format Reactions
<ol>
	<li>Thaw Solutions A and B, DNA template, BL21(DE3) extract (if frozen), T7RNAP, and an aliquot of molecular grade water. 
	NOTE: CFPS reaction template can be found in the Supplementary Information. Solutions A and B recipes are provided in the Supplementary Information and correspond to specific concentrations for numerous reagents to support the PANOx-SP based energy system for CFPS. The role of each reagent and acceptable variation in these reagent concentrations that can support CFPS have been determined50. A T7RNAP purification protocol can be found in the Supplementary Information51. Supplemental T7RNAP can increase volumetric yields but is not necessary if T7RNAP is induced during cell growth. Plasmid DNA template (pJL1-sfGFP) can be prepared using a maxiprep kit with two washes using the wash buffer in the kit, followed by a post-processing DNA-cleanup using a PCR purification kit (Figure 2B). Linear DNA templates can also be used in CFPS reactions.</li>
	<li>Label the necessary amount of microfuge tubes needed for CFPS reactions. 
	NOTE: Reactions can be performed in various vessel sizes, but a smaller vessel can decrease volumetric protein yields (Figure 2C). Scaling up a reaction in the same size vessel may also reduce volumetric yields, as a function of decreasing the oxygen exchange, due to a decrease in the surface area to volume ratio. When increasing reaction volume above 100 &#956;L, it is recommended to use flat bottom well plates 31,37,52.</li>
	<li>Add 2.2 &#181;L of Solution A, 2.1 &#181;L of Solution B, 5 &#181;L of BL21*(DE3) extract, 0.24 &#956;g of T7RNAP (16 &#956;g/mL final concentration), 0.24 ng of DNA template (16 ng/&#181;L final concentration), and water to bring the final volume to 15 &#181;L. 
	NOTE: Vortex Solutions A and B frequently during reaction setup to avoid sedimentation of components and ensure that each reaction receives a homogenous aliquot of each solution. Avoid vortexing the extract, instead invert the tube to mix.</li>
	<li>After all reagents have been added to the reaction, mix each tube by pipetting up and down or gently vortexing while ensuring that the final reaction mixture is combined into a single 15 &#181;L bead at the bottom of the 1.5 mL microfuge tube.</li>
	<li>Place each reaction into the 37 &#176;C incubator without shaking for 4 h, or 30 &#176;C overnight. 
	NOTE: Successful reactions can be qualitatively assessed visually based on the green color of the sfGFP product within the CFPS reaction mixture (Figure 3D). Expression of the protein of interest can also be confirmed by SDS-PAGE (Supplemental Figure 2).</li>
</ol>
4. Quantification of the Reporter Protein, [sfGFP]
<ol>
	<li>Load 48 &#181;L of 0.05 M HEPES, pH 8, into each well needed for quantification (usually performed in triplicate per reaction tube).</li>
	<li>Remove reactions from incubator. Pipette up and down to mix each reaction, then transfer 2 &#181;L of reaction into the 48 &#181;L of 0.05 M HEPES, pH 8. Pipet up and down again in the well to mix.</li>
	<li>Once all reactions are loaded and mixed, place the 96 well plate into the fluorometer and measure the sfGFP endpoint fluorescence. 
	NOTE: Excitation and emission wavelengths for sfGFP fluorescence quantification are 485 nm and 510 nm, respectively.</li>
	<li>Using a previously generated standard curve, determine the [sfGFP] from the obtained fluorescence readings. 
	NOTE: Instructions for generating a standard curve of sfGFP concentration versus fluorescence intensity are provided in Supplementary Information (Supplementary Figure 3). Users will need to establish a standard curve for their instrument since instrument sensitivity may vary.</li>
</ol>

The authors declare that they have no competing financial interests or other conflicts of interest.

Cell-free protein synthesis has emerged as a powerful and enabling technology for a variety of applications ranging from biomanufacturing to rapid prototyping of biochemical systems. The breadth of applications is supported by the capacity to monitor, manipulate, and augment cellular machinery in real-time. In spite of the expanding impact of this platform technology, broad adaptation has remained slow due to technical nuances in the implementation of the methods. Through this effort, we aim to provide simplicity and cla...

Cell-free Protein Synthesis (CFPS) has emerged as a technology that has unlocked a number of new opportunities for protein production, functional genomics, metabolic engineering, and more within the last 50 years1,2. Compared to standard in vivo protein expression platforms, CFPS provides three key advantages: 1) the cell-free nature of the platform enables the production of proteins that would be potentially toxic or foreign to the cell3,4,5,6; 2) inactivation of genomic DNA and the introduction of a template DNA encoding the gene(s) of interest channel all of the systemic energy within the reaction to the production of the protein(s) of interest; and 3) the open nature of the platform enables the user to modify and monitor the reaction conditions and composition in real time7,8. This direct access to the reaction supports the augmentation of biological systems with expanded chemistries and redox conditions for the production of novel proteins and the tuning of metabolic processes2,9,10. Direct access also allows the user to combine the CFPS reaction with activity assays in a single-pot system for more rapid design-build-test cycles11. The capacity to perform the CFPS reaction in small volume droplets or on paper-based devices further supports high-throughput discovery efforts and rapid prototyping12,13,14,15,16. As a result of these advantages and the plug and play nature of the system, CFPS has uniquely enabled a variety of biotechnology applications such as the production of proteins that are difficult to solubly express in vivo17,18,19,20, detection of disease21,22,23, on demand biomanufacturing18,24,25,26,27, and education28,29, all of which show the flexibility and utility of the cell-free platform.
CFPS systems can be generated from a variety of crude lysates from both prokaryotic and eukaryotic cell lines. This allows for diverse options in the system of choice, each of which have advantages and disadvantages depending on the application of interest. CFPS systems also vary greatly in preparation time, cost, and productivity. The most commonly utilized cell extracts are produced from wheat germ, rabbit reticulocyte, insect cells, and Escherichia coli cells, with the latter being the most cost-effective to date while producing the highest volumetric yields of protein30. While other CFPS systems can be advantageous for their innate post-translational modification machinery, emerging applications using the E. coli-based machinery are able to bridge the gap by generating site-specifically phosphorylated and glycosylated proteins on demand31,32,33,34,35.
CFPS reactions can be run in either batch, continuous-exchange cell-free (CECF) or continuous-flow cell-free (CFCF) formats. The batch format is a closed system whose reaction lifetime is limited due to diminishing quantities of reactants and the accumulation of inhibitory byproducts of the reaction. CECF and CFCF methods increase the lifetime of the reaction, and thereby result in increased volumetric protein yields compared to the batch reaction. This is accomplished by allowing the byproducts of protein synthesis to be removed from the reaction vessel while new reactants are supplied throughout the course of the reaction2. In the case of CFCF, the protein of interest can also be removed from the reaction chamber, while in CECF, the protein of interest remains in the reaction chamber comprised of a semi-permeable membrane36,37. These methods are especially valuable in overcoming poor volumetric yields of difficult-to-express proteins of interest38,39,40,41,42,43. The challenges in implementing the CECF and CFCF approaches are that 1) while they result in more efficient use of the bio machinery responsible for transcription and translation, they require notably larger quantities of reagents that increases overall cost and 2) they require more complex reaction setups and specialized equipment compared to the batch format44. In order to ensure accessibility for new users, the protocols described herein focus on the batch format at reaction volumes of 15 &#181;L with specific recommendations for increasing the reaction volume to the milliliter scale.
The methods presented herein enable non-experts with basic laboratory skills (such as undergraduate students) to implement cell growth, extract preparation, and batch format reaction setup for an E. coli-based CFPS system. This approach is cost-effective compared to commercially available kits without sacrificing the ease of kit-based reaction setup. Furthermore, this approach enables applications in the laboratory and in the field. When deciding to implement CFPS, new users should thoroughly evaluate the efficacy of conventional protein expression systems for startup investment, as CFPS may not be superior in every case. The CFPS methods described here enable the user to directly implement a variety of applications, including functional genomics, high-throughput testing, the production of proteins that are intractable for in vivo expression, as well as field applications including biosensors and educational kits for synthetic biology. Additional applications such as metabolic engineering, tuning of protein expression conditions, disease detection, and scale-up using CECF or CFCF methods are still possible but may require experience with the CFPS platform for further modification of reaction conditions. Our methods combine growth in enriched media and baffled flasks, with relatively rapid and reproducible methods of cell lysis through sonication, followed by a simplified CFPS reaction setup that utilizes optimized premixes45. While the cellular growth methods have become somewhat standardized within this field, methods for cell lysis vary widely. In addition to sonication, common lysis methods include utilization of a French press, a homogenizer, bead beaters, or lysozyme and other biochemical and physical disruption methods46,47,48,49. Using our methods, approximately 2 mL of crude cell extract are obtained per 1 L of cells. This quantity of cell extract can support four hundred 15 &#181;L CFPS reactions, each producing ~900 &#181;g/mL of reporter sfGFP protein from the template plasmid pJL1-sfGFP. This method costs $0.021/&#181;g of sfGFP produced ($.019/&#181;L of reaction), excluding the cost of labor and equipment (Supplemental Figure 1). Starting from the scratch, this method can be implemented in 4 days by a single person and repeat CFPS reactions can be completed within hours (Figure 1). Additionally, the protocol can be scaled up in volume for larger batches of reagent preparation to suit the user&#39;s needs.Importantly, the protocol presented here can be implemented by laboratory trained non-experts such as undergraduate students, as it only requires basic laboratory skills. The procedures described below, and the accompanying video have been specifically developed to improve accessibility of the E. coli CFPS platform for broad usage.

<table><tbody><tr><td>Luria Broth</td><td>ThermoFisher</td><td>12795027</td><td></td></tr><tr><td>Tryptone</td><td>Fisher Bioreagents</td><td>73049-73-7</td><td></td></tr><tr><td>Yeast Extract</td><td>Fisher Bioreagents</td><td>1/2/8013</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S3014-1KG</td><td></td></tr><tr><td>Potassium Phosphate Dibasic</td><td>Sigma-Aldrich</td><td>60353-250G</td><td></td></tr><tr><td>Potassium Phosphate Monobasic</td><td>Sigma-Aldrich</td><td>P9791-500G</td><td></td></tr><tr><td>D-Glucose</td><td>Sigma-Aldrich</td><td>G8270-1KG</td><td></td></tr><tr><td>KOH</td><td>Sigma-Aldrich</td><td>P5958-500G</td><td></td></tr><tr><td>IPTG</td><td>Sigma-Aldrich</td><td>I6758-1G</td><td></td></tr><tr><td>Mg(OAc)2</td><td>Sigma-Aldrich</td><td>M5661-250G</td><td></td></tr><tr><td>K(OAc)</td><td>Sigma-Aldrich</td><td>P1190-1KG</td><td></td></tr><tr><td>Tris(OAc)</td><td>Sigma-Aldrich</td><td>T6066-500G</td><td></td></tr><tr><td>DTT</td><td>ThermoFisher</td><td>15508013</td><td></td></tr><tr><td>tRNA</td><td>Sigma-Aldrich</td><td>10109541001</td><td></td></tr><tr><td>Folinic Acid</td><td>Sigma-Aldrich</td><td>F7878-100MG</td><td></td></tr><tr><td>NTPs</td><td>ThermoFisher</td><td>R0481</td><td></td></tr><tr><td>Oxalic Acid</td><td>Sigma-Aldrich</td><td>P0963-100G</td><td></td></tr><tr><td>NAD</td><td>Sigma-Aldrich</td><td>N8535-15VL</td><td></td></tr><tr><td>CoA</td><td>Sigma-Aldrich</td><td>C3144-25MG</td><td></td></tr><tr><td>PEP</td><td>Sigma-Aldrich</td><td>860077-250MG</td><td></td></tr><tr><td>K(Glu)</td><td>Sigma-Aldrich</td><td>G1501-500G</td><td></td></tr><tr><td>NH4(Glu)</td><td>MP Biomedicals</td><td>02180595.1</td><td></td></tr><tr><td>Mg(Glu)2</td><td>Sigma-Aldrich</td><td>49605-250G</td><td></td></tr><tr><td>Spermidine</td><td>Sigma-Aldrich</td><td>S0266-5G</td><td></td></tr><tr><td>Putrescine</td><td>Sigma-Aldrich</td><td>D13208-25G</td><td></td></tr><tr><td>HEPES</td><td>ThermoFisher</td><td>11344041</td><td></td></tr><tr><td>Molecular Grade Water</td><td>Sigma-Aldrich</td><td>7732-18-5</td><td></td></tr><tr><td>L-Aspartic Acid</td><td>Sigma-Aldrich</td><td>A7219-100G</td><td></td></tr><tr><td>L-Valine</td><td>Sigma-Aldrich</td><td>V0500-25G</td><td></td></tr><tr><td>L-Tryptophan</td><td>Sigma-Aldrich</td><td>T0254-25G</td><td></td></tr><tr><td>L-Phenylalanine</td><td>Sigma-Aldrich</td><td>P2126-100G</td><td></td></tr><tr><td>L-Isoleucine</td><td>Sigma-Aldrich</td><td>I2752-25G</td><td></td></tr><tr><td>L-Leucine</td><td>Sigma-Aldrich</td><td>L8000-25G</td><td></td></tr><tr><td>L-Cysteine</td><td>Sigma-Aldrich</td><td>C7352-25G</td><td></td></tr><tr><td>L-Methionine</td><td>Sigma-Aldrich</td><td>M9625-25G</td><td></td></tr><tr><td>L-Alanine</td><td>Sigma-Aldrich</td><td>A7627-100G</td><td></td></tr><tr><td>L-Arginine</td><td>Sigma-Aldrich</td><td>A8094-25G</td><td></td></tr><tr><td>L-Asparagine</td><td>Sigma-Aldrich</td><td>A0884-25G</td><td></td></tr><tr><td>Glycine</td><td>Sigma-Aldrich</td><td>G7126-100G</td><td></td></tr><tr><td>L-Glutamine</td><td>Sigma-Aldrich</td><td>G3126-250G</td><td></td></tr><tr><td>L-Histadine</td><td>Sigma-Aldrich</td><td>H8000-25G</td><td></td></tr><tr><td>L-Lysine</td><td>Sigma-Aldrich</td><td>L5501-25G</td><td></td></tr><tr><td>L-Proline</td><td>Sigma-Aldrich</td><td>P0380-100G</td><td></td></tr><tr><td>L-Serine</td><td>Sigma-Aldrich</td><td>S4500-100G</td><td></td></tr><tr><td>L-Threonine</td><td>Sigma-Aldrich</td><td>T8625-25G</td><td></td></tr><tr><td>L-Tyrosine</td><td>Sigma-Aldrich</td><td>T3754-100G</td><td></td></tr><tr><td>Fisherbrand Premium Microcentrifuge Tubes: 2.0 mL</td><td>Fisher Scientific</td><td>05-408-138</td><td></td></tr><tr><td>Fisherbrand Premium Microcentrifuge Tubes: 1.5 mL</td><td>Fisher Scientific</td><td>05-408-129</td><td></td></tr><tr><td>Fisherbrand Premium Microcentrifuge Tubes: 0.6 mL</td><td>Fisher Scientific</td><td>05-408-120</td><td></td></tr><tr><td>PureLink HiPure Plasmid Prep Kit</td><td>ThermoFisher</td><td>K210007</td><td></td></tr><tr><td>Ultrasonic Processor</td><td>QSonica</td><td>Q125-230V/50Hz</td><td>3.175 mm diameter probe</td></tr><tr><td>Avanti J-E Centrifuge</td><td>Beckman Coulter</td><td>369001</td><td></td></tr><tr><td>JLA-8.1000 Rotor</td><td>Beckman Coulter</td><td>366754</td><td></td></tr><tr><td>1L Centrifuge Tube</td><td>Beckman Coulter</td><td>A99028</td><td></td></tr><tr><td>Tunair 2.5L Baffeled Shake Flask</td><td>Sigma-Aldrich</td><td>Z710822</td><td></td></tr><tr><td>Microfuge 20</td><td>Beckman Coulter</td><td>B30134</td><td></td></tr><tr><td>New Brunswick Innova 42/42R Incubator</td><td>Eppendorf</td><td>M1335-0000</td><td></td></tr><tr><td>Cytation 5</td><td>BioTek</td><td></td><td></td></tr><tr><td>Strep-Tactin XT Starter Kit</td><td>IBA</td><td>2-4998-000</td><td></td></tr><tr><td>pJL1-sfGFP</td><td>Addgene</td><td>69496</td><td></td></tr><tr><td>BL21(DE3)</td><td>New England BioLabs</td><td></td><td></td></tr></tbody></table>

escherichia coli-based cell-free protein synthesis: protocols for a robust, flexible, and accessible platform technology

Over the last 50 years, Cell-Free Protein Synthesis (CFPS) has emerged as a powerful technology to harness the transcriptional and translational capacity of cells within a test tube. By obviating the need to maintain the viability of the cell, and by eliminating the cellular barrier, CFPS has been foundational to emerging applications in biomanufacturing of traditionally challenging proteins, as well as applications in rapid prototyping for metabolic engineering, and functional genomics. Our methods for implementing an E. coli-based CFPS platform allow new users to access many of these applications. Here, we describe methods to prepare extract through the use of enriched media, baffled flasks, and a reproducible method of tunable sonication-based cell lysis. This extract can then be used for protein expression capable of producing 900 &#181;g/mL or more of super folder green fluorescent protein (sfGFP) in just 5 h from experimental setup to data analysis, given that appropriate reagent stocks have been prepared beforehand. The estimated startup cost of obtaining reagents is $4,500 which will sustain thousands of reactions at an estimated cost of $0.021 per &#181;g of protein produced or $0.019 per &#181;L of reaction. Additionally, the protein expression methods mirror the ease of the reaction setup seen in commercially available systems due to optimization of reagent pre-mixes, at a fraction of the cost. In order to enable the user to leverage the flexible nature of the CFPS platform for broad applications, we have identified a variety of aspects of the platform that can be tuned and optimized depending on the resources available and the protein expression outcomes desired.

We have presented a sonication-based E. coli extract preparation protocol that can be completed over a four-day span, with Figure 1 demonstrating the procedural breakdown over each day. There is malleability to the steps that can be completed in each day with various pausing points, but we have found this workflow to be the most effective to execute. Additionally, both the cell pellets (step 1.3.18) and fully prepared extract (step 2.10) are stable a...

Watch this Scientific Journal Video about Escherichia coli-Based Cell-Free Protein Synthesis: Protocols for a robust, flexible, and accessible platform technology at JoVE.com

Escherichia coli-Based Cell-Free Protein Synthesis: Protocols for a robust, flexible, and accessible platform technology

This protocol details the steps, costs, and equipment necessary to generate E. coli-based cell extracts and implement in vitro protein synthesis reactions within 4 days or less. To leverage the flexible nature of this platform for broad applications, we discuss reaction conditions that can be adapted and optimized.

Escherichia coli-Based Cell-Free Protein Synthesis: Protocols for a robust, flexible, and accessible platform technology

Over the last 50 years, Cell-Free Protein Synthesis (CFPS) has emerged as a powerful technology to harness the transcriptional and translational capacity ...

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Research

JoVE Journal

Chemistry

34.6K Views.  California Polytechnic State University, San Luis Obispo. Dit protocol vereenvoudigt en verduidelijkt de methoden voor de implementatie van zelfhereiïteïnesynthese door niet-deskundigen. Verbeterde toegang tot deze methoden zal helpen bij het ontmarkten van het platform en de brede set van toepassingen die het mogelijk maakt. De belangrijkste voordelen van deze techniek zijn de snelheid, kosteneffectiviteit, en het gemak van reactie opgezet in vergelijking met andere zelf re-eiwit synthese platforms.Ons platform kan een aantal toepassingen ...

Video: Escherichia coli-Based Cell-Free Protein Synthesis: Protocols for a robust, flexible, and accessible platform technology

Particles without a Box: Brush-first Synthesis of Photodegradable PEG Star Polymers under Ambient Conditions

Convenient methods for the rapid, parallel synthesis of diversely functionalized nanoparticles will enable discovery of novel formulations for drug delivery, biological imaging, and supported catalysis. In this report, we demonstrate parallel synthesis of brush-arm star polymer (BASP) nanoparticles by the &#34;brush-first&#34; method. In this method, a norbornene-terminated poly(ethylene glycol) (PEG) macromonomer (PEG-MM) is first polymerized via ring-opening metathesis polymerization (ROMP) to generate a living brush macroinitiator. Aliquots of this initiator stock solution are added to vials that contain varied amounts of a photodegradable bis-norbornene crosslinker. Exposure to crosslinker initiates a series of kinetically-controlled brush+brush and star+star coupling reactions that ultimately yields BASPs with cores comprised of the crosslinker and coronas comprised of PEG. The final BASP size depends on the amount of crosslinker added. We carry out the synthesis of three BASPs on the benchtop with no special precautions to remove air and moisture. The samples are characterized by gel permeation chromatography (GPC); results agreed closely with our previous report that utilized inert (glovebox) conditions. Key practical features, advantages, and potential disadvantages of the brush-first method are discussed.

Poly(ethylene glycol) (PEG) brush-arm star polymers (BASPs) with narrow mass distributions and tunable nanoscopic sizes are synthesized in via ring opening metathesis polymerization (ROMP) of a PEG-norbornene macromonomer followed by transfer of portions of the resulting living brush initiator to vials containing varied amounts of a rigid, photo-cleavable bis-norbornene crosslinker.

Convenient methods for the rapid, parallel synthesis of diversely functionalized nanoparticles will enable discovery of novel formulations for drug ...

Synthesis of Infectious Bacteriophages in an E. coli-based Cell-free Expression System

A new generation of cell-free transcription-translation (TXTL) systems, engineered to have a greater versatility and modularity, provide novel capabilities to perform basic and applied sciences in test tube reactions. Over the past decade, cell-free TXTL has become a powerful technique for a broad range of novel multidisciplinary research areas related to quantitative and synthetic biology. The new TXTL platforms are particularly useful to construct and interrogate biochemical systems through the execution of synthetic or natural gene circuits. In vitro TXTL has proven convenient to rapidly prototype regulatory elements and biological networks as well as to recapitulate molecular self-assembly mechanisms found in living systems. In this article, we describe how infectious bacteriophages, such as MS2 (RNA), &#934;&#935;174 (ssDNA), and T7 (dsDNA), are entirely synthesized from their genome in one-pot reactions using an all Escherichia coli, cell-free TXTL system. Synthesis of the three coliphages is quantified using the plaque assay. We show how the yield of synthesized phage depends on the biochemical settings of the reactions. Molecular crowding, emulated through a controlled concentration of PEG 8000, affects the amount of synthesized phages by orders of magnitudes. We also describe how to amplify the phages and how to purify their genomes. The set of protocols and results presented in this work should be of interest to multidisciplinary researchers involved in cell-free synthetic biology and bioengineering.

Synthesis of Infectious Bacteriophages in an E. coli-based Cell-free Expression System

A new generation of cell-free transcription-translation platforms has been engineered to construct biochemical systems in vitro through the execution of gene circuits. In this article, we describe how bacteriophages, such as MS2, &#934;&#935;174, and T7, are synthesized from their genome using an all E. coli cell-free TXTL system.

A new generation of cell-free transcription-translation (TXTL) systems, engineered to have a greater versatility and modularity, provide novel ...

Bioengineering

CFPS Protocols for Micro-Compartmentalized Synthetic Cells

Cell-Free Protein Synthesis System for Building Synthetic Cells

The Cell-Free Protein Synthesis (CFPS) system has been widely employed to facilitate the bottom-up assembly of synthetic cells. It serves as the host for the core machinery of the Central Dogma, standing as an optimal chassis for the integration and assembly of diverse artificial cellular mimicry systems. Despite its frequent use in the fabrication of synthetic cells, establishing a tailored and robust CFPS system for a specific application remains a nontrivial challenge. In this methods paper, we present a comprehensive protocol for the CFPS system, routinely employed in constructing synthetic cells. This protocol encompasses key stages in the preparation of the CFPS system, including the cell extract, template preparation, and routine expression optimization utilizing a fluorescent reporter protein. Additionally, we show representative results by encapsulating the CFPS system within various micro-compartments, such as monolayer droplets, double-emulsion vesicles, and chambers situated atop supported lipid bilayers. Finally, we elucidate the critical steps and conditions necessary for the successful assembly of these CFPS systems in distinct environments. We expect that our approach will facilitate the establishment of good working practices among various laboratories within the continuously expanding synthetic cell community, thereby accelerating progress in the field of synthetic cell development.

Author Spotlight: Optimizing CFPS Systems for Synthetic Cell Construction

This protocol describes the Cell-Free Protein Synthesis (CFPS) system used in constructing synthetic cells. It outlines key stages with representative results in different micro-compartments. The protocol aims to establish best practices for diverse laboratories in the synthetic cell community, advancing progress in synthetic cell development.

The Cell-Free Protein Synthesis (CFPS) system has been widely employed to facilitate the bottom-up assembly of synthetic cells. It serves as the host for ...

Biochemistry

Cell-free Protein Expression Using the Rapidly Growing Bacterium Vibrio natriegens

The marine bacterium Vibrio natriegens has garnered considerable attention as an emerging microbial host for biotechnology due to its fast growth rate. A general protocol is described for the preparation of V. natriegens crude cell extracts using common laboratory equipment. This high yielding protocol has been specifically optimized for user accessibility and reduced cost. Cell-free protein synthesis (CFPS) can be carried out in small scale 10 &#956;L batch reactions in either a 96- or 384-well format and reproducibly yields concentrations of &#62; 260 &#956;g/mL super folder GFP (sfGFP) within 3 h. Overall, crude cell extract preparation and CFPS can be achieved in 1&#8722;2 full days by a single user. This protocol can be easily integrated into existing protein synthesis pipelines to facilitate advances in bio-production and synthetic biology applications.

Cell-free Protein Expression Using the Rapidly Growing Bacterium Vibrio natriegens

Cell-free expression systems are powerful and cost-efficient tools for the high-throughput synthesis and screening of important proteins. Here, we describe the preparation of cell-free protein expression system using Vibrio natriegens for the rapid protein production using plasmid DNA, linear DNA, and mRNA template.

The marine bacterium Vibrio natriegens has garnered considerable attention as an emerging microbial host for biotechnology due to its fast growth rate. A ...

Preparing Protein Producing Synthetic Cells using Cell Free Bacterial Extracts, Liposomes and Emulsion Transfer

The bottom-up assembly approach for construction of synthetic cells is an effective tool for isolating and investigating cellular processes in a cell mimicking environment. Furthermore, the development of cell-free expression systems has demonstrated the ability to reconstitute the protein production, transcription and translation processes (DNA&#8594;RNA&#8594;protein) in a controlled manner, harnessing synthetic biology. Here we describe a protocol for preparing a cell-free expression system, including the production of a potent bacterial lysate and encapsulating this lysate inside cholesterol-rich lipid-based giant unilamellar vesicles (GUVs) (i.e., stable liposomes), to form synthetic cells. The protocol describes the methods for preparing the components of the synthetic cells including the production of active bacterial lysates, followed by a detailed step-by-step preparation of the synthetic cells based on a water-in-oil emulsion transfer method. These facilitate the production of millions of synthetic cells in a simple and affordable manner with a high versatility for producing different types of proteins. The obtained synthetic cells can be used to investigate protein/RNA production and activity in an isolated environment, in directed evolution, and also as a controlled drug delivery platform for on-demand production of therapeutic proteins inside the body.

This protocol describes the method, materials, equipment and steps for bottom-up preparation of RNA and protein producing synthetic cells. The inner aqueous compartment of the synthetic cells contained the S30 bacterial lysate encapsulated within a lipid bilayer (i.e., stable liposomes), using a water-in-oil emulsion transfer method.

The bottom-up assembly approach for construction of synthetic cells is an effective tool for isolating and investigating cellular processes in a cell ...

Rapid, Enzymatic Methods for Amplification of Minimal, Linear Templates for Protein Prototyping using Cell-Free Systems

This protocol describes the design of a minimal DNA template and the steps for enzymatic amplification, enabling rapid prototyping of assayable proteins in less than 24 h using cell-free expression. After receiving DNA from a vendor, the gene fragment is PCR-amplified, cut, circularized, and cryo-banked. A small amount of the banked DNA is then diluted and amplified significantly (up to 106x) using isothermal rolling circle amplification (RCA). RCA can yield microgram quantities of the minimal expression template from picogram levels of starting material (mg levels if all starting synthetic fragment is used). In this work, a starting amount of 20 pg resulted in 4 &#181;g of the final product. The resulting RCA product (concatemer of the minimal template) can be added directly to a cell-free reaction with no purification steps. Due to this method being entirely PCR-based, it may enable future high-throughput screening efforts when coupled with automated liquid handling systems.

The study describes a protocol for creating large (&#181;g-mg) quantities of DNA for protein screening campaigns from synthetic gene fragments without cloning or using living cells. The minimal template is enzymatically digested and circularized and then amplified using isothermal rolling circle amplification. Cell-free expression reactions could be performed with the unpurified product.

This protocol describes the design of a minimal DNA template and the steps for enzymatic amplification, enabling rapid prototyping of assayable proteins ...

Rapid Characterization of Genetic Parts with Cell-Free Systems

Characterizing and cataloging genetic parts are critical to the design of useful genetic circuits. Having well-characterized parts allows for the fine-tuning of genetic circuits, such that their function results in predictable outcomes. With the growth of synthetic biology as a field, there has been an explosion of genetic circuits that have been implemented in microbes to execute functions pertaining to sensing, metabolic alteration, and cellular computing. Here, we show a rapid and cost-effective method for characterizing genetic parts. Our method utilizes cell-free lysate, prepared in-house as a medium to evaluate parts via the expression of a reporter protein. Template DNA is prepared by PCR amplification using inexpensive primers to add variant parts to the reporter gene, and the template is added to the reaction as linear DNA without cloning. Parts that can be added in this way include promoters, operators, ribosome binding sites, insulators, and terminators. This approach, combined with the incorporation of an acoustic liquid handler and 384-well plates, allows the user to carry out high-throughput evaluations of genetic parts in a single day. By comparison, cell-based screening approaches require time-consuming cloning and have longer testing times due to overnight culture and culture density normalization steps. Further, working in cell-free lysate allows the user to exact tighter control over the expression conditions through the addition of exogenous components and DNA at precise concentrations. Results obtained from cell-free screening can be used directly in applications of cell-free systems or, in some cases, as a way to predict function in whole cells.

Well-characterized genetic parts are necessary for the design of novel genetic circuits. Here we describe a cost-effective, high-throughput method for rapidly characterizing genetic parts. Our method reduces cost and time by combining cell-free lysates, linear DNA to avoid cloning, and acoustic liquid handling to increase throughput and reduce reaction volumes.

Characterizing and cataloging genetic parts are critical to the design of useful genetic circuits. Having well-characterized parts allows for the ...

From Cells to Cell-Free Protein Synthesis within 24 Hours Using Cell-Free Autoinduction Workflow

Cell-free protein synthesis (CFPS) has grown as a biotechnology platform that captures transcription and translation machinery in vitro. Numerous developments have made the CFPS platform more accessible to new users and have expanded the range of applications. For lysate based CFPS systems, cell extracts can be generated from a variety of organisms, harnessing the unique biochemistry of that host to augment protein synthesis. Within the last 20 years, Escherichia coli (E. coli) has become one of the most widely used organisms for supporting CFPS due to its affordability and versatility. Despite numerous key advances, the workflow for E. coli cell extract preparation has remained a key bottleneck for new users to implement CFPS for their applications. The extract preparation workflow is time-intensive and requires technical expertise to achieve reproducible results. To overcome these barriers, we previously reported the development of a 24 hour cell-free autoinduction (CFAI) workflow that reduces user input and technical expertise required. The CFAI workflow minimizes the labor and technical skill required to generate cell extracts while also increasing the total quantities of cell extracts obtained. Here we describe that workflow in a step-by-step manner to improve access and support the broad implementation of E. coli based CFPS.

This work describes the preparation of cell extract from Escherichia coli (E. coli) followed by cell-free protein synthesis (CFPS) reactions in under 24 hours. Explanation of the cell-free autoinduction (CFAI) protocol details improvements made to reduce researcher oversight and increase quantities of cell extract obtained.

Cell-free protein synthesis (CFPS) has grown as a biotechnology platform that captures transcription and translation machinery in vitro. Numerous ...

Cell-Free Protein Synthesis from Exonuclease-Deficient Cellular Extracts Utilizing Linear DNA Templates

Cell-free protein synthesis (CFPS) has recently become very popular in the field of synthetic biology due to its numerous advantages. Using linear DNA templates for CFPS will further enable the technology to reach its full potential, decreasing the experimental time by eliminating the steps of cloning, transformation, and plasmid extraction. Linear DNA can be rapidly and easily amplified by PCR to obtain high concentrations of the template, avoiding potential in vivo expression toxicity. However, linear DNA templates are rapidly degraded by exonucleases that are naturally present in the cell extracts. There are several strategies that have been proposed to tackle this problem, such as adding nuclease inhibitors or chemical modification of linear DNA ends for protection. All these strategies cost extra time and resources and are yet to obtain near-plasmid levels of protein expression. A detailed protocol for an alternative strategy is presented here for using linear DNA templates for CFPS. By using cell extracts from exonuclease-deficient knockout cells, linear DNA templates remain intact without requiring any end-modifications. We present the preparation steps of cell lysate from Escherichia coli BL21 Rosetta2 &#916;recBCD strain by sonication lysis and buffer calibration for Mg-glutamate (Mg-glu) and K-glutamate (K-glu) specifically for linear DNA. This method is able to achieve protein expression levels comparable to that from plasmid DNA in E. coli CFPS.

Presented here is a protocol for the preparation and buffer calibration of cell extracts from exonuclease V knockout strains of Escherichia coli BL21 Rosetta2 (&#916;recBCD and &#916;recB). This is a fast, easy, and direct approach for expression in cell-free protein synthesis systems using linear DNA templates.

Cell-free protein synthesis (CFPS) has recently become very popular in the field of synthetic biology due to its numerous advantages. Using linear DNA ...

Protocols for Implementing an Escherichia coli Based TX-TL Cell-Free Expression System for Synthetic Biology

Ideal cell-free expression systems can theoretically emulate an in vivo cellular environment in a controlled in vitro platform.1 This is useful for expressing proteins and genetic circuits in a controlled manner as well as for providing a prototyping environment for synthetic biology.2,3 To achieve the latter goal, cell-free expression systems that preserve endogenous Escherichia coli transcription-translation mechanisms are able to more accurately reflect in vivo cellular dynamics than those based on T7 RNA polymerase transcription. We describe the preparation and execution of an efficient endogenous E. coli based transcription-translation (TX-TL) cell-free expression system that can produce equivalent amounts of protein as T7-based systems at a 98% cost reduction to similar commercial systems.4,5 The preparation of buffers and crude cell extract are described, as well as the execution of a three tube TX-TL reaction. The entire protocol takes five days to prepare and yields enough material for up to 3000 single reactions in one preparation. Once prepared, each reaction takes under 8 hr from setup to data collection and analysis. Mechanisms of regulation and transcription exogenous to E. coli, such as lac/tet repressors and T7 RNA polymerase, can be supplemented.6 Endogenous properties, such as mRNA and DNA degradation rates, can also be adjusted.7 The TX-TL cell-free expression system has been demonstrated for large-scale circuit assembly, exploring biological phenomena, and expression of proteins under both T7- and endogenous promoters.6,8 Accompanying mathematical models are available.9,10 The resulting system has unique applications in synthetic biology as a prototyping environment, or "TX-TL biomolecular breadboard."

Protocols for Implementing an Escherichia coli Based TX-TL Cell-Free Expression System for Synthetic Biology

This five-day protocol outlines all steps, equipment, and supplemental software necessary for creating and running an efficient endogenous Escherichia coli based TX-TL cell-free expression system from scratch. With reagents, the protocol takes 8 hours or less to setup a reaction, collect, and process data.

Ideal cell-free expression systems can theoretically emulate an in vivo cellular environment in a controlled in vitro platform.1 This is useful for ...

Escherichia coli-Based Cell-Free Protein Synthesis: Protocols for a robust, flexible, and accessible platform technology

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