The authors thank Zachary Sun and Richard Murray (California Institute of Technology) for kindly providing the plasmid P_araBAD-gamS, and Kaeko Kamei (Kyoto Institute of Technology) for kindly providing a high-speed cooling centrifuge. This work was supported by grants from the National Institutes of Health and from the ARO MURI program and was partly supported by the Leading Initiative for Excellent Young Researchers, MEXT, Japan.

1. Prepare media and buffers.
<ol>
	<li>Prepare 2xYTPG medium.
	<ol>
		<li>Mix 62 g 2xYT powder, 5.99 g potassium phosphate monobasic, 13.93 g potassium phosphate dibasic, and deionized water to 2 L.</li>
		<li>Autoclave on liquid cycle with an exposure time of 30 min14.</li>
		<li>To 400 mL of 2xYTP media from 1.1.2, add 7.2 g D-glucose (dextrose) and mix until dissolved.</li>
		<li>Filter-sterilize through a 0.2 &#181;m filter.</li>
	</ol>
	</li>
	<li>Prepare S30A buffer.
	<ol>
		<li>Mix Tris-HCl (pH 7.7, 50 mM final concentration), potassium glutamate (60 mM final), and magnesium glutamate (14 mM final).</li>
		<li>Adjust the pH to 7.7 using 10 M KOH.</li>
	</ol>
	</li>
	<li>Prepare Solution 1 (see Table 1).
	<ol>
		<li>Resuspend 4-(2-hydroxyethyl)-1- piperazineethanesulfonic acid (HEPES) in 2 mL of water.</li>
		<li>Adjust the pH to 8.0 using KOH.</li>
		<li>Add all other components from Table 1.</li>
		<li>Adjust the pH to 7.6 using 10 M KOH. Filter-sterilize.</li>
	</ol>
	</li>
	<li>Prepare 2.5x premix solution (see Table 2).
	<ol>
		<li>Mix all the components in Table 2.</li>
		<li>Adjust the pH to 7.5 using KOH.</li>
		<li>Aliquot and freeze at -80 &#176;C. 
		&#8203;NOTE: A 20 &#181;L reaction uses 8.9 &#181;L of premix.</li>
	</ol>
	</li>
</ol>
2. Prepare cells.
<ol>
	<li>Streak the autolysate cells onto LB agar plates containing 50 &#181;g/mL ampicillin using an inoculating loop and grow at 37 &#176;C (see Note 1).</li>
	<li>Pick a single colony into a starter culture of LB/ampicillin medium using a pipet tip and grow at 37 &#176;C overnight.</li>
	<li>Inoculate 400 mL of 2xYTPG medium containing 50 &#181;g/mL ampicillin with 400 &#181;L of starter culture, and grow at 37 &#176;C in a 1 L Erlenmeyer flask, shaking at 300 rpm.</li>
	<li>Periodically measure the culture&#39;s optical density at 600 nm (OD600) using a spectrophotometer to read an optical cuvette with a 1 cm path length. When the OD600 exceeds 1, begin diluting the culture 5-fold before measurements to ensure that the measurements remain within the linear range of a typical laboratory spectrophotometer. Continue growing the cells until the 5-fold diluted culture reaches OD600 of 0.3 (corresponding to a culture OD600 of 1.5).</li>
</ol>
3. Prepare the lysate.
<ol>
	<li>Prepare S30A buffer supplemented with 2 mM dithiothreitol (DTT). Mix 3&#160;mL of S30A buffer with 6&#160;&#181;L of DTT stock solution at 1 M. Place on ice for use in step 3.7.</li>
	<li>Harvest the cells by centrifuging at 1800 &#215; g for 15 min at room temperature.</li>
	<li>Discard the supernatant by pouring it off and using a pipet to remove any remaining liquid.</li>
	<li>Resuspend the pellet in 45 mL of cold (4-10 &#176;C) S30A buffer using a vortex mixer.</li>
	<li>Weigh an empty 50 mL centrifuge tube, transfer the cells into it, and repeat steps 3.2-3.3 to wash the cells.</li>
	<li>Weigh the pellet, subtracting the weight of an empty 50 mL tube. Make sure to carefully aspirate any remaining supernatant to ensure an accurate measurement of pellet weight. 
	NOTE: A typical yield is ~1.3 g of cell pellet from 400 mL of production culture.</li>
	<li>Add 2 volumes of cold S30A buffer supplemented with 2 mM dithiothreitol, i.e., 2 mL of buffer for every 1 g of cell pellet, and resuspend the cells by vigorously vortex mixing.</li>
	<li>Freeze the cells. Place the 50 mL tube containing the&#160;cells in a -20 &#176;C or -80 &#176;C freezer until the pellet is thoroughly frozen. 
	NOTE: The freezing step is a good stopping point for the day.</li>
	<li>Thaw the cells in a room temperature water bath.</li>
	<li>Vortex vigorously for 2-3 min.</li>
	<li>Incubate at 37 &#176;C for 45 min with shaking at 300 rpm.</li>
	<li>Clear the sample of heavy cellular debris by centrifuging in transparent centrifuge tubes at 30,000 &#215; g for 45 min at 4 &#176;C. 
	NOTE: If a centrifuge capable of 30,000 x g is not available, centrifuge for 45 min at 21,000 &#215; g, and use additional caution in step 3.13, as the pellet will be less compact.</li>
	<li>Carefully transfer the supernatant to a new tube with a pipet, avoiding disturbing the pellet as much as possible. If the transferred supernatant is contaminated with material from the pellet, repeat the previous step.</li>
	<li>Transfer the supernatant to 1.5 mL centrifuge tubes and centrifuge once more at 21,000 &#215; g (or the maximum speed of a tabletop centrifuge) for 5 min.</li>
	<li>Aliquot the cleared autolysate into the desired volumes, carefully avoiding any remaining pellet, and freeze at -80 &#176;C or use immediately. 
	&#8203;NOTE: A single 20 &#181;L reaction uses 8 &#181;L of autolysate.</li>
</ol>
4. Cell-free gene expression
NOTE: The autolysate is now ready for any desired end-use. The following is an example standard protocol for cell-free gene expression.
<ol>
	<li>For a 20 &#181;L reaction, mix on ice 8 &#181;L of autolysate and 8.9 &#181;L of premix. See&#160;NOTE&#160;at the end of the protocol section regarding the optimization of magnesium glutamate and PEG 8000 concentrations.</li>
	<li>Add DNA (e.g., pBEST-OR2-OR1-Pr-UTR1-deGFP-T500 to a final concentration of 8 nM), any other reagents, and water to 20 &#181;L.</li>
	<li>Place the reaction in a 384-well microplate and measure the fluorescence time course and/or endpoints using a plate reader. For green fluorescent protein (GFP), use an excitation wavelength of 485 nm and an emission wavelength of 520 nm.</li>
</ol>
5. Protocol modifications
NOTE: The following modifications of the protocol enable it to serve other applications.
<ol>
	<li>Cell-free gene expression using linear DNA templates
	<ol>
		<li>Perform the steps in section 4, supplementing the reaction with 2.2 &#181;M purified GamS protein (expressed and purified as described12) before the addition of the linear DNA.</li>
	</ol>
	</li>
	<li>Cell-free gene expression incorporating protein degradation
	<ol>
		<li>In step 2.1, use autolysate cells containing the plasmid pACYC-FLAG-dN6-His (see the Table of Materials). In all growth media, additionally include 34 &#181;g/mL chloramphenicol.</li>
		<li>In step 2.3, include 40 &#181;M isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) in the growth medium to induce expression from the plasmid.</li>
		<li>Repeat steps 3.2-3.4 (washing) two additional times (for a total of three washes) to ensure the complete removal of chloramphenicol, which is a translation inhibitor. For the first two washes (step 3.4), substitute S30A buffer with phosphate-buffered saline (pH 7.4).</li>
		<li>In step 4.2, supplement with an additional 3 mM ATP (added from a stock solution of 100 mM ATP in water, pH 7.2) and 4.5 mM magnesium glutamate (using a 1 M stock solution in water) (final concentrations) to compensate for high ATP use by ClpXP, as well as chelation of magnesium by the additional ATP.</li>
	</ol>
	</li>
	<li>Cell-free gene expression using LacZ-based readouts (including colorimetric)
	<ol>
		<li>In step 2.1, use autolysate cells that do not natively express LacZ (see the Table of Materials).</li>
	</ol>
	</li>
	<li>Alternatively, prepare the lysate directly from freeze-dried cells.
	<ol>
		<li>Perform all steps from 1.1 to 3.7.</li>
		<li>Mix 8 &#181;L of cell suspension with 8.9 &#181;L of premix.</li>
		<li>Add plasmid DNA (if desired), other custom reagents, and water to reach a final volume of 20 &#181;L.</li>
		<li>Transfer the reaction to a 384-well microplate and freeze-dry it. 
		NOTE: Freeze-dried samples can be stored for up to a week and possibly longer.</li>
		<li>To begin the reaction, rehydrate it with 18 &#181;L of deionized water supplemented with any desired DNA or other reagents.</li>
		<li>Follow the fluorescence dynamics in a plate reader.</li>
	</ol>
	</li>
</ol>
NOTE&#160;1: Autolysate cells are designed to lyse upon a freeze-thaw cycle, so it is particularly important to use cryoprotectant when making frozen stocks. We froze stocks in 24% wt/vol glycerol and stored them at -80 &#176;C.
NOTE 2: Magnesium ions and PEG 8000 are critical for lysate performance. The base 2.5x premix, based on previously published data, contains 6 mM magnesium glutamate and 4.8% wt/vol PEG 8000, which become 2.4 mM and 1.9%, respectively, in the final reaction. The autolysate prepared with the protocol here typically performs best with an additional 5 mM Mg-glutamate and 1.5% PEG 8000 in the final reaction. However, this can be optimized in the range of an additional 0-10 mM Mg glutamate and an additional 0-3% PEG 8000 (compared to the base premix). To prepare the premix with the recommended additional 5 mM Mg-glutamate and 1.5% PEG 8000 (final concentrations), mix 380 &#181;L of premix with 4.75 &#181;L of magnesium glutamate at 1 M and 36.1 &#181;L of PEG 8000 at 40% weight/volume.

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	<li>Dudley, Q. M., Karim, A. S., Jewett, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-free+metabolic+engineering:+biomanufacturing+beyond+the+cell.">Cell-free metabolic engineering: biomanufacturing beyond the cell.</a> Biotechnology Journal. 10 (1), 69-82 (2015).</li><li>Smith, M. T., Wilding, K. M., Hunt, J. M., Bennett, A. M., Bundy, B. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+emerging+age+of+cell-free+synthetic+biology.">The emerging age of cell-free synthetic biology.</a> FEBS Letters. 588 (17), 2755-2761 (2014).</li><li>Casteleijn, M. G., Urtti, A., Sarkhel, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+without+boundaries:+cell-free+protein+synthesis+in+pharmaceutical+research.">Expression without boundaries: cell-free protein synthesis in pharmaceutical research.</a> International Journal of Pharmaceutics. 440 (1), 39-47 (2013).</li><li>He, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-free+protein+synthesis:+applications+in+proteomics+and+biotechnology.">Cell-free protein synthesis: applications in proteomics and biotechnology.</a> New Biotechnology. 25 (2-3), 126-132 (2008).</li><li>Pardee, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Paper-based+synthetic+gene+networks.">Paper-based synthetic gene networks.</a> Cell. 159 (4), 940-954 (2014).</li><li>Pardee, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=low-cost+detection+of+Zika+virus+using+programmable+biomolecular+components.">low-cost detection of Zika virus using programmable biomolecular components.</a> Cell. 165 (5), 1255-1266 (2016).</li><li>Takahashi, M. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+and+prototyping+genetic+networks+with+cell-free+transcription-translation+reactions.">Characterizing and prototyping genetic networks with cell-free transcription-translation reactions.</a> Methods. 86, 60-72 (2015).</li><li>Siegal-Gaskins, D., Tuza, Z. A., Kim, J., Noireaux, V., Murray, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+circuit+performance+characterization+and+resource+usage+in+a+cell-free+"breadboard".">Gene circuit performance characterization and resource usage in a cell-free "breadboard".</a> ACS Synthetic Biology. 3 (6), 416-425 (2014).</li><li>Niederholtmeyer, H., Chaggan, C., Devaraj, N. 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J.H. is a co-founder of GenCirq Inc, which focus on cancer therapeutics. He is on the Board of Directors and has equity in GenCirq.

The protocol described here yields highly active bacterial lysate for cell-free gene expression. The key is to use cells carrying the plasmid pAD-LyseR, which expresses the lambda phage endolysin cytosolically. These cells are potentiated to lyse themselves upon permeabilization of the inner membrane, allowing the endolysin access to the cell wall, which the method achieves through a simple freeze-thaw cycle. Because the cells effectively lyse themselves, the product is referred to as autolysate. After the cells have lys...

Gene expression in cell-free lysates has several advantages over using live cells1,2,3,4. Lysates can be easily modified biochemically and used in conditions that could be detrimental to or impossible to achieve in live cells. Gene expression circuits do not have to contend or compete with host biological processes, and testing new genetic circuits is as simple as adding DNA. For these reasons, cell-free gene expression has found various applications, from biosensors5,6 to rapidly prototyping synthetic gene circuits7,8 to developing artificial cells9. Most cell-free gene expression utilizes cellular lysates that have been highly processed, generally requiring long and complex protocols, specialized equipment, and/or sensitive steps that can lead to significant variation between users and batches10,11.
This paper describes a simple, efficient method for producing cell-free lysate that requires minimal processing and expertise (Figure 1A)12. The method relies on E. coli cells that are engineered to lyse following a simple freeze-thaw cycle. The cells express an endolysin from phage lambda that degrades the cell wall. As the cells are growing, this endolysin remains in the cytoplasm, sequestered from the cell wall. However, a simple freeze-thaw cycle disrupts the cytoplasmic membrane, releasing the endolysin into the periplasm, where it degrades the cell wall, resulting in rapid cell lysis. The protocol can be completed with only a few hours of hands-on work and requires only a freezer, a centrifuge capable of 30,000 &#215; g (for optimal results; lower speeds can be used with more care not to disturb the pellet), a vortex mixer, and a simple buffer solution. Functional lysate can even be produced by freeze-drying the cells and rehydrating them in situ. However, this method produces lysates with lower activity, presumably&#160;due to the remaining cell debris.
The lysates are highly active for cell-free gene expression, and they can be enhanced in various ways depending on the end use. The rate of protein synthesis can be further increased by concentrating the lysate using standard spin concentrators. Linear DNA can be protected from degradation by adding purified GamS protein. Protein degradation, necessary for more complex circuit dynamics such as oscillation, can be achieved by co-expressing a ClpX hexamer in the autolysate-producing strain13. Finally, LacZ-based visual readouts are enabled by using an autolysate strain lacking lacZ. Overall, this method produces highly active cell-free lysate that is suitable for a wide range of applications.

<table><tbody><tr><td>2xYT media</td><td>EMD Millipore</td><td>4.85008</td><td>or equivalent</td></tr><tr><td>3-PGA</td><td>Sigma Aldrich</td><td>P8877</td><td>or equivalent</td></tr><tr><td>Amicon Ultra-15 centrifugal filter unit, 3 kDa cutoff</td><td>Millipore Sigma</td><td>UFC900308</td><td>optional, can be used to concentrate lysate, select concentrator capacity appropriate for the volume to be concentrated</td></tr><tr><td>ampicillin</td><td>Sigma Aldrich</td><td>A0166-5G</td><td>or carbenicillin, a more stable variant</td></tr><tr><td>ATP</td><td>Sigma Aldrich</td><td>A8937</td><td>or equivalent</td></tr><tr><td>cAMP</td><td>Sigma Aldrich</td><td>A9501</td><td>or equivalent</td></tr><tr><td>CoA</td><td>Sigma Aldrich</td><td>C4282</td><td>or equivalent</td></tr><tr><td>CTP</td><td>United States Biosciences</td><td>14121</td><td>or equivalent</td></tr><tr><td>D-glucose (dextrose)</td><td>Fisher Scientific</td><td>AAA1749603</td><td>or equivalent</td></tr><tr><td>dithiothreitol (DTT)</td><td>Sigma Aldrich</td><td>D0632-1G</td><td>or equivalent</td></tr><tr><td>E. coli BL21-Gold (DE3) carrying pAD-LyseR</td><td>Addgene</td><td>99244</td><td></td></tr><tr><td>E. coli BL21-Gold (DE3) &amp;Delta;lacZ carrying pAD-LyseR</td><td>Addgene</td><td>99245</td><td></td></tr><tr><td>Folinic acid</td><td>Sigma Aldrich</td><td>F7878</td><td>or equivalent</td></tr><tr><td>GTP</td><td>United States Biosciences</td><td>16800</td><td>or equivalent</td></tr><tr><td>HEPES</td><td>Sigma Aldrich</td><td>H3375-25G</td><td>or equivalent</td></tr><tr><td>LB media</td><td>Fisher Scientific</td><td>DF0446075</td><td>or equivalent</td></tr><tr><td>magnesium glutamate</td><td>Sigma Aldrich</td><td>49605-250G</td><td>or equivalent</td></tr><tr><td>NAD</td><td>Sigma Aldrich</td><td>N6522</td><td>or equivalent</td></tr><tr><td>potassium glutamate</td><td>Sigma Aldrich</td><td>G1501-100G</td><td>or equivalent</td></tr><tr><td>potassium hydroxide (KOH)</td><td>Sigma Aldrich</td><td>221473-25G</td><td>for adjusting pH</td></tr><tr><td>potassium phosphate dibasic</td><td>Fisher Scientific</td><td>BP363-500</td><td>or equivalent</td></tr><tr><td>potassium phosphate monobasic</td><td>Fisher Scientific</td><td>BP362-500</td><td>or equivalent</td></tr><tr><td>Spermidine</td><td>Sigma Aldrich</td><td>85558</td><td>or equivalent</td></tr><tr><td>Tris-HCl</td><td>Fisher Scientific</td><td>9310500GM</td><td>or equivalent</td></tr><tr><td>tRNA mix</td><td>Roche</td><td>10109541001</td><td>or equivalent</td></tr><tr><td>UTP</td><td>United States Biosciences</td><td>23160</td><td>or equivalent</td></tr></tbody></table>

rapid, affordable, and uncomplicated production of bacterial cell-free lysate

Cell-free gene expression offers the power of biology without the complications of a living organism. Although many such gene expression systems exist, most are quite expensive to buy and/or require special equipment and finely honed expertise to produce effectively. This protocol describes a method to produce bacterial cell-free lysate that supports high levels of gene expression, using only standard laboratory equipment and requiring minimal processing. The method uses an Escherichia coli strain producing an endolysin that does not affect growth, but which efficiently lyses a harvested cell pellet following a simple freeze-thaw cycle. The only further processing required is a brief incubation followed by centrifugation to clear the autolysate of cellular debris. Dynamic gene circuits can be achieved through heterologous expression of the ClpX protease in the cells before harvesting. An E. coli strain lacking the lacZ gene can be used for high-sensitivity, cell-free biosensing applications using a colorimetric or fluorescent readout. The entire protocol requires as few as 8-9 hours, with only 1-2 hours of hands-on labor from inoculation to completion. By reducing the cost and time to obtain cell-free lysate, this method should increase the affordability of cell-free gene expression for various applications.

Representative results can be observed by using autolysate to express GFP from a constitutively expressing plasmid, here pBEST-OR2-OR1-Pr-UTR1-deGFP-T500, and recording a time course of GFP fluorescence in a plate reader (Figure 1B). A dilution series of plasmid DNA found strong expression even at 1 nM DNA. Compared to a commercially available lysate, the autolysate can produce a greater total yield and achieved a greater maximum production rate, calculated as the time derivative of the GFP ...

Watch this Scientific Journal Video about Rapid, Affordable, and Uncomplicated Production of Bacterial Cell-free Lysate at JoVE.com

Rapid, Affordable, and Uncomplicated Production of Bacterial Cell-free Lysate

This protocol describes a rapid and simple method to produce bacterial lysate for cell-free gene expression, using an engineered strain of Escherichia coli and requiring only standard laboratory equipment.

Cell-free gene expression offers the power of biology without the complications of a living organism. Although many such gene expression systems exist, ...

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4.2K Views.  University of California, San Diego. This protocol describes a rapid and simple method to produce bacterial lysate for cell-free gene expression, using an engineered strain of Escherichia coli and requiring only standard laboratory equipment.

Video: Rapid, Affordable, and Uncomplicated Production of Bacterial Cell-free Lysate

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.

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.

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

Chemistry

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 ...

Bioengineering

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 ...

A Convenient and General Expression Platform for the Production of Secreted Proteins from Human Cells

Recombinant protein expression in bacteria, typically E. coli, has been the most successful strategy for milligram quantity expression of proteins. However, prokaryotic hosts are often not as appropriate for expression of human, viral or eukaryotic proteins due to toxicity of the foreign macromolecule, differences in the protein folding machinery, or due to the lack of particular co- or post-translational modifications in bacteria. Expression systems based on yeast (P. pastoris or S. cerevisiae) 1,2, baculovirus-infected insect (S. frugiperda or T. ni) cells 3, and cell-free in vitro translation systems 2,4 have been successfully used to produce mammalian proteins. Intuitively, the best match is to use a mammalian host to ensure the production of recombinant proteins that contain the proper post-translational modifications. A number of mammalian cell lines (Human Embryonic Kidney (HEK) 293, CV-1 cells in Origin carrying the SV40 larget T-antigen (COS), Chinese Hamster Ovary (CHO), and others) have been successfully utilized to overexpress milligram quantities of a number of human proteins 5-9. However, the advantages of using mammalian cells are often countered by higher costs, requirement of specialized laboratory equipment, lower protein yields, and lengthy times to develop stable expression cell lines. Increasing yield and producing proteins faster, while keeping costs low, are major factors for many academic and commercial laboratories.
Here, we describe a time- and cost-efficient, two-part procedure for the expression of secreted human proteins from adherent HEK 293T cells. This system is capable of producing microgram to milligram quantities of functional protein for structural, biophysical and biochemical studies. The first part, multiple constructs of the gene of interest are produced in parallel and transiently transfected into adherent HEK 293T cells in small scale. The detection and analysis of recombinant protein secreted into the cell culture medium is performed by western blot analysis using commercially available antibodies directed against a vector-encoded protein purification tag. Subsequently, suitable constructs for large-scale protein production are transiently transfected using polyethyleneimine (PEI) in 10-layer cell factories. Proteins secreted into litre-volumes of conditioned medium are concentrated into manageable amounts using tangential flow filtration, followed by purification by anti-HA affinity chromatography. The utility of this platform is proven by its ability to express milligram quantities of cytokines, cytokine receptors, cell surface receptors, intrinsic restriction factors, and viral glycoproteins. This method was also successfully used in the structural determination of the trimeric ebolavirus glycoprotein 5,10.
In conclusion, this platform offers ease of use, speed and scalability while maximizing protein quality and functionality. Moreover, no additional equipment, other than a standard humidified CO2 incubator, is required. This procedure may be rapidly expanded to systems of greater complexity, such as co-expression of protein complexes, antigens and antibodies, production of virus-like particles for vaccines, or production of adenoviruses or lentiviruses for transduction of difficult cell lines.

In the post-human genomics era, the availability of recombinant proteins in native conformations is crucial to structural, functional and therapeutic research and development. Here, we describe a test- and large-scale protein expression system in human embryonic kidney 293T cells that can be used to produce a variety of recombinant proteins.

Recombinant protein expression in bacteria, typically E. coli, has been the most successful strategy for milligram quantity expression of proteins. ...

OnePot PURE Cell-Free System

The defined PURE (protein synthesis using recombinant elements) transcription-translation system provides an appealing chassis for cell-free synthetic biology. Unfortunately, commercially available systems are costly, and their tunability is limited. In comparison, a home-made approach can be customized based on user needs. However, the preparation of home-made systems is time-consuming and arduous due to the need for ribosomes as well as 36 medium scale protein purifications. Streamlining protein purification by coculturing and co-purification allows for minimizing time and labor requirements. Here, we present an easy, adjustable, time- and cost-effective method to produce all PURE system components within 1 week, using standard laboratory equipment. Moreover, the performance of the OnePot PURE is comparable to commercially available systems. The OnePot PURE preparation method expands the accessibility of the PURE system to more laboratories due to its simplicity and cost-effectiveness.

We present a fast and cost-effective method to produce the recombinant PURE cell-free TX-TL system using standard laboratory equipment.

The defined PURE (protein synthesis using recombinant elements) transcription-translation system provides an appealing chassis for cell-free synthetic ...

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 ...

Rapid, Affordable, and Uncomplicated Production of Bacterial Cell-free Lysate

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

Cell-Free Protein Synthesis System for Building Synthetic Cells

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

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

A Convenient and General Expression Platform for the Production of Secreted Proteins from Human Cells

OnePot PURE Cell-Free System

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

Rapid Characterization of Genetic Parts with Cell-Free Systems

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

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