Name: Video: Rapid Characterization of Genetic Parts with Cell-Free Systems
Uploaded: 2021-08-30T14:00:00
Duration: 5 min

This work was made possible by the Office of the Secretary of Defense&#39;s Applied Research for the Advancement of Science and Technology Priorities program. We thank Scott Walper (Naval Research Laboratory) for providing the stock of sfGFP used, and Zachary Sun and Abel Chiao (Tierra Biosciences) for fruitful discussions related to prototyping with cell-free systems and related troubleshooting of acoustic liquid handling.

1. Preparation of cell extract
<ol>
	<li>Preparation of media
	<ol>
		<li>For 2xYT media: Add 16 g of tryptone, 10 g of yeast extract, and 5 g of NaCl to 900 mL of deionized water and adjust the pH to 7.0 with 5 M NaOH. Raise the solution volume to 1 L using deionized water and autoclave or filter sterilize. Alternatively, purchase 2xYT media.</li>
		<li>For S30B buffer: Prepare a solution of 14 mM Mg-glutamate, 60 mM K-glutamate, and 5 mM Tris in 2 L of deionized water. Use 2 M Tris to adjust the pH to 8.2, autoclave,&#160;and store at 4 &#176;C. Complete the solution by adding dithiothreitol (DTT) to a final concentration of 1 mM just before use.</li>
	</ol>
	</li>
	<li>Preparation of cells
	<ol>
		<li>Streak Escherichia coli BL21(DE3) Rosetta2 cells or other cell line of choice (see Cole et al.44 for a recent comprehensive review) onto an LB (Lysogeny Broth) agar plate and incubate at 37 &#176;C for 10-14 h.</li>
		<li>Use a single E. coli colony to inoculate 3 mL of 2xYT medium in a 10 mL culture tube. Incubate this tube at 37 &#176;C with shaking at 250 rpm for 8 h.</li>
		<li>Use 50 &#181;L from the 3 mL culture to inoculate 50 mL of 2xYT medium in a 500 mL flask. Incubate this flask at 37 &#176;C with shaking at 250 rpm for 8 h.</li>
		<li>Use 7.5 mL from the 50 mL culture to inoculate each of four 4 L baffled flasks containing 0.75 L of 2xYT medium. Incubate these flasks at 37 &#176;C with shaking at 220 rpm until they have reached an optical density at 600 nm of 2 to 4, after approximately 3-4 h.</li>
		<li>Harvest the cells from each flask by transferring them to 1 L containers and centrifuging at 5,000 x g for 12 min at 4&#160;&#176;C. Discard the supernatant by decanting into a waste container.</li>
		<li>Wash each cell pellet with 150 mL of ice-cold S30B buffer by completely resuspending them using a pipette to disrupt the cell mass, and then collect the cells again by centrifugation at 5,000 x g for 12 min. Discard the supernatant.</li>
		<li>Wash each cell pellet again in 40 mL of ice-cold S30B buffer by completely resuspending them and disrupting the cell mass using a pipette. Transfer the cells to pre-weighed 50 mL conical tubes and collect the cells again by centrifugation at 2,000 x g for 8 min&#160;at 4&#160;&#176;C. Discard the supernatant by decanting.</li>
		<li>Weigh the wet cell pellets. Flash-freeze the cell pellets by placing the tubes directly into liquid nitrogen and store at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Cell lysis
	<ol>
		<li>Thaw the cell pellets on ice.</li>
		<li>Resuspend each cell pellet in 1.4 mL of S30B buffer per 1 g of the cell pellet by vortexing.</li>
		<li>Lyse the cells by French pressure cell at 640 psi at 4 &#176;C. Collect the lysate in microcentrifuge tubes on ice and add 3 &#181;L of 1 M DTT per 1 mL of lysate immediately after lysis. 
		NOTE: It is best to tap the French press release valve with a small metal rod to maintain even pressure and avoid sudden drops in pressure.</li>
		<li>Clear the lysate by centrifugation at 30,000 x g for 30 min at 4 &#176;C and discard the pellet after pipetting the supernatant to a new ice-cold microcentrifuge tube, taking care not to disrupt the pellet.</li>
		<li>Centrifuge the supernatant a second time at 30,000 x g for 30 min at 4 &#176;C. Pipette the resulting supernatant into an ice-cold microcentrifuge tube. Discard the pellet.</li>
		<li>Incubate the supernatant in a 37 &#176;C water bath for 1 h.</li>
		<li>Clear the supernatant by centrifugation at 15,000 x g for 15 min at 4 &#176;C and transfer the resulting supernatant to an ice-cold microcentrifuge tube, taking care not to disrupt the pellet.</li>
		<li>Centrifuge the supernatant a second time at 15,000 x g for 15 min at 4 &#176;C and transfer the resulting supernatant to an ice-cold microcentrifuge tube, taking care not to disrupt any remaining pellet.</li>
		<li>Distribute the supernatant in 100 &#181;L aliquots into 1.5 mL microcentrifuge tubes and flash freeze them by placing directly into liquid nitrogen. Store the supernatant at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Linear template preparation
<ol>
	<li>Primer design
	<ol>
		<li>Choose a core sequence as the PCR template. Include at a minimum a reporter sequence, such as sfGFP (superfolder Green Fluorescent Protein), LacZ, or Spinach aptamer. Include other parts that will be fixed across screened variants, such as terminators, promoters or RBSs, as appropriate for the design. 
		NOTE: Inclusion of a terminator is not always required for expression from linear DNA in cell-free systems.</li>
		<li>For the forward primers, choose a minimum of 20 bp matching the 5&#697; end of the core sequence as the 3&#697; end of the primer. If adding parts to the 5&#8242; end of the construct, design the remainder of the 5&#697; end of the primer to add the genetic parts of interest to the core sequence via PCR amplification (Figure 1A and Figure 2). 
		NOTE: Since primers above ~60 bp frequently increase dramatically in cost, multiple overlapping primers can be designed to add longer sequences or multiple parts. While multiple primers can be used in a single PCR reaction, performing multiple rounds of PCR is recommended.</li>
		<li>For the reverse primers, choose a minimum of 20 bp to match the 3&#8242; end of the core sequence as the 3&#697; end of the primer. If adding parts to the 3&#8242; end of the construct, design the remainder of the 5&#697; end of the primer to add the genetic parts of interest to the core sequence via PCR amplification (Figure 1A and Figure 2). Ensure that the reverse primer&#39;s annealing temperature is within 5 &#176;C of the annealing temperature of the entire forward primer.</li>
	</ol>
	</li>
	<li>Linear template amplification
	<ol>
		<li>Determine the number of PCR reactions to be performed based on the number of core sequences and calculate the amount of each component required using Table 1.</li>
		<li>Prepare the master mix according to Table 1 and store it on ice. Aliquot 30 or 40 &#181;L (see Table 1) of the master mix into the determined number of PCR tubes and add 10 &#181;L of each variable primer (i.e., primers encoding a part change, see Table 1) at 5 &#181;M to appropriately labeled PCR tubes.</li>
		<li>Place the PCR tubes into the thermocycler and run the following PCR program: 98 &#176;C for 3 min; 30 cycles of 98 &#176;C for 15 s, XX &#176;C for 20 s, 72 &#176;C for YY min;&#160;final extension at 72 &#176;C for 10 min. Then, hold the reaction at 4 &#176;C. 
		Where, XX represents the annealing temperature for the primer with the lower annealing temperature and YY represents the extension time calculated for the length of the amplicon based on the manufacturer&#39;s recommendations for the high-fidelity polymerase used. Optimize these conditions as needed for different primers and/or templates.</li>
		<li>(Optional) Add 1 &#181;L of DpnI restriction enzyme to digest the original template. Incubate the reaction at 37 &#176;C for 1 h. Perform this step only if the original template is plasmid DNA.</li>
		<li>Analyze 5 &#181;L of each PCR product by gel electrophoresis. Separate the product using a 1% agarose gel at 180 V for 20 min. Check for the correct band size, which will vary with the chosen core sequence and the length of the parts added.</li>
	</ol>
	</li>
	<li>Purify the linear template using a commercial PCR purification kit or by the preferred PCR cleanup method. If multiple bands were present by gel electrophoresis analysis, either optimize the PCR conditions or purify the correct molecular weight bands using a commercial gel extraction kit as per the manufacturer&#39;s recommendation.</li>
	<li>Quantify each DNA template using a spectrophotometer. Assess the DNA template quality by checking that the 260 nm/280 nm ratio is approximately 1.8.</li>
	<li>(Optional) Again separate a portion of the DNA template using a 1% agarose gel at 180 V for 20 min and ensure that any unwanted bands were removed during template purification.</li>
	<li>Use purified DNA templates immediately or store at -20 &#176;C.</li>
</ol>
3. Purified protein preparation
<ol>
	<li>Protein expression
	<ol>
		<li>For each protein to be expressed, assemble an appropriate expression construct. Codon-optimize the gene for expression in E. coli. Insert the gene into a pET-22b expression vector or other appropriate expression vector via the preferred plasmid assembly method. Transform the expression plasmid into BL21(DE3) Rosetta2 expression cells or other appropriate cell line.</li>
		<li>For each protein, use a single colony to inoculate 3 mL of LB medium in a 10 mL culture tube. Incubate these tubes at 37 &#176;C with shaking at 250 rpm overnight.</li>
		<li>Inoculate a 2 L flask containing 750 mL of LB medium with 1 mL of the overnight culture. Incubate these flasks at 37 &#176;C with shaking at 250 rpm until they reach an OD600 of 0.6-1.0.</li>
		<li>Induce protein expression by adding 0.75 mL of 1 M isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) in water to each flask and continue to incubate these flasks at 37 &#176;C with shaking at 250 rpm for 4 h.</li>
		<li>Harvest the cells from each flask, using a 1 L centrifuge bottle, by centrifugation at 5,000 x g for 12 min. Discard the supernatant.</li>
		<li>Transfer the pellets to a 50 mL conical tube and weigh each pellet. Flash-freeze the cells in liquid nitrogen and store them at -80 &#176;C or proceed to step 3.2. 
		NOTE: 2-5 g of cells per 0.75 mL are expected to result from this step.</li>
	</ol>
	</li>
	<li>Protein purification by nickel affinity column chromatography
	<ol>
		<li>Prepare the lysis buffer by combining 50 mM Tris-Cl, 500 mM NaCl, and 5 mM imidazole. Adjust to pH 8.0.</li>
		<li>Prepare the wash buffer by combining 50 mM Tris-Cl, 500 mM NaCl, and 25 mM imidazole. Adjust to pH 8.0.</li>
		<li>Prepare the elution buffer by combining 50 mM Tris-Cl, 500 mM NaCl, and 250 mM imidazole. Adjust to pH 8.0.</li>
		<li>Prepare the dialysis buffer by combining 50 mM NaHPO4, 100 mM NaCl, and 2% DMSO. Adjust to pH 7.5.</li>
		<li>Thaw the cell pellets by placing the tubes in room temperature water. Add 5 mL of the lysis buffer per 1 g of the cell pellet and resuspend by vortexing.</li>
		<li>Lyse the cells by sonication. Separate the cell homogenate so that there is no more than 30 mL per 50 mL conical tube and keep each tube on ice. Lyse the cells using a sonicator with a 0.16 cm diameter probe in 15 s rounds with 30 s breaks, 10 times. 
		NOTE: Avoid foaming, as this denatures the protein. The formation of foam can be avoided by keeping the sonicator tip at least 2/3 submerged in the lysate while it is operational. Other methods of cell lysis besides sonication are also possible44.</li>
		<li>Clear the lysate by centrifugation at 15,000 x g for 30 min at 4 &#176;C and place the supernatant into a new 50 mL conical tube.</li>
		<li>Add 1 mL of nickel-nitrilotriacetic acid (Ni-NTA) resin for each 5 mL of supernatant. Divide the cell lysate/Ni-NTA slurry so that there is no more than 36 mL per 50 mL conical tube. Incubate at 4 &#176;C on a tube rotator at 10 rpm for 1 h.</li>
		<li>Load the resin by decanting the cell lysate/Ni-NTA slurry into a 2 mL bed volume chromatography column and collect the eluant if needed for further analysis, otherwise, discard. Wash the resin with 10 resin bed volumes of wash buffer.</li>
		<li>Collect the protein by adding three resin bed volumes of elution buffer to the column and concentrate the volume to 1.5 mL using a centrifugal concentrator with the appropriate molecular weight cut-off membrane for each protein.</li>
		<li>Dialyze the protein against 2 L of dialysis buffer at 4 &#176;C for 1 h. Dialyze the protein again against 2 L of dialysis buffer overnight at 4 &#176;C.</li>
		<li>Quantify the protein using its molar extinction coefficient and absorbance at 280 nm. Analyze the protein for purity by separating it using sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). Store the protein at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
4. Cell-free protein synthesis
<ol>
	<li>Preparation of CFPS reaction mixture
	<ol>
		<li>Prepare the Supplement Mix by following the Amino Acid Solution Preparation, Energy Solution Preparation, and Buffer Preparation steps in Sun et al.42. Store separately or combined at -80 &#176;C in aliquots. Ensure that final concentrations match those described in Sun et al.42 in the Experimental Execution of a TX-TL Reaction section.</li>
		<li>Prepare an additive to protect linear DNA from degradation. If using GamS33,45, prepare via steps in section 3 above, or obtain from a commercial vendor; for other approaches, check the corresponding literature46,47,48. Alternatively, use a CFPS system that does not require additives49.</li>
		<li>Prepare T7 polymerase, repressor proteins, and other additives using the steps in section 3 above or obtain from a commercial vendor.</li>
		<li>Determine the number of CFPS reactions to be performed and calculate the amount of each component required using Table 2. Modify the concentrations of the components, including adding or removing components as needed, and adjust the amount of water such that the final volume of each reaction mixture is always 10 &#181;L. Similarly, modify the master mix to facilitate dispensing of other components by acoustic liquid handling as desired (see the Discussion section).</li>
		<li>Thaw all the components on ice and prepare a master mix by mixing each component as calculated above. Mix all the components thoroughly by pipette. Pay careful attention to avoid precipitation, especially for the amino acid mixture. Keep the master mix on ice.</li>
		<li>Chill a 384-well plate on ice and distribute the master mix in 9 &#181;L aliquots into each well. 
		NOTE: It is possible to distribute these components by acoustic liquid handling, though care should be taken to ensure proper dispensing (see the Discussion section for troubleshooting).</li>
	</ol>
	</li>
	<li>Distribution of additional components by acoustic liquid handling
	<ol>
		<li>Calculate the amount of repressor protein (and other optional components) required for all the CFPS reactions.</li>
		<li>Thaw the repressor protein on ice and distribute it into an acoustic liquid handling source plate or other appropriate plate. Ensure that the appropriate amount of dead volume required for the type of source plate used is included.</li>
		<li>Distribute the repressor protein in 1 &#181;L volumes into the appropriate wells via the liquid handler. For more information on distribution troubleshooting, see the Discussion section.</li>
	</ol>
	</li>
	<li>Standard curves
	<ol>
		<li>Include a serial dilution of the purified reporter (see section 3 for protein purification)41 or appropriate chemical standard50&#160;on the plate to enable comparison of results with other studies and other labs. Choose a range of concentrations appropriate for the reporter used and expected expression range of the experiments.</li>
	</ol>
	</li>
	<li>Running CFPS reactions
	<ol>
		<li>Pre-warm the plate reader to 30 &#176;C. Set the plate reader to read at settings appropriate for the reporter used in the core sequence without&#160;shaking steps. 
		NOTE: While 30 &#176;C is used here, 29 &#176;C and 37 &#176;C are also commonly used and work well with this protocol. Other temperatures may be preferred for alternative cell-free reaction preparations. For read intervals, 10 min is sufficient to achieve a good resolution for the representative data presented here; however, other resolutions may be better depending on the reporter protein and the particular CFPS recipe.</li>
		<li>(Optional) Run a test reaction first to set the appropriate gain or sensitivity setting to capture the change in fluorescence without signal overflow.</li>
		<li>Seal the 384-well plate with an impermeable plastic sealable lid to prevent evaporation. If possible, on the instrument, set a 1 &#176;C vertical temperature gradient to limit condensation on the seal. Place the 384-well plate on the plate holder and begin reading.</li>
	</ol>
	</li>
</ol>

RMM has a financial stake in Tierra Biosciences, a private company that makes use of cell-free technologies such as those described in this article for protein expression and screening.
The other authors have nothing to disclose.

The protocols described here provide a cost-effective and rapid means to screen genetic parts via the expression of a reporter protein by CFPS. Well-characterized genetic parts are crucial to the design of predictable genetic circuits with useful function. This methodology increases throughput and decreases the time needed to screen new genetic parts by removing the requirement to work in living cells, while retaining functionality that mirrors the cellular environment by retaining the metabolic process of protein expression in the cell lysate. Our protocol can be performed in 1 day after receipt of primers (~2.5-6 h for reaction preparation, 2-16 h for CFPS reaction; Figure 1), compared to at least 3 days for traditional cloning (1 day each for construct assembly and transformation, sequence verification of clones, and culturing of cells for assessment). We further estimate that the cost per construct using linear DNA is roughly one-third of the traditional cloning ($78 vs. $237; Supplementary Table 1) methods. Commercial synthesis services currently quote a minimum of 4 business days depending on size, though they would have similar costs to our method if linear fragments are screened directly in CFPS ($78 vs. $91); we have not verified this approach. The cost to evaluate a part with CFPS is small compared to the generation of the template DNA ($0.05/reaction22 vs. $78 per template), though it should be noted that the startup costs for bulk reagents and lysis equipment is at least several thousand dollars. The use of an acoustic liquid handler only marginally improves costs by enabling smaller volumes down to 0.5 &#181;L40; the more significant advantage is the reduction of time to prepare reactions (~10 min vs. up to 1 h, depending on the number of reactions), especially when preparing a large number of reactions raises concerns of prepared reaction sitting for extended times before incubation.
While rapid and cost-effective, the limitations on when CFPS prototyping adequately predicts in vivo function remain to be seen. For example, any cross-reactivity with genomic DNA will not be detected due to removal of the host genome during the production of the CFPS system. Also, component concentrations can be 1-2 orders of magnitude lower in CFPS than in cells51, which is likely to affect the behavior of some parts as a result of different macromolecular crowding conditions. Further, the ability of linear DNA to predict in vivo function may be limited, for example, when DNA secondary structure plays an important role. A final limitation is that constructs are not sequence-verified before testing for functions. There may be cases where the part characterized is not actually aligned with the intended theoretical sequence. All of these limitations can be mitigated by validating a subset of the parts screened by this method in the intended in vivo application.
We originally developed this methodology to investigate the effects of changing the operator position on hybrid T7-tetO promoters32. We have presented the protocols here in a more generic format, such that they can be applied to promoters, operators, ribosome binding sequences, insulators, and terminators. These genetic parts can be added to the 5&#697; or 3&#697; end of the reporter gene by PCR using primers for each design, obviating the need for synthesis or cloning of each variant to test. The resulting PCR products serve as template DNA for evaluation via the expression of a reporter protein. In our work, the affinity purification protocol provided here was used for TetR and GamS. The same procedure can be used for the expression and purification of other repressors, activators, polymerases, sigma factors, and other proteins cognate to a genetic part of interest, although modifications may be needed for the desired protein being expressed. Purification and titration of these proteins into CFPS reactions enables a more detailed characterization of a particular genetic part. Finally, numerous alternative CFPS protocols exist and each should be amenable to the parts screening portion of methodology. As an example, we do not include a dialysis step in this protocol, which others have found to be important for expression from native bacterial promoters22. Varying the concentrations of underlying constituent components of the CFPS is also possible. The use of liquid handling enhances the ability to test the myriad conditions by increasing throughput and decreasing the materials required34,35.
One area that can require significant troubleshooting is optimization of the acoustic liquid handler. Acoustic liquid handler dispensing should be optimized for each component being transferred and it is strongly recommended to run controls to verify proper distribution and reproducibility before collecting data. The ideal source plate type and liquid class setting will depend on the specific liquid to be dispensed and its components. It is not recommended to use amine-coated plates to dispense DNA, as the amine coating may interact with the DNA. It should also be noted that the ability to dispense higher concentrations of certain components may depend on the acoustic liquid handler model. A test liquid transfer may be conducted by dispensing onto a foil plate seal to visualize successful droplet formation; however, this test provides limited information and droplets from different settings may appear identical. The use of a water-soluble dye, such as tartrazine, may be used to more accurately verify the correct volume is dispensed with a given setting or workflow (see Representative Results). Optimal programming of liquid transfers can also influence the accuracy and consistency of data generated; for transfers &#62;1 &#181;L from one source well to one destination well, we have found that sequential transfers of &#8804;1 &#181;L should be programmed to reduce systematic well-to-well variability (Figure 4). Lastly, theoretical and actual source well dead volumes can vary dramatically depending on the source plate type, liquid class setting, and components of the specific liquid; using the acoustic liquid handler survey function to assess the well volumes prior to running a program may help gauge how accurately the instrument is able to measure a particular liquid.
CFPS reaction performance can vary when comparing results between different users, batches of materials, plate readers, and laboratories41. For instances where such comparisons are required while prototyping genetic circuits, we recommend including internal control reactions with standard constitutive promoters in each reaction plate to help normalize results across experimental setups. The method of DNA preparation can also contribute majorly to CFPS activity; the inclusion of an ethanol precipitation step is recommended. In addition, the optimal reaction composition can vary by the batch of extract34. Optimal magnesium glutamate and potassium glutamate concentrations, in particular, have been shown to vary by batch42 or with the promoter or reporter protein used24. Concentrations of these components should be optimized by screening across several concentrations of each component per genetic construct and per cell extract preparation to determine the optimal conditions for protein expression. Finally, best practices for consistent CFPS reaction performance include thorough mixing, careful pipetting, and consistency in the preparation of each reagent component.
Beyond characterization of individual parts, the same method can be used to screen combinations of parts that form complex circuits, such as logic circuits16 or oscillators52,53. This method can also be applied to screening and optimizing biosensors for applications in epidemiological diagnostics54,55,56,57 or hazard detection and quantification3,58,59. The application of AI-driven techniques such as active learning34 can also be paired with the high-throughput nature of this method to drive rapid exploration of complex biological design spaces. Ultimately, we envision this approach supporting accelerated development times for new genetic designs in synthetic biology.

A core effort of synthetic biology is to develop genetic tool kits containing well-characterized parts, which can be used to construct genetic circuits1 that carry out useful functions when deployed in microbes or cell-free lysates. Areas in which such genetic circuits have gained traction are sensing2,3,4, human performance5,6, biofuels7,8, materials production9,10, and cellular computing11. Registries of standardized genetic parts have been established12 to catalog new and existing parts into categories such as promoters, operators, coding sequences, and terminators, to name just a few. Efforts such as the iGEM (international Genetically Engineered Machines) competition13 have been instrumental in characterizing and cataloging these genetic parts. Many methods have been developed to facilitate the rapid assembly of these parts into useful genetic circuits14,15. Software has even been developed to automate the composition of well-characterized parts into circuits that achieve a desired function16. However, the assembly of useful genetic circuits with predictable functions rests on the presumption that the genetic tool kits contain well-characterized genetic parts. Due to the necessity of these tool kits toward the advancement of synthetic biology, numerous efforts to better catalog circuits and parts with appropriate characterization data have been described17,18,19,20,21.
One approach to characterizing genetic components makes use of cell-free protein synthesis (CFPS) systems, which reconstitute cellular functions such as transcription and translation ex vivo22. Several studies have demonstrated the potential of CFPS for prototyping genetic components23,24,25,26,27,28,29,30,31,32 whether for direct applications in cell-free systems or to predict the function of genetic constructs in cells, such as the relative activity of parts within a library29, metabolic pathway optimization27, and cellular burden30. Advantages to prototyping in CFPS versus cells highlighted by these studies include avoiding time-consuming cloning, precise control over the concentration of DNA and other reaction components, and the ability to easily mix and match multiple DNA constructs. The advantage of avoiding cloning is especially apparent when using linear DNA templates, which enables new constructs to be assembled by in vitro methods that take hours instead of days33. The ability to manipulate the concentration of DNA constructs and other components simply by pipetting makes the approach even more attractive by enabling high-throughput experimentation powered by liquid handling robots34,35. While successes using CFPS for prototyping have been reported, it is important to note that it remains to be seen under what contexts CFPS results can reliably predict functionality in cells.
Here, we present a method for CFPS prototyping that emphasizes the advantages in speed, throughput, and cost compared to traditional cell-based approaches. The approach is derived from our previous work where we used CFPS to rapidly characterize a library of T7 promoter variants regulated by the transcription factor TetR32, significantly expanding on the small handful of regulated T7 promoter variants that were available in the literature at the time36,37. Others have, since then, further expanded the range of such promoters38. In our method, genetic construct assembly is accelerated by using PCR to amplify template DNA via primers that add variant genetic parts to a reporter gene. Acoustic liquid handling in 384-well plates is used to increase throughput and decrease the volume of materials required. Previous work has demonstrated successful use of acoustic liquid handling at significantly lower volumes39,40 with variability comparable to manual pipetting of larger volumes41. In addition to the method, we provide troubleshooting information and an assessment of potential cost and time savings. Note that while we include a protocol for producing cell-free lysates based on Sun et al.42 here, numerous other commercial kits and protocols43,44 should also work. Similarly, while we demonstrate the method for the characterization of promoter variants32, other parts can be interchanged by PCR amplification, such as riboregulators, Ribosome Binding Sites (RBSs), insulators, protein tags, and terminators. We hope that this methodology can help the synthetic biology community continue to grow the number of characterized parts for the assembly of predictable genetic circuits with useful function.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2x YT medium</td>
			<td>Sigma-Aldrich</td>
			<td>Y2377-250G</td>
			<td>Alternative to making 2xYT media</td>
		</tr>
		<tr>
			<td>Acetic Acid</td>
			<td>J.T. Baker</td>
			<td>9508-01</td>
			<td>S30 Buffer B</td>
		</tr>
		<tr>
			<td>Agar</td>
			<td>Bacto</td>
			<td>214010</td>
			<td>For plating cells</td>
		</tr>
		<tr>
			<td>Chromatography column (5 cm diameter)</td>
			<td>BIO-RAD</td>
			<td>731-1550</td>
			<td>Used for protein purification.</td>
		</tr>
		<tr>
			<td>Destination plate</td>
			<td>Thermo Scientific Nunc plate</td>
			<td>142761</td>
			<td>For CFPS reactions</td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma-Aldrich</td>
			<td>D2650</td>
			<td>For dialysis buffer</td>
		</tr>
		<tr>
			<td>DpnI</td>
			<td>NEB</td>
			<td>R0176L</td>
			<td>For digestion of plasmid templates</td>
		</tr>
		<tr>
			<td>DTT</td>
			<td>Roche</td>
			<td>20871723</td>
			<td>S30 Buffer B</td>
		</tr>
		<tr>
			<td>E. coli BL21(DE3) Rosetta2</td>
			<td>Novagen</td>
			<td>70954</td>
			<td>Cell line used for production of lysate and purified proteins</td>
		</tr>
		<tr>
			<td>Echo acoustic liquid handler</td>
			<td>Labcyte</td>
			<td>525</td>
			<td>Acoustic liquid handler</td>
		</tr>
		<tr>
			<td>French pressure cell</td>
			<td>Thermo Spectronic</td>
			<td>FA-078</td>
			<td>For lysing cells for CFPS</td>
		</tr>
		<tr>
			<td>Imidazole</td>
			<td>Sigma-Aldrich</td>
			<td>56750</td>
			<td>For buffers</td>
		</tr>
		<tr>
			<td>Impermeable plastic sealable lid</td>
			<td>Thermo</td>
			<td>232702</td>
			<td>Plate seal</td>
		</tr>
		<tr>
			<td>IPTG</td>
			<td>RPI</td>
			<td>I56000-25.0</td>
			<td>Used for protein induction.</td>
		</tr>
		<tr>
			<td>K-Glu</td>
			<td>Sigma-Aldrich</td>
			<td>g1501-500G</td>
			<td>S30 Buffer B</td>
		</tr>
		<tr>
			<td>Labcyte Echo source plate</td>
			<td>Labcyte</td>
			<td>PL-05525</td>
			<td>For use with Echo acoustic liquid handler</td>
		</tr>
		<tr>
			<td>Mg-Glu</td>
			<td>Sigma-Aldrich</td>
			<td>49605-250G</td>
			<td>S30 Buffer B</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S7653-250G</td>
			<td>For buffers</td>
		</tr>
		<tr>
			<td>NaHPO4</td>
			<td>Sigma-Aldrich</td>
			<td>71505</td>
			<td>For dialysis buffer</td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Mallinckrodt Chemicals</td>
			<td>7708-10</td>
			<td>For making 2xYT media. Currently not produced by Mallinckrodt. Alternate: Sigma-Aldrich S0899</td>
		</tr>
		<tr>
			<td>Ni-NTA resin</td>
			<td>Invitrogen</td>
			<td>R901-15</td>
			<td>For production of purified proteins</td>
		</tr>
		<tr>
			<td>PCR H2O</td>
			<td>Ambion</td>
			<td>AM9937</td>
			<td>PCR of linear templates</td>
		</tr>
		<tr>
			<td>Plate Reader</td>
			<td>BioTek</td>
			<td>H10</td>
			<td>Plate reader used</td>
		</tr>
		<tr>
			<td>Q5 PCR Master Mix</td>
			<td>NEB</td>
			<td>M0494S</td>
			<td>PCR of linear templates</td>
		</tr>
		<tr>
			<td>QIAquick Gel Extraction Kit</td>
			<td>Qiagen</td>
			<td>28606</td>
			<td>PCR of linear templates</td>
		</tr>
		<tr>
			<td>QIAquick PCR Purification Kit</td>
			<td>Qiagen</td>
			<td>28004</td>
			<td>PCR of linear templates</td>
		</tr>
		<tr>
			<td>QSonica Ultrasonic Processor</td>
			<td>Qsonica</td>
			<td>Q700</td>
			<td>Cell disruption during protein purification</td>
		</tr>
		<tr>
			<td>RTS Amino Acid Sampler</td>
			<td>biotechrabbit</td>
			<td>BR1401801</td>
			<td>Updated supplier from Sun et al.</td>
		</tr>
		<tr>
			<td>Tris</td>
			<td>MP</td>
			<td>819623</td>
			<td>S30 Buffer B</td>
		</tr>
		<tr>
			<td>Tris-Cl</td>
			<td>Sigma-Aldrich</td>
			<td>T5941</td>
			<td>For buffers</td>
		</tr>
		<tr>
			<td>Tryptone</td>
			<td>Fluka</td>
			<td>T7293</td>
			<td>For making 2xYT media</td>
		</tr>
		<tr>
			<td>Yeast Extract</td>
			<td>Bacto</td>
			<td>212750</td>
			<td>For making 2xYT media</td>
		</tr>
	</tbody>
</table>

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.

To demonstrate the utility of our methods, we present results that describe the effects of proximity of the tetO sequence to the T7 promoter on the regulation of T7 RNAP-driven expression. The full results and their implications can be found in the work of McManus et al.32. The workflow is described in Figure 1. Fifteen linear templates, varying only in the distance of the T7 promoter relative to the tetO sequence, were prepared by PCR-amplifying the sfGFP reporter with primers designed to add each promoter variant (Figure 2) as described in section 2 of the protocol. CFPS reaction components and reactions were prepared following the protocol. The expression of sfGFP was measured from each template with a titration of 12 different concentrations of the TetR protein, in triplicate, using an acoustic liquid handler. At 36 CFPS reactions per template and 15 templates, a total of 540 reactions for the entire set of T7-tetO combinations were performed. The entire evaluation was carried out on two plates in two plate readers. Analysis of this data showed that the T7 RNAP downregulates T7-driven expression equally up through 13 bp downstream from the start of the T7 transcript (Figure 3). This result has implications for the future design of regulatable T7-driven gene circuits by describing a putative window for an effective repression of T7 by other repressors. Comparison of results from the protocol described here with DNA prepared by traditional cloning revealed a small but statistically significant difference in the degree of TetR repression between formats. We hypothesized that non-specific binding of TetR to the vector DNA could explain the observed difference. Experimental results showed that addition of linear vector DNA to reactions with linear template DNA reduced the difference to non-statistical significance, though it did not rule out contributions from other factors, such as differences in periodicity of the DNA helix for linear vs. circular formats, which, in turn, could affect TetR binding. Depending upon the application, the use of linear template may require additional validation.
We further include representative data on potential issues with accurate dispensing using acoustic liquid handling (Figure 4). A solution of 1x phosphate buffered saline (PBS), pH 7.4 containing 0.25 mM tartrazine dye was used to evaluate two methods of programming an acoustic liquid handler to dispense volumes &#62;1 &#181;L. Following liquid dispensing, the destination plate was sealed and centrifuged at 1,500 x g for 1 min, and the absorbance at 425 nm measured with a plate reader. Representative results of nine experiments are shown and demonstrate more consistent dispensing across the series of eight destination wells when the 5 &#181;L transfer is divided into separate 1 &#181;L dispenses. Based on these observations, it is recommended that transfers &#62;1 &#181;L be broken down into multiple transfers of &#8804;1 &#181;L.&#160;See the Discussion section for more details on troubleshooting this important aspect of the protocol.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62816/62816fig01.jpg" /> 
Figure 1: Single-day workflow for the evaluation of promoter parts in cell-free extract. (A) A reporter is PCR-amplified using primers containing genetic parts to be evaluated (2-5 h). (B) The cell-free reaction mix is prepared as detailed in the protocol and distributed into a 384-well plate with the PCR-amplified templates (30 min). (C) Acoustic liquid handling is used to distribute additional components, which can include repressor proteins, effector molecules, and any other conditional effectors (10 min). (D) Reporter protein expression from each reaction is measured in a plate reader (2-16 h, depending on the CFPS recipe and construct). <a href="https://www.jove.com/files/ftp_upload/62816/62816fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62816/62816fig02.jpg" /> 
Figure 2: Primer design for adding genetic parts to a reporter gene by PCR amplification. (A) The sfGFP reporter gene (green) will be amplified to add an RBS (red) and a T7 promoter (blue) by PCR. (B) The sfGFP (green) and an RBS (red) will be amplified to add a tetO sequence (gold) and a T7 promoter (blue) by PCR. <a href="https://www.jove.com/files/ftp_upload/62816/62816fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62816/62816fig03.jpg" /> 
Figure 3: The effect of tetO position on the regulation of a T7-driven expression. Normalized maximum repression values for linear and circular template as a function of tetO position. Traces represent the mean and standard deviations for three replicates. This figure has been modified from McManus et al.32 under a Creative Commons CC-BY license. <a href="https://www.jove.com/files/ftp_upload/62816/62816fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62816/62816fig04.jpg" /> 
Figure 4: Using tartrazine dye to validate liquid dispensing with an acoustic liquid handler. Black bars indicate dispensing 5 &#181;L of tartrazine solution from a single source well into each of the eight consecutive destination wells of a 384-well plate using a single programming command. Gray bars indicate dispensing 1 &#181;L from a single source well into each of eight consecutive destination wells using a single programming command, and then repeating this step four times for a total of 5 &#181;L dispensed in each destination well. <a href="https://www.jove.com/files/ftp_upload/62816/62816fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component Name</td>
			<td>Volume for 1 reaction (&#181;L)</td>
			<td>Volume for 110% of X number of reactions (&#181;L)</td>
		</tr>
		<tr>
			<td>Q5 PCR Premix</td>
			<td>25</td>
			<td></td>
		</tr>
		<tr>
			<td>Water</td>
			<td>4</td>
			<td></td>
		</tr>
		<tr>
			<td>Template (1&#8211;3 ng/&#181;L)</td>
			<td>1</td>
			<td></td>
		</tr>
		<tr>
			<td>(if fixed1) Forward Primer (5 &#181;M)</td>
			<td>0 or 10</td>
			<td></td>
		</tr>
		<tr>
			<td>(if fixed1) Reverse Primer (5 &#181;M)</td>
			<td>0 or 10</td>
			<td></td>
		</tr>
		<tr>
			<td>Master Mix Total:</td>
			<td>30 or 40</td>
			<td></td>
		</tr>
		<tr>
			<td>(if variable1) Forward Primer (5 &#181;M)</td>
			<td>0 or 10</td>
			<td></td>
		</tr>
		<tr>
			<td>(if variable1) Reverse Primer (5 &#181;M)</td>
			<td>0 or 10</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1: Worksheet for the preparation of reagents for PCR reactions. Values in the rightmost column can be filled in by users depending on the intended number of reactions. 1Variable primers contain a specific part to be added in the PCR reaction and can be the forward primer, reverse primer, or both. Fixed primers do not add a part and can be the forward primer or reverse primer but not both.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component Name</td>
			<td>Volume for 1 reaction (&#181;L)</td>
			<td>Volume for 110% of X number of reactions (&#181;L)</td>
		</tr>
		<tr>
			<td>Cell Extract</td>
			<td>4.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Supplement Mix</td>
			<td>3.3</td>
			<td></td>
		</tr>
		<tr>
			<td>GamS Protein (207 &#181;M)</td>
			<td>0.15</td>
			<td></td>
		</tr>
		<tr>
			<td>Template DNA (20 nM)</td>
			<td>1</td>
			<td></td>
		</tr>
		<tr>
			<td>T7 Polymerase (13 mg/mL)</td>
			<td>0.12</td>
			<td></td>
		</tr>
		<tr>
			<td>Water</td>
			<td>0.73 (this number may vary)</td>
			<td></td>
		</tr>
		<tr>
			<td>Master Mix Total:</td>
			<td>9</td>
			<td></td>
		</tr>
		<tr>
			<td>Repressor Protein:</td>
			<td>1</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 2: Worksheet for the preparation of reagents for CFPS reactions. Values in the rightmost column can be filled in by users depending on the intended number of reactions.
Supplementary Table. <a href="https://www.jove.com/files/ftp_upload/62816/Table S1 - cost estimates_REV EDITS.xlsx" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Rapid Characterization of Genetic Parts with Cell-Free Systems at JoVE.com

Rapid Characterization of Genetic Parts with Cell-Free Systems

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 ...

rapid-characterization-of-genetic-parts-with-cell-free-systems

Research

JoVE Journal

Bioengineering

1.7K Views.  US Army Combat Capabilities Development Command Army Research Laboratory. 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.

Video: Rapid Characterization of Genetic Parts with Cell-Free Systems

High efficiency, Site-specific Transfection of Adherent Cells with siRNA Using Microelectrode Arrays (MEA)

The discovery of RNAi pathway in eukaryotes and the subsequent development of RNAi agents, such as siRNA and shRNA, have achieved a potent method for silencing specific genes1-8 for functional genomics and therapeutics. A major challenge involved in RNAi based studies is the delivery of RNAi agents to targeted cells. Traditional non-viral delivery techniques, such as bulk electroporation and chemical transfection methods often lack the necessary spatial control over delivery and afford poor transfection efficiencies9-12. Recent advances in chemical transfection methods such as cationic lipids, cationic polymers and nanoparticles have resulted in highly enhanced transfection efficiencies13. However, these techniques still fail to offer precise spatial control over delivery that can immensely benefit miniaturized high-throughput technologies, single cell studies and investigation of cell-cell interactions. 
Recent technological advances in gene delivery have enabled high-throughput transfection of adherent cells14-23, a majority of which use microscale electroporation. Microscale electroporation offers precise spatio-temporal control over delivery (up to single cells) and has been shown to achieve high efficiencies19, 24-26. Additionally, electroporation based approaches do not require a prolonged period of incubation (typically 4 hours) with siRNA and DNA complexes as necessary in chemical based transfection methods and lead to direct entry of naked siRNA and DNA molecules into the cell cytoplasm. As a consequence gene expression can be achieved as early as six hours after transfection27. Our lab has previously demonstrated the use of microelectrode arrays (MEA) for site-specific transfection in adherent mammalian cell cultures17-19. In the MEA based approach, delivery of genetic payload is achieved via localized micro-scale electroporation of cells. An application of electric pulse to selected electrodes generates local electric field that leads to electroporation of cells present in the region of the stimulated electrodes. The independent control of the micro-electrodes provides spatial and temporal control over transfection and also enables multiple transfection based experiments to be performed on the same culture increasing the experimental throughput and reducing culture-to-culture variability. 
Here we describe the experimental setup and the protocol for targeted transfection of adherent HeLa cells with a fluorescently tagged scrambled sequence siRNA using electroporation. The same protocol can also be used for transfection of plasmid vectors. Additionally, the protocol described here can be easily extended to a variety of mammalian cell lines with minor modifications. Commercial availability of MEAs with both pre-defined and custom electrode patterns make this technique accessible to most research labs with basic cell culture equipment.

High efficiency, Site-specific Transfection of Adherent Cells with siRNA Using Microelectrode Arrays MEA

The article details the protocol for site-specific transfection of scrambled sequence of siRNA in an adherent mammalian cell culture using a microelectrode array (MEA).

The discovery of RNAi pathway in eukaryotes and the subsequent development of RNAi agents, such as siRNA and shRNA, have achieved a potent method for ...

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

Characterization of Complex Systems Using the Design of Experiments Approach: Transient Protein Expression in Tobacco as a Case Study

Plants provide multiple benefits for the production of biopharmaceuticals including low costs, scalability, and safety. Transient expression offers the additional advantage of short development and production times, but expression levels can vary significantly between batches thus giving rise to regulatory concerns in the context of good manufacturing practice. We used a design of experiments (DoE) approach to determine the impact of major factors such as regulatory elements in the expression construct, plant growth and development parameters, and the incubation conditions during expression, on the variability of expression between batches. We tested plants expressing a model anti-HIV monoclonal antibody (2G12) and a fluorescent marker protein (DsRed). We discuss the rationale for selecting certain properties of the model and identify its potential limitations. The general approach can easily be transferred to other problems because the principles of the model are broadly applicable: knowledge-based parameter selection, complexity reduction by splitting the initial problem into smaller modules, software-guided setup of optimal experiment combinations and step-wise design augmentation. Therefore, the methodology is not only useful for characterizing protein expression in plants but also for the investigation of other complex systems lacking a mechanistic description. The predictive equations describing the interconnectivity between parameters can be used to establish mechanistic models for other complex systems.

We describe a design of experiments approach that can be used to determine and model the influence of transgene regulatory elements, plant growth and development parameters, and incubation conditions on the transient expression of monoclonal antibodies and reporter proteins in plants.

Plants provide multiple benefits for the production of biopharmaceuticals including low costs, scalability, and safety. Transient expression offers the ...

Sealable Femtoliter Chamber Arrays for Cell-free Biology

Cell-free systems provide a flexible platform for probing specific networks of biological reactions isolated from the complex resource sharing (e.g., global gene expression, cell division) encountered within living cells. However, such systems, used in conventional macro-scale bulk reactors, often fail to exhibit the dynamic behaviors and efficiencies characteristic of their living micro-scale counterparts. Understanding the impact of internal cell structure and scale on reaction dynamics is crucial to understanding complex gene networks. Here we report a microfabricated device that confines cell-free reactions in cellular scale volumes while allowing flexible characterization of the enclosed molecular system. This multilayered poly(dimethylsiloxane) (PDMS) device contains femtoliter-scale reaction chambers on an elastomeric membrane which can be actuated (open and closed). When actuated, the chambers confine Cell-Free Protein Synthesis (CFPS) reactions expressing a fluorescent protein, allowing for the visualization of the reaction kinetics over time using time-lapse fluorescent microscopy. Here we demonstrate how this device may be used to measure the noise structure of CFPS reactions in a manner that is directly analogous to those used to characterize cellular systems, thereby enabling the use of noise biology techniques used in cellular systems to characterize CFPS gene circuits and their interactions with the cell-free environment.

A microfabricated device with sealable femtoliter-volume reaction chambers is described. This report includes a protocol for sealing cell-free protein synthesis reactants inside these chambers for the purpose of understanding the role of crowding and confinement in gene expression.

Cell-free systems provide a flexible platform for probing specific networks of biological reactions isolated from the complex resource sharing (e.g., ...

Rapid Subtractive Patterning of Live Cell Layers with a Microfluidic Probe

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one particular implementation of the MFP, the hierarchical hydrodynamic flow confinement (hHFC), multiple liquids are simultaneously brought in contact with a substrate. Local chemical action and liquid shaping using the hHFC, is exploited to create cell patterns by locally lysing and removing cells. By utilizing the scanning ability of the MFP, user-defined patterns of cell monolayers are created. This protocol enables rapid, real-time and spatially controlled cell patterning, which can allow selective cell-cell and cell-matrix interaction studies.

We present a protocol to perform subtractive patterning of live cell monolayers on a surface. This is achieved by local and selective lysis of adherent cells using a microfluidic probe (MFP). The cell lysate retrieved from local regions can be used for downstream analysis, enabling molecular profiling studies.

The microfluidic probe (MFP) facilitates performing local chemistry on biological substrates by confining nanoliter volumes of liquids. Using one ...

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate (LAL) Assays

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause inflammation, fever, hypo- or hypertension, and, in extreme cases, can lead to tissue and organ damage that may become fatal. The amounts of endotoxin in pharmaceutical products, therefore, are strictly regulated. Among the methods available for endotoxin detection and quantification, the Limulus Amoebocyte Lysate (LAL) assay is commonly used worldwide. While any pharmaceutical product can interfere with the LAL assay, nano-formulations represent a particular challenge due to their complexity. The purpose of this paper is to provide a practical guide to researchers inexperienced in estimating endotoxins in engineered nanomaterials and nanoparticle-formulated drugs. Herein, practical recommendations for performing three LAL formats including turbidity, chromogenic and gel-clot assays are discussed. These assays can be used to determine endotoxin contamination in nanotechnology-based drug products, vaccines, and adjuvants.

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate LAL Assays

Detection of endotoxins in engineered nanomaterials represents one of the grand challenges in the field of nanomedicine. Here, we present a case study that describes the framework composed of three different LAL formats to estimate potential endotoxin contamination in nanoparticles.

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause ...

A Multilayer Microfluidic Platform for the Conduction of Prolonged Cell-Free Gene Expression

The limitations of cell-based synthetic biology are becoming increasingly apparent as researchers aim to develop larger and more complex synthetic genetic regulatory circuits. The analysis of synthetic genetic regulatory networks in vivo is time consuming and suffers from a lack of environmental control, with exogenous synthetic components interacting with host processes resulting in undesired behavior. To overcome these issues, cell-free characterization of novel circuitry is becoming more prevalent. In vitro transcription and translation (IVTT) mixtures allow the regulation of the experimental environment and can be optimized for each unique system. The protocols presented here detail the fabrication of a multilayer microfluidic device that can be utilized to sustain IVTT reactions for prolonged durations. In contrast to batch reactions, where resources are depleted over time and (by-) products accumulate, the use of microfluidic devices allows the replenishment of resources as well as the removal of reaction products. In this manner, the cellular environment is emulated by maintaining an out-of-equilibrium environment in which the dynamic behavior of gene circuits can be investigated over extended periods of time. To fully exploit the multilayer microfluidic device, hardware and software have been integrated to automate the IVTT reactions. By combining IVTT reactions with the microfluidic platform presented here, it becomes possible to comprehensively analyze complex network behaviors, furthering our understanding of the mechanisms that regulate cellular processes.

The fabrication process of a PDMS-based, multilayer, microfluidic device that allows in vitro transcription and translation (IVTT) reactions to be performed over prolonged periods is described. Furthermore, a comprehensive overview of the hardware and software required to automate and maintain these reactions for prolonged durations is provided.

The limitations of cell-based synthetic biology are becoming increasingly apparent as researchers aim to develop larger and more complex synthetic genetic ...

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 ...

Liquid Chromatography Coupled to Refractive Index or Mass Spectrometric Detection for Metabolite Profiling in Lysate-based Cell-free Systems

Engineering cellular metabolism for targeted biosynthesis can require extensive design-build-test-learn (DBTL) cycles as the engineer works around the cell&#39;s survival requirements. Alternatively, carrying out DBTL cycles in cell-free environments can accelerate this process and alleviate concerns with host compatibility. A promising approach to cell-free metabolic engineering (CFME) leverages metabolically active crude cell extracts as platforms for biomanufacturing and for rapidly discovering and prototyping modified proteins and pathways. Realizing these capabilities and optimizing CFME performance requires methods to characterize the metabolome of lysate-based cell-free platforms. That is, analytical tools are necessary for monitoring improvements in targeted metabolite conversions and in elucidating alterations to metabolite flux when manipulating lysate metabolism. Here, metabolite analyses using high-performance liquid chromatography (HPLC) coupled with either optical or mass spectrometric detection were applied to characterize metabolite production and flux in E. coli S30 lysates. Specifically, this report describes the preparation of samples from CFME lysates for HPLC analyses using refractive index detection (RID) to quantify the generation of central metabolic intermediates and by-products in the conversion of low-cost substrates (i.e., glucose) to various high-value products. The analysis of metabolite conversion in CFME reactions fed with 13C-labeled glucose through reversed-phase liquid chromatography coupled to tandem mass spectrometry (MS/MS), a powerful tool for characterizing specific metabolite yields and lysate metabolic flux from starting materials, is also presented. Altogether, applying these analytical methods to CFME lysate metabolism enables the advancement of these systems as alternative platforms for executing faster or novel metabolic engineering tasks.

The protocols describe high-performance liquid chromatography methods coupled to refractive index or mass spectrometric detection for studying metabolic reactions in complex lysate-based cell-free systems.

Engineering cellular metabolism for targeted biosynthesis can require extensive design-build-test-learn (DBTL) cycles as the engineer works around the ...