This work was supported by the NSF&#8217;s Research Triangle MRSEC (DMR-1121107).

1. ELP Expression
<ol>
	<li>Inoculate 3 ml of sterile Terrific Broth media with a DMSO stock or agar plate colony of E. coli suitable for protein expression that contains the desired ELP-encoding plasmid under control of the T7-lac promoter. Add the appropriate antibiotic and incubate at 37 &#176;C overnight (O/N) with continuous shaking at 200 rpm.</li>
	<li>Add 1 ml of the O/N&#160;culture to 1 L of Terrific Broth sterile media in a 4-L flask. Add the appropriate antibiotic and incubate at 37 &#176;C for 24 hr with continuous shaking at 200 rpm. NOTE: The &#8220;leaky&#8221; baseline expression seen with the T7-lac promoter is sufficient for high level expression of many ELPs if the culture is incubated for 24 hr. However, a more conventional approach can be used, wherein protein expression is further induced by the addition of 0.2-1 mM isopropyl-beta-D-thiogalactoside (IPTG) when the optical density of the culture reaches 0.6.</li>
	<li>Transfer the culture from the 4-L flask to a 1-L centrifuge bottle and centrifuge at 4 &#176;C for 15 min at 2,000 x g.</li>
	<li>Discard the supernatant. NOTE: If more than 1 L of culture is grown they can be condensed by adding another liter of culture to the pellet and repeating step 1.3. Up to 2 L of culture can be condensed into a single cell pellet.</li>
	<li>Resuspend the pellet in phosphate buffered saline (PBS), water, or other desired buffer to reach 45 ml of cell suspension and transfer to a 50&#160;ml conical tube.</li>
</ol>
2. ELP Purification: Preliminary Steps and First Round of Inverse Transition Cycling
<ol>
	<li>Sonicate the resuspended cell pellet for a total of 9 min in cycles of 10 sec &#8216;on&#8217; and 20 sec &#8216;off&#8217; at an output power of 85 W, while keeping the sample on ice. Briefly allow the sample to cool on ice and then repeat the 9 min cycle once more. NOTE: If the ELP is anticipated to have a very low Tt the &#8216;off&#8217; interval can be increased up to 40 sec to avoid heating of the solution above the Tt.</li>
	<li>Transfer the lysate to a 50 ml round bottom centrifuge tube.</li>
	<li>Add 2 ml of 10% (w/v) polyethyleneimine (PEI) for each liter of culture and shake to mix. NOTE: PEI is a positively charged polymer that aids in the condensation of negatively charged genetic contaminants. Do not perform this step if the ELP is negatively charged, as the PEI may also condense and remove the ELP product.</li>
	<li>Centrifuge at 4 &#176;C for 10 min at 16,000 x g.</li>
	<li>Transfer the supernatant to a clean 50&#160;ml round bottom centrifuge tube and discard the pellet.</li>
	<li>Optional &#8220;bake out&#8221; step: Incubate the sample at 60 &#176;C for 10 min to denature protein contaminants and then transfer to 4 &#176;C or to ice until the ELP is completely resolubilized. Centrifuge the sample at 4 &#176;C for 10 min at 16,000 x g. Transfer the supernatant to a clean 50&#160;ml round bottom centrifuge tube and discard the pellet. NOTE: Do not perform this step if a peptide or protein is fused to the ELP and is expected to denature in the condition of elevated heat. This may cause the ELP transition to become heat-irreversible or may destroy the activity of the fused moiety.</li>
	<li>Induce the ELP transition with the addition of crystalline NaCl (not exceeding 3 M). Alternatively, sodium citrate (not exceeding 0.3 M) can be used for ELPs that have higher Tts or low yields. NOTE: The solution should turn cloudy once the salt has dissolved, indicating the phase separation of ELP from solution. Very hydrophilic ELPs with high Tts may require some degree of heating, in combination with the addition of salt, to induce the phase separation of the ELP in this preliminary induction of the ELP transition.</li>
	<li>Centrifuge at room temperatue (RT) for 10 min at 16,000 x g (hot spin). NOTE: An ELP pellet should be observed, whose size depends on the expression yield of the ELP. This ELP pellet may at first appear opaque, but after cooling it will appear&#160;translucent and may be brown in color. The ELP pellet will become more colorless as purification proceeds and contaminants are removed.</li>
	<li>Discard the supernatant and add 1-5 ml of desired buffer to the pellet. Resuspend the pellet by pipetting or setting the tube to rotate at 4 &#176;C. NOTE: The required volume of buffer will depend on the size of the ELP pellet, and thus the yield of ELP expression, where a sufficient amount of buffer should be added to allow for resolubilization of the ELP pellet. ELPs with high cysteine content benefit from purification in water with 10 mM Tris(2-carboxyethyl)phosphine hydrochloride (TCEP-HCl) at pH 7 to eliminate disulfide interactions with contaminating proteins. ELPs with high charge content benefit from purification at a pH adjusted to neutralize their charge and reduce electrostatic interactions with contaminating proteins.</li>
	<li>Transfer the resuspended ELP solution in 1 ml aliquots into clean 1.5-ml microcentrifuge tubes.</li>
	<li>Centrifuge at 4 &#176;C for 10 min at 16,000 x g (cold spin).&#160;A&#160;small pellet of insoluble contaminants should be observed.</li>
	<li>Transfer the supernatant to a clean tube and discard the pellet.</li>
</ol>
3. ELP Purification: Subsequent Rounds of Inverse Transition Cycling
<ol>
	<li>Induce the ELP transition with heat and/or the addition of NaCl. Samples can be incubated on a heat block or in a water bath for 15 min at a temperature above the Tt. If, however, the ELP is fused to a protein (precluding the use of heat) or if the ELP has a high Tt that cannot be reached with heat alone, a 5 M solution of NaCl can be added drop-wise to induce the ELP transition. NOTE: The solution should turn cloudy, indicating the phase separation of ELP from solution.</li>
	<li>Centrifuge for 10 min at 16,000 x g (hot spin). If heat was used in step 3.1, perform this centrifugation step at a temperature above that which was required to induce the ELP transition. If NaCl alone was used in step 3.1, perform this centrifugation step at RT. NOTE: An ELP pellet should be observed.</li>
	<li>Discard the supernatant, add 500-750 &#956;l of desired buffer, and resuspend the pellet by pipetting or setting the tube to rotate at 4 &#176;C.</li>
	<li>Centrifuge at 4 &#176;C for 10 min at 16,000 x g (cold spin). A small pellet of insoluble contaminants should be observed.</li>
	<li>Transfer the supernatant to a clean tube and discard the pellet.</li>
	<li>Repeat steps 3.1 to 3.5 until no contaminant pellet is observed during step 3.4.</li>
</ol>
4. Post-purification Processing (optional)
<ol>
	<li>If desired, dialyze the sample against a suitable buffer to remove excess salt from the purified ELP solution. NOTE: ELPs to be lyophilized should be dialyzed against ddH2O O/N at 4 &#176;C.</li>
	<li>If desired, lyophilize the ELP for long-term storage by freezing the solution at -80 &#176;C for 30 min prior to lyophilizing O/N. NOTE: Lyophilization should not be used for ELPs with appended proteins.</li>
	<li>If desired, recover free peptide or protein from a peptide or protein ELP fusion by removing the ELP tag:
	<ol>
		<li>Digest the peptide or protein ELP fusion with biotinylated protease (as recommended by the protease supplier) or with a protease ELP fusion corresponding to the protease site genetically designed between the peptide or protein and ELP.</li>
		<li>If applicable, remove the biotinylated protease using streptavidin-modified beads (as recommended by the protease supplier).</li>
		<li>Induce the transition of the cleaved ELP and/or protease ELP fusion by the drop-wise addition of a 5 M NaCl solution.</li>
		<li>Centrifuge at RT for 10 min at 16,000 x g. NOTE: An ELP pellet should be observed.</li>
		<li>Collect the supernatant and discard the ELP pellet. Dialyze the supernatant against a desired buffer or use a centrifugal filter to perform a buffer exchange in order to remove the high salt concentration from the pure peptide or protein solution.</li>
		<li>Confirm the complete cleavage of free peptide or protein from the ELP tag by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE).</li>
	</ol>
	</li>
</ol>
5. Characterization: SDS-PAGE
<ol>
	<li>Prepare ELP samples for SDS-PAGE by mixing 10 &#956;l of ELP diluted in water with 10 &#956;l of sample buffer containing 2-mercaptoethanol. NOTE: 40 &#956;g of ELP is often sufficient to evaluate the size and purity of the ELP product.</li>
	<li>Incubate the sample at 100 &#176;C for 2 min and then load onto a polyacrylamide gel along with an appropriately sized protein ladder.</li>
	<li>Run the gel at 180 V until the dye approaches the bottom of the gel (approximately 45 min).</li>
	<li>Stain the gel with a 0.5 M CuCl2 solution for 5 min, while rocking. NOTE: ELPs are typically not visualized by Coomassie Brilliant Blue stain, as some ELP sequences do not interact with this dye. Coomassie Brilliant Blue interacts primarily with arginine residues and interacts weakly with other basic residues&#8211;histidine and lysine&#8211;and aromatic residues&#8211;tryptophan, tyrosine, and phenylalanine26. Thus ELPs and peptide or protein ELP fusions can be stained with Coomassie Brilliant Blue only if their primary sequence includes a sufficient number of these specific residues.</li>
	<li>Verify that the MW of the ELP band corresponds to that of the theoretical MW expected from the ELP gene and that no extraneous contaminant bands are present. NOTE: Some ELPs migrate with an apparent MW up to 20% higher than their expected MW9,27.</li>
</ol>
6. Characterization: Matrix-assisted Laser Desorption/ionization Time-of-Flight Mass Spectrometry (MALDI-TOF-MS)
<ol>
	<li>Prepare an ELP solution in 50% aqueous acetonitrile containing 0.1% trifluoroacetic acid to achieve an ELP concentration of approximately 25 &#956;M. NOTE: This solution should be free of salt, so ELP must be dialyzed against H2O following purification.</li>
	<li>Prepare a matrix solution of saturated sinapinic acid in 50% aqueous acetonitrile containing 0.1% trifluoroacetic acid.</li>
	<li>Add 1 &#956;l of the ELP solution to 9 &#956;l of the matrix solution to achieve an ELP concentration of approximately 2.5 pmol/&#956;l. NOTE: This mixture can be further diluted with matrix solution to optimize the MALDI-TOF signal.</li>
	<li>Deposit 1-2 &#956;l of ELP-matrix mixture onto a metal MALDI plate and let the spot dry completely.</li>
	<li>Obtain 5-10 spectra with MALDI-TOF to determine the mass to charge ratio (m/z) of the ELP and infer its measured MW.</li>
</ol>
7. Characterization: Temperature-programmed Turbidimetry and Dynamic Light Scattering
<ol>
	<li>Determine the Tt of the ELP by temperature-programmed turbidimetry:
	<ol>
		<li>Prepare a dilution series (e.g., from 5-100 &#956;M) of ELP solutions in the desired buffer.</li>
		<li>Using a temperature-controlled UV-Vis spectrophotometer, measure the optical density (O.D.) at 350 nm over a range of temperatures (e.g., 20-80 &#176;C) at a heating rate of 1 &#176;C/min. NOTE: If no increase in O.D. is seen the Tt may exceed the upper temperature limit of the instrument. The apparent Tt can be decreased by the addition of NaCl. Start at 1 M NaCl and increase in steps of 0.5 M until the ELP transition is detected within a measurable temperature range.</li>
		<li>Verify reversibility of the ELP transition by measuring the decrease in O.D. over the same range of temperature at a cooling rate of 1 &#176;C/min.</li>
		<li>Determine the Tt of the homopolymer ELP by identifying the inflection point of the turbidity profile (O.D. plotted versus temperature). NOTE: This temperature can be easily defined from the derivative of the O.D. curve, where the temperature corresponding to the maximum of the derivative is defined as the Tt. More complex ELP architectures will display more complicated turbidity profiles and should be further analyzed as described in section 7.2.</li>
	</ol>
	</li>
	<li>Additional characterization of ELPs that display more complex thermal properties (e.g., ELP diblock copolymers that self-assemble into spherical micelles) is achieved by dynamic light scattering (DLS):
	<ol>
		<li>Prepare an ELP solution in the desired buffer and pass the sample through a filter with a 20-450 nm pore size. NOTE: The choice of filter pore size will depend on the presence, size, and stability of any ELP assemblies in solution. The smallest feasible filter pore size is preferred to eliminate contaminants, such as dust, that diminish the quality of DLS measurements. A larger filter pore size should be used when significant resistance is experienced when passing an ELP solution through smaller filter pore sizes.</li>
		<li>Measure the hydrodynamic radius (RH) over a range of temperatures that corresponds to a region of interest identified in the turbidity profile. NOTE: ELP unimers typically have a RH&#60;10 nm, nanoparticle assemblies have a RH~20-100 nm, and aggregates have a RH&#62;500 nm. Each increase in RH should correspond to an increase in O.D. at 350 nm measured by UV-Vis spectrophotometry as a function of temperature, which is proportional to the size of the particle.</li>
	</ol>
	</li>
</ol>

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The authors have no competing financial interests to disclose.

ELPs offer an inexpensive and chromatography-free means of purification by exploitation of their stimulus-responsive phase behavior. This approach takes advantage of the LCST behavior of ELPs and peptide or protein ELP fusions to eliminate both soluble and insoluble contaminants after expression of genetically encoded ELPs in E. coli. This ease of purification can be used to produce ELPs for a variety of applications, or can be exploited for purification of recombinant peptides or proteins in which the ELP can act as a purification tag that may be removed with post-purification processing.
ELP purification involves preliminary steps to lyse the E. coli and remove genomic and insoluble cell debris from the crude culture lysate (Figure 1), followed by removal of residual soluble and insoluble contaminants by ITC (Figure 2). Centrifugation after triggering the ELP transition by the addition of heat or salt separates the ELP from soluble contaminants in the supernatant, in a step termed a hot spin. After resolubilizing the ELP, the solution is centrifuged again at a temperature below the Tt to remove the insoluble contaminant pellet, in a step termed a cold spin. Alternating hot and cold spins improves the purity of the ELP solution with each cycle, at a small cost to yield. Purification yields vary depending on the ELP Tt, length, and fused peptides or proteins. Typically, this protocol yields 100 mg of purified ELP per liter of E. coli culture, however yields can reach up to 500 mg/L. The final purity of the ELP product is confirmed by SDS-PAGE (Figure 3). The MW of the purified ELP should match closely with the theoretical MW encoded by the ELP gene. However, some ELPs migrate in SDS-PAGE with an apparent MW up to 20% higher than their expected MW9,27. More precise analysis of the ELP MW can be achieved by MALDI-TOF-MS, which can also provide additional information on the purity of the ELP product along with orthogonal analytic techniques such as high-performance liquid chromatography (HPLC).
Following purification, the ELP Tt is measured by temperature-programmed turbidimetry. This technique monitors the O.D. of an ELP solution while the temperature is increased. The Tt is concentration dependent so it is advisable to characterize a concentration series relevant to the intended application of the ELP. For ELP homopolymers the turbidity profile exhibits a single sharp increase that corresponds to the ELP transition from unimer to micron-scale aggregates (Figure 4A). The Tt is defined as the temperature corresponding to the inflection point in the turbidity profile, precisely determined as the maximum of the first derivative of the O.D. with respect to temperature. The reversibility of the ELP phase transition is confirmed by a decrease in O.D. to baseline as the temperature is lowered below the Tt (Figure 4B). The turbidity profile with increasing and decreasing temperature ramps will differ in magnitude and kinetics due to settling of ELP coacervates and variable hysteresis of ELP resolubilization. Peptide or protein ELP fusions similarly exhibit LCST behavior in this way, where the peptide or protein fused to the ELP affects the Tt. For protein ELP fusions the transition is reversible below the melting temperature of the protein. While temperature-programmed turbidimetry is an excellent method for the initial thermal characterization of ELP products, alternative techniques, such as differential scanning calorimetry (DSC), can also be used to measure the ELP Tt.
ELPs with more complex architectures exhibit more complicated thermal behaviors that can also be characterized by temperature-programmed turbidimetry. ELP diblock copolymers, for example, exhibit a characteristic turbidity profile corresponding to their temperature-triggered self-assembly into spherical micelles at their critical micellization temperature. For such ELP diblock copolymers the O.D. typically first increases 0.1-0.5 units above baseline indicating the transition from unimers to micelles, after which a sharp increase in O.D (up to 2.0 units above baseline) at a higher temperature indicates the formation of micron-scale aggregates (Figure 5A). Additional information about temperature-triggered self-assembled ELP structures is obtained with DLS, a technique that measures the RH of ELP assemblies in solution. Changes in RH agree closely with changes in O.D. measured with turbidimetry (Figure 5B). ELP unimers typically exhibit a RH&#60;10 nm while nanoparticle assemblies exhibit a RH~20-100 nm and aggregates exhibit a RH&#62;500 nm. Additional information about self-assembled ELP nanoparticles, such as aggregation number and morphology, can be obtained by static light scattering or cryogenic transmission electron microscopy17,23,29.
Due to the tunability of ELP thermal properties, a range of Tts is obtained by various ELP designs. It is important to keep in mind that the inherent Tt will influence the optimization of the purification protocol for each ELP, where extremely low or high Tts will require the most modification to this standard protocol. ELPs with extremely high transition temperatures may be unsuitable for purification with this approach. If the design of novel ELPs and peptide or protein ELP fusions may compromise the thermal response of the ELP, a simple histidine tag can be included for alternative purification by immobilized metal affinity chromatography. Additionally, characteristics of the ELP sequence may require modification of this protocol if the guest residue is charged. Manipulation of the buffer pH can be used as a method to change the overall charge of the ELP in an effort to eliminate&#160;electrostatic interactions with contaminants17. Furthermore, this protocol is appropriate for the special circumstance of ELP fusions with peptides and proteins when appropriate measures are taken to ensure the purification process does not perturb the activity of the fused moiety. Notes throughout the protocol on such modifications serve to direct the purification of ELPs that may present these challenges with respect to Tt, charge, or fusion concerns.
The purification of ELPs by means of their LCST behavior presents a simple and chromatography-free approach to purify the majority of ELPs and peptide or protein ELP fusions expressed in E. coli. The protocol summarized here permits purification of ELPs in a single day using equipment that is common to most biology laboratories. The ease of purification of ELPs and their fusions will, we hope, encourage an ever-growing diversity of ELP designs for new applications in materials science, biotechnology, and medicine.

Elastin-like polypeptides (ELPs) are biopolymers composed of the repeating pentapeptide VPGXG where X, the guest residue, is any amino acid except proline. ELPs exhibit lower critical solution temperature (LCST) phase transition behavior, such that a homogeneous ELP solution will separate into two phases upon heating to its LCST, which is commonly called the inverse transition temperature (Tt) in the ELP literature1. The two phases are composed of a very dilute ELP globule phase and an ELP rich sediment phase. The ELP rich sediment is formed on a short timescale upon aggregation of the ELP chains into micron-sized particles that subsequently coalesce. This behavior occurs over a range of a few degrees Celsius and is typically reversible, as a homogeneous solution is recovered upon returning to a temperature below the Tt.
ELPs are typically synthesized in Escherichia coli (E. coli) from an artificial gene that is ligated into an expression plasmid. This plasmid is then transformed into an E. coli cell line that is optimal for protein expression. We have exclusively used the T7-lac pET vector system for expression of a large variety of ELPs in E. coli, although other expression systems in yeast2-4, fungi5, and plants6-8 have also been utilized by other investigators. A number of approaches exist to genetically construct repetitive ELP genes, including recursive directional ligation (RDL)9, recursive directional ligation by plasmid reconstruction (PRe-RDL)10, and overlap extension rolling circle amplification (OERCA)11. The ability to engineer ELPs at the genetic level affords the opportunity to use recombinant DNA techniques to create ELPs with a variety of architectures (e.g.,&#160;monoblocks, diblocks, triblocks, etc.), which can be further appended with functional peptides and proteins. Control at the genetic level also ensures that each ELP is expressed with the exact length and composition dictated by its genetic plasmid template, providing perfectly monodisperse biopolymer products.
The thermal properties of each ELP depend on parameters intrinsic to the biopolymer such as its molecular weight (MW) and sequence, as well as extrinsic factors including its concentration in solution and the presence of other cosolutes, such as salts. The length of the ELP12 and its guest residue composition1,13,14 are two orthogonal parameters in ELP design that can be used to control the Tt, where hydrophobic guest residues and longer chain lengths result in lower Tts, while hydrophilic guest residues and shorter chain lengths result in higher Tts. &#160;ELP concentration is inversely related to the Tt, where solutions of greater ELP concentration have lower Tts12. The type and concentration of salts also influence the ELP Tt, where the effect of salts follows the Hofmeister series15. Kosmotropic anions (Cl- and higher on the Hofmeister series) lower the ELP Tt and increasing salt concentration enhances this effect. These intrinsic and extrinsic parameters can be tuned to obtain thermal behavior within a target temperature range that is required for a specific application of an ELP.
The stimulus-responsive behavior of ELPs is useful for a diverse range of applications including biosensing16,17, tissue engineering18, and drug delivery19,20. Additionally, when ELPs are fused to peptides or proteins at the genetic level, the ELP can serve as a simple purification tag to provide an inexpensive batch method for purification of recombinant peptides or proteins that requires no chromatography21. Modification of the purification process ensures that the activity of the peptide or protein fused to the ELP is maintained. Peptide or protein ELP fusions can be purified for applications in which the ELP tag is useful22,23 or alternatively, when the free peptide or protein is required, a protease recognition site can be inserted between the peptide or protein and ELP. Removal of the ELP tag can then be achieved by digestion with free protease or a protease ELP fusion, where the latter provides the additional ease of processing as a final round of ELP purification can separate the ELP tag and protease ELP fusion from the target peptide or protein in a single step24,25. The following protocol describes the purification procedure for ELPs and peptide or protein ELP fusions by means of their thermal properties and discusses basic techniques for characterizing the thermal response of ELP products.

<table border="0" cellpadding="0" cellspacing="0" width="1503">
	<tbody>
		<tr>
			<td>pET-24a(+) 
			pET-25b(+)</td>
			<td>Novagen</td>
			<td>69749-3 
			69753-3</td>
			<td>T7-lac expression vectors with resistance to kanamycin (pET-24) or ampicillin (pET-25).</td>
		</tr>
		<tr>
			<td>Ultra BL21 (DE3) competent cells</td>
			<td>Edge BioSystems</td>
			<td>45363</td>
			<td>Competent E. coli for recombinant protein expression.</td>
		</tr>
		<tr>
			<td>Terrific Broth (TB) Dry Powder Growth Media</td>
			<td>MO BIO Laboratories</td>
			<td>12105</td>
			<td>Media is reconstituted in DI H2O and autoclaved before use.</td>
		</tr>
		<tr>
			<td>Isopropyl-beta-D-thiogalactoside (IPTG)</td>
			<td>Gold Biotechnology</td>
			<td>I2481C</td>
			<td>IPTG is reconstituted in DI H2O, sterile filtered, and added to cultures to induce enhanced expression.</td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS) tablet</td>
			<td>Calbiochem</td>
			<td>524650</td>
			<td>These PBS tablets, when dissolved in 1 L of DI H2O, yield a 10 mM phosphate buffer with 140 mM NaCl, and 3 mM KCl with a pH of 7.4 at 25 &#176;C.</td>
		</tr>
		<tr>
			<td>Polyethyleneimine (PEI) Solution (~50% w/v)</td>
			<td>MP Biomedicals</td>
			<td>195444</td>
			<td>PEI is prepared as a 10% (w/v) solution in deionized H2O.</td>
		</tr>
		<tr>
			<td>Nalgene Oak Ridge high-speed centrifuge tubes, 50 mL</td>
			<td>Thermo Scientific</td>
			<td>3138-0050</td>
			<td>These round-bottom tubes withstand high-speed centrifugation of 30-50 ml.</td>
		</tr>
		<tr>
			<td>Tris(2-carboxyethyl)phosphine hydrochloride (TCEP-HCl)</td>
			<td>Thermo Scientific</td>
			<td>20491</td>
			<td>A stock solution of TCEP-HCl is prepared at 100 mM in DI H2O and adjusted to pH 7.0. For ELPs with a high cysteine content the stock solution of this reducing agent is added to the ELP pellet to reach 10 mM in H2O.</td>
		</tr>
		<tr>
			<td>Ready Gel Tris-HCL gel, 4-20% linear gradient polyacrylamide gel, 10 well, 30 &#956;l</td>
			<td>Bio-Rad Laboratories</td>
			<td>161-1105</td>
			<td>These linear gradient gels offer good visualization of ELPs with a range of MWs.</td>
		</tr>
		<tr>
			<td>Cupric chloride dihydrate (CuCl2-2H2O)</td>
			<td>Fisher Scientific</td>
			<td>C454-500</td>
			<td>A filtered 0.5 M solution is prepared for negative staining of Tris-HCL polyacrylamide gels.</td>
		</tr>
		<tr>
			<td>Anotop 10 syringe filter: 
			0.02 &#956;m 
			0.1 &#956;m 
			0.2 &#956;m</td>
			<td>Whatman</td>
			<td>6809-1002 
			6809-1012 
			6809-1022</td>
			<td>These 10 mm diameter syringe filters allow preparation of small volumes for DLS measurements.</td>
		</tr>
		<tr>
			<td>Millex-HV filter: 
			0.45 &#956;m</td>
			<td>EMD Millipore</td>
			<td>SLHVX13NK</td>
			<td>These 13 mm diameter syringe filters allow preparation of small volumes of solutions with large nanoparticle assemblies for DLS measurements.</td>
		</tr>
	</tbody>
</table>

non-chromatographic purification of recombinant elastin-like polypeptides and their fusions with peptides and proteins from escherichia coli

Elastin-like polypeptides are repetitive biopolymers that exhibit a lower critical solution temperature phase transition behavior, existing as soluble unimers below a characteristic transition temperature and aggregating into micron-scale coacervates above their transition temperature. The design of elastin-like polypeptides at the genetic level permits precise control of their sequence and length, which dictates their thermal properties. Elastin-like polypeptides are used in a variety of applications including biosensing, tissue engineering, and drug delivery, where the transition temperature and biopolymer architecture of the ELP can be tuned for the specific application of interest. Furthermore, the lower critical solution temperature phase transition behavior of elastin-like polypeptides allows their purification by their thermal response, such that their selective coacervation and resolubilization allows the removal of both soluble and insoluble contaminants following expression in Escherichia coli. This approach can be used for the purification of elastin-like polypeptides alone or as a purification tool for peptide or protein fusions where recombinant peptides or proteins genetically appended to elastin-like polypeptide tags can be purified without chromatography. This protocol describes the purification of elastin-like polypeptides and their peptide or protein fusions and discusses basic characterization techniques to assess the thermal behavior of pure elastin-like polypeptide products.

Here we describe representative results for the purification and characterization of an ELP homopolymer and an ELP diblock copolymer. Following 24 hr of expression in E. coli, cells are collected by centrifugation and lysed by sonication. Typically, a subtle change in color occurs after sonication, confirming sufficient cell lysis (Figure 1A). After the addition of PEI, genomic components and cellular debris are collected by centrifugation, separating a large pellet of insoluble contaminants from the soluble ELP in the supernatant (Figure 1B). ELP is further purified by inverse transition cycling (ITC), which exploits the LCST behavior to separate the ELP from both soluble and insoluble contaminants (Figure 2)28. The phase transition of the ELP is triggered by the addition of heat and/or salt. Centrifugation results in the formation of a pellet at the bottom of the tube that consists of ELP and some insoluble contaminants (Figure 1C). This step&#173;&#173; &#8211;termed a hot spin&#8211; discards soluble contaminants in the supernatant while the ELP pellet is retained and resuspended in cold buffer. The resolubilized ELP is centrifuged again at 4 &#176;C. This step &#8211;termed a cold spin&#8211; discards the pellet of insoluble contaminants while the supernatant containing soluble ELP is retained. Repeating the cycle of alternating hot and cold spins increases the purity of the ELP, but at the cost of slightly decreasing yield, as residual ELP is lost during each ITC round in centrifuge tubes and on pipette tips.
Successful purification of ELPs and peptide or protein ELP fusions is confirmed by SDS-PAGE. Small aliquots taken throughout the purification process demonstrate the progressive purification of ELP from the E. coli lysate. ELP is overexpressed in the crude E. coli lysate and contaminants decrease with successive rounds of ITC (Figure 3). Purified ELP appears as a single band near the predicted theoretical MW.
The ELP Tt is characterized by temperature-programmed turbidimetry. The turbidity profile of an ELP homopolymer exhibits a single sharp increase in O.D. at 350 nm (approximately 2.0 units above baseline for an ELP concentration of 25 &#956;M) (Figure 4A). Cooling turbidity scans confirm the reversible nature of the ELP transition as the O.D. returns to baseline when the temperature is lowered below the Tt (Figure 4B). ELP diblock copolymers exhibit a more complex turbidity profile, as compared to homopolymer ELPs. The O.D. typically first increases to 0.1-0.5 units above baseline, after which the O.D. increases sharply to 2.0 units above baseline (Figure 5A). DLS measurements provide information about the change in RH with respect to temperature, corroborating the information gained from the turbidity profile (Figure 5B).
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51583/51583fig1highres.jpg" width="600" /> 
Figure 1. Preliminary ELP purification. After 24 hr&#160;of expression, E. coli cells are collected by centrifugation and resuspended in PBS. A) Cells are lysed by sonication, which is accompanied by a subtle change in color such that the lysate darkens after sonication. B) After addition of PEI to condense DNA contaminants, the lysate is centrifuged and a large pellet of insoluble debris is separated from soluble ELP in the supernatant. C) The ELP transition is induced with the addition of heat and/or salt and centrifugation forms a translucent ELP pellet. <a href="https://www.jove.com/files/ftp_upload/51583/51583fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51583/51583fig2highres.jpg" width="600" /> 
Figure 2.&#160;Inverse transition cycling. ELP is purified by means of its LCST behavior from both soluble and insoluble contaminants. Starting with the ELP-rich lysate (1) the ELP transition is induced with the addition of heat and/or salt and the ELP is separated by centrifugation (hot spin) (2). The ELP pellet is retained (3a) while the supernatant containing soluble contaminants is discarded (3b). The ELP is resuspended in cold buffer (4) and centrifuged again at 4 &#176;C (cold spin) (5). The pellet containing insoluble contaminants is discarded (6a) while the supernatant containing soluble ELP is retained (6b). Cycles of alternating hot and cold spins are repeated until the desired purity is reached, as determined with SDS-PAGE. <a href="https://www.jove.com/files/ftp_upload/51583/51583fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51583/51583fig3highres.jpg" width="600" /> 
Figure 3.&#160;SDS-PAGE analysis of ELP purification products. Successful purification of an ELP is confirmed by SDS-PAGE. Overexpressed ELP is evident in the crude E. coli lysate after sonication (2). After the addition of PEI and centrifugation the soluble ELP is largely retained in the supernatant (3), although some ELP is lost in the discarded pellet of insoluble contaminants (4). Supernatant following the first hot spin contains significant levels of soluble contaminants (5). Supernatant following the first cold spin indicates good separation of ELP from insoluble contaminants (6). Additional cycles of hot (7) and cold spins (8) remove residual soluble and insoluble contaminants, respectively, until final hot (9) and cold spin (10) supernatants exhibit no extraneous contaminant bands, confirming satisfactory purity of the ELP product. This ELP homopolymer, with a sequence of SKGPG-(VGVPG)80-Y, has an expected molecular weight of 33.37 kDa. <a href="https://www.jove.com/files/ftp_upload/51583/51583fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51583/51583fig4highres.jpg" width="600" /> 
Figure 4.&#160;Characterization of ELP homopolymer by temperature-programmed turbidimetry. The ELP Tt is determined by monitoring the O.D. at 350 nm while increasing the temperature at a rate of 1 &#176;C/min. A) ELP homopolymers exhibit a single sharp increase in O.D. that defines their Tt at a given concentration. B) The reversibility of the ELP transition is verified by monitoring the O.D. at 350 nm of a 25 &#956; M solution while decreasing the temperature, where reversibility is confirmed by the return of the O.D. to baseline. This ELP homopolymer has the sequence SKGPG-(VGVPG)80-Y. <a href="https://www.jove.com/files/ftp_upload/51583/51583fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51583/51583fig5highres.jpg" width="600" /> 
Figure 5.&#160;Characterization of ELP diblock copolymer by temperature-programmed turbidimetry and dynamic light scattering. ELPs that undergo nano-scale self-assembly, such as ELP diblock copolymers that self-assemble into spherical micelles, exhibit more complex turbidity profiles. A) Moderate increase in O.D. indicates the transition from unimers to micelles, prior to a rapid increase in O.D. that indicates complete aggregation of the ELP into micron-scale aggregates. B) DLS measurements confirm the behavior inferred from the turbidity profile for a solution at 25 &#956;M by providing information about the change in RH with respect to temperature. Here the ELP unimer has a RH ~8 nm and the micelle has a RH ~25 nm. This ELP diblock copolymer has the sequence GCGWPG-(VGVPG)60-(AGVPGGGVPG)30-PGGS. <a href="https://www.jove.com/files/ftp_upload/51583/51583fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli at JoVE.com

Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli

Elastin-like polypeptides are stimulus-responsive biopolymers with applications ranging from recombinant protein purification to drug delivery. This protocol describes the purification and characterization of elastin-like polypeptides and their peptide or protein fusions from Escherichia coli using their lower critical solution temperature phase transition behavior as a simple alternative to chromatography.

Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli

Elastin-like polypeptides are repetitive biopolymers that exhibit a lower critical solution temperature phase transition behavior, existing as soluble ...

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21.5K Views.  Duke University. Elastin-like polypeptides are stimulus-responsive biopolymers with applications ranging from recombinant protein purification to drug delivery. This protocol describes the purification and characterization of elastin-like polypeptides and their peptide or protein fusions from Escherichia coli using their lower critical solution temperature phase transition behavior as a simple alternative to chromatography.

Video: Non-chromatographic Purification of Recombinant Elastin-like Polypeptides and their Fusions with Peptides and Proteins from Escherichia coli

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted into fuel alcohols1. The potential for enzymatic hydrolysis of cellulosic biomass to provide renewable energy has intensified efforts to engineer cellulases for economical fuel production2. Of particular interest are fungal cellulases3-8, which are already being used industrially for foods and textiles processing.
Identifying active variants among a library of mutant cellulases is critical to the engineering process; active mutants can be further tested for improved properties and/or subjected to additional mutagenesis. Efficient engineering of fungal cellulases has been hampered by a lack of genetic tools for native organisms and by difficulties in expressing the enzymes in heterologous hosts. Recently, Morikawa and coworkers developed a method for expressing in E. coli the catalytic domains of endoglucanases from H. jecorina3,9, an important industrial fungus with the capacity to secrete cellulases in large quantities. Functional E. coli expression has also been reported for cellulases from other fungi, including Macrophomina phaseolina10 and Phanerochaete chrysosporium11-12. 
We present a method for high throughput screening of fungal endoglucanase activity in E. coli. (Fig 1) This method uses the common microbial dye Congo Red (CR) to visualize enzymatic degradation of carboxymethyl cellulose (CMC) by cells growing on solid medium. The activity assay requires inexpensive reagents, minimal manipulation, and gives unambiguous results as zones of degradation (&#x201C;halos&#x201D;) at the colony site. Although a quantitative measure of enzymatic activity cannot be determined by this method, we have found that halo size correlates with total enzymatic activity in the cell. Further characterization of individual positive clones will determine , relative protein fitness. 
Traditional bacterial whole cell CMC/CR activity assays13 involve pouring agar containing CMC onto colonies, which is subject to cross-contamination, or incubating cultures in CMC agar wells, which is less amenable to large-scale experimentation. Here we report an improved protocol that modifies existing wash methods14 for cellulase activity: cells grown on CMC agar plates are removed prior to CR staining. Our protocol significantly reduces cross-contamination and is highly scalable, allowing the rapid screening of thousands of clones. In addition to H. jecorina enzymes, we have expressed and screened endoglucanase variants from the Thermoascus aurantiacus and Penicillium decumbens (shown in Figure 2), suggesting that this protocol is applicable to enzymes from a range of organisms.

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

We describe a low cost, high throughput method to screen for fungal endoglucanase activity in E. coli. The method relies on a simple visual readout of substrate degradation, does not require enzyme purification, and is highly scalable. This allows for the rapid screening of large libraries of enzyme variants.

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has become the method of choice for introducing DNA into prokaryotes that are not naturally competent. Electroporation is a rapid, efficient, and streamlined transformation method that, in addition to purified DNA and competent bacteria, requires commercially available gene pulse controller and cuvettes. In contrast to the pulsing step, preparation of electrocompetent cells is time consuming and labor intensive involving repeated rounds of centrifugation and washes in decreasing volumes of sterile, cold water, or non-ionic buffers of large volumes of cultures grown to mid-logarithmic phase of growth. Time and effort can be saved by purchasing electrocompetent cells from commercial sources, but the selection is limited to commonly employed E. coli laboratory strains. We are hereby disseminating a rapid and efficient method for preparing electrocompetent E. coli, which has been in use by bacteriology laboratories for some time, can be adapted to V. cholerae and other prokaryotes. While we cannot ascertain whom to credit for developing the original technique, we are hereby making it available to the scientific community.

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation is a commonly employed method for introducing DNA into bacteria in a process known as transformation. Traditional protocols for the preparation of electrocompetent cells are time consuming and labor intensive. This article describes an alternate, rapid, and efficient method for the preparation of electrocompetent cells presently employed by some laboratories.

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has ...

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. However, purifying proteins is sometimes challenging since many proteins are expressed in an insoluble form. When working with difficult or multiple targets it is therefore recommended to use high throughput (HTP) protein expression screening on a small scale (1-4&#160;ml cultures) to quickly identify conditions for soluble expression. To cope with the various structural genomics programs of the lab, a quantitative (within a range of 0.1-100 mg/L culture of recombinant protein) and HTP protein expression screening protocol was implemented and validated on thousands of proteins. The protocols were automated with the use of a liquid handling robot but can also be performed manually without specialized equipment.
Disulfide-rich venom proteins are gaining increasing recognition for their potential as therapeutic drug leads. They can be highly potent and selective, but their complex disulfide bond networks make them challenging to produce. As a member of the FP7 European Venomics project (<a href="http://www.venomics.eu">www.venomics.eu</a>), our challenge is to develop successful production strategies with the aim of producing thousands of novel venom proteins for functional characterization. Aided by the redox properties of disulfide bond isomerase DsbC, we adapted our HTP production pipeline for the expression of oxidized, functional venom peptides in the E. coli cytoplasm. The protocols are also applicable to the production of diverse disulfide-rich proteins. Here we demonstrate our pipeline applied to the production of animal venom proteins. With the protocols described herein it is likely that soluble disulfide-rich proteins will be obtained in as little as a week. Even from a small scale, there is the potential to use the purified proteins for validating the oxidation state by mass spectrometry, for characterization in pilot studies, or for sensitive micro-assays.

High Throughput Quantitative Expression Screening and Purification Applied to Recombinant Disulfide-rich Venom Proteins Produced in E. coli

A protocol for the quantitative, high throughput expression screening and analytical purification of fusion proteins from small-scale Escherichia coli cultures is described and applied to the expression of disulfide-rich animal venom protein targets.

Escherichia coli (E. coli) is the most widely used expression system for the production of recombinant proteins for structural and functional studies. ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

The Multifaceted Benefits of Protein Co-expression in Escherichia coli

We report here that the expression of protein complexes in vivo in Escherichia coli can be more convenient than traditional reconstitution experiments in vitro. In particular, we show that the poor solubility of Escherichia coli DNA polymerase III &#949; subunit (featuring 3&#8217;-5&#8217; exonuclease activity) is highly improved when the same protein is co-expressed with the &#945; and &#952; subunits (featuring DNA polymerase activity and stabilizing &#949;, respectively). We also show that protein co-expression in E. coli can be used to efficiently test the competence of subunits from different bacterial species to associate in a functional protein complex. We indeed show that the &#945; subunit of Deinococcus radiodurans DNA polymerase III can be co-expressed in vivo with the &#949; subunit of E. coli. In addition, we report on the use of protein co-expression to modulate mutation frequency in E. coli. By expressing the wild-type &#949; subunit under the control of the araBAD promoter (arabinose-inducible), and co-expressing the mutagenic D12A variant of the same protein, under the control of the lac promoter (inducible by isopropyl-thio-&#946;-D-galactopyranoside, IPTG), we were able to alter the E. coli mutation frequency using appropriate concentrations of the inducers arabinose and IPTG. Finally, we discuss recent advances and future challenges of protein co-expression in E. coli.

The Multifaceted Benefits of Protein Co-expression in Escherichia coli

Protein co-expression is a powerful alternative to the reconstitution in vitro of protein complexes, and is of help in performing biochemical and genetic tests in vivo. Here we report on the use of protein co-expression in Escherichia coli to obtain protein complexes, and to tune the mutation frequency of cells.

We report here that the expression of protein complexes in vivo in Escherichia coli can be more convenient than traditional reconstitution experiments in ...

Residue-specific Incorporation of Noncanonical Amino Acids into Model Proteins Using an Escherichia coli Cell-free Transcription-translation System

The canonical set of amino acids leads to an exceptionally wide range of protein functionality. Nevertheless, the set of residues still imposes limitations on potential protein applications. The incorporation of noncanonical amino acids can enlarge this scope. There are two complementary approaches for the incorporation of noncanonical amino acids. For site-specific incorporation, in addition to the endogenous canonical translational machineries, an orthogonal aminoacyl-tRNA-synthetase-tRNA pair must be provided that does not interact with the canonical ones. Consequently, a codon that is not assigned to a canonical amino acid, usually a stop codon, is also required. This genetic code expansion enables the incorporation of a noncanonical amino acid at a single, given site within the protein. The here presented work describes residue-specific incorporation where the genetic code is reassigned within the endogenous translational system. The translation machinery accepts the noncanonical amino acid as a surrogate to incorporate it at canonically prescribed locations, i.e., all occurrences of a canonical amino acid in the protein are replaced by the noncanonical one. The incorporation of noncanonical amino acids can change the protein structure, causing considerably modified physical and chemical properties. Noncanonical amino acid analogs often act as cell growth inhibitors for expression hosts since they modify endogenous proteins, limiting in vivo protein production. In vivo incorporation of toxic noncanonical amino acids into proteins remains particularly challenging. Here, a cell-free approach for a complete replacement of L-arginine by the noncanonical amino acid L-canavanine is presented. It circumvents the inherent difficulties of in vivo expression. Additionally, a protocol to prepare target proteins for mass spectral analysis is included. It is shown that L-lysine can be replaced by L-hydroxy-lysine, albeit with lower efficiency. In principle, any noncanonical amino acid analog can be incorporated using the presented method as long as the endogenous in vitro translation system recognizes it.

Residue-specific Incorporation of Noncanonical Amino Acids into Model Proteins Using an Escherichia coli Cell-free Transcription-translation System

An easy-to-use, cell-free expression protocol for the residue-specific incorporation of noncanonical amino acid analogs into proteins, including downstream analysis, is presented for medical, pharmaceutic, structural and functional studies.

The canonical set of amino acids leads to an exceptionally wide range of protein functionality. Nevertheless, the set of residues still imposes ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...