In most cases, conventional site-directed mutagenesis is not appropriate to study the functional role of prolines in proteins. Here, we present a method to incorporate proline analogues as non-canonical amino acids into ribosomally-synthesized proteins to investigate the role of proline residues on protein folding and function. The substitution of prolines by standard amino acids is a risky approach.
Incorporation of proline analogs permits a kind of molecular surgery. In other words, it introduces subtle molecular changes to rationally influence and manipulate protein properties. Since proline residues play an essential role in the stability of protein scaffolds, our technology could be helpful to understand the reasons behind protein folding deficits, which underlie certain pathological states in humans.
The protein alterations induced by proline replacements have implications for the biotechnological engineering of enzymes and provide opportunities for enhancement of ribosomal protein translation or folding. This method does not require special equipment and expensive instrumentation. Special care should be taken with protocol steps, in which native proline is depleted from E.coli cultures prior to the addition of proline analogues.
To begin the production of the fluorescent proteins with native proline, inoculate 200 milliliters of fresh NMM medium in a two-liter Erlenmeyer flask with two milliliters of the overnight culture. Incubate the cells at 37 degrees Celsius in an orbital shaker at 220 RPM for approximately 3.5 hours. During the incubation, measure the optical density at 600 nanometers in a spectrophotometer every 30 minutes.
When the optical density reaches 0.7, spin down the cell suspension and gently decant the supernatant into waste. Then, wash the cells by resuspending in 50 milliliters of ice-cold NMM without any amino acids or NMM without proline. Spin the cells again and decant the supernatant into waste.
Next, resuspend the cell pellet by gentle pipetting in 200 milliliters of NMM without proline and supplemented with ampicillin in a two-liter Erlenmeyer flask, and incubate the cells in an orbital shaker for 30 minutes to allow for the complete depletion of proline. After incubation, add an appropriate volume of either L-proline or the proline analogues from the 50-millimolar stock solutions to obtain a one-millimolar final concentration in the cell suspension. Next, induce target protein expression by adding 500-micromolar IPTG from a one-molar stock solution.
Express the target protein overnight at 37 degrees Celsius in an orbital shaker and measure the optical density the next day. Collect the bacterial cells by centrifugation and decant the supernatant into waste. Then, wash the cells by careful pipetting in 50 milliliters of binding buffer containing 10%glycerol.
Spin the cells again, decant the supernatant, and store the cell pellet in a 50-milliliter tube at minus 20 or 80 degrees Celsius until further use. Before evaluating fluorescence emission, perform an SDS-PAGE to ensure that the sample purity is above 95%Then, adjust the samples of each purified protein variant to a concentration of 0.3 micromoles, taking the calculated absorbance value at the appropriate wavelength as a reference. Ensure that the approximate final sample volume is 200 microliters.
Let the diluted samples equilibrate at room temperature. After one hour, transfer the samples into a one-centimeter quartz cuvette and measure the fluorescence emission spectrum of the samples using a fluorescence spectrometer. To induce denaturation, dilute the samples of each protein variant twenty-fold by adding the two microliter of 300-micromolar samples to 18 microliters of 1.11X PBS buffer containing 8.89-molar urea and 5.56-millimolar DTT.
Incubate the samples at 95 degrees Celsius for five minutes. Then, dilute the samples 100-fold by adding 1, 980 microliters of PBS containing five-millimolar DTT to induce renaturation and immediately transfer 200 microliters of the samples into a one-centimeter quartz cuvette. Insert the quartz cuvette into an appropriate fluorescence spectrometer and monitor protein refolding by acquiring a fluorescence spectrum every three seconds over 30 minutes.
Transfer the refolding samples into 1.5-milliliter microcentrifuge tubes, then close the lid and store the samples at room temperature in the dark to allow complete refolding of EGFP variants. After 24 hours, measure fluorescence emission of the refolded protein samples using the same excitation wavelength as before to capture the temporal endpoint of fluorescence recovery. Pellets from cells expressing native protein and variants bearing S-fluoroproline and dehydroproline had a typical bright color due to the intact chromophore.
In contrast, variants containing R-fluoroproline and difluoroproline remained colorless, indicating misfolding and deposition of unfolded protein in inclusion bodies. SDS-PAGE analysis verified the presence of insoluble R-fluoroproline-containing proteins. In contrast, native proteins and S-fluoroproline and dehydroproline-bearing variants were found in the soluble fractions.
In liquid chromatography/mass spectrometry analysis, each proline replacement with S-fluoroproline produced a positive 18-dalton shift per proline residue, while replacement with dehydroproline produced a negative shift of two daltons per residue. In light absorption and emission spectra, chromophore absorbance was found at 488 nanometers for EGFP and 493 nanometers for NowGFP. In KillerOrange, the chromophore absorbance region comprised two bands corresponding to two possible configurational and charge states of the complex chromophore.
Further, fluorescence spectra recorded upon excitation at the corresponding maximum absorbance wavelengths implied that the analogues did not alter the chemical environment of the chromophores. In refolding experiments, the native EGFP chromophore fluorescence recovered partially, although the tryptophan-specific fluorescence was larger after renaturation. Similar behavior was observed for the variant containing dehydroproline.
In EGFP-containing S-fluoroproline, a similar result was observed with a 295-nanometer excitation, while fluorescence recovered to a much higher extent at 488 nanometers. Only the EGFP variants showed fast refolding kinetics with tryptophan emission recovery being twice as fast compared to the chromophore emission. Fresh bacterial cultures with suitable growth media must be used.
Sterile techniques and regular monitoring of culture growth are also important prerequisites for the success of the experiment. This method is applicable to protein engineering, as it allows to study the role of other canonical amino acids. For this, suitable auxotrophic E.coli strains and amino acid analogues are needed.
Mass spectrometry of whole proteins is required as analytical evidence to confirm the incorporation of non-canonical amino acid analogues. The SPI method for incorporating proline analogues allows us to manipulate the protein backbone directly, and offers advantages for rational design and facilitated production of proteins through global stabilization and improvement of ribosomal translation or folding.
