Targeting Cysteine Thiols for in Vitro Site-specific Glycosylation of Recombinant Proteins

The overall goal of this procedure is to incorporate cysteine residues in proteins by site directed mutagenesis at natively glycosylated sites and employ in-vitro glucose attachment and assessment approaches to evaluate the efficiency and structural effects of glycosylation. This method can help answer key questions in the Structural Biology field such as, what are the structural consequences of protein glycosylation and how do these modifications affect the underlying mechanisms of function? The main advantages of this technique are that it is easily adaptable to study or handling glycosylation.

It is highly efficient and both structure perturbations and efficiency can be rapidly assessed. The implications of this technique extend towards the therapy of diseases because altered glycosylation patterns have been associated with several cancers and neurodegenerative disorders. Though this method can provide insight into the effects of natively glycosylated proteins, it could also provide much needed information on how to alter glycosylation patterns may contribute to this range of diseases.

To introduce a single cysteine mutation in the expression vector, prepare two individual PCR reaction tubes. In the first tube, add the forward mutagenic primer and in the second tube, add the reverse mutagenic primer. Add a final concentration of 1X PCR buffer to both the tubes.

Then, set up a five cycle program for the two separate PCR reaction tubes. Add an extension time of seven and a half minutes at 72 degrees Celsius at the end of the fifth cycle. After the five cycles are completed, mix the two separate PCR reactions in a single tube, centrifuge the tube for about three seconds, and repeat an additional 20 cycles following the same PCR conditions.

Next, run 15 microliters of the sample on a 1%DNA agarose gel at 120 Volts for 40 minutes. Then, immerse the gel in water containing 0.5 milligrams per milliliter ethidium bromide and shake it at room temperature for 30 minutes. Then, confirm the amplification of the template under UV light at 302 nanometers wavelength.

After that, add 0.5 microliters of DPN1 restriction enzyme to the remaining 25 microliters of PCR reaction mix. Incubate the mix for two and a half hours at 37 degrees Celsius to digest the methylated template DNA. Then, add approximately five to 10 microliters of the digested mixture to 100 microliters of heat shock competent E.coli cells and incubate on ice for 60 minutes.

Then, heat shock the cell-DNA mixture at 42 degrees Celsius for 45 seconds on a dry heat block. Then, immediately incubate the mixture on ice for three minutes. Next, add 900 microliters of LB broth to the cells and transfer the suspension into a sterile 14 milliliter round bottomed tube.

Incubate at 37 degrees Celsius with constant shaking at 190 rpm for 90 minutes. Then, spread the centrifuged cell pellet on antibiotic supplemented agar plates. Incubate the plates at 37 degrees Celsius for 16 hours.

The following day, aseptically inoculate a single colony from the plate into five milliliters of antibiotic supplemented LB medium. Then, grow the culture overnight at 37 degrees Celsius with constant shaking. Transform the cysteine residue mutated plasmid into E.coli cells via heat shock treatment, followed by growing the culture overnight in antibiotic supplemented LB medium.

The next day, directly spread 150 microliters of cell suspension on kanamycin supplemented LB agar plates, and keep the plates for overnight incubation at 37 degrees Celsius. The following day, aseptically transfer a single colony in 20 milliliters of LB medium in an Erlenmeyer flask, and leave for overnight incubation at 37 degrees Celsius. The next day, transfer the overnight culture to a 50 milliliter conical tube.

Then, pellet the cells by centrifuging the overnight liquid culture at 2, 400 times G for 15 minutes. Next, add 10 milliliters of M9 minimal medium to the cell pellet and then suspend the cellular mixture in one liter of M9 medium containing kanamycin. Keep growing the M9 liquid culture at 37 degrees Celsius with constant shaking at approximately 190 rpm until the optical density at 600 nanometers reaches approximately 0.6 to 0.8.

Then, add 200 micromolar of IPTG to the M9 culture for inducing protein expression and leave the culture to grow overnight. The following day, pellet the bacterial cells from the overnight culture by centrifuging at approximately 10, 000 times G at four degrees Celsius for 30 minutes. Using a 10 milliliter motorized transfer pipette, homogenize the frozen bacterial cell pellet in approximately 40 milliliters of homogenization buffer.

Then, incubate the homogenate in a hybridization oven for 90 minutes with constant rotation. Centrifuge the mixture at approximately 15, 000 times G at eight degrees Celsius for 40 minutes to separate the soluble protein fraction from the insoluble pellet. Then, add 50%volume by volume nickel nitrilotriacetic acid agarose bead slurry to the supernatant and incubate by inversion in the hybridization oven for 90 minutes at room temperature.

Next, use a gravity flow protein purification column to collect the agarose beads with hexahistidine tagged protein attached to the nickel. Finally, wash the collected agarose beads three times. Elute the proteins in a series of two milliliter fractions.

Next, confirm purified hexahistidine tagged protein in the eluted fractions by running a Coomassie blue stained SDS-PAGE gel. Pool the eluted protein fractions into a dialysis membrane with a 3, 500 daltons molecular weight cutoff and incubate in refolding buffer at four degrees Celsius overnight with magnetic stirring. Then, add approximately one unit of thrombin per milligram of protein to the pooled protein in the dialysis membrane and again incubate for one day at four degrees Celsius.

The next day, remove the protein from the dialysis bag and concentrate it approximately tenfold using an ultrafiltration centrifugal concentrator with a 10, 000 daltons molecular weight cutoff. Subsequently, re-dilute the protein concentrate approximately 20-fold in sodium chloride-free refolding buffer. To purify the protein, load it on a prepacked anion-exchange column equilibrated with the same sodium chloride-free buffer.

Begin eluting the proteins in a zero to 60%gradient of one molar sodium chloride containing refolding buffer. Transfer the pooled protein fractions into a dialysis bag. Conduct a dialysis step at four degrees Celsius overnight to exchange the protein into the experimental refolding buffer with the optimal molar concentration of sodium chloride.

First, dialyze 1.5 milliliters of approximately 60 micromolar protein into one liter of modification buffer for glycosylation of protein. After 24 hours, at four degrees Celsius, transfer the modified protein into a microcentrifuge tube and add DMSO solubilized glucose-5-MTS to a final concentration of two millimolar. Gently tap the microcentrifuge tube every 10 minutes while incubating the sample in the dark for an hour in ambient temperature.

After incubation, perform dialysis at four degrees Celsius to re-exchange the protein into the final experimental buffer. Measure the accurate mass of the protein using electrospray ionization mass spectrometry. Finally, use solution NMR to calculate the modification efficiency and identify the structural changes of glycosylated cys residues in the protein.

A successful mutational PCR amplified template DNA is shown in this representative 1%DNA agarose gel. DNA ladder and an equivalent amount of template DNA that has not been amplified by PCR are used as controls in the gel. The topmost band with high intensity ethidium bromide staining shows the amplified template of the desired size.

Next, the protein was purified using an anion-exchange column. The chromatogram shows an elution profile observed at 280 nanometers in blue exhibiting three peaks eluting against a sodium chloride gradient indicated in red with increasing concentration until 0.6 molar. The bottom panel shows a Coomassie blue stained SDS-PAGE gel of the protein fractions.

Glycosylation of the cysteine residues was determined by electrospray ionization mass spectrometry. The chromatogram shows the masses of the unmodified and modified protein before and after glycosylation of the cysteine residue. For confirmation, solution NMR was performed showing the HSQC spectral overlay of the residue specific amide chemical shifts, indicating both modifications and structural perturbations of glycosylated protein.

When it’s mastered, all stages from mutagenesis to NMR assesment can be completed in approximately 12 days, excluding the wait time for primer synthesis, DNA sequencing, and mass spectroscopy services. While attempting this procedure, it’s important to remember to assess the structural and biophysical effects of the cysteine mutations in the absence of the thiol modification as base line control measurements. Following this procedure, other methods such as stability, secondary structure, ion binding, protein-protein interaction assessments, can be performed to reveal how the biophysical properties of underlying protein functions are affected by glycosylation.

After its development, this technique paved the way for researchers in the field of calcium signaling to individually and cumulatively assess the roles of asparagine 131 and asparagine 171 and glycosylation in stromal interaction molecule one calcium signaling associated with store operated calcium entry.