This research was supported by the Natural Sciences and Engineering Research Council of Canada (05239 to P.B.S.), Canadian Foundation for Innovation/Ontario Research Fund (to P.B.S.), Prostate Cancer Fight Foundation - Telus Ride for Dad (to P.B.S.) and Ontario Graduate Scholarship (to Y.J.C. and N.S.).

1. Polymerase chain reaction (PCR)-mediated site-directed mutagenesis for the incorporation of Cys into a bacterial pET-28a expression vector.
<ol>
	<li>Determine the concentration of the pET-28a vector (i.e. double stranded DNA) using a ultraviolet (UV) extinction coefficient of 0.020 (&#956;g/mL) cm-1 at 260 nm.</li>
	<li>Synthesize a pair of complementary mutagenic primers for each Cys mutation such that i) there are a minimum of 15 nucleotides complementary to the template prior to the first base mismatch and 15 nucleotides complementary to the template after the final base mismatch, ii) total primer length does not exceed 45 nucleotides, and iii) a guanine or cytosine is located at the first and last nucleotide position of each primer (Table 1). Ensure primer synthesis is performed using a 0.025 &#956;mol scale and cartridge purification.</li>
	<li>Using a high fidelity DNA polymerase, set up two 20 &#181;L PCR reaction mixtures: one containing the forward primer and the second containing the reverse primer. Prepare each mixture to contain final concentrations of 1x&#160;PCR buffer with 1.5 mM MgCl2, 0.2 mM dNTPs, 0.5 &#956;M primer, 0.4 &#956;L DMSO, 1.25 ng/&#956;L template DNA, 0.02 U/&#956;L high fidelity DNA polymerase.</li>
	<li>Thermally cycle the separate mixtures using a three-step protocol: 98 &#176;C for 30 s (denaturing), 53 - 56 &#176;C for 30 s (annealing), 72 &#176;C for 30 s kilobase(kb)-1 (extension) of the template DNA. Repeat the temperature program for 5 cycles and add a final 72 &#176;C extension step for 7.5 min.</li>
	<li>After the initial PCR with forward and reverse primers in separate tubes, combine the products into a single tube (i.e. 40 &#956;L total) and continue the PCR reaction for an additional 20 cycles using the same cycling parameters as described in step 1.4.</li>
	<li>Electrophorese 15 &#956;L of the PCR reaction mixture on a 1% (w/v) agarose gel using 0.5x Tris, acetic acid, ethylene diamine tetra acetic acid (EDTA) running buffer (TAE). As controls, electrophorese an equivalent amount of template DNA which has not been amplified by PCR and an aliquot of reference DNA ladder which contains marker bands both greater and less than the expected PCR product size.</li>
	<li>After electrophoresis at 120 V for 40 min, submerge the gel in water containing 0.5 &#956;g/mL ethidium bromide and shake for 30 min at room temperature. Confirm the full-length template has been amplified by the mutagenic primers as an increase in the relative ethidium bromide fluorescent intensity of the amplified band compared to the control template band under UV light (302 nm).
	<ol>
		<li>If no amplification is apparent, repeat the PCR after adjusting annealing temperature in 0.5 &#176;C increments between the 53 - 56 &#176;C temperature range.</li>
	</ol>
	</li>
	<li>Upon confirmation of amplification of the template by the mutagenic primers, treat the remaining ~25 &#956;L of the PCR reaction mixture with the DpnI restriction enzyme to digest the methylated template DNA. Use DpnI at 0.5 &#181;L (10 units) per 25 &#956;L PCR reaction mixture and a final concentration of 1&#215; DpnI reaction buffer. Incubate for 2.5 h at 37 &#176;C.</li>
	<li>Following template digestion, add ~5 - 10 &#956;L of the digested mixture to 100 &#181;L of heat shock competent DH5&#945; Esherichia coli cells in a 1.75 mL microcentrifuge tube. Incubate the cell-DNA mixture on ice for 60 min.</li>
	<li>Heat shock the cell-DNA mixture in the microcentrifuge tube at 42 &#176;C for 45 s on a dry heat block. After incubating the mixture on ice for 3 min, add 900 &#956;L of room temperature Luria-Bertani broth (LB) to the cells and transfer the total cell suspension into a sterile 14 mL round-bottom tube.</li>
	<li>Incubate the cell suspension at 37 &#176;C for 90 min with constant shaking at 190 rpm.</li>
	<li>Subsequently, transfer the cell suspension back into a 1.75 mL microcentrifuge tube and centrifuge at 10,000 x g for 5 min at room temperature.</li>
	<li>After the centrifugation, remove 900 &#956;L of the supernatant and resuspend the bacterial cells by gentle pipetting in the remaining 100 &#956;L of LB.</li>
	<li>Transfer the resultant concentrated cell suspension onto an LB-agar plate containing antibiotic which is selective for the expression vector (i.e. 60 &#956;g/mLKanamycin). Aseptically spread the suspension evenly on the agar plate and incubate for ~16 h at 37 &#176;C.</li>
	<li>The following day, inoculate a single colony from the plate into 5 mL of liquid LB containing the antiobiotic selection pressure (i.e. 60 &#956;g/mL Kanamycin). Grow the liquid culture overnight at 37 &#176;C with constant shaking at 37 &#176;C.</li>
	<li>Isolate and purify the propagated plasmid from the E. coli cells using a commercially available kit based on the alkaline lysis procedure 45.</li>
	<li>Confirm the mutation of interest is present and in the proper reading frame by Sanger DNA sequencing of the plasmid 46.</li>
</ol>
2. Uniform 15N-labeled protein expression in BL21 &#916;E3 Escherichia coli .
NOTE: Different recombinant proteins require different expression conditions. The following is the optimized procedure for expression of the human STIM1 EFSAM protein.
<ol>
	<li>Transform the expression vector harboring the Cys mutations (i.e. pET-28a-EFSAM) into BL21 &#916;E3 codon (+) heat shock competent cells and plate on LB-agar plates containing the antibiotic selection pressure as described in steps 1.9-1.14) with the following modifications: directly plate a 150 &#956;L aliquot out of the ~1,000 &#956;L total cell suspension on the LB-agar plate without the need for concentrating the cells in a microcentrifuge tube by centrifugation.</li>
	<li>The following day, aseptically transfer a single colony into a 200 mL Erlenmeyer flask containing 20 mL of LB supplemented with the appropriate antibiotic (i.e. 60 &#956;g/mL kanamycin for pET-28a-EFSAM). Grow this liquid starter culture overnight (i.e. ~16 h) at 37 &#176;C with constant shaking at ~ 190 rpm.</li>
	<li>On the same day as step 2.2, prepare M9 medium for 15N-labeled protein expression by autoclaving 1 L of M9 buffer salts (i.e. 42 mM Na2HPO4, 22 mM KH2PO4, 8.6 mM NaCl, pH 7.4) in a 4 L Erlenmeyer flask. Once cool, filter a mixture of 20% (w/v) D-glucose, 1 M CaCl2, 1 M thiamine, 1 M MgSO4, 1 mg/mL biotin and 0.2 g/mL15N-NH4Cl through a 0.2 &#956;m sterile syringe filter into the 1 L sterile M9 salt solution so that the final concentrations of these components are 0.2% (w/v) D-glucose, 100 &#956;M CaCl2, 50 &#956;M thiamine, 1 mM MgSO4, 1 &#956;g/mL biotin and 1 mg/mL15N-NH4Cl.</li>
	<li>The next day, aseptically transfer the 20 mL overnight liquid starter culture into a 50 mL sterile conical tube and centrifuge at 2,400 &#215;g for 15 min to pellet the cells.</li>
	<li>After decanting the LB medium, resuspend the resulting cell pellet in 10 mL of M9 minimal medium and transfer the resuspended pellet mixture into 1 L of M9 minimal medium with antibiotic (i.e. 60 &#956;g/mL kanamycin).</li>
	<li>Grow the 1 L of M9 minimal medium containing the bacterial starter culture at 37 &#176;C and ~190 rpm constant shaking until the optical density at 600 nm (OD600) reaches ~0.6-0.8.</li>
	<li>When the specfified OD600 range is reached, add 200 &#956;M of isopropyl &#946;-D-1-thiogalactopyranoside (IPTG) to induce protein expression.</li>
	<li>After IPTG addition, continue incubating the cells for protein expression at ambient temperature with constant shaking at ~190 rpm for ~16 h (i.e. overnight).</li>
	<li>The next day, harvest the bacteria by centrifugation at ~10,000 &#215;g, 4 &#176;C for 30 min.</li>
	<li>Decant the LB and transfer the cell pellet into a 50 mL conical tube. Store the pellet at -80 &#176;C until purification.</li>
</ol>
3. Purification of recombinant protein from E. coli.
NOTE: Different recombinant proteins require distinct purification procedures. The following is the protocol for 6&#215;His-tagged EFSAM purification from inclusion bodies expressed from the pET-28a construct.
<ol>
	<li>Manually homogenize the frozen bacterial cell pellet in 6 M guanidine-HCl, 20 mM Tris-HCl (pH 8) and 5 mM &#946;-mercaptoethanol using a motorized 10 mL transfer pipette. Add approximately 40 mL of guanidine-HCl per 5 mL of wet cell pellet for this step.</li>
	<li>Following a 90 min incubation at ambient temperature with constant rotation in a hybridization oven, centrifuge the mixture at ~15,000 &#215;g, 8 &#176;C for 40 min to separate the insoluble cell debris (i.e.&#160;pellet) from the soluble protein mixture (i.e. supernatant).</li>
	<li>Add 750 &#181;L of a 50% (v/v) Ni2+-nitrilotriacetic acid agarose bead slurry to the clarified lysate and incubate for another 90 min at room temperature with inversion in a hybridization oven.</li>
	<li>Subsequently, capture the 6&#215;His-tagged protein bound to the Ni2+ by collecting the agarose beads in a gravity flow protein purification column. Allow the lysate to flow through the column completely prior to moving to step 3.5.</li>
	<li>Wash the collected beads three times with 10 mL of 6 M urea, 20 mM Tris-HCl pH 8 and 5 mM &#946;-mercaptoethanol. Ensure that the entire 10 mL passes through the column prior to each subsequent 10 mL wash.</li>
	<li>Elute the proteins in a series of 2 mL fractions using 6 M urea, 20 mM Tris-HCl pH 8, 300 mM imidazole, and 5 mM &#946;-mercaptoethanol with a 90 s incubation time between fractions. Ensure that the entire 2 mL passes through the column prior to each subsequent elution step.</li>
	<li>At this stage, confirm the protein of interest is present in the eluted fractions by Coomassie blue-stained sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) using the method of Laemmli 47. Assess the protein size, quantity and purity by comparison against standard molecular weight marker bands which are both less than and greater than the expected molecular weight of the protein of interest.</li>
	<li>Pool the eluted protein fractions into a dialysis membrane with a 3,500 Da molecular weight cutoff and incubate in 1 L refolding buffer (20 mM Tris, 300 mM NaCl, 1 mM DTT, 5 mM CaCl2, pH 8) at 4 &#176;C overnight while the buffer is being stirred by a magnetic stirrer.</li>
	<li>After ~16 h of refolding time, add ~1 U of thrombin per mg of protein directly to the dialysis bag and incubate at 4 &#176;C for an additional ~24 h.</li>
	<li>Verify the extent of the 6&#215;His tag cleavage by Coomassie-blue staining of ~15 &#956;L protein aliquots taken from the dialysis bag before and after incubation with thrombin which are electrophoresed on denaturing polyacrylamide gels (SDS-PAGE) using the method of Laemmli 47. If a ~2 kDa shift in the migration is observed corresponding to the molecular weight of the cleaved 6&#215;His tag, continue to step 3.11; if a fraction of undigested protein remains that is detectable by Coomassie-blue staining, add ~0.2 U of thrombin per mg of protein directly to the dialysis bag and incubate at 4 &#176;C for an additional ~24 h.</li>
	<li>Use size exclusion or ion exchange chromatography to further purify the protein. For anion exchange chromatography of EFSAM, remove the protein solution from the dialysis bag and concentrate ~10-fold using an ultrafiltration centrifugal concentrator with a 10,000 Da molecular weight cutoff. Subsequently, re-dilute the solution ~20-fold in a NaCl-free buffer (20 mM Tris, 5 mM CaCl2, 1 mM DTT, pH 8).</li>
	<li>Equilibrate a prepacked anion exchange column with 10 column volumes of the NaCl-free buffer described in step 3.11. Equilibrate using buffer loaded into a luer-lock syringe containing no air bubbles by pushing the solution through the column in a dropwise manner and avoiding syringe pressures which cause steady streams of solution exiting the column. Use a strong anion exchanger (e.g. crosslinked agarose with quaternary ammonium functional groups).</li>
	<li>Load the protein solution diluted into NaCl-free buffer (step 3.11) onto the column as described in step 3.12.</li>
	<li>Elute the proteins&#160;in a gradient [i.e. 0 - 60% (v/v)] of increasing NaCl buffer (20 mM Tris, 1 M NaCl, 5 mM CaCl2, 1 mM DTT, pH 8) using a two-pump fast protein liquid chromatography (FPLC) system. Set the FPLC system to collect ~1-1.5 mL fractions and monitor the protein elution profile using UV 280 nm absorbance and a flow rate 0.5 mL/min.</li>
	<li>Identify the elution peaks and fractions containing the protein of interest as well as protein purity by Coomassie blue-stained SDS-PAGE gels using the method of Laemmli 47.</li>
	<li>Pool fractions showing &#62; 95% (i.e. taken as fractions which show only a single protein band on Coomassie-blue stained gels) into a dialysis bag and exchange into experimental buffer of interest by dialysis as described in step 3.8.</li>
</ol>
4. Chemical attachment of glucose-5-MTS to protein by dialysis.
<ol>
	<li>Prepare a 55 mM stock solution of N-(&#946;-D-glucopyranosyl)-N&#39;-[2-methanethiosulfonyl)ethyl]urea (glucose-5-MTS) by dissolving 10 mg of the compound in 500 &#956;l of 100% (v/v) DMSO. Store unused glucose-5-MTS solubilized in DMSO at -20 &#176;C.</li>
	<li>Prepare the protein sample for modification by dialyzing 1.5 mL of ~60 &#956;M protein into 1 L of modification buffer made up of 20 mM MOPS, 150 mM NaCl, 5 mM CaCl2 and 0.1 mM TCEP, pH 8.3. Use a dialysis membrane molecular weight cutoff which is smaller than the size of protein being modified (e.g. use a 3,500 Da cutoff for the ~17,500 Da EFSAM).</li>
	<li>After 24 h at 4 &#176;C, transfer the sample from the dialysis bag into a microcentrifuge tube. Add the DMSO-solubilized glucose-5-MTS to a final concentration of 2 mM.</li>
	<li>Incubate the sample in the dark for 1 h at ambient temperature. During the 1 h incubation period, mix the solution by gentle tapping of the tube every 10 min.</li>
	<li>Subsequently, re-exchange the protein into the final experimental buffer containing no reducing agent by dialysis at 4 &#176;C as described in step 4.2 or by centrifugal ultrafiltration. For the ultrafiltration procedure, concentrate the ~1.5 mL protein sample to &#60; 0.5 mL and subsequently dilute in the same concentrator with the experimental buffer. Repeat this concentration-dilution step two additional times so that the total exchange is a minimum of 30&#215;30&#215;30 = 27,000-fold. For EFSAM, use 20 mM Tris, 150 mM NaCl, 5 mM CaCl2, pH 7.5 as the experimental buffer.</li>
	<li>Prepare the sample for electrospray ionization mass spectrometry by dialysis or ultrafiltration exchange as described in steps 4.2 and 4.5, respectively, into 25 mM ammonium bicarbonate or 25 mM ammonium acetate. If using dialysis, ensure you exchange at least three times to remove any residual NaCl and CaCl2 salts.</li>
	<li>Determine the accurate mass (i.e. &#177; 1 Da) of the protein of interest using electrospray ionization mass spectrometry 48,49. Expect each covalent glucose addition to a Cys thiol via the methanethiosulfonate chemical to add 281.3 Da to the protein mass (i.e. add 360.4 Da for the glucose-5-MTS and subtract 79.1 Da for the CH3SO2 leaving group during covalent attachment).</li>
</ol>
5. Solution NMR assessment of modification efficiency and structural perturbations.
<ol>
	<li>Ensure the concentration of the modified protein is &#62; 100 &#956;M after the glucose attachment and final buffer exchange. For EFSAM, estimate protein concentration using a UV extinction coefficient at 280 nm of 1.54 (mg mL-1) cm-1.</li>
	<li>Supplement the protein solution with 60 &#956;M of 4,4-dimethyl-4-silapentane-1-sulfonic acid (DSS) for shimming and pulse calibration and 10% (v/v) D2O for the signal lock. For high signal-to-noise use 600 &#956;L samples in frequency-matched 5 mm NMR tubes, inserted into a minimum 600 MHz spectrometer equipped with a triple resonance HCN cryogenic probe.</li>
	<li>Collect standard 1H-15N HSQC spectra as previously detailed 50,51 at temperature, 1H and 15N sweep widths, transient and increment settings suitable for the particular sample. For EFSAM spectra, use 20 &#176;C, 256 1H transients, 64 15N dimension increments and 1H and 15N sweep widths set to 8,000 and 1,800 Hz, respectively.</li>
	<li>Following the acquisition of the glycosylated protein spectrum, add dithiothreitol (DTT) to the NMR sample from a 1 M stock to a final concentration of 15 mM. The DTT removes the glucose moiety from the protein by reduction of the disulfide-mediated attachment.</li>
	<li>Acquire a second 1H-15N HSQC under these reduced/unmodified conditions, providing a reference spectrum to assess modification efficiency and structural perturbations caused by the glucose attachment.</li>
	<li>Process the NMR data using NMRPipe as detailed previously 52. Ensure that processing minimally includes data conversion, phasing, solvent suppression, Fourier transform and initial visualization of the spectra.</li>
	<li>Assess modification efficiency by measuring the amide peak intensities and chemical shift values in the modified and reduced spectra using the NEASY plugin on CARA 53. Ensure to evaluate the peak intensity of the Cys amide both in the glucose attached and reduced spectra. If the Cys amide cannot be reliably located in both spectra, use the intensities of residues adjacent to the Cys as a readout.</li>
	<li>Calculate the efficiency as the intensity of the amide from the Cys-modified spectrum divided by the intensity of the amide from the Cys-reduced (i.e. DTT-treated) spectrum, multiplied by 100: 
	<img alt="Equation 1" src="/files/ftp_upload/56302/56302eq1.jpg" />, where IM is the intensity of the amide in the Cys-modified spectrum and IR is the intensity of the amide in the Cys-reduced spectrum. Alternatively, evaluate the mean efficiency over several amide peaks: 
	<img alt="Equation 2" src="/files/ftp_upload/56302/56302eq2.jpg" />, where Efficiencyi is the separately determined efficiency calculated for each residue, i, and n is the total number of residues used in the calculation.</li>
	<li>Calculate chemical shift perturbations (CSP) from the chemical shift differences between the two spectra observed in the 15N and 1H dimensions of each peak and normalizing for the larger 15N chemical shift range using the following equation: 
	<img alt="Equation 3" src="/files/ftp_upload/56302/56302eq3.jpg" />, where &#916;H is the ppm change in the proton dimension and &#916;N is the ppm change in the nitrogen dimension.</li>
</ol>

Protein glycosylation is a post-translational modification where sugars are covalently attached to polypeptides primarily through linkages to amino acid side chains. As many as 50% of mammalian proteins are glycosylated 54, where the glycosylated proteins can subsequently have a diverse range of effects from altering biomolecular binding affinity, influencing protein folding, altering channel activity, targeting molecules for degradation and cellular trafficking, to name a few (reviewed in<sup cla...

Store operated calcium (Ca2+) entry (SOCE) is the major pathway by which immune cells take up Ca2+ from the extracellular space into the cytosol. In T lymphocytes, T cell receptors located on the plasma membrane (PM) bind antigens which activate protein tyrosine kinases (reviewed in 1,2,3). A phosphorylation cascade leads to the activation of phospholipase-&#947; (PLC&#947;) which subsequently mediates the hydrolysis of membrane phosphatidylinositol 4,5-bisphosphate (PIP2) into diacylglycerol and inositol 1,4,5-trisphosphate (IP3). IP3 is a small diffusible messenger which binds to IP3 receptors (IP3R) on the endoplasmic reticulum (ER) thereby opening this receptor channel and permitting Ca2+ to flow down the concentration gradient from the ER lumen to the cytosol (reviewed in 4). Receptor signaling from G protein coupled and tyrosine kinase receptors in a variety of other excitable and non-excitable cell types lead to the same production of IP3 and activation of IP3Rs.
Due to the finite Ca2+ storage capacity of the ER, the IP3-mediated release and resultant increase in cytosolic Ca2+ is only transient; however, this depletion of the ER luminal Ca2+ profoundly effects stromal interaction molecule-1 (STIM1), a type-I transmembrane (TM) protein mostly found on the ER membrane 5,6,7. STIM1 contains a lumen-oriented Ca2+ sensing domain made up of an EF-hand pair and sterile &#945;-motif (EFSAM). Three cytosolic-oriented coiled-coil domains are separated from EFSAM by the single TM domain (reviewed in 8). Upon ER luminal Ca2+ depletion, EFSAM undergoes a destabilization-coupled oligomerization 7,9 which causes structural rearrangements of the TM and coiled-coil domains 10. These structural changes culminate in a trapping of STIM1 at ER-PM junctions 11,12,13,14 through interactions with PM phosphoinositides 15,16 and Orai1 subunits 17,18. Orai1 proteins are the PM subunits which assemble to form Ca2+ channels 19,20,21,22. The STIM1-Orai1 interactions at ER-PM junctions facilitate an open Ca2+ release activated Ca2+ (CRAC) channel conformation which enables the movement of Ca2+ into the cytosol from the high concentrations of the extracellular space. In immune cells, the sustained cytosolic Ca2+ elevations via CRAC channels induce the Ca2+-calmodulin/calcineurin dependent dephosphorylation of the nuclear factor of activated T-cells which subsequently enters the nucleus and begins transcriptional regulation of genes promoting T-cell activation 1,3. The process of CRAC channel activation by STIM1 23,24 via agonist-induced ER luminal Ca2+ depletion and the resulting sustained cytosolic Ca2+ elevation is collectively termed SOCE 25. The vital role of SOCE in T-cells is evident by studies demonstrating that heritable mutations in both STIM1 and Orai1 can cause severe combined immunodeficiency syndromes 3,19,26,27. EFSAM initiates SOCE after sensing ER-luminal Ca2+ depletion via the loss of Ca2+ coordination at the canonical EF-hand, ultimately leading to the destabilization-coupled self-association 7,28,29.
Glycosylation is the covalent attachment and processing of oligosaccharide structures, also known as glycans, through various biosynthetic steps in the ER and Golgi (reviewed in 30,32,33). There are two predominant types of glycosylation in eukaryotes: N-linked and O-linked, depending on the specific amino acid and atom bridging the linkage. In N-glycosylation, glycans are attached to the side chain amide of Asn, and in most cases, the initiation step occurs in the ER as the polypeptide chain moves into the lumen 34. The first step of N-glycosylation is the transfer of a fourteen-sugar core structure made up of glucose (Glc), mannose (Man), and N-acetylglucosamine (GlcNAc) (i.e. Glc3Man9GlcNAc2) from an ER membrane lipid by an oligosaccharyltransferase 35,36. Further steps, such as cleavage or transfer of glucose residues, are catalyzed in the ER by specific glycosidases and glycosyltransferases. Some proteins that leave the ER and move into the Golgi can be further processed 37. O-glycosylation refers to the addition of glycans, usually to the side chain hydroxyl group of Ser or Thr residues, and this modification occurs entirely in the Golgi complex 33,34. There are several O-glycan structures which can be made up of N-acetylglucosamine, fucose, galactose, and sialic acid with each monosaccharide added sequentially 33.
While no specific sequence has been identified as prerequisite for many types of O-glycosylation, a common consensus sequence has been associated with the N-linked modification: Asn-X-Ser/Thr/Cys, where X can be any amino acid except Pro 33. STIM1 EFSAM contains two of these consensus N-glycosylation sites: Asn131-Trp132-Thr133 and Asn171-Thr172-Thr173. Indeed, previous studies have shown that EFSAM can be N-glycosylated in mammalian cells at Asn131 and Asn171 38,39,40,41. However, previous studies of the consequences of N-glycosylation on SOCE have been incongruent, suggesting suppressed, potentiated or no effect by this post-translational modification on SOCE activation 38,39,40,41. Thus, research on the underlying biophysical, biochemical, and structural consequences of EFSAM N-glycosylation is vital to comprehending the regulatory effects of this modification. Due to the requirement for high levels of homogeneous proteins in these in vitro experiments, a site-selective approach to covalently attach glucose moieties to EFSAM was applied. Interestingly, Asn131 and Asn171 glycosylation caused structural changes that converge within the EFSAM core and enhance the biophysical properties which promote STIM1-mediated SOCE 42.
The chemical attachment of glycosyl groups to Cys thiols has been well-established by a seminal work which first demonstrated the utility of this enzyme-free approach to understanding the site-specific effects of glycosylation on protein function 43,44. More recently and with respect to STIM1, the Asn131 and Asn171 residues were mutated to Cys and glucose-5-(methanethiosulfonate) [glucose-5-(MTS)] was used to covalently link glucose to the free thiols 42. Here, we describe this approach which not only uses mutagenesis to incorporate site specific Cys residues for modification, but also applies solution nuclear magnetic resonance (NMR) spectroscopy to rapidly assess both modification efficiency and structural perturbations as a result of the glycosylation. Notably, this general methodology is easily adaptable to study the effects of either O- or N-glycosylation of any recombinantly produced protein.

<table><tbody><tr><td>Phusion DNA Polymerase</td><td>Thermo Fisher Scientific</td><td>F530S</td><td>Use in step 1.3.</td></tr><tr><td>Generuler 1kb DNA Ladder</td><td>Thermo Fisher Scientific</td><td>FERSM1163</td><td>Use in step 1.6.</td></tr><tr><td>DpnI Restriction Enzyme</td><td>New England Biolabs, Inc.</td><td>R0176</td><td>Use in step 1.8.</td></tr><tr><td>Presto Mini Plasmid Kit</td><td>GeneAid, Inc.</td><td>PDH300</td><td>Use in step 1.16.</td></tr><tr><td>BL21 DE3 codon (+) E. coli</td><td>Agilent Technologies, Inc.</td><td>230280</td><td>Use in step 2.1.</td></tr><tr><td>DH5a E. coli</td><td>Invitrogen, Inc.</td><td>18265017</td><td>Use in step 1.9.</td></tr><tr><td>0.22 mm Syringe Filter</td><td>Millipore, Inc.</td><td>SLGV033RS</td><td>Use in step 2.3.</td></tr><tr><td>HisPur Ni&lt;sup&gt;2+&lt;/sup&gt;-NTA Agarose Resin</td><td>Thermo Fisher Scientific</td><td>88221</td><td>Use in step 3.3.</td></tr><tr><td>3,500 Da MWCO Dialysis Tubing</td><td>BioDesign, Inc.</td><td>D306</td><td>Use in step 3.8, 3.16, 4.2, 4.5 and 4.6.</td></tr><tr><td>Bovine Thrombin</td><td>BioPharm Laboratories, Inc.</td><td>SKU91-055</td><td>Use in step 3.9.</td></tr><tr><td>5 mL HiTrap Q FF Anion Exchange Column</td><td>GE Healthcare, Inc.</td><td>17-5156-01</td><td>Use in step 3.11.</td></tr><tr><td>Glucose-5-MTS</td><td>Toronto Research Chemicals, Inc.</td><td>G441000</td><td>Use in step 4.1.</td></tr><tr><td>Vivaspin 20 Ultrafiltration Centrifugal Concentrators</td><td>Sartorius, Inc.</td><td>VS2001</td><td>Use in step 3.11, 4.2, 4.5 and 4.6.</td></tr><tr><td>PageRuler Unstained Broad Protein Ladder</td><td>Thermo Fisher Scientific</td><td>26630</td><td>Use in step 3.7, 3.10 and 3.15</td></tr><tr><td>HiTrap Q FF Anion Exchange Column</td><td>GE Healthcare, Inc.</td><td>17-5053-01</td><td>Use in step 3.12.</td></tr><tr><td>AKTA Pure Fast Protein Liquid Chromatrography System</td><td>GE Healthcare, Inc.</td><td>29018224</td><td>Use in step 3.14.</td></tr><tr><td>600 MHz Varian Inova NMR Spectrometer</td><td>Agilent Technologies, Inc.</td><td></td><td>Use in step 5.2 and 5.5.</td></tr></tbody></table>

targeting cysteine thiols for in vitro site-specific glycosylation of recombinant proteins

Stromal interaction molecule-1 (STIM1) is a type-I transmembrane protein located on the endoplasmic reticulum (ER) and plasma membranes (PM). ER-resident STIM1 regulates the activity of PM Orai1 channels in a process known as store operated calcium (Ca2+) entry which is the principal Ca2+ signaling process that drives the immune response. STIM1 undergoes post-translational N-glycosylation at two luminal Asn sites within the Ca2+ sensing domain of the molecule. However, the biochemical, biophysical, and structure biological effects of N-glycosylated STIM1 were poorly understood until recently due to an inability to readily obtain high levels of homogeneous N-glycosylated protein. Here, we describe the implementation of an in vitro chemical approach which attaches glucose moieties to specific protein sites applicable to understanding the underlying effects of N-glycosylation on protein structure and mechanism. Using solution nuclear magnetic resonance spectroscopy we assess both efficiency of the modification as well as the structural consequences of the glucose attachment with a single sample. This approach can readily be adapted to study the myriad glycosylated proteins found in nature.

The first step of this approach requires the mutagenesis of the candidate glycosylation residues to Cys residues which can be modifiable using the glucose-5-MTS. EFSAM has no endogenous Cys residues, so no special considerations need to be made prior to the mutagenesis. However, native Cys residues must be mutated to non-modifiable residues prior to performing the described chemistry. To minimally effect the native structure, we suggest performing a global sequence alignment of the protei...

Watch this Scientific Journal Video about Targeting Cysteine Thiols for in Vitro Site-specific Glycosylation of Recombinant Proteins at JoVE.com

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

Biochemical and structural analyses of glycosylated proteins require relatively large amounts of homogeneous samples. Here, we present an efficient chemical method for site-specific glycosylation of recombinant proteins purified from bacteria by targeting reactive Cys thiols.

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

Stromal interaction molecule-1 (STIM1) is a type-I transmembrane protein located on the endoplasmic reticulum (ER) and plasma membranes (PM). ER-resident ...

targeting-cysteine-thiols-for-vitro-site-specific-glycosylation

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Immunology and Infection

6.5K Views.  University of Western Ontario. Biochemical and structural analyses of glycosylated proteins require relatively large amounts of homogeneous samples. Here, we present an efficient chemical method for site-specific glycosylation of recombinant proteins purified from bacteria by targeting reactive Cys thiols.

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

Synthesis of Protein Bioconjugates via Cysteine-maleimide Chemistry

The chemical linking or bioconjugation of proteins to fluorescent dyes, drugs, polymers and other proteins has a broad range of applications, such as the development of antibody drug conjugates (ADCs) and nanomedicine, fluorescent microscopy and systems chemistry. For many of these applications, specificity of the bioconjugation method used is of prime concern. The Michael addition of maleimides with cysteine(s) on the target proteins is highly selective and proceeds rapidly under mild conditions, making it one of the most popular methods for protein bioconjugation.
We demonstrate here the modification of the only surface-accessible cysteine residue on yeast cytochrome c with a ruthenium(II) bisterpyridine maleimide. The protein bioconjugation is verified by gel electrophoresis and purified by aqueous-based fast protein liquid chromatography in 27% yield of isolated protein material. Structural characterization with MALDI-TOF MS and UV-Vis is then used to verify that the bioconjugation is successful. The protocol shown here is easily applicable to other cysteine - maleimide coupling of proteins to other proteins, dyes, drugs or polymers.

Synthesis of Protein Bioconjugates via Cysteine-maleimide Chemistry

This protocol details the important steps required for the bioconjugation of a cysteine containing protein to a maleimide, including reagent purification, reaction conditions, bioconjugate purification and bioconjugate characterization.

The chemical linking or bioconjugation of proteins to fluorescent dyes, drugs, polymers and other proteins has a broad range of applications, such as the ...

Chemistry

Characterization of Glycoproteins with the Immunoglobulin Fold by X-Ray Crystallography and Biophysical Techniques

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of these glycoproteins possess immunoglobulin (Ig) folds and are central to immune recognition and regulation. Here, we present a platform for the design, expression and biophysical characterization of the extracellular domain of human B cell receptor CD22. We propose that these approaches are broadly applicable to the characterization of mammalian glycoprotein ectodomains containing Ig domains. Two suspension human embryonic kidney (HEK) cell lines, HEK293F and HEK293S, are used to express glycoproteins harbouring complex and high-mannose glycans, respectively. These recombinant glycoproteins with different glycoforms allow investigating the effect of glycan size and composition on ligand binding. We discuss protocols for studying the kinetics and thermodynamics of glycoprotein binding to biologically relevant ligands and therapeutic antibody candidates. Recombinant glycoproteins produced in HEK293S cells are amenable to crystallization due to glycan homogeneity, reduced flexibility and susceptibility to endoglycosidase H treatment. We present methods for soaking glycoprotein crystals with heavy atoms and small molecules for phase determination and analysis of ligand binding, respectively. The experimental protocols discussed here hold promise for the characterization of mammalian glycoproteins to give insight into their function and investigate the mechanism of action of therapeutics.

We present approaches for the biophysical and structural characterization of glycoproteins with the immunoglobulin fold by biolayer interferometry, isothermal titration calorimetry, and X-ray crystallography.

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of ...

Biochemistry

Analysis of the Solvent Accessibility of Cysteine Residues on Maize rayado fino virus Virus-like Particles Produced in Nicotiana benthamiana Plants and Cross-linking of Peptides to VLPs

Mimicking and exploiting virus properties and physicochemical and physical characteristics holds promise to provide solutions to some of the world's most pressing challenges. The sheer range and types of viruses coupled with their intriguing properties potentially give endless opportunities for applications in virus-based technologies. Viruses have the ability to self- assemble into particles with discrete shape and size, specificity of symmetry, polyvalence, and stable properties under a wide range of temperature and pH conditions. Not surprisingly, with such a remarkable range of properties, viruses are proposed for use in biomaterials 9, vaccines 14, 15, electronic materials, chemical tools, and molecular electronic containers4, 5, 10, 11, 16, 18, 12.
In order to utilize viruses in nanotechnology, they must be modified from their natural forms to impart new functions. This challenging process can be performed through several mechanisms including genetic modification of the viral genome and chemically attaching foreign or desired molecules to the virus particle reactive groups 8. The ability to modify a virus primarily depends upon the physiochemical and physical properties of the virus. In addition, the genetic or physiochemical modifications need to be performed without adversely affecting the virus native structure and virus function. Maize rayado fino virus (MRFV) coat proteins self-assemble in Escherichia coli producing stable and empty VLPs that are stabilized by protein-protein interactions and that can be used in virus-based technologies applications 8. VLPs produced in tobacco plants were examined as a scaffold on which a variety of peptides can be covalently displayed 13. Here, we describe the steps to 1) determine which of the solvent-accessible cysteines in a virus capsid are available for modification, and 2) bioconjugate peptides to the modified capsids. By using native or mutationally-inserted amino acid residues and standard coupling technologies, a wide variety of materials have been displayed on the surface of plant viruses such as, Brome mosaic virus 3, Carnation mottle virus 12, Cowpea chlorotic mottle virus 6, Tobacco mosaic virus 17, Turnip yellow mosaic virus 1, and MRFV 13.

Analysis of the Solvent Accessibility of Cysteine Residues on Maize rayado fino virus Virus-like Particles Produced in Nicotiana benthamiana Plants and Cross-linking of Peptides to VLPs

A method to analyze the solvent accessibility of the thiol group of cysteine residues of Maize rayado fino virus (MRFV)-virus-like particles (VLPs) followed by a peptide cross-linking reaction is described. The method takes advantage of the availability of several chemical groups on the surface of the VLPs that can be targets for specific reactions.

Mimicking  and exploiting virus properties and physicochemical and physical  characteristics holds promise to provide solutions to some of  the world's ...

Synthesis and Bioconjugation of Thiol-Reactive Reagents for the Creation of Site-Selectively Modified Immunoconjugates

Maleimide-bearing bifunctional probes have been employed for decades for the site-selective modification of thiols in biomolecules, especially antibodies. Yet maleimide-based conjugates display limited stability in vivo because the succinimidyl thioether linkage can undergo a retro-Michael reaction. This, of course, can lead to the release of the radioactive payload or its exchange with thiol-bearing biomolecules in circulation. Both of these processes can produce elevated activity concentrations in healthy organs as well as decreased activity concentrations in target tissues, resulting in reduced imaging contrast and lower therapeutic ratios. In 2018, we reported the creation of a modular, stable, and easily accessible phenyloxadiazolyl methyl sulfone reagent &#8212; dubbed &#8216;PODS&#8217; &#8212; as a platform for thiol-based bioconjugations. We have clearly demonstrated that PODS-based site-selective bioconjugations reproducibly and robustly create homogenous, well-defined, highly immunoreactive, and highly stable radioimmunoconjugates. Furthermore, preclinical experiments in murine models of colorectal cancer have shown that these site-selectively labeled radioimmunoconjugates exhibit far superior in vivo performance compared to radiolabeled antibodies synthesized via maleimide-based conjugations. In this protocol, we will describe the four-step synthesis of PODS, the creation of a bifunctional PODS-bearing variant of the ubiquitous chelator DOTA (PODS-DOTA), and the conjugation of PODS-DOTA to the HER2-targeting antibody trastuzumab.

In this protocol, we will describe the synthesis of PODS, a phenyoxadiazolyl methyl sulfone-based reagent for the site-selective attachment of cargos to the thiols of biomolecules, particularly antibodies. In addition, we will describe the synthesis and characterization of a PODS-bearing bifunctional chelator and its conjugation to a model antibody.

Maleimide-bearing bifunctional probes have been employed for decades for the site-selective modification of thiols in biomolecules, especially antibodies. ...

Preparation of Site-Specific Cytotoxic Protein Conjugates via Maleimide-thiol Chemistry and Sortase A-Mediated Ligation

Cancer is currently the second most common cause of death worldwide. The hallmark of cancer cells is the presence of specific marker proteins such as growth factor receptors on their surface. This feature enables development of highly selective therapeutics, the protein bioconjugates, composed of targeting proteins (antibodies or receptor ligands) connected to highly cytotoxic drugs by a specific linker. Due to very high affinity and selectivity of targeting proteins the bioconjugates recognize marker proteins on the cancer cells surface and utilize receptor-mediated endocytosis to reach the cell interior. Intracellular vesicular transport system ultimately delivers the bioconjugates to the lysosomes, where proteolysis separates free cytotoxic drugs from the proteinaceous core of the bioconjugates, triggering drug-dependent cancer cell death. Currently, there are several protein bioconjugates approved for cancer treatment and large number is under development or clinical trials.
One of the main challenges in the generation of the bioconjugates is a site-specific attachment of the cytotoxic drug to the targeting protein. Recent years have brought a tremendous progress in the development of chemical and enzymatic strategies for protein modification with cytotoxic drugs. Here we present the detailed protocols for the site-specific incorporation of cytotoxic warheads into targeting proteins using a chemical method employing maleimide-thiol chemistry and an enzymatic approach that relies on sortase A-mediated ligation. We use engineered variant of fibroblast growth factor 2 and fragment crystallizable region of human immunoglobulin G as an exemplary targeting proteins and monomethyl auristatin E and methotrexate as model cytotoxic drugs. All the described strategies allow for highly efficient generation of biologically active cytotoxic conjugates of defined molecular architecture with potential for selective treatment of diverse cancers.

Here we provide detailed protocols for a site-specific labeling of proteins with cytotoxic drugs using maleimide-thiol reaction and sortase A-mediated ligation.

Cancer is currently the second most common cause of death worldwide. The hallmark of cancer cells is the presence of specific marker proteins such as ...

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed procedure of a quantitative redox proteomics method that utilizes resin-assisted capture (RAC) in combination with tandem mass tag (TMT) isobaric labeling and liquid chromatography-tandem mass spectrometry (LC-MS/MS) to allow multiplexed stochiometric quantification of oxidized protein thiols at the proteome level. The site-specific quantitative information on oxidized cysteine residues provides additional insight into the functional impacts of such modifications. 
The workflow is adaptable across many sample types, including cultured cells (e.g., mammalian, prokaryotic) and whole tissues (e.g., heart, lung, muscle), which are initially lysed/homogenized and with free thiols being alkylated to prevent artificial oxidation. The oxidized protein thiols are then reduced and captured by a thiol-affinity resin, which streamlines and simplifies the workflow steps by allowing the proceeding digestion, labeling, and washing procedures to be performed without additional transfer of proteins/peptides. Finally, the labeled peptides are eluted and analyzed by LC-MS/MS to reveal comprehensive stoichiometric changes related to thiol oxidation across the entire proteome. This method greatly improves the understanding of the role of redox-dependent regulation under physiological and pathophysiological states related to protein thiol oxidation.

Protein thiol oxidation has significant implications under normal physiological and pathophysiological conditions. We describe the details of a quantitative redox proteomics method, which utilizes resin-assisted capture, isobaric labeling, and mass spectrometry, enabling site-specific identification and quantification of reversibly oxidized cysteine residues of proteins.

Reversible oxidative modifications on protein thiols have recently emerged as important mediators of cellular function. Herein we describe the detailed ...

Nonspecific Proteolysis-Based Glycomics Strategy for Bacterial Glycoproteins

Protein glycosylation is one of the most common and complex post-translational modifications. Many techniques have been developed to characterize the specific roles of glycans, the relationship between their structures and their impact on the functions of proteins. A common method for glycan analysis is to employ exoglycosidase cleavage to release N-linked glycans from glycoproteins or glycopeptides using Peptide-N-Glycosidase F (PNGase F). However, the glycan-protein linkages in bacteria are different and there is no enzyme available to release glycans from bacterial glycoproteins. In addition, free glycans have also been described in mammalian cells, bacteria, yeast, plants, and fish. In this article, we present a method that can characterize the N-linked glycosylation system in Campylobacter jejuni by detecting asparagine (Asn)-linked and free glycans that are not attached to their target proteins. In this method, total proteins from C. jejuni were digested by Pronase E with a higher enzyme to protein ratio (2:1&#8722;3:1) and a longer incubation time (48&#8722;72 h). The resulted Asn-glycans and free glycans were then purified using porous graphitic carbon cartridges, permethylated, and analyzed by mass spectrometry.

This study presents a method for glycomics analysis of glycoproteins by a combination of pronase E digestion, permethylation, and mass spectrometry analysis. This method is capable of analyzing all types of N-linked glycans, including bacterial N-glycans.

Protein glycosylation is one of the most common and complex post-translational modifications. Many techniques have been developed to characterize the ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Biology

Homogeneous Glycoconjugate Produced by Combined Unnatural Amino Acid Incorporation and Click-Chemistry for Vaccine Purposes

Genetic code expansion is a powerful tool to introduce unnatural amino acids (UAAs) into proteins to modify their characteristics, to study or create new protein functions or to have access to protein conjugates. Stop codon suppression, in particular amber codon suppression, has emerged as the most popular method to genetically introduce UAAs at defined positions. This methodology is herein applied to the preparation of a carrier protein containing an UAA harboring a bioorthogonal functional group. This reactive handle can next be used to specifically and efficiently graft a synthetic oligosaccharide hapten to provide a homogeneous glycoconjugate vaccine. The protocol is limited to the synthesis of glycoconjugates in a 1:1 carbohydrate hapten/carrier protein ratio but amenable to numerous pairs of biorthogonal functional groups. Glycococonjugate vaccine homogeneity is an important criterion to ensure complete physico-chemical characterization, thereby, satisfying more and more demanding drug regulatory agency recommendations, a criterion which is unmet by classical conjugation strategies. Moreover, this protocol makes it possible to finely tune the structure of the actual conjugate vaccine, giving rise to tools to address structure-immunogenicity relationships.

Genetic code expansion is applied for the introduction of an unnatural amino acid bearing a biorthogonal functional group on a carrier protein at a defined site. The biorthogonal function is further used for the site-selective coupling of a carbohydrate antigen to provide a homogeneous glycoconjugate vaccine.

Genetic code expansion is a powerful tool to introduce unnatural amino acids (UAAs) into proteins to modify their characteristics, to study or create new ...

Bioengineering

Rapid Antibody Glycoengineering in Chinese Hamster Ovary Cells

Recombinant monoclonal antibodies bind specific molecular targets and, subsequently, induce an immune response or inhibit the binding of other ligands. However, monoclonal antibody functionality and half-life may be reduced by the type and distribution of host-specific glycosylation. Attempts to produce superior antibodies have inspired the development of genetically modified producer cells that synthesize glyco-optimized antibodies. Glycoengineering typically requires the generation of a stable knockout or knockin cell line using methods such as clustered regularly interspaced short palindromic repeats (CRISPR)-associated protein 9. Monoclonal antibodies produced by engineered cells are then characterized using mass spectrometric methods to determine if the desired glycoprofile has been obtained. This strategy is time-consuming, technically challenging, and requires specialists. Therefore, an alternative strategy that utilizes streamlined protocols for genetic glycoengineering and glycan detection may assist endeavors toward optimal antibodies. In this proof-of-concept study, an IgG-producing Chinese hamster ovary cell served as an ideal host to optimize glycoengineering. Short interfering RNA targeting the Fut8 gene was delivered to Chinese hamster ovary cells, and the resulting changes in FUT8 protein expression were quantified. The results indicate that knockdown by this method was efficient, leading to a ~60% reduction in FUT8. Complementary analysis of the antibody glycoprofile was performed using a rapid yet highly sensitive technique: capillary gel electrophoresis and laser-induced fluorescence detection. All knockdown experiments showed an increase in afucosylated glycans; however, the greatest shift achieved in this study was ~20%. This protocol simplifies glycoengineering efforts by harnessing in silico design tools, commercially synthesized gene targeting reagents, and rapid quantification assays that do not require extensive prior experience. As such, the time efficiencies offered by this protocol may assist investigations into new gene targets.

The glycosylation pattern of an antibody determines its clinical performance, thus industrial and academic efforts to control glycosylation persist. Since typical glycoengineering campaigns are time- and labor-intensive, the generation of a rapid protocol to characterize the impact of glycosylation genes using transient silencing would prove useful.

Recombinant monoclonal antibodies bind specific molecular targets and, subsequently, induce an immune response or inhibit the binding of other ligands. ...

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

Summary

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Chapters in this video

Synthesis of Protein Bioconjugates via Cysteine-maleimide Chemistry

Characterization of Glycoproteins with the Immunoglobulin Fold by X-Ray Crystallography and Biophysical Techniques

Analysis of the Solvent Accessibility of Cysteine Residues on Maize rayado fino virus Virus-like Particles Produced in Nicotiana benthamiana Plants and Cross-linking of Peptides to VLPs

Synthesis and Bioconjugation of Thiol-Reactive Reagents for the Creation of Site-Selectively Modified Immunoconjugates

Preparation of Site-Specific Cytotoxic Protein Conjugates via Maleimide-thiol Chemistry and Sortase A-Mediated Ligation

Resin-Assisted Capture Coupled with Isobaric Tandem Mass Tag Labeling for Multiplexed Quantification of Protein Thiol Oxidation

Nonspecific Proteolysis-Based Glycomics Strategy for Bacterial Glycoproteins

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Homogeneous Glycoconjugate Produced by Combined Unnatural Amino Acid Incorporation and Click-Chemistry for Vaccine Purposes

Rapid Antibody Glycoengineering in Chinese Hamster Ovary Cells