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. Author manuscript; available in PMC: 2024 Apr 3.
Published in final edited form as: Methods Mol Biol. 2023;2674:147–167. doi: 10.1007/978-1-0716-3243-7_10

Mass Spectrometry-Based Methods to Determine the Substrate Specificities and Kinetics of N-Linked Glycan Hydrolysis by Endo-β-N-Acetylglucosaminidases

Jonathan J Du 1, Diego Sastre 2, Beatriz Trastoy 3, Blaine Roberts 2, Daniel Deredge 4, Erik H Klontz 5,6, Maria W Flowers 2, Nazneen Sultana 2, Marcelo E Guerin 3,7, Eric J Sundberg 8
PMCID: PMC10988651  NIHMSID: NIHMS1969275  PMID: 37258966

Abstract

Glycosylation is a common posttranslational modification of proteins and refers to the covalent addition of glycans, chains of polysaccharides, onto proteins producing glycoproteins. The glycans influence the structure, function, and stability of proteins. They also play an integral role in the immune system, and aberrantly glycosylated proteins have wide ranging effects, including leading to diseases such as autoimmune conditions and cancer. Carbohydrate-active enzymes (CAZymes) are produced in bacteria, fungi, and humans and are enzymes which modify glycans via the addition or subtraction of individual or multiple saccharides from glycans. One of the hurdles in studying these enzymes is determining the types of substrates each enzyme is specific for and the kinetics of enzymatic activity. In this chapter, we discuss methods which are currently used to study the substrate specificity and kinetics of CAZymes and introduce a novel mass spectrometry-based technique which enables the specificity and kinetics of CAZymes to be determined accurately and efficiently.

Keywords: LC-MS, Kinetics, N-glycans, ENGases

1. Introduction

N-glycosylation is a common posttranslational modification of proteins and refers to the covalent addition of glycans at asparagine (Asn) residues on proteins that contain the consensus glycosylation motif Asn-X-Ser/Thr/Cys where X is any amino acid except proline. These modifications markedly influence the structure, function, stability, and serum half-lives of proteins [1–4]. Glycans play a major role in many biological systems and are particularly important in immunity, to which they contribute to lymphocyte development, cell adhesion, immune signaling, and host-pathogen interactions [5–11]. As such, aberrantly glycosylated proteins have wide-ranging effects, altering cell signaling cascades that can lead to diseases such as autoimmunity and cancer.

The N-glycan is initially synthesized on dolichol-phosphate (Dol-P), a lipid-like polyisoprenoid molecule. During the translocation of newly synthesized proteins into the endoplasmic reticulum (ER), the pre-assembled common oligosaccharide Glc3Man9GlcNAc2 is transferred en bloc onto the protein. In the lumen of the ER and Golgi, a variety of glycoside hydrolases (GHs) and glycosyltransferases (GTs) remove and add various sugar residues generating chemically and structurally different N-glycans [12, 13].The final types of N-glycan produced are dependent on the organism species, cell type, protein, and even glycosylation sites [14].

There are three major types of N-glycans in eukaryotic cells: complex type (CT), high mannose (HM), and hybrid-type (Hy) as shown in Fig. 1. All N-glycans contain a common chitobiose core composed of Manα1–6(Manα1–3)Manβ1–4GlcNAcβ1–4-GlcNAcβ1. CT glycans, beyond the pentasaccharide core, have elongated branches that are made by GlcNAc, galactose (Gal), and sialic acid (also termed neuraminic acid, Neu5Ac) residues toward their terminal ends. CT glycans can be bi-, tri-, or tetraantennary [14]. Additionally, these glycans can contain a fucose (Fuc) attached to the Asn-proximal GlcNAc of the chitobiose core and also a bisecting GlcNAc located on the core Man (Fig. 1). The Hy-type N-glycan has a dual nature: its α(1, 3) antenna is equivalent to the α(1, 3) antenna of the CT-type N-glycans, while its α(1, 6) antenna is covered into the α(1, 6) antenna of HM-type N-glycans.

Fig. 1.

Fig. 1

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Common N-glycan structures including the structure of biantennary, triantennary, and tetraantennary complex-type (CT) glycans and high mannose (HM)- and hybrid (Hy)-type glycans

1.1. Endo-β-N-Acetylglucos-aminidases

GHs catalyze the hydrolysis of glycosidic bonds in complex sugars. The diversity of GHs is extraordinary with 173 GH families whose members hydrolyze distinct glycan bonds. Endoglycosidases cleave glycosidic bonds internal to a larger oligosaccharide glycan structure (e.g., endo-β-N-acetylglucosamines (ENGases) belonging to the glycoside hydrolase 18 (GH18) and 85 (GH85) families hydrolyze the bond between the two GlcNAc residues of the chitobiose core in N-linked glycans). In contrast, exoglycosidases, including most sialidases, galactosidases, β-N-acetylglucosaminidases, and fucosidases, remove terminal residues from glycans (e.g., sialidases, α-L-fucosidases, and α-1,2-L-fucosidases belonging to GH33 [15], GH29 [16] and GH95 [17] families, respectively). The majority of GHs currently known derive from bacteria, which mainly use these enzymes to liberate and metabolize glycans from their environment as a food source and to remodel their cell envelope. Some bacteria also leverage GHs to modify glycans on key proteins, providing the organism a mechanism for immune evasion [18, 19]. Nearly all of these enzymes are protein agnostic, only exhibiting specificity in recognizing their substrate glycan structures. However, some enzymes from Streptococcus pyogenes, including the GH18 family endoglycosidases EndoS and EndoS2, have been shown to be strictly protein specific, in this case to immunoglobulin G (IgG) antibodies [20–25].

The GHs produced in nature exhibit distinct substrate specificities and mechanisms of action, with the ability to remove combinations or individual sugar residues from each glycan. Structural studies revealed that many of these enzymes contain conserved catalytic domains that adopt a (β/α)8 TIM-barrel fold, such as for GH18 [23–26], GH85 [27–29], GH29 [16], and GH20 [30–32] families, or β-propeller folds for most sialidases [33, 34]. In each of these cases, the primary catalytic domain is decorated with loops that connect the secondary structural elements and mediate substrate specificity. Some GHs may contain additional domains, including carbohydrate binding modules (CBMs) believed to play myriad functional roles, such as increasing the local concentration of substrate for the GH domain and aiding enzymes in adhering to molecules and/or cell surfaces [35, 36].

One class of proteins that are especially influenced by glycans are IgG antibodies, which play an integral role in immunity, directly by neutralizing pathogens and indirectly by stimulating other immune responses against them. The latter, known as antibody-mediated effector functions, occurs when antibodies bind to Fc γ receptors (FcγRs) or C1q complement, triggering signaling cascades that either stimulate or repress the immune system. The glycan attached to Asn297 of the fragment crystallizable (Fc) region IgG antibodies influences the structure and dynamics of the Fc region and is the key molecular determinant in initiating this signaling cascade [37]. Different IgG Fc domain glycoforms influence the type and intensity of signaling and subsequent downstream effects. For instance, the removal of the core fucose moiety increases binding to the activating FcγR3a by ~100-fold, producing marked increases in antibody-dependent cellular cytotoxicity (ADCC) [38, 39], while the presence of galactosylation is associated with both pro-[40] and anti-inflammatory [41] effects, and sialylation is associated with anti-inflammatory effects [42–44]. The glycan composition is heterogeneous when produced in the body or in cell culture systems used for recombinant glycoprotein production, such as human embryonic kidney (HEK) or Chinese hamster ovary (CHO) cells. Although IgG monoclonal antibodies (mAbs) are highly efficacious therapeutics, the varied glycan compositions linked to Asn297 result in inter-batch variability in the immune responses that they induce. In clinical practice, the use of therapeutics such as mAbs with custom glycoprofiles can improve therapeutic outcomes [45, 46].

1.2. Chemoenzymatic Remodeling of Glycoproteins

One method for producing homogenously glycosylated glycoproteins is through chemoenzymatic synthesis. Research by Wang and coworkers has shown that mutation of the residue that stabilizes the reaction intermediate, adjacent to the acid/base in the catalytic site of GH18 [47, 48] and GH85 [49, 50] enzymes, produces variants capable of transferring glycans with an activated donor, such as an oxazoline-derivatized glycan, back onto glycoproteins. Initially, wild-type GH18 or GH85 enzymes are used to remove the heterogeneous glycans originally attached to the protein. The glycoprotein, now with only a GlcNAc linked to the Asn, is incubated with a glycosynthase variant of the enzyme and an oxazoline-linked substrate to produce a homogeneously glycosylated glycoprotein. This reaction can be monitored using a variety of methods as described below, such as LC-MS [51]. A schematic of chemoenzymatic remodeling pathways is shown in Fig. 2.

Fig. 2.

Fig. 2

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Chemoenzymatic remodeling schematic. ENGases from the GH18 and GH85 family hydrolyze the heterogenous IgG glycoforms, and glycosynthase mutants can transfer a defined glycan from an oxazoline donor to the GlcNAc still linked to the Asn297 of the antibody, according to the N-glycan specificity of the enzyme

2. SEAK/C-SEAK Protocol

2.1. Materials

2.1.1. Bacterial Strains and Media

  1. E. coli strains – BL21, Neb5α

  2. Luria Bertani broth.

  3. Ampicillin 100 mg/mL.

2.1.2. Eukaryotic Expression Strains

  1. HEK293F cells.

2.1.3. Plasmids and Cloning

  1. Rituximab heavy chain (RitHc) in pcDNA3.4-TOPO vector (Thermo).

  2. Rituximab light chain (RitLc) in pcDNA3.4-TOPO vector (Thermo).

  3. Primers to produce Fc.

2.1.4. Proteins

  1. Transferrin (Sigma 90910).

  2. RNase B (NEB P7817S).

2.1.5. Protein Purification

  1. PBS pH 7.4.

  2. Phytic acid (Sigma P8810) (1 mg/mL) in PBS pH 7.4.

  3. Ni-NTA agarose.

  4. HiTrap Protein A column.

2.1.6. Materials for Eukaryotic Expression

  1. Expi293 Expression System (Thermo 14635).

2.1.7. Mass Spectrometry Materials

  1. Buffer A: 0.1% formic acid in H2O.

  2. Buffer B: 0.1% formic acid in acetonitrile (ACN).

  3. Mass spectrometry sample vials.

  4. PLRP-S 1000 Å, 2.1 × 50 mm, 5 μm column (Agilent PL1912-1502)

2.1.8. Instruments

  1. Agilent 6545XT Advance Bio LC/Q-TOF equipped with an Agilent 1290 Infinity II UHPLC.

2.1.9. Software

  1. Agilent Acquisition Software.

  2. Agilent BioConfirm Software.

3. Methods

3.1. Cloning to Produce Fc Plasmid

  1. The primers Fc forward primer (Fcfp) (5′-GAGCCTAAGAG CTGCGAC-3′) and Fc reverse primer (Fcrp) (5′-TCCTGTAG ATCCGGGCAC-3′) are diluted to 10 μM with deionized water. Set up the PCR reaction following the Q5 Site-Directed Mutagenesis Kit Quick Protocol (NEB). Mix 1 ng of DNA with 6.3 μL of Q5 Hot Start High-Fidelity 2X master mix, 0.6 μL each of the forward and reverse primers (10 μM) and deionized water added for a final reaction volume of 13 μL. The Ta is 67C, with a 30 s annealing time and an extension time of 3.4 min was used at 72C.

  2. Mix 1 μL of the PCR product with 5 μL of 2X KLD Reaction buffer, 1 μL of KLD enzyme mix, and 3 μL of deionized water. Incubate the reaction mixture at room temperature for 5 min.

  3. Transform 5 μL of the reaction mixture into chemically competent E. coli Neb5α cells, plated on agar plates with ampicillin (100 μg/mL).

  4. Pick single colonies from the plate and grow overnight at 37 C in 5 mL LB containing ampicillin (100 μg/mL).

  5. Follow the protocol in a commercial Miniprep Kit to extract the DNA.

  6. Confirmation of the new plasmid is verified by DNA sequencing. The forward sequencing is conducted using the commercially universal CMV primer (5′-CGCAAATGGGCGGTAGG CGTG-3′) and reverse sequencing is performed using the custom primer (5′-GGTAGGGATCGAACCCTTACCGG-3′).

3.2. Expression and Purification of EndoS/EndoS2

  1. Transform 100 ng of plasmid into chemically competent E. coli BL21 cells, and plate on agar plates supplemented with ampicillin (100 μg/mL) and leave to grow overnight at 37 C.

  2. Pick a single colony from the plate to inoculate 100 mL of LB broth containing ampicillin (100 μg/mL) and grow overnight on an orbital shaker at 37 C.

  3. Inoculate 1 L of LB with ampicillin (100 μg/mL) with the overnight culture at a ratio of 1:50. Grow the cells at 37 C until the OD600 is ~0.6–0.8 and induce with IPTG for a final concentration of 0.5 mM. Grow the culture at 22 C overnight in an orbital shaker.

  4. Harvest the cells by spinning them at 4500G for 20 min, discard the supernatant and resuspend the pellet with PBS + 10% glycerol.

  5. Lyse the cells via sonication or chemical lysis and clarify by spinning the lysate at 18000RPM for 45 min.

  6. Load the supernatant onto NiNTA resin in a gravity column pre-equilibrated with PBS pH 7.4. Incubate the supernatant with the NiNTA resin for 2 h and wash the column with 10 column volumes of PBS pH 7.4.

  7. Induce CPD cleavage by adding 2CVof a 1 mg/mL solution of phytic acid in PBS pH 7.4 to the gravity column and incubate with the resin for 1 h on a rotator or rocking device.

  8. Collect the flowthrough in 1 mL fractions and run the samples on an SDS-PAGE to confirm purity. The enzyme should be sufficiently pure.

3.3. Transfection and Expression of HM-Rituximab and Fc

  1. Transfection protocols were carried out per manufacturer’s instructions.

3.4. Purification of IgG Antibodies and Fc

  1. Pre-equilibrate a HiTrap Protein A column with PBS pH 7.4.

  2. Flow the harvested serum over the Protein A column using either an Akta or peristaltic pump at 1 mL/min until all the serum has been passed over the resin.

  3. Elute the protein by attaching the column to an Akta system and wash the column with 10CV of PBS pH 7.4.

  4. To elute the protein, inject 5 mL of 100 mM citrate pH 3 into the column at 1 mL/min to elute the protein and collect in 1 mL fractions. Add 100 μL of 1 M Tris pH 9 to each fraction to neutralize the citrate and return the buffer to a neutral pH.

  5. To confirm the purity of the sample, run the IgG or Fc protein on an SDS-PAGE gel. Run 2 samples, one with reducing buffer and one without.

3.5. LC-MS Setup for Glycoprotein Analysis

Table 1 shows the parameters for intact glycoprotein analysis using the Agilent 1290 Infinity II UHPLC system and the Agilent Dual Jet Stream ESI source. Table 2 shows the binary pump gradient which is programmed into the analysis method (see Note 1) – gradient can be modified to change sampling rate.

Table 1.

LC-MS parameters

LC 1290 Infinity II UHPLC
Column compartment 80 °C
Flow rate 0.4 mL/min
Method runtime 4.5 min
Posttime 1 min
Source Agilent Dual Jet Stream
Gas temperature 350 °C
Gas flow 12 L/min
Nebulizer 60 psig
400 °C
Sheath gas flow 11 L/min
Vcap 5.5 kV
Nozzle voltage 2 kV
Fragmentor 380 kV
Skimmer 140 kV
Acquisition rate 1 spectrum/sec
Acquisition mode Positive
Mass range 100–3200, 100–10,000 m/z
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Table 2.

LC Binary Pump gradient

Time(min) %A %B
0 98 2
1.25 80 20
2.6 50 50
3 50 50
4 10 90
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3.6. SEAK with EndoS

  1. Separately, buffer exchange the EndoS and rituximab into PBS pH 7.4 using Amicon Centrifugation units with appropriate molecular weight cutoffs. Adjust the final concentration of rituximab to 30 μM and EndoS2 to 10 nM (see Note 2).

  2. Set up worklist in Agilent Acquisition software to repeatedly sample the same reaction vial.

  3. Add 5 μL of rituximab to 22 μL of PBS pH 7.4 in a microcentrifuge tube.

  4. Start the reaction next to the instrument by adding 3 μL of EndoS to the reaction mixture (final reaction volume of 30 μL) and mix well. Transfer the reaction mixture to a mass spec vial and place into the autosampler.

  5. The LC-MS method runs takes approximately 4.5 min to run. Inject 0.1 μL each time.

  6. Using BioConfirm, deconvolute the mass spectra for each run and export the deconvoluted spectra as a csv file.

3.7. SEAK with EndoBT-3987

  1. Separately, buffer exchange the EndoBT-3987 and RNase B into PBS pH 7.4 using Amicon Centrifugation units with appropriate molecular weight cutoffs. Adjust the final concentration of RNase B to 10 μM and EndoBT-3987 to 500 nM (see Note 2).

  2. Set up worklist in Agilent Acquisition software to repeatedly sample the same reaction vial.

  3. Start the reaction next to the instrument by adding 3 μL of EndoBT-3987 to 27 μL of RNase B in a microcentrifuge tube the reaction mixture (final reaction volume of 30 μL) and mix well. Transfer the reaction mixture to a mass spec vial and place into the autosampler.

  4. The LC-MS method runs takes approximately 4.5 min to run. Inject 0.1 μL each time.

  5. Using BioConfirm, deconvolute the mass spectra for each run and export the deconvoluted spectra as a csv file. The deconvolution parameters for each component of the SEAK cocktail are summarized in Table 3.

Table 3.

Deconvolution parameters for SEAK cocktail

Protein m/z range Mass range (kDa)
RNase B 1000–2400 13–16
Transferrin 1800–3000 75–81
Rituximab 2000–7000 144–150
Fc 1250–3000 52–57
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3.8. C-SEAK with EndoS2

  1. Separately, buffer exchange the EndoS2, HM-IgG1WT (HM-Rituximab), IgG1-FcWT, RNase B, and Transferrin into PBS pH 7.4 using Amicon Centrifugation units with appropriate molecular weight cutoffs. Adjust the final concentration of the substrates to be 30 μM and EndoS2 to be at 10 nM (see Note 2).

  2. Set up worklist in Agilent Acquisition software to repeatedly sample the same reaction vial.

  3. Add 1 μL of each glycoprotein substrate to 23 μL of PBS pH 7.4.

  4. Start the reaction next to the instrument by adding 3 μL of EndoS2 to the reaction mixture (final reaction volume of 30 μL) and mix well. Transfer the reaction mixture to a mass spec vial and place into the autosampler.

  5. The LC-MS method runs takes approximately 4.5 min to run. Inject 0.1 μL each time.

  6. Using BioConfirm, deconvolute the mass spectra for each run individually for each protein, and export the deconvoluted spectra as a csv file. The deconvolution parameters for each component of the SEAK cocktail are summarized in Table 3.

3.9. C-SEAK with EndoBT-3987

  1. Separately, buffer exchange the EndoBT-3987, HM-IgG1WT (HM-Rituximab), IgG1-FcWT, RNase B, and Transferrin into PBS pH 7.4 using Amicon Centrifugation units with appropriate molecular weight cutoffs. Adjust the final concentration of the substrates to be 30 μM and EndoBT-3987 to be 500 Nm (see Note 2).

  2. Set up worklist in Agilent Acquisition software to repeatedly sample the same reaction vial.

  3. Add 2 μL of each glycoprotein substrate to 19 μL of PBS pH 7.4.

  4. Start the reaction next to the instrument by adding 3 μL of EndoBT-3987 to the reaction mixture (final reaction volume of 30 μL) and mix well. Transfer the reaction mixture to a mass spec vial and place into the autosampler.

  5. The LC-MS method runs takes approximately 4.5 min to run. Inject 0.1 μL each time.

  6. Using BioConfirm, deconvolute the mass spectra for each run individually for each protein, and export the deconvoluted spectra as a csv file. The deconvolution parameters for each component of the SEAK cocktail are summarized in Table 3.

3.10. Importing Data into Excel

To import all the extracted data into Excel for semi-quantitation, the power query feature of Excel is used.

3.11. Exporting Data from Excel into KinTek

  1. To prepare the data for use in KinTek, rearrange the data in the Excel sheet to be organized by columns such as time, pGG, pG, and p.

  2. Export the table as a tab-delimited file.

  3. See KinTek manual instructions for how to design a reaction model and fit data.

4. Measuring ENGase Activities and Specificities

One challenging aspect of studying ENGases and other glycan-modifying enzymes is the determination of the enzyme’s substrate specificity and the kinetics of its deglycosylation and transglycosylation reactions. The substrate specificities of enzymes can sometimes be predicted using bioinformatic approaches and comparison of amino acid sequences to previously characterized enzymes; however, experimental validation is often still required. A multitude of analytical methods exists for experimental determination of glycan specificities of enzymes, such as SDS-PAGE, TLC, liquid chromatography, and mass spectrometry. All of these methods can be used for either qualitative or quantitative analyses. In addition to quantification of the formation of reaction products, the kinetic constants (e.g., KM and Vmax) of these enzyme-catalyzed reactions can be determined by analyzing the reactions over time. These constants can be determined in multiple ways. One method is to make initial velocity measurements in which varying substrate concentrations are incubated with a constant amount of enzyme, and all other variables are kept constant. The reactions are quenched after a period, and the quantified products can be fitted to a Michaelis-Menten kinetic model to obtain the rate constants. An alternative method is to analyze these reactions over time, tracking the relative populations of the substrate and products. The resulting data can then be fitted to kinetic models with software such as KinTek Explorer [52] in order to obtain reaction rate constants. In each case, the main requirement is the ability to accurately quantify the amount of substrate and products with methods including densitometric analysis of chromatographic separations, or spectroscopic methods including UV absorbance, fluorescence, or mass spectrometry. A broad range of methods have been used to measure carbohydrate active enzyme (CAZyme) activities and specificities, a sampling of which is listed in Table 4. These methods are described below from the more traditional and lower technology-dependent approaches to the more recently developed approaches that typically rely on state-of-the-art instrumentation. Finally, we describe a novel method for measuring the activities and specificities of ENGases and provide a detailed protocol for this approach.

Table 4.

Summary of analytical methods to study the activity of CAZymes

Analytical method Qualitative Quantitative Pros Cons
SDS-PAGE X X Easy to setup and quick to run
Quantification achieved by densitometric analyses
Kinetic constants can be obtained by fitting to kinetic models such as Michelis Menten		 Low resolution, can resolve removal of whole glycans such as ENGases and PNGaseF
Glycosylation causes smearing of the sample, further hindering resolution
TLC X X Requires use of additional stains
Capillary Electrophoresis X X Glycan identification achieved through online or offline enzymatic or lectin treatment
Can be coupled with MS methods for glycan identification		 Sample preparation requires coupling to fluorophores
Proteomics/glycomics X X Highly accurate quantitation Sample preparation is lengthy and cumbersome
CUPRA-ZYME X X Kinetic constants can be obtained by fitting to kinetic models such as Michaelis Menten Chemical synthesis may be rate limiting step
SEAK and C-SEAK X X Requires prolonged periods of time on the instrument
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4.1. Chromatographic Methods Combined with Densitometry Analysis

4.1.1. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE)

SDS-PAGE is a commonly used technique for protein separation and visualization [53]. The proteins are denatured by the running buffer (Tris/glycine/SDS), surrounding them with negative charges, and are then loaded into a polyacrylamide gel. An applied voltage causes the proteins to migrate through the gel away from the negatively charged cathode to the positively charged anode. The structure of the gel acts like a sieve, allowing smaller proteins to migrate more quickly through the gel. While larger proteins migrate much more slowly resulting in the separation of proteins or glycoproteins based on their size. However, this technique has limited resolution, made worse by heterogeneous glycosylation of a given glycoprotein, which often results in a smear rather than a distinct band on the gel. As a result, its potential application in studying CAZymes is largely limited to those that hydrolyze polysaccharide portions of a large glycan, such as ENGases and PNGaseF.

4.1.2. Thin-Layer Chromatography (TLC)

TLC requires that the samples be loaded onto thin plates of silica, usually with a glass capillary. The spotted silica plates are then placed in a tank containing a small amount of solvent typically consisting of a mixture of n-butanol, acetic acid, and water. The solvent is allowed to migrate up the silica plate, and sugar residues liberated from the glycoprotein will also migrate up the plate along with the solvent based on their polarity relative to the solvent. After the migration is complete, glycans can be visualized using a combination of stains such as diphenylamine-aniline-phosphoric acid and heat [54, 55].

Densitometry-based analyses can be coupled with SDS-PAGE and TLC for quantification of enzyme activity. In both cases, the gel or TLC plate is loaded with the samples of interest and the relevant controls, including proteins of known concentration for SDS-PAGE, or characterized glycans for TLC. The gel or plate can then be imaged and analyzed with software such as the gel analyzer tool within the ImageJ software package [56].

4.1.3. Capillary Electrophoresis

Capillary electrophoresis (CE) is a technique that can be used to separate mixtures of N-glycans based on their charge-to-size ratio [57]. This technique is based on a superimposition of electroosmotic flow and electrophoretic mobility. The electroosmotic flow is based on the flow of the background electrolyte, while the electrophoretic mobility is determined by the size and charge of the glycans. Glycans are injected into a fused capillary tube and an electric field is applied, resulting in the glycans to be attracted to either the anode or cathode. The size of the glycan determines the speed of the transport, with larger glycans moving more quickly. If the glycans are injected into a bare fused capillary tube, the interior surface of the tube is negatively charged, and thus the background electrolyte moves from the negatively charged anode toward the positively charged anode. However, except for N-acetylneuraminic acid, all the other saccharides found in N-glycans are uncharged. This prevents the high-resolution separation of glycans. Additionally, N-glycans do not contain chromophores, meaning they cannot be detected using UV absorbance techniques. To address these issues, a common approach is to label the glycan with a fluorophore (e.g., 8-aminopyrene-1,3,6-trisulfonate; APTS). As most fluorophores are negatively charged, the electric field is applied in reverse polarity, with the negatively charged cathode being placed at the end of the capillary where the glycans are injected. The labeled glycans, which are now negatively charged, migrate toward the positively charged anode, with larger glycans moving more quickly.

To characterize the size of the resolved glycans, two approaches have been developed: (1) a complete glycan ladder can be injected in an additional run; or (2) three internal standards – maltose (DP2), maltotriose (DP3), and maltopentadecaose (DP15) – can be injected with the sample. In the second method, DP2 and DP15 are the smallest and largest glycans in the sample, respectively, and DP3 will run between the samples. This approach allows various glycan sizes to be estimated, providing an estimate of the hydrodynamic size of the glycan.

The glycans resolved with CE can also be partly identified using exoglycosidases or lectins with known specificities. Exoglycosidases remove terminal sugar residues, resulting in a smaller glycan, which results in slower migration. The use of lectins will significantly increase the size of the glycan, making it disappear from the electropherogram. These techniques can be used on- or offline.

Coupling CE with mass spectrometry (MS) methods discussed below can be used to determine the structures of, and even to quantify, glycans. These approaches are termed CE-MS analyses. These methods can either be performed in-line, where the capillary directly feeds into an ionization source for electrospray ionization (ESI) or spots samples for matrix-assisted laser desorption ionization (MALDI), or offline where the samples are manually transferred. The structures of the glycans can be determined either by analysis of the glycans before and after exoglycosidase treatment or by using mass spectrometry-based fragmentation approaches such as collision-induced dissociation (CID) [58, 59].

4.2. Spectroscopic Methods

4.2.1. UV and Fluorescence Spectroscopy

Ultraviolet and fluorescence spectroscopy [60] can be used to investigate the substrate specificity of CAZymes. In these methods, the saccharide substrates are attached to visible compounds such as 4-nitrophenol (4-NP) or fluorescence-active compounds such as 4-methylumbelliferone (4-MeU). These molecules can only be detected once the glycan has been removed. The data can then be fit (e.g., with commercially available software such as GraphPad Prism) to a Michaelis-Menten model and kinetic parameters derived. Substrates such as 4-NP-Man and 4-NP-GlcNAc are commercially available; however, such substrates are only available for a limited number of glycosidic bonds. Compounds not commercially available can be custom made, although cost and effort can be prohibitive.

4.2.2. Mass Spectrometry-Based Approaches

MS is one of the most sensitive techniques available for glycan analyses. The limits of detection are currently in the femtomolar to attomolar range. MS has the added benefit of providing a highly accurate mass, essential for differentiating glycan and non-glycan peaks. In addition to determining the rate constants of the CAZyme, determining the structure of the released glycan is also important for complete characterization of enzymatic specificity.

Various types of ionization sources and mass analyzers have been used to determine the structures and saccharide compositions of glycans. The most used ionization techniques are MALDI and ESI. In MALDI, the samples are suspended in a matrix on a sample plate, which is then vaporized by a laser to produce ions. In ESI, ions are produced by applying a high voltage to a liquid to produce aerosols which contain the ions [61]. These are coupled to mass analyzers such as time of flight (TOF), quadrupole time of flight (QTOF), Fourier-transform ion cyclotron resonance (FT-ICR), and Orbitraps. FT-ICR and Orbitrap analyzers are more accurate compared to the TOF analyzers. MALDI and ESI coupled to time-of-flight analyzers such as a TOF and QTOF are often used for glycoproteomic analyses to provide quantitation and to identify site-specific glycosylation. The high scan rates in TOF and QTOF analyzers enable high-throughput analyses to provide reliable quantitative data. These techniques can be used in bottom-up or top-down approaches. Additionally, these systems can also be coupled to chromatography systems, such as high-performance liquid chromatography (HPLC), often equipped with a C8 or C18 column to separate glycans in solution before mass analysis. In addition to characterizing intact glycans, these methods can also be used to obtain structural information about the glycans through fragmentation methods such as collision-induced dissociation (CID), UV photodissociation (UVPD), electron transfer dissociation (ETD), and electron transfer and higher energy collision dissociation (EThcD) [58, 59].

Bottom-Up Approaches

These approaches have been used to study the substrate specificity of GHs. The process involves analyzing the glycan substrate before and after incubation with the GH of interest. The sample preparation differs slightly depending on whether endoglycosidases or exoglycosidases are being investigated. In the cases of endoglycosidases, the released glycans can be enriched from the reaction mixture using either porous graphitized carbon chromatography (PGC) for unlabeled glycans or labeled via reductive amination with compounds such as 2-aminobenzamide (2-AB), 2-aminobenzoic acid, 2-aminopyridine, 2 aminonaphthalene trisulfonic acid, and 1-aminopyrene-3,6,8-trisulfonic acid [62]. These compounds add a chromophore or fluorophore to the glycan which can aid with detection on UV or fluorescence detectors present on the liquid chromatography system. Derivatization of glycans can be a time-consuming process and requires optimization to ensure selective and complete labeling and may not be amenable to kinetic studies. For exoglycosidases, the process is similar; however, sample preparation may require an extra step to release the glycans from the glycoprotein with enzymes such as PNGaseF or previously characterized endoglycosidases.

Intact Mass Spectrometry Approaches

Intact MS is an alternative method to study the substrate specificity and kinetics of these GHs. As mentioned above, MS has previously been used to study the release of glycans from glycoproteins, previously by tracking the released glycan or saccharide or by trypsinization of the glycoprotein and characterizing the peptides using a proteomics approach. Both methods can be cumbersome and time-consuming.

Competitive Universal Proxy Receptor Assay (CUPRA-ZYME)

CUPRA-ZYME is a recently developed method to study the glycan specificities and activities of CAZymes [63]. In this approach, compounds composed of a ligand for the universal proxy receptor joined via a linker to the oligosaccharides of interest are synthesized. An example of the CUPRA-ligand pairing is streptavidin and biotin. In the assay, the glycan modifying enzyme of interest is incubated with the CUPRA and the bifunctional ligand. The reaction can be analyzed after periods of time or continuously to produce kinetic readouts of a reaction. Intact mass specrometry is used to analyze the reaction mixture looking for the CUPRA bound ligand in its intact and cleaved forms and quantifying these. This approach allows both the substrate specificity of an enzyme to be determined and its kinetic parameters. However, this approach requires the chemical synthesis of the ligand which may be nontrivial and time-consuming.

SEAK and C-SEAK

Many of the methods discussed above can be used to determine the substrate activity and specificity of glycan-modifying enzymes; however, each of them has some drawbacks. SDS-PAGE and TLC have limited resolution, while CE and TLC require the glycan to be isolated separately from the glycoprotein substrate, usually requiring additional time and processing steps. Spectroscopic methods using 4-NP or 4-MeU may require custom synthesis of reagents which may be quite costly. Finally, bottom-up mass spectrometry approaches are time-consuming and require optimization of glycan derivatization processes, and the CUPRA-ZYME approach, although versatile, requires the synthesis of custom bifunctional ligands which is a rate-limiting step (Table 4).

We have developed an MS-based assay to study the activities of ENGases that address some of the drawbacks seen with the other methods. In this method, the ENGase of interest is mixed with the substrate and loaded into a sample vial. The reaction is sampled regularly (e.g., once every 5 min), and the intact glycoprotein products are analyzed by LC-MS until the reaction reaches completion. This approach enables automated data collection and provides information about the reaction over time. As a result, this data can be fitted to kinetic models to obtain reaction rate constants for the enzyme. We have termed this approach specificity of enzymatic activity and kinetics (SEAK). A schematic and examples of this approach are shown in Fig. 3.

Fig. 3.

Fig. 3

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The overall workflow of the SEAK assay; the reaction is set up and loaded into an LC-MS system for automated reaction sampling and subsequent data processing, (a) deconvoluted mass spectra of IgG at various timepoints throughout the reaction showing deglycosylation by EndoS, (b) modeling from KinTek Explorer showing the rates of removal of the first and second glycans from IgG, (c) deconvoluted mass spectra of RNase B at various timepoints throughout the reaction showing the deglycosylation of RNase B by EndoBT-3987, (d) modeling from KinTek Explorer showing the rates of removal of the first and second glycans from RNase B

A similar assay can be used to determine the substrate specificities of ENGases. In this case, a cocktail of glycoproteins, each bearing a different glycan, is mixed and incubated with the enzyme of interest. The reaction is sampled at regular intervals. The mass spectra are deconvoluted, and the masses of the proteins can be used to identify the glycan specificities of the enzyme and the kinetic constants. This approach is versatile and the characteristics of the glycoprotein cocktail can be customized by varying the substrates used in order to suit the question being asked. We refer to this variation of the SEAK assay in which combinations of glycoprotein substrates are used as Competitive-SEAK (C-SEAK). The C-SEAK assay can be used to simulate a simplified biological environment. The primary requirement for whether a glycoprotein can be used in the assay is whether it can be successfully ionized for MS analysis. As an example of C-SEAK to test the specificity of an enzyme, a reaction mixture can be produced by combining four unique glycoprotein substrates: HM-IgG1WT (recombinantly produced Rituximab in the presence of kifunensine (an α-mannosidase inhibitor)) to produce IgG1 bearing HM-glycans, recombinantly produced Fc bearing CT-glycans, commercially available transferrin (non-IgG glycoprotein bearing CT-glycans), and commercially available RNase B (non-IgG glycoprotein bearing HM-glycans). After incubation of the glycoprotein cocktail with enzyme, the reaction mixture can be analyzed by LC-MS. In the case of EndoS2 (which is IgG-specific – it is only capable of removing glycans from full-length IgG antibodies or Fcs – but can hydrolyze both CT and HM glycans), the analyses show that the recombinant HM-IgG1WT and IgG1-FcWT have been deglycosylated, while the other substrates are intact throughout the reaction. In contrast, in the case of EndoBT-3987 (which is an HM-specific ENGase that exhibits no protein specificity), the analyses show that only the two HM-glycosylated substrates – HM-IgG1WT and RNase B – have been deglycosylated. A schematic and examples of this approach are shown in Fig. 4. Although we developed the SEAK and C-SEAK methods to measure the hydrolytic activities and substrate specificities of ENGases, the utility of these methods could potentially be extended to the analysis of transglycosylation reactions by ENGases or of reactions conducted by other CAZymes.

Fig. 4.

Fig. 4

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C-SEAK assay to study the substrate specificity of EndoS2 (a) the overall workflow of the C-SEAK assay, the reaction is set up with multiple substrates and loaded into an LC-MS system for automated reaction sampling and subsequent data processing, (b) deconvoluted mass spectra of HM-IgG1, IgG1-Fc, RNase B, and Transferrin at the beginning and 1 h into the reaction showing mass shifts associated with deglycosylation by EndoS2 and EndoBT-3987, (c) the glycosylated fraction of each of the substrates during incubation with EndoS2, and (d) the glycosylated fraction of each of the substrates during incubation with EndoBT-3987

5. Notes

  1. The LC Binary Pump gradient can be modified to change the sampling rate. They can be changed based on the retention time of the substrate on the column.

  2. These stock concentrations will need to be initially determined for each enzyme-substrate pair with a preliminary endpoint assay.

Acknowledgments

We would like to thank Professor Kenneth A. Johnson for invaluable discussions and advice on the use and application of the KinTek Explorer Software.

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