key: cord-0753993-xeoe3daq
authors: Chen, Shao‐Yung; Clark, David J.; Zhang, Hui
title: High‐Throughput Analyses of Glycans, Glycosites, and Intact Glycopeptides Using C4‐and C18/MAX‐Tips and Liquid Handling System
date: 2021-07-07
journal: Curr Protoc
DOI: 10.1002/cpz1.186
sha: 1d4769d4d82d7e6337f6fc7df26b43bb4149ae1d
doc_id: 753993
cord_uid: xeoe3daq

Protein glycosylation is one of the most common and diverse modifications. Aberrant protein glycosylation has been reported to associate with various diseases. High‐throughput and comprehensive characterization of glycoproteins is crucial for structural and functional studies of altered glycosylation in biological, physiological, and pathological processes. In this protocol, we detail a workflow for comprehensive analyses of intact glycopeptides (IGPs), glycosylation sites, and glycans from N‐linked glycoproteins. By utilizing liquid handling systems, our workflow could enrich IGPs in a high‐throughput manner while reducing sample processing time and human error involved in traditional proteomics sample processing techniques. Together, our workflow enables a high‐throughput enrichment of glycans, glycosites, and intact glycopeptides from complex biological or clinical samples. © 2021 The Authors. Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Enzymatic digestion of glycoproteins using C4‐tips Basic Protocol 2: Intact glycopeptide analysis using C18/MAX‐tips Basic Protocol 3: Glycan and glycosite analysis

Protein glycosylation is one of the most common and diverse post translational modifications, with as many as half of the proteins in the human body reported being glycosylated (Bennun et al., 2016; Khoury, Baliban, & Floudas, 2011) . Aberrant glycosylation has also been reported to relate to different diseases, including various cancer types (An et al., 2006; Hu et al., 2020) , heart disease (Nagai-Okatani & Minamino, 2016) , and rheumatoid arthritis (Ercan et al., 2010) . Protein glycosylation has also been found to play pivotal roles in the pharmacokinetics of monoclonal antibodies and Fc fusion proteins (Higel, Seidl, Sörgel, & Friess, 2016) . Recently, it was reported that afucosylated IgGs could potentially promote the exacerbation of coronavirus disease 2019 response (Larsen et al., 2020) . Deciphering the mystery of protein glycosylation is crucial and necessary to improve our understanding for glycoprotein functions.

For mass spectrometry-based glycoproteomic analysis, investigation of N-linked glycosylation involves multiple steps, including proteolytic digestion, followed by enrichment and subsequent downstream identification of glycopeptides. Various methods have been utilized to characterize N-linked glycosylation on different levels, which can be roughly separated into three categories: (1) glycosite analysis, (2) glycan profiling, or (3) site-specific intact glycopeptide analysis. Glycosite analysis can be achieved using hydrazide chemistry and its automated method (Tian, Zhou, Elliott, Aebersold, & Zhang, 2007; Zhang, Li, Martin, & Aebersold, 2003) , lectin enrichment (Kaji et al., 2003; Zielinska, Gnad, Wiśniewski, & Mann, 2010) , or hydrophilic enrichment (Wada, Tajiri, & Yoshida, 2004) . Glycan profiling can be analyzed on a native glycan level (Fujitani et al., 2013) via sialic acid derivatization or permethylation (Kang, Mechref, Klouckova, & Novotny, 2005) . Site-specific intact glycopeptide analysis can be achieved using strategies such as solid-phase extraction of N-linked glycans and glycosite-containing peptides (Sun et al., 2016) , hydrophilic interaction liquid chromatography (HILIC) enrichment combined with spectral library search (Shu et al., 2020) , and rapid analysis of glycopeptides by permethylation (Shajahan, Supekar, Heiss, Ishihara, & Azadi, 2017) . For intact glycopeptide analysis, a mass spectrometry-based method such as electron-transfer/high-energy collision dissociation (EThcD; Yu et al., 2017) , stepped-normalized collision energy (NCE) during higher-energy collisional dissociation tandem mass spectrometry (HCD MS/MS) fragmentation (Yang, Yang, & Sun, 2018) , intact O-glycopeptide analysis using data-independent acquisition (DIA) mode (Ye, Mao, Clausen, & Vakhrushev, 2019) , or field asymmetric wave ion mobility spectrometry (FAIMS; Creese & Cooper, 2012) have also been reported to be efficient glycopeptide analysis strategies.

Site-specific intact glycopeptide analysis is considered to be the most promising strategy to comprehensively characterize glycoproteins. However, it is challenging to readily analyze intact glycopeptides because enrichment methods, mass spectrometric analysis, and data analysis software still need to be further developed to ensure glycoproteome coverage and correct glycoform annotation. Furthermore, several challenges need to be addressed in order to increase sample processing throughput for large-scale glycoproteomics characterization, including reducing the time span for sample processing, limiting sample loss during sample enrichment procedures (i.e, peptide desalting, sample dry down to reconstitute in appropriate liquid composition), and sample processing reproducibility.

In this protocol, we present a comprehensive intact glycopeptide, glycan, and glycosite analysis workflow that can obtain the glycoproteomic information from standard protein, cell lysates, or even bodily fluids like human urine or serum. For urinary glycoproteomic analysis, a raw urine sample typically needs to undergo sample pre-processing, for instance, ultracentrifugation, ultrafiltration, or buffer exchange, in order to concentrate proteins and remove contaminants from urine for subsequent enzymatic digestion. However, the presented C4-tip and C18/MAX-tip workflows do not need additional sample pre-processing techniques; raw urine can be acidified and directly bound to the C4 resin digested into peptide mixtures. Utilizing a liquid handling system, intact glycopeptides can be enriched in a high-throughput and highly reproducible Figure 1 Workflow for C4-tip and C18/MAX-tip sample processing. Starting at the glycoprotein level, glycoproteins are first bound to C4-tip, which undergo enzymatic digestion. Digested peptides are then loaded onto C18/MAX-tip for intact glycopeptide (IGP) enrichment. (1) Non-glycosylated peptides and (2) intact glycopeptides are sequentially eluted. Subsequently, an aliquot of the IGPs fraction can be subjected to PNGaseF digest. (3) Deglycosylated peptides, and (4) glycans are separated by C18-stage tips. This figure is cited from our previous publication, Chen et al., 2020. manner, while allowing for orthogonal verification of glycosylation site identification via analysis of intact glycopeptides (IGPs) and deglycosylated N-linked glycopeptides (Fig.  1) . The presented enrichment workflow offers the flexibility of analyzing various levels of N-linked glycoproteins (i.e., glycosite, glycans, and intact glycopeptides). Once intact glycopeptides are readily enriched, users can choose to further process the intact glycopeptides by PNGaseF into de-N-glycosylated peptides and glycans. Further separation using reverse-phase C18 resin would allow separation of de-N-glycosylated peptides from glycans. By analysis of (1) intact glycopeptides, (2) de-N-glycosylated peptides, and (3) glycans, users would achieve (1) site-specific intact glycopeptide analysis, (2) glycosite analysis, and (3) glycan profiling altogether by using this single workflow. Being a tip-format, the workflow also offers a high degree of user flexibility, by allowing the users to process samples using a single pipet, a multichannel pipet, or liquid handling systems.

Basic Protocol 1 provides detailed instruction on C4-tip production and on-tip enzymatic digestion of proteins using the C4-tip. Basic Protocol 2 outlines instruction on C18/MAX-tip production and intact glycopeptide enrichment using the C18/MAX-tip. Basic Protocol 3 describes steps for de-N-glycopeptide sample processing by PNGaseF, glycan clean up by a porous graphitic carbon (PGC) NuTip, and also glycan profiling by matrix-assisted laser desorption/ionization (MALDI).

CAUTION: Basic Protocols 1 and 2 describe steps to produce resin-based tips; users are advised to operate in a fume hood or wear a mask to prevent inhalation of resins.

NOTE: Basic Protocols 1 and 2 are designed for users with liquid handling systems; users need to compile a procedure that matches the processing format of their individual liquid handling system (see Commentary, Time Considerations to construct instrument specific procedures). If users prefer to follow this protocol using a pipet or multichannel pipet, still follow Basic Protocols 1 and 2 but maintain the number of aspiration and dispersion amounts, as well as the overall reaction time. 

While reverse-phase separation resins such as C18 have been extensively used for peptide binding and subsequent desalting, the long alkyl chains of C18 would impede downstream protein elution steps. In our previous publication (Clark et al., 2019) , we evaluated the use of reverse-phase resin with shorter alkyl chains (i.e., C4) for use in urinary protein isolation paired with "on-tip" protease digestion. Herein, we describe detailed steps for C4-tip production and "on-tip" protease digestion. Similar to traditional proteomics sample processing, Basic Protocol 1 should be used for enzymatic digestion of protein mixtures into peptides, with the benefits that on-tip digestion can provide: Direct binding of urinary proteins, without doing sample preparation methods such as protein precipitation, ultrafiltration, or analytical ultracentrifugation. Users should skip Basic Protocol 1 and proceed with Basic Protocol 2 if focused on glycopeptide enrichment from peptide mixtures.

NOTE: For C4-tips, we previously evaluated the binding capacity of the C4 resin and it is ∼400 μg of protein material per 30 mg of C4 resin (Clark et al., 2019) . Users are advised to adjust the total amount of C4 resin according to the protein material being processed. 2. Use a Harris Uni-core disposable punch (I.D. 2 mm) to punch 2-mm holes in the 1/16 in. fine sheet and shoot in the D.A.R.T.S tips; make sure the flat side of the filter stays horizontal with the tip. Use a sterilized needle or iron wire to pack the filter firmly into the D.A.R.T.S tips; do not pack filter too tightly because it will affect downstream liquid aspiration and dispersion (Fig. 2) . The resin should stay in the middle during the entire sample process procedure. If either the 5-or 2-mm sheet is not placed horizontally, the resin may leak during the sample processing procedure.

3. Add 300 μl MeOH into Eppendorf tubes that contain C4 resin beads.

4. Vortex thoroughly and inject MeOH/C4 resin mixture into the D.A.R.T.S tips that contain 2-mm filters. Rinse the Eppendorf tube again using 100 μl MeOH and inject into D.A.R.T.S tips to ensure C4 resin is all transferred to the D.A.R.T.S tip.

If using other liquid handling systems, the D.A.R.T.S tip might not be compatible with certain liquid handlers but the same packing method can still be implemented.

Punch 5-mm holes using the Harris Uni-core disposable punch (I.D. 5 mm) in the 1/16 in. fine sheet; the resin should be in between the 2-and 5-mm filters.

6. Use tweezers or a flat shaped rod to push the 5-mm filter firmly against the upper level of the resin.

We recommend marking a specific position on the flat shaped rod to push down the 5-mm filter to the same depth across production of all tips.

7. Allow MeOH to flow through the bottom of the D.A.R.T.S tips naturally.

Steps 1 to 7 describe the production of a single C4-tip. Conditioning C4-tips 8. Take out two racks, add 30 ml 50% ACN + 0.1% TFA, and 30 ml 0.1% TFA, separately.

The volume specification in step 8 is just to ensure C4-tips can aspirate enough liquid, as long as the C4-tip can be submerged to aspirate liquids; the specific volume should not be critical.

9. Use the liquid handling system to aspirate and disperse the C4-tips in 50% ACN + 0.1% TFA for ten cycles.

10. Aspirate and disperse C4-tips in 0.1% TFA for another ten cycles.

For the liquid handling system (i.e., Versette), one cycle (aspiration and dispersion) will take ∼2 min. If using other liquid handling systems or a multichannel pipet, procedures need to be changed accordingly to follow the overall processing time of specific steps.

Chen et al.

Current Protocols Sample loading on C4-tips 11. If the protein samples are dried, reconstitute sample into 300 μl with 1% formic acid.

For C4-tips, we previously evaluated the binding capacity of the C4 resin as ∼400 μg of protein material per 30 mg of C4 resin (Clark et al., 2019) . Users are advised to adjust the total amount of C4 resin according to the protein material being processed.

12. Vortex samples thoroughly.

13. Centrifuge samples at 900 × g for 10 min to remove insoluble proteins and precipitates.

14. Aliquot 300 μl supernatant into the 96-well Deepwell plate; be cautious and do not aliquot precipitates.

15. Acidify samples with 20% formic acid until final concentration of formic acid is 1% to keep pH <3.

If samples are already at pH <3, this step can be skipped.

16. Using the liquid handling system, aspirate and disperse acidified protein solution using C4-tips for 30 cycles.

For the sample loading procedure, we added an extra delay time of 1 min between aspiration and dispersion; therefore, one cycle would be ∼3 min. If using other liquid handling systems, procedures need to be changed accordingly to follow the overall processing time of specific steps.

17. Remove contaminants and inorganic salts using 0.1% TFA; run for ten cycles.

18. Adjust the pH in each of the tips using 50 mM TEAB and run for ten cycles.

For downstream on-tip digestion using a C4-tip, the user must adjust the pH in each tip in order to maintain enzymatic activity, i.e., pH ∼7.6-8.0. If TEAB is not compatible for the user's analysis, we recommend other pH buffers like ammonium bicarbonate.

"On-tip" protein digestion using C4-tip 19. Reduce protein disulfide bonds on bound proteins using 10 mM TCEP and run for twenty cycles.

20. Alkylate reduced cysteine residues on bound proteins using 15 mM IAA in the dark and run for twenty cycles.

IAA must be added in the dark because IAA is light sensitive.

21. Make a digestion buffer consisting of 50 mM TEAB in 30% ACN.

The composition of digestion buffer has been evaluated in our previous publication (Clark et al., 2019) . We have found inclusion of 30% ACN in our digestion buffer resulted in a higher peptide recovery and also a higher trypsin enzyme activity.

(1:40 enzyme/protein) for 30 cycles.

23. Perform subsequent protease digestion on bound proteins by adding trypsin in digestion buffer (1:40 enzyme/protein) for another 120 cycles.

24. Elute digested peptide on a C4-tip using 150 μl 50% ACN + 0.1% TFA and run for ten cycles.

25. Repeat step 24 to ensure higher recovery of digested peptides.

26. Pool solutions from steps 22, 23, 24, and 25 from the same C4-tip.

Chen et al.

Current Protocols 27. Dry down digested peptides using a SpeedVac concentrator.

For SpeedVac processing to dry down the samples, we use the SpeedVac SC210A concentrator with settings of concentrator mode: "ON" and drying rate mode: "Low."

28. Store samples at -20°C until ready to start Basic Protocol 2. 

With the advancement of mass spectrometry, high-throughput sample preparation has been the focus of proteomics (Clark et al., 2019; Fu et al., 2018) , while high-throughput glycoproteomics has been somewhat limited to glycosite analysis using hydrazide chemistry (Berven, Ahmad, Clauser, & Carr, 2010; Chen, Shah, & Zhang, 2013) . Traditionally, intact glycopeptides are enriched by hydrophilic chromatography (Sun et al., 2016; or lectin (Guo et al., 2015; Zhou et al., 2017) , while sialylated glycopeptides are typically enriched by TiO 2 (Kawahara et al., 2018; Palmisano et al., 2010) , strong cationic exchange (SCX; Lewandrowski, Zahedi, Moebius, Walter, & Sickmann, 2007) , immobilized metal affinity chromatography (IMAC) based on charge selection (Hu, Shah, Clark, Ao, & Zhang, 2018) , or periodate-oxidation of sialylated glycopeptides (Halim, Nilsson, Rüetschi, Hesse, & Larson, 2012; Nilsson et al., 2009) . For periodate-oxidation, sialic acid information is lost after de-sialylation and release from hydrazide beads. With the limits described above, we developed a highthroughput intact glycopeptide enrichment platform that would combine two distinct proteomics enrichment strategies (i.e., hydrophilic and hydrophobic) for simultaneous desalting and enrichment of glycopeptides. We've previously compared the enrichment efficiency and specificity of multiple enrichment methods (W. , and reported mixed-mode anionic exchange (MAX) showed higher yield of enrichment and great enrichment specificity. Hence, we further developed our glycopeptide enrichment platform using MAX resin . Basic Protocol 2 provides a step-bystep introduction of glycopeptide enrichment using a combination of hydrophilic and hydrophobic peptide enrichment strategies, i.e., C18 resin plus MAX resin. Utilizing liquid handling systems, users should expect enrichment of intact glycopeptide after ∼6 hr of processing time.

Peptide samples ( 10. Punch 5-mm holes using the Harris Uni-core disposable punch (I.D. 5 mm) in the 1/16 in. fine sheet.

11. Use tweezers or a flat shaped rod to push the 5-mm filter firmly against the upper level of the resin.

12. Allow MeOH to flow through the bottom of the D.A.R.T.S tips naturally.

13. Take out four racks, add 30 ml 100% ACN, 100 mM TAAB, 95% ACN + 1% TFA, and 0.1% TFA, respectively.

Chen et al.

The volume specification is just to ensure C18/MAX-tips can aspirate enough liquid; as long as C18/MAX-tips can be submerged to aspirate liquids, the specific volume should not be critical.

14. Use the Versette liquid handling system to aspirate and disperse the C18/MAX-tips sequentially in 100% ACN, 100 mM TAAB, 95% ACN + 1% TFA, and 0.1% TFA for ten cycles.

We have reported that for a column-based elution, the related position of C18 and MAX resin are optimized if they are stacked with C18 on top (G. Yang et al., 2020) . However, because liquid handling systems would aspirate and disperse in an up-down movement (unlike column-based flow, which generally uses gravity to allow the elution wash to flow naturally). The relative position of C18 and MAX should not be critical. 95% ACN + 1% TFA is the intact glycopeptide sample loading condition which was evaluated in our previous publication (W. . Simultaneously, 95% ACN will Chen et al.

Current Protocols also allow for non-glycosylated peptide to elute from the C18 material, while glycosylated peptide will still be retained on the MAX material via hydrophilic interactions. 

Basic Protocols 1 and 2 describe details for enzymatic digestion of protein mixtures and glycopeptide enrichment from global peptides. Basic Protocol 3 provides step-by-step instruction for N-linked glycan cleavage using PNGaseF and downstream separation of glycans from de-N-glycosylated peptides as well as glycan clean up by a (PGC) NuTip. Following Basic Protocol 3, users can expect to receive (1) glycan information and (2) glycosite information by analyzing glycans and de-N-glycosylated peptides, separately. When comparing the intact glycopeptide results with glycosite results, a user can achieve an orthogonal verification of glycosylation sites by observing the 0.98 Da mass shift after deamidation of asparagine (N) into aspartic acid (D).

Acetonitrile (ACN), Optima ® LC/MS (Thermo Fisher Scientific, cat. no. A955-4) Formic Acid (FA), Optima ® LC/MS (Thermo Fisher Scientific, cat. no. A117-50) Triethylammonium bicarbonate (TEAB; MilliporeSigma, cat. no. T7408-500ML) Peptide-N-glycosidase F (PNGase F; New England BioLabs, cat. no. P0704L) NuTip Carbon (Hypercarb), large 10-200 μl (Glygen, cat. no 

Current Protocols 3. Aliquot 20 μl out into a new 96-well Deepwell plate.

The purpose of this step is to preserve half of the samples as intact glycopeptides for intact-glycopeptide analysis. If users are not interested in intact-glycopeptide analysis, this step can be omitted. However, for the following steps, reaction volumes will have to be doubled.

According to protocols provided by New England BioLabs, 1 μl of PNGaseF contains 500 units of PNGaseF enzyme.

5. Add the PNGaseF/TEAB mixture in the original Deepwell plate that contains 20 μl of intact-glycopeptide sample.

6. Let react at room temperature overnight using a shaker.

We recommend this reaction be performed in the original 96-well Deepwell plate to save time and minimize sample loss during multiple sample transfer procedures.

There are multiple C18 stage tip protocols and manufacturers available to users. We pack our own C18 stage tips using a solid phase extraction disk (CDS Empore TM ).

7. Condition C18 stage tips with 100 μl MeOH, twice. Centrifuge at 1,500 rpm (211 × g) for 2 min.

8. Centrifuge 2 min at 1,800 rpm (304 × g) with 100 μl 50% ACN + 0.1% FA, twice.

9. Centrifuge 2 min under 1,800 rpm (304 × g) with 100 μl 0.1% FA, twice.

10. Load glycan/de-N-glycopeptide samples and centrifuge for 5 min at 1,000 rpm (94 × g), twice.

11. Wash for 2 min by centrifuging at 1,800 rpm (304 × g) with 100 μl 0.1% FA, twice.

12. Collect flow through from steps 10 and 11; this will be the glycan sample.

13. Elute de-N-glycopeptides using 75 μl 50% ACN + 0.1% FA, twice.

The specific parameters (time and spin rate) during stage tip procedures will vary according to the centrifuge and the tightness of the C18 packing material being packed. We highly recommend that users test out specific parameters prior to operating. We recommend not increasing the spin rate but increase the time of centrifuging, if parameters needed to be optimized. 

Recently, the glycoproteomics field has drastically advanced, from initial discovery of several hundred intact glycopeptides to the detection of >20,000 unique intact glycopeptides (Shu et al., 2020) . The mass spectrometry (MS) instrumentation, sample preparation techniques, and data analysis software for intact glycopeptide analysis are all coming of age, which have made characterization of large-scale cohorts possible. With isobaric tags like isobaric tag for relative and absolute quantitation (iTRAQ), tandem mass tag (TMT), or downstream intact glycopeptide analysis software like GPQuest, Byonic, or MAGIC emerging, the rate-limiting step seems to shift towards the sample preparation aspect. Therefore, implementation of high-throughput sample processing for intact glycopeptide is a must. Over the years, high-throughput glycoproteomics has been somewhat limited to glycosite analysis using hydrazide chemistry (Berven et al., 2010; Chen et al., 2013) or glycan profiling (Yang, Clark, Liu, Li, & Zhang, 2017) . This protocol introduces a comprehensive workflow by combining two of our previous publications Clark et al., 2019) , such that users can process protein samples of intact glycopeptides in a high-throughput manner. This protocol can offer rapid sample processing, while still retaining highly reproducible characteristics. Users can expect to obtain intact glycopeptide samples starting from protein within 3 days. The C4-tip and C18/MAX-tip can also be utilized for standard protein, bodily fluids, cell lysates, or tissue samples.

To successfully enrich glycopeptide samples, users must make sure of the following: During the sample binding procedure, users must be mindful that the pH of the binding condition is <3 to ensure optimum binding for both C4 and C18/MAX. When processing bodily fluids or tissue samples, it is Comparison of (A) non-glycosylated peptide, (B) de-N-glycopeptide, and (C) intact glycopeptide for standard protein fetuin, peptide: VVHAVEVALATFNAESN#GSYLQLVEISR. The site of glycosylation and Asn to Asp transition are indicated by #. Glycosylation site can be determined by observing +0.98 Da shift after PNGaseF digest, as seen in y12 + ion of these spectrums. Intactglycopeptide spectrum can provide orthogonal verification of a glycosylation site, while further providing glycan information. This figure is cited from our previous publication Chen et al., 2020. critical to do a rigorous centrifuge step-3,000 rpm (900 × g) for 10 min-to remove any sediments that would impede downstream sample processing. It is also crucial to consider the evaporation of organic solvents during the liquid handling system operation, especially during the intact-glycopeptide enrichment steps (when 95% ACN + 1% TFA is used). Do not let high volumes of organic solvents remain excessively long in the liquid handling system; this can be avoided by us-ing fresh eluant multiple times during Basic Protocol 2, steps 22 to 25. (For example, by using 3 × 5 cycles during these steps, organic solvents will evaporate more than if eluted in 1 × 15 cycles, thus users are encouraged to use 3 × 5 cycles instead of 1 × 15 cycles.) Furthermore, liquid will evaporate at a faster rate due to the temperature rising during the liquid handling system operation; this can be avoided by attaching a heater-cooler thermoblock inside the liquid handling system.

Current Protocols

Glycan, glycosite, and intact-glycopeptide analysis on the standard protein fetuin can be used for orthogonal verification of a glycosylation site using the C4-tip and C18/MAX-tip workflow.

To demonstrate the feasibility of the C4tip and C18/MAX-tip workflow, we first used the standard protein fetuin, started from the protein level, to non-glycosylated peptide, intact glycopeptide, glycan, and de-Nglycopeptide. After PNGaseF digest of Nlinked glycans, users can expect a deamidation change from asparagine (N) to aspartic acid (D). By comparison of de-N-glycopeptide with non-glycosylated peptide, the glycosylation site can be further verified via comparison of an intact-glycopeptide spectrum and a de-glycopeptide spectrum (Fig. 3) .

Urine is an attractive sample source for urological diseases because of its proximity, sample availability, and non-invasiveness during sample collection. However, urine also contains several components that will confound proteomic analysis, such as a high concentration of urea, inorganic salts, or other biomolecules. Unlike traditional urine proteomics sample processing, use of C4 resin does not need additional sample preprocessing, such as protein precipitation, buffer exchange, ultrafiltration, or ultracentrifugation. The above-mentioned sample preprocess methodologies cannot be readily adapted for high-throughput urinary protein sample preparation; C4 resin, on the other hand, can be adapted into liquid handling systems with ease.

Besides standard protein, the C4-tip and C18/MAX-tip workflow can also be used for human bodily fluids, such as human serum, blood, or urine. Similar to fetuin, the user can expect to obtain non-glycosylated peptides and intact glycopeptides by using the C4tip and C18/MAX-tip workflow, while further PNGaseF digestion could provide orthogonal verification of glycosylation sites.

However, since normal urine protein concentration can vary from 0 to 14 mg/dL, one might expect the peptide or intact glycopeptide identification rate to vary according to the initial urine protein concentration.

The C4-tip (Basic Protocol 1) can be completed in ∼15 hr, which includes 1 hr of C4-tip conditioning, 1.5 hr of sample loading, 1 hr of sample washing, 1 hr of reduction using TCEP, 1 hr of alkylation using IAA: Take out the tips that contain the bound proteins from the liquid handling systems, aliquot ∼200 μl of 50 mM TEAB on top of each C4-tip, seal the C4-tips that contain the bound proteins using Parafilm, and store at 4°C. TEAB and IAA can be dispersed the next morning right before enzymatic digestion.

The C18/MAX (Basic Protocol 2) can be completed in ∼6 hr, which includes 2 hr of C18/MAX-tip conditioning, 1 hr of sample loading, 1 hr of sample washing and desalting, 0.5 hr of non-glycosylated peptide elution, and 0.5 hr of intact glycopeptide elution.

Basic Protocol 3 can be achieved in ∼15 hr, which includes 12 hr of overnight PNGaseF digestion, 2 hr of C18 stage tips procedure for glycan/de-N-glycopeptide separation, and 1 hr of glycan purification.

Basic Protocols 1, 2, and 3 all include multiple procedures to dry samples using a Speed-Vac concentrator. These procedures are not included in the time consideration calculations because different reaction volume, instrumentation, and sample size can contribute to the overall processing time.

Systems glycobiology: Integrating glycogenomics, glycoproteomics, glycomics, and other 'omics data sets to characterize cellular glycosylation processes

Optimizing performance of glycopeptide capture for plasma proteomics

Solid phase extraction of N-linked glycopeptides using hydrazide tip

Glycans, glycosite, and intact glycopeptide analysis of Nlinked glycoproteins using liquid handling systems

Simple tip-based sample processing method for urinary proteomic analysis

Separation and identification of isomeric glycopeptides by high field asymmetric waveform ion mobility spectrometry

Aberrant IgG galactosylation precedes disease onset, correlates with disease activity, and is prevalent in autoantibodies in rheumatoid arthritis

Highly reproducible automated proteomics sample preparation workflow for quantitative mass spectrometry

Total cellular glycomics allows characterizing cells and streamlining the discovery process for cellular biomarkers

Differential urinary glycoproteome analysis of type 2 diabetic nephropathy using 2D-LC-MS/MS and iTRAQ quantification

Human urinary glycoproteomics; attachment site specific analysis of Nand O-linked glycosylations by CID and ECD

N-glycosylation heterogeneity and the influence on structure, function and pharmacokinetics of monoclonal antibodies and Fc fusion proteins

Integrated proteomic and glycoproteomic characterization of human high-grade serous ovarian carcinoma

Reanalysis of global proteomic and phosphoproteomic data identified a large number of glycopeptides

Lectin affinity capture, isotope-coded tagging and mass spectrometry to identify N-linked glycoproteins

Solid-phase permethylation of glycans for mass spectrometric analysis

Distinct urinary glycoprotein signatures in prostate cancer patients

Proteome-wide post-translational modification statistics: Frequency analysis and curation of the swiss-prot database

Afucosylated IgG characterizes enveloped viral responses and correlates with COVID-19 severity

Enhanced Nglycosylation site analysis of sialoglycopeptides by strong cation exchange prefractionation applied to platelet plasma membranes

Aberrant glycosylation in the left ventricle and plasma of rats with cardiac hypertrophy and heart failure

Enrichment of glycopeptides for glycan structure and attachment site identification

Selective enrichment of sialic acidcontaining glycopeptides using titanium dioxide chromatography with analysis by HILIC and mass spectrometry

Mass spectrometric analysis of sialylated glycans with use of solid-phase labeling of sialic acids

Tool for rapid analysis of glycopeptide by permethylation via onepot site mapping and glycan analysis

Large-scale identification of n-linked intact glycopeptides in human serum using HILIC enrichment and spectral library search

Comprehensive analysis of protein glycosylation by solid-phase extraction of N-linked glycans and glycosite-containing peptides

Solid-phase extraction of Nlinked glycopeptides

Hydrophilic affinity isolation and MALDI multiple-stage tandem mass spectrometry of glycopeptides for glycoproteomics

One-step enrichment of intact glycopeptides from glycoengineered chinese hamster ovary cells. Frontiers in Chemistry

Characterization of glycopeptides using a stepped higherenergy C-trap dissociation approach on a hybrid quadrupole orbitrap

High-throughput analysis of N-glycans using AutoTip via glycoprotein immobilization

Comparison of enrichment methods for intact N-and O-linked glycopeptides using strong anion exchange and hydrophilic interaction liquid chromatography

Glyco-DIA: A method for quantitative O-glycoproteomics with in silico-boosted glycopeptide libraries

Electrontransfer/higher-energy collision dissociation (EThcD)-enabled intact glycopeptide/glycoproteome characterization

Identification and quantification of Nlinked glycoproteins using hydrazide chemistry, stable isotope labeling and mass spectrometry

Site-specific fucosylation analysis identifying glycoproteins associated with aggressive prostate cancer cell lines using tandem affinity enrichments of intact glycopeptides followed by mass spectrometry, Analytical Chemistry

Precision mapping of an in vivo N-glycoproteome reveals rigid topological and sequence constraints

This work was supported in part by grants from the National Institutes of Health, National Cancer Institute, the Early Detection Research Network (EDRN, U01CA152813), and the Clinical Proteomic Tumor Analysis Consortium (CPTAC, U24CA210985).

Shao-Yung Chen: Conceptualization, investigation, methodology, project administration, writing: original draft, David Clark: Conceptualization, methodology, Hui Zhang: Conceptualization, funding acquisition, resources, supervision

The authors declare no competing financial interest.

Data sharing is not applicable to this article as no new data were created or analyzed in this study.

An, H. J., Miyamoto, S., Lancaster, K. S., Kirmiz, C., Li, B., Lam, K. S., … Lebrilla, C. B. (2006) . Profiling of glycans in serum for the discovery of potential biomarkers for ovarian Chen et al.

Current Protocols