Targeted Identification of Glycosylated Proteins in the Gastric Pathogen Helicobacter pylori (Hp)

Abstract

Virulence of the gastric pathogen Helicobacter pylori (Hp) is directly linked to the pathogen's ability to glycosylate proteins; for example, Hp flagellin proteins are heavily glycosylated with the unusual nine-carbon sugar pseudaminic acid, and this modification is absolutely essential for Hp to synthesize functional flagella and colonize the host's stomach. Although Hp's glycans are linked to pathogenesis, Hp's glycome remains poorly understood; only the two flagellin glycoproteins have been firmly characterized in Hp. Evidence from our laboratory suggests that Hp synthesizes a large number of as-yet unidentified glycoproteins. Here we set out to discover Hp's glycoproteins by coupling glycan metabolic labeling with mass spectrometry analysis. An assessment of the subcellular distribution of azide-labeled proteins by Western blot analysis indicated that glycoproteins are present throughout Hp and may therefore serve diverse functions. To identify these species, Hp's azide-labeled glycoproteins were tagged via Staudinger ligation, enriched by tandem affinity chromatography, and analyzed by multidimensional protein identification technology. Direct comparison of enriched azide-labeled glycoproteins with a mock-enriched control by both SDS-PAGE and mass spectrometry-based analyses confirmed the selective enrichment of azide-labeled glycoproteins. We identified 125 candidate glycoproteins with diverse biological functions, including those linked with pathogenesis. Mass spectrometry analyses of enriched azide-labeled glycoproteins before and after cleavage of O-linked glycans revealed the presence of Staudinger ligation-glycan adducts in samples only after beta-elimination, confirming the synthesis of O-linked glycoproteins in Hp. Finally, the secreted colonization factors urease alpha and urease beta were biochemically validated as glycosylated proteins via Western blot analysis as well as by mass spectrometry analysis of cleaved glycan products. These data set the stage for the development of glycosylation-based therapeutic strategies, such as new vaccines based on natively glycosylated Hp proteins, to eradicate Hp infection. Broadly, this report validates metabolic labeling as an effective and efficient approach for the identification of bacterial glycoproteins.

Helicobacter pylori (Hp)¹ infection poses a significant health risk to humans worldwide. The Gram-negative, pathogenic bacterium Hp colonizes the gastric tract of more than 50% of humans (1). Approximately 15% of infected individuals develop duodenal ulcers and 1% of infected individuals develop gastric cancer (2). Current treatment to clear infection requires “triple therapy” (3), a combination of multiple antibiotics that is often associated with negative side effects (4). Because of poor patient compliance and the evolution of antibiotic resistance, existing antibiotics are no longer effective at eradicating Hp infection (4). New treatment methods are needed to eliminate Hp from the human gastric tract.

Recent work has focused on gaining insights into the pathogenesis of Hp to aid the development of new treatments. The most recent findings in this area have conclusively revealed that glycosylation of proteins in Hp is required for pathogenesis. Hp use complex flagella, comprised of flagellin proteins, to navigate the host's gastric mucosa (5, 6). The flagellin proteins are heavily glycosylated with the unusual nine-carbon sugar pseudaminic acid, found exclusively in mucosal-associated pathogens (Hp (7), Campylobacter jejuni (8) and Pseudomonas aeruginosa (9)). This modification is absolutely essential for the formation of functional flagella on Hp (7, 10). Deletion of any one of the enzymes in the pseudaminic acid biosynthetic pathway results in Hp that lack flagella, are nonmotile, and are unable to colonize the host's stomach (7). Although pseudaminic acid is critical for Hp virulence, it is absent from humans (11, 12). Therefore, insights into Hp's pathogenesis have revealed that Hp's glycan pseudaminic acid is a bona fide target of therapeutic intervention. This is one of a number of examples linking protein glycosylation to virulence in medically significant bacterial pathogens (13, 14).

Despite these findings, Hp's glycome remains poorly understood overall. Only the two flagellin glycoproteins have been firmly characterized in Hp (7) to date. Nine other candidate glycoproteins have been identified in Hp, but their glycosylation status has not been biochemically confirmed (15). The relative paucity of information regarding Hp's glycoproteins is due in part to the previously held belief that protein glycosylation could not occur in bacteria (13, 16, 17). However, even after Szymanski (18, 19), Koomey (20), Guerry (21), Logan (7), Comstock and others (13, 16, 17) disproved this belief by firmly establishing the synthesis of glycoproteins in bacteria, the study of bacterial glycoproteins has presented unique challenges for analytical study (14, 22). For example, the unusual structures of bacterial glycans, which often contain amino- and deoxy-carbohydrates exclusively found in bacteria (12, 23–25), hampers their identification using existing tools. Though methods such as the use of glycan-binding reagents (20, 24, 26, 27) and periodic acid/hydrazide glycan labeling (15) have successfully detected glycoproteins in a range of bacteria, they present limitations. Glycan binding-based methods are often limited because of the unavailability of lectins or antibodies with binding specificity for glycosylated proteins in the bacteria of interest (14, 22). Periodic acid/hydrazide-based labeling is plagued by a lack of specificity for glycosylated proteins (15). Thus, an efficient and robust approach to discover Hp's glycoproteins is needed.

In previous work, we established that the chemical technique known as metabolic oligosaccharide engineering (MOE), which was developed by Bertozzi (28, 29), Reutter (30), and others for the study of mammalian glycoproteins, is a powerful approach to label and detect Hp's glycoproteins (31). Briefly, Hp metabolically processes the unnatural, azide-containing sugar peracetylated N-azidoacetylglucosamine (Ac₄GlcNAz) (32), an analog of the common metabolic precursor N-acetylglucosamine (GlcNAc), into cellular glycoproteins (Fig. 1). Elaboration of azide-labeled glycoproteins via Staudinger ligation (33) with a phosphine probe conjugated to a FLAG peptide (Phos-FLAG) (34) followed by visualization with an anti-FLAG antibody (Fig. 1) revealed a glycoprotein fingerprint containing a large number of as-yet unidentified Hp glycoproteins that merit further investigation (31).

Fig. 1.

Metabolic oligosaccharide engineering facilitates labeling and detection of Hp's glycoproteins. Supplementation of Hp with Ac₄GlcNAz leads to metabolic labeling of Hp's N-linked and O-linked glycoproteins with azides. Azide-modified glycoproteins covalently labeled with Phos-FLAG can be detected via Western blot analysis with anti-FLAG antibody to yield Hp's glycoprotein fingerprint, which contains a large number of as-yet unidentified glycoproteins.

Here we describe a glycoproteomic identification strategy for the selective detection, isolation, and discovery of Hp's glycoproteins. In particular, we demonstrate that glycan metabolic labeling coupled with mass spectrometry analysis is an efficient and robust chemical approach to identify novel glycoproteins in Hp. This work characterizes glycosylated virulence factors in Hp, thus opening the door to new vaccination and antibiotic therapies to eradicate Hp infection. Broadly, this work validates metabolic oligosaccharide engineering as a complementary method to discover bacterial glycoproteins.

EXPERIMENTAL PROCEDURES

Materials

Protease inhibitor mixture, antibiotics, anti-FLAG antibodies and anti-FLAG agarose were purchased from Sigma (St. Louis, MO). Sequencing grade trypsin was from Promega (Madison, WI), and fetal bovine serum (FBS) was from Invitrogen (Carlsbad, CA). Nitrocellulose paper, Ni-NTA resin, zinc stain, and 4–16% Ready Gel Tris-HCl gels were from BioRad (Hercules, CA). GlcNAz, Ac₄GlcNAc, Ac₄GlcNAz, Phos-FLAG (Phos-DYKDDDDK) and Phos-FLAG-His₆ (Phos-DYKDDDDKHHHHHH) were prepared as previously described (29). Hp strain 26695 was provided by Dr. Manuel Amieva (Stanford, CA). Azidoacetyl ethyl ester was a gift from Dr. Jennifer Prescher (Irvine, CA).

Metabolic Labeling of Hp

Hp strain 26695 was grown on horse blood agar plates for 3–4 days in a microaerophilic environment (14% CO₂) at 37 °C. The bacteria were then transferred to Brucella broth containing 10% FBS, 6 μg/ml vancomycin, and 1 mm peracetylated N-acetylglucosamine (Ac₄GlcNAc) or Ac₄GlcNAz, as previously described (31). Alternatively, the bacteria were transferred to Brucella broth containing 10% FBS and 6 μg/ml vancomycin supplemented with 5 mm sodium azidoacetate or 5 mm azidoacetyl ethyl ester. Liquid cultures were grown for 3–5 days in a microaerophilic environment with gentle rocking at 37 °C, then harvested as described below.

Harvesting Hp to Probe for Total Cellular Azide-Labeled Glycoproteins

Hp grown in the presence of Ac₄GlcNAc, Ac₄GlcNAz, sodium azidoacetate, or azidoacetyl ethyl ester were centrifuged at 3500 rpm using a Sorvall Legend RT⁺ centrifuge (Thermo Scientific) and washed two times with PBS. The cells were resuspended in ice cold lysis buffer (20 mm Tris-HCl, pH 7.4; 1% Igepal; 150 mm NaCl; 1 mm EDTA) containing protease inhibitors (protease inhibitor mixture, Sigma) for 5 mins at room temperature. Protein concentration of the samples was measured using BioRad's D_C protein concentration assay (BioRad) per manufacturer's instructions. All samples were standardized to 3.0 mg/ml. To probe for azide incorporation, the lysates were diluted 1:1 with 500 μm Phos-FLAG (34), reacted overnight at room temperature, and then analyzed via Western blot with anti-FLAG antibody (Sigma) or via SDS-PAGE with zinc stain (BioRad).

Subcellular Fractionation of Glycoproteins

Hp grown in the presence of Ac₄GlcNAc or Ac₄GlcNAz were harvested from Brucella broth by centrifugation at 6000 × g, and the conditioned medium was set aside to access secreted proteins. The harvested cells were fractionated as described by Hoshino et al. to obtain periplasmic, inner membrane-associated, and cytoplasmic protein fractions (35). Secreted proteins were isolated from the conditioned Brucella broth via TCA-precipitation as described by Bumann et al. (36). An acid-glycine extraction was performed as described previously to isolate Hp's surface-associated proteins (37). An N-lauroylsarcosine extraction was performed as reported by Hopf et al. to access outer-membrane Hp proteins (15, 38). The protein concentration of subcellular fractions was standardized to 3.0 mg/ml using a BioRad D_C Protein Assay (BioRad). Standardized samples were incubated 1:1 with 500 μm Phos-FLAG (34) overnight at room temperature to label azide-containing glycoproteins, and then analyzed by Western blot analysis with anti-FLAG antibody (Sigma) or via SDS-PAGE with zinc stain (BioRad). In addition, standardized periplasmic and cytoplasmic samples were assayed for malate dehydrogenase (35) activity alongside a positive control (malate dehydrogenase, Sigma) according to the manufacturer's (Sigma's) instructions to confirm the efficacy of subcellular fractionation.

Preparation and Analysis of Enriched Azide-Labeled Glycoproteins

Hp metabolically labeled with 1 mm Ac₄GlcNAc or Ac₄GlcNAz were centrifuged at 3000 × g and washed two times with phosphate buffered saline (PBS). The cells were lysed according to Koenigs et al. and centrifuged at 3000 × g for 10 min to remove insoluble debris (31). Following the protocol by Laughlin et al., Phos-FLAG-His₆ was added as a solid directly to the lysate at a final concentration of 500 μm, and Staudinger ligation was run at room temperature for 24 h under argon (29, 39). The unreacted Phos-FLAG-His₆ was removed by Bio-Rad P-10 size-exclusion column according to the manufacturer's protocol. Flow-through fractions with A₂₈₀ > 0.050 were combined and concentrated using a centrifugal filter device with 30 kDa molecular weight cutoff (Millipore). FLAG-His₆-tagged glycoproteins were enriched using 2 ml of α-FLAG agarose beads (Sigma) followed by 5 ml of nickel-nitrilotriacetic acid (Ni-NTA)-agarose resin (Bio-Rad), as previously described (29, 39). After rinsing away nonbinders, the bound glycoproteins were eluted from the nickel column with one column volume of a solution of 8 m urea, 0.1 m Na₂HPO₄, 10 mm Tris (pH 8.0), and 250 mm imidazole. Samples containing 15 μg of material from Ac₄GlcNAc and Ac₄GlcNAz-treated Hp both before purification (input) and after purification (eluent) were analyzed by SDS-PAGE to confirm successful enrichment. In addition, 150 μg of Hp's enriched azide-labeled glycoproteins were suspended in 10 mm Tris (pH 8) and focused on 11 cm pH 3–10 nonlinear IPG strips (GE Healthcare) using GE Healthcare's Multiphor^™ II system. These strips were reduced, alkylated and loaded onto 8–18% polyacrylamide gels (GE Healthcare), which were run at 10 °C and stained with zinc stain (BioRad).

Protein Identification by MudPIT

An azide-labeled enriched glycoprotein sample containing 150 μg of protein and a mock enriched control (which was labeled with Ac₄GlcNAc, reacted with Phos-FLAG-His₆, and enriched via anti-FLAG and Ni-NTA affinity chromatography, as described above) were concentrated using a Microcon centrifugal filter device (Millipore) to ∼10 μl, suspended in 8 m urea and 100 mm Tris-HCl (pH 8.5), reduced with 5 mm tris(2-carboxyethyl)phosphine (TCEP), and reacted with 10 mm iodoacetamide. The samples were diluted by a factor of four with 100 mm Tris-HCl (pH 8.5) and 1 mm CaCl₂, and then digested with 0.25 μg of trypsin (Promega) overnight at 37 °C. The digests were mixed with formic acid to a final concentration of 5%, desalted using C₁₈ Spec tips (Varian), fully dried by speed vacuum at room temperature, and analyzed by multidimensional protein identification technology (mudPIT) (40). The Vincent J. Coates Proteomics/Mass Spectrometry Laboratory at University of California Berkeley performed mass spectrometry analyses of the enriched, azide-labeled sample and mock-enriched control sample. A nano LC column was packed in a 100 μm inner diameter glass capillary with an emitter tip. The column consisted of 10 cm of Polaris C18 5 μm packing material (Varian), followed by 4 cm of Partisphere 5 SCX (Whatman). The column was loaded by use of a pressure bomb and washed extensively with buffer A (see below). The column was then directly coupled to an electrospray ionization source mounted on a Thermo LTQ XL linear ion trap mass spectrometer. An Agilent 1200 HPLC equipped with a split line so as to deliver a flow rate of 30 nl/min was used for chromatography. Peptides were eluted using an 8-step MudPIT procedure (40). Buffer A was 5% acetonitrile/0.02% heptaflurobutyric acid (HBFA); buffer B was 80% acetonitrile/0.02% HBFA. Buffer C was 250 mm ammonium acetate/5% acetonitrile/0.02% HBFA; buffer D was same as buffer C, but with 500 mm ammonium acetate.

The programs SEQUEST, as embodied in BIOWORKS BROWSER (version 3.3.1 SP1), (41) and DTASELECT (version 1.9) (42) were used to identify peptides and proteins from a database consisting of all proteins (1560 total) from Hp 26695 plus a collection of 160 common contaminants (1720 protein entries in the database actually searched). The Hp database was the newest available from NCBI RefSeq on October 7, 2010. Search parameters were altered to DTA generation 600.00–4000.00 with an absolute threshold of 500. The search was restricted to tryptic peptides and allowed three missed cleavages. Carboxyamidomethylation of cysteines as a fixed modification and oxidation of methionines as a variable modification were specified. The mass tolerance for precursor ions was 1.4 and for fragment ions was 1.0. Individual peptides were accepted with Xcorr of 1.8 for charge state +1, 2.2 for charge state +2 and 3.5 for charge state +3. These values have given < 1% false positives over many data sets. Proteins were further required to have at least two peptides meeting acceptance criteria. Data was considered high confidence if assigned spectra were present in the Ac₄GlcNAz samples but absent from the mock-enriched Ac₄GlcNAc control samples.

Prediction of Subcellular Localization

Initial predictions of the subcellular localization of Hp proteins identified by mudPIT were based on literature reports, where available. Because of the shortage of subcellular location information for hypothetical proteins and functionally uncharacterized proteins, each identified protein was also subjected to PSORTb v.3.0 to predict subcellular location (43).

Accessing Synthetic Staudinger Ligation-Glycan Standards

Staudinger ligation was performed overnight at 37 °C by mixing 1:1 solutions of GlcNAz (100 mm in H₂O) and Phos-FLAG-His₆ (100 mm in PBS) to produce GlcNAz-Phos-FLAG-His₆. Similarly, overnight incubation at 37 °C of solutions of GlcNAz (100 mm in H₂O) and Phos-FLAG-His₆ (500 μm in PBS) mixed 1:1 produced GlcNAz-Phos-FLAG-His₆. Successful synthesis was confirmed by high-resolution mass spectrometry analysis as described below (see HPLC-Chip/Q-TOF mass spectrometric analysis).

Phos-FLAG Labeling to Produce “Tagged Lysates”

Hp metabolically labeled with 1 mm Ac₄GlcNAc or Ac₄GlcNAz were centrifuged at 3000 × g and washed two times with phosphate buffered saline (PBS). The cells were lysed according to Koenigs et al. (31). Lysates were rinsed extensively with PBS using a Microcon centrifugal filter device (Millipore, 10000 MWCO) to remove residual Ac₄GlcNAc or Ac₄GlcNAz. Lysates were reacted with 250 μm Phos-FLAG at room temperature overnight. The unreacted Phos-FLAG was removed by extensive rinsing with ddH₂O and passage of reacted lysates through a Microcon centrifugal filter device (Millipore, 10000 MWCO) to yield samples of “tagged lysates.” These lysates were then subjected to beta-elimination, as described below.

Biochemical Purification of Azide-Labeled Urease

Urease was purified from Hp strain 26695 using ion exchange chromatography (IEC) and size exclusion chromatography (SEC) based on previously described purification schemes by Hu and Mobley (44, 45) and Rokita et al. (46). A 250 ml culture of Hp metabolically labeled with 1 mm Ac₄GlcNAz was inoculated and grown as described above. Cells were harvested by centrifugation at 10,000 × g and the resultant pellet was frozen at −20 °C. The cells were re-suspended in ice-cold buffer (20 mm sodium phosphate, pH 6.9) containing protease inhibitors (protease inhibitor mixture, Sigma) and 10 μg/ml DNase and then sonicated to achieve complete cell lysis. Concentrated stock solutions of EDTA and β-mercaptoethanol (βME) were added to the lysis solution to make it consistent with the initial IEC buffer (20 mm sodium phosphate, pH 6.9, 1 mm EDTA, 1 mm βME). Cellular debris was removed from the sample through centrifugation at 14,000 × g. The supernatant was loaded onto a Q-Sepharose Fast Flow column (25 ml, 3 cm diameter, GE Healthcare) that had been pre-equilibrated with the above IEC buffer. Urease was eluted from the column at ∼150 mm NaCl using a linear gradient from 0 m to 0.5 m NaCl in IEC buffer. The presence of urease in the elution fractions was detected qualitatively using a phenol-red colorimetric assay (44). Fractions that exhibited urease activity were further analyzed by SDS-PAGE with zinc-stain (BioRad) and via Western with anti-ureA and anti-ureB antibodies (Santa Cruz Biotechnology) to verify the presence of ureA and ureB. Fractions with the most urease and fewest contaminants were pooled for subsequent purification by SEC. The pooled fractions were exchanged into the SEC buffer (20 mm sodium phosphate, pH 6.9, 150 mm NaCl, 1 mm EDTA, 1 mm βME) using a 9000 Da MWCO filter (Pierce) and concentrated to a final volume of 7 ml. This concentrated sample was loaded onto a pre-equilibrated Sepharose CL-6B (GE Healthcare) SEC column (200 ml total volume, 3 cm diameter) and eluted at 1 ml/min. Fractions containing urease were identified using the phenol-red assay and subsequently analyzed by SDS-PAGE. The purest urease-containing fractions (see supplemental Fig. S7) were pooled, concentrated to 0.5 mg/ml via ultrafiltration (Millipore, 100,000 MWCO), and mixed 1:1 with 500 μm Phos-FLAG at room temperature overnight. The unreacted Phos-FLAG was removed by extensive rinsing with 50% MeOH in ddH₂O, 15% acetonitrile in ddH₂O, and 100% ddH₂O over a Microcon centrifugal filter device (Millipore, 10000 MWCO) to yield samples of “tagged urease.” This sample was then subjected to beta-elimination, as described below.

Beta-Elimination of Hp's O-Linked Glycans

Hp's enriched azide-labeled glycoproteins, Phos-FLAG-reacted Ac and Az lysates (“tagged lysates”), and purified azide-labeled urease (“tagged urease”) were subjected to beta-elimination to remove glycans overnight at 4 °C using GlycoProfile Beta-elimination Kit (Sigma-Aldrich) according to manufacturer's instructions. Released glycans were then analyzed by mass spectrometry, as described below.

HPLC-Chip/Q-TOF Mass Spectrometric Analysis

Samples of Hp's enriched azide-labeled glycoproteins, Phos-FLAG-reacted Ac and Az lysates (“tagged lysates”), and purified azide-labeled urease (“tagged urease”) were analyzed before and after beta-elimination of O-linked glycans using a 6530 High Performance Liquid Chromatographic-Chip Quadrupole Time-of-Flight Mass Spectrometer (HPLC-Chip/Q-TOFMS; Agilent Technologies, Santa Clara, CA). Chromatographic separation and nano-electrospray ionization (nanoESI) was performed with a 1260 Chip Cube system (Agilent Technologies) using a ProtID-chip with a 40 nL enrichment column and a 150 mm x 75 μm analytical column (Agilent Technologies). The enrichment and analytical columns were packed with 300 Å, 5 μm particles with C18 stationary phase. The mobile phases were 0.1% formic acid/H₂O (A) and 0.1% formic acid/acetonitrile (B). Samples (0.2–2 μl) were loaded on the enrichment column using 98:2 (A:B) at 4 μl/min and eluted through the analytical column using a gradient from 98:2 (A:B) to 80:20 (A:B) over 2 min to 2:98 (A:B) over a period of 8 min at 0.3 μl/min. Mass spectra (MS and MS/MS) were collected in positive ion mode; the ionization voltage ranged from 1850–1950 V and the ion source temperature was held at 350 °C. The fragmentor and collision cell voltages, as well as other tuning parameters, were optimized to minimize metastable water losses associated with ion sampling and transmission to the TOF analyzer. This was of particular importance for the analysis of HexNAz-Phos-FLAG-His₆ (1), for which metastable water losses became increasingly prominent for higher charge state ions. A similar effect, impacting metastable losses of sulfate from sulfated oligosaccharides, has been reported by Zaia and co-workers (47). Spectra were internally calibrated using methyl stearate (C₁₇H₃₅CO₂CH₃) or dibutyl phthalate (C₁₆H₂₂O₄) and hexakis(1H, 1H, 4H-hexafluorobutyloxy)phosphazine (HP-1221; C₂₄H₁₈O₆N₃P₃F₃₆), continuously introduced and detected as [M+H]⁺. Collision-induced dissociation (CID)-MS/MS experiments were executed with precursor selection determined using a targeted or data-dependent approach. Precursor ions were subjected to CID using nitrogen as the target gas with collision energies ranging from 20–35 V.

Immunopurification of Hp's ureA and ureB

Lysate from metabolically labeled Hp was standardized to 4.4 mg/ml and incubated 1:1 with 500 μm Phos-FLAG overnight at room temperature to label azide-containing glycoproteins. Insoluble membrane material was separated from soluble proteins by centrifugation at 12,000 × g for 5 min. To the soluble protein samples, 1 μl of α-Hp ureA or ureB (Santa Cruz Biotechnology) was added per 100 μl of protein volume. The samples were incubated at 4 °C overnight with gentle shaking. After completion of the overnight incubation, 100 μl of a 50% Protein-G agarose suspension in HNTG washing buffer (20 mm HEPES buffer pH 7.5, containing 150 mm NaCl, 0.1% (w/v) Triton X-100, and 10% (w/v) glycerol) was added to Hp lysates and allowed to incubate for 90 min at 4 °C with gentle shaking. The immunoprecipated complexes were collected via centrifugation at 3000 × g for 5 min at 4 °C and the pellet was washed three times with ice-cold HNTG washing buffer. The final pellet was re-suspended in 40 μl of ice-cold HNTG washing buffer, diluted with an equal volume of 2 × SDS-loading buffer, and heated at 95 °C for 5 min. The boiled samples were centrifuged at 12,000 × g for 30 s to separate the agarose beads from the immunopreciptated proteins. Immunoprecipitated proteins were then analyzed by Western blot with anti-FLAG (Sigma) or anti-urease antibody (Santa Cruz Biotechnology).

Gel and Western Blot Analyses

All lanes were loaded with 10–15 μg of protein sample unless otherwise noted. Western blots were treated with HRP-conjugated anti-FLAG antibody (1:3000, Sigma), rabbit polyclonal anti-Hp ureA (1:3000, Santa Cruz Biotechnology), or rabbit polyclonal anti-Hp ureB antibody (1:3000, Santa Cruz Biotechnology), followed by goat α-rabbit IgG-HRP (1:10000, Santa Cruz Biotechnology), then developed with chemiluminescent substrate (Pierce). Polyacrylamide gels were visualized with zinc stain (BioRad).

RESULTS AND DISCUSSION

Metabolic Labeling of Hp's Proteins With Azides Does Not Appear to be Catabolic

Here we employed metabolic glycan labeling to characterize Hp's glycoproteins and unveil molecules that have the potential to serve as the basis of novel anti-Hp treatments. In previous work, our laboratory demonstrated that supplementation of Hp's media with 1 mm Ac₄GlcNAz for three to 5 days leads to incorporation of azide-dependent signal into a large number of glycoproteins (31). The majority, though not all, of this azide-dependent signal can be removed by glycosidases that catalyze the cleavage of certain N-linked and O-linked glycans (31). Thus, we sought to assess whether the glycosidase-resistant azide signal is because of an alternative metabolic fate of Ac₄GlcNAz in Hp; specifically, catabolism of the azidosugar to the azidoacetyl unit followed by subsequent activation and covalent addition to proteins. This type of azidosugar catabolism has been observed in some other organisms. If this process were to occur in Hp, then supplementation of Hp's media with the azidoacetyl moiety would likely lead to metabolic labeling of Hp's proteins with azides. To assess this possibility, Hp cells were grown in media supplemented with 5 mm sodium azidoacetate or with the more bioavailable azidoacetyl ethyl ester (Fig. 2A); Hang and coworkers demonstrated that structurally analogous compounds are cell permeable (48, 49). Ac₄GlcNAz or the azide-free sugar peracetylated GlcNAc (Ac₄GlcNAc) were employed as positive and negative controls of metabolic labeling, respectively. Ac₄GlcNAz treatment resulted in robust azide-dependent signal in lysates, whereas no azide-labeled proteins were observed in lysates from cells supplemented with Ac₄GlcNAc, with sodium azidoacetate (Fig. 2B), or with azidoacetyl ethyl ester (Fig. 2C). SDS-PAGE analysis confirmed the presence of equivalent protein levels in all lanes (supplemental Fig. S1). These data suggest that azide-dependent labeling of Hp's proteins is not because of catabolic incorporation of the azidoacetyl moiety into proteins. The remaining azide-dependent signal could be glycosidase resistant because of the presence of glycans that are not substrates of the enzymes, because similar glycosidase-resistance, and even resistance to chemical cleavage, has been observed for glycoproteins produced by C. jejuni (24). With the confidence that the vast majority of Hp's azide-labeled proteins are glycosylated, we undertook experiments to study and identify these labeled glycoproteins.

Fig. 2.

Metabolic labeling of Hp's proteins with azides. A, Azidoacetyl compounds used in metabolic feeding experiments with Hp. B, Western blot analysis of lysate from Hp treated with 5 mm sodium azidoacetate (acetate), 1 mm Ac₄GlcNAz (Az), or with 1 mm of the azide-free control sugar Ac₄GlcNAc (Ac) for 4 days. After lysis, samples were reacted with Phos-FLAG and visualized by Western blot with anti-FLAG antibody. C, Western blot analysis of lysate from Hp treated with 5 mm azidoacetyl ethyl ester (ester), 1 mm Ac₄GlcNAz (Az), or with 1 mm of the azide-free control sugar Ac₄GlcNAc (Ac) for 4 days. After lysis, samples were reacted with Phos-FLAG and visualized by Western blot with anti-FLAG antibody.

Assessment of the Subcellular Distribution of Hp's Azide-Labeled Glycoproteins Reveals Glycoproteins Throughout Hp, Including on the Cell Surface

To study Hp's glycoproteins, we began by assessing the subcellular distribution of these species. In particular, we were curious to see whether any of Hp's secreted and cell surface proteins are glycosylated, as these proteins are likely to directly modulate interactions with the host and could be novel therapeutic targets. Hp cells supplemented with the azidosugar Ac₄GlcNAz or with the azide-free control Ac₄GlcNAc (31) were fractionated to acquire secreted (36), periplasmic, inner membrane-associated, cytoplasmic (35), and surface associated proteins (37). Further, fractions containing outer membrane proteins were also obtained (15, 38). Western blot visualization of azide-labeled glycoproteins (Fig. 3) and SDS-PAGE analysis of total protein (see supplemental Fig. S2A–C) demonstrated that protein composition varied between fractions, consistent with successful subcellular separation of proteins. Further, malate dehydrogenase assays (50) of fractions revealed activities consistent with successful separation of cytoplasmic and periplasmic proteins (35) (supplemental Fig. S2D). No azide-labeled proteins were observed in fractions from cells supplemented with Ac₄GlcNAc, whereas Ac₄GlcNAz treatment resulted in robust azide-dependent signal in all subcellular fractions (Fig. 3). Our data demonstrate the remarkable selectivity of the Staudinger ligation between phosphines and azides, as these abiotic functional groups are nonreactive with endogenous functional groups yet display exquisitely selective reactivity with each other to form a covalent adduct (51, 52). Moreover, these data indicate that glycoproteins are present throughout Hp and may serve diverse functions. For example, modification of internal glycoproteins may be important for the physiology of Hp, whereas glycosylation of secreted and surface proteins may be crucial for Hp to survive and cause harm within the gastric mucosa.

Fig. 3.

Subcellular distribution of Hp's azide-labeled glycoproteins. Western blot analysis of subcellular fractions from Hp treated with 1 mm Ac₄GlcNAz (Az) or with the azide-free control sugar Ac₄GlcNAc (Ac) for 4 days. After subcellular fractionation, samples were reacted with Phos-FLAG and visualized by Western blot with anti-FLAG antibody. A, Glycoprotein profiles of four subcellular fractions are shown: secreted (sec), inner membrane (memb), periplasmic (peri), cytoplasmic (cyto). B, Glycoprotein profile of Hp's surface-associated proteins. C, Glycoprotein profile of Hp's outer membrane fraction.

The prominent display of glycans on a subset of Hp's cell surface proteins might be harnessed for novel anti-Hp therapeutic strategies. For example, these glycosylated proteins might provide the basis of a carbohydrate-based vaccine; analogous glycoconjugate vaccines have been remarkably successful in the clinic (14, 53). Moreover, if glycan modifications are important for the interactions of Hp with its host, small molecule inhibitors could target glycan synthesis to inactivate Hp. Finally, Hp's glycans could be selectively labeled with azides and targeted with phosphines to disrupt their function (14, 54, 55). Realizing these strategies would first require identification of Hp's glycoproteins.

Targeted Isolation of Hp's Azide-Labeled Glycoproteins was Facilitated by the Staudinger Ligation

To identify Hp's glycoproteins, we began by taking advantage of the metabolically incorporated azide, in conjunction with the Staudinger ligation, to enrich these species. Lysates from Hp supplemented with Ac₄GlcNAz or Ac₄GlcNAc were subjected to Staudinger ligation with a phosphine comprising a tandem FLAG-hexahistidine peptide (Phos-FLAG-His₆, Fig. 4A) (29, 56). Tagged glycoproteins containing both the FLAG and His₆ epitopes were enriched by purification via anti-FLAG chromatography followed by Ni²⁺ affinity chromatography (29, 56) (Fig. 4A). SDS-PAGE analysis enabled comparison of protein profiles from samples before purification (input) versus after enrichment of azide-labeled species (eluent). A large number of proteins were present in both the azide-labeled and acetyl-labeled samples before purification (Fig. 4B). In contrast, after purification, proteins were present in substantially higher abundance in the enriched, azide-labeled sample relative to the mock-enriched control (Fig. 4B). These data highlight the selectivity of the Staudinger ligation between phosphines and azides. The ladder of proteins observed in the enriched, azide-labeled fraction is consistent with the large number of azide-labeled proteins observed by Western blot analysis of lysate from Ac₄GlcNAz-treated cells (Figs. 2B, 2C). These data indicate a dramatic enrichment of Hp's azide-labeled glycoproteins using Phos-FLAG-His₆. Analysis of the enriched, azide-labeled glycoprotein sample by two-dimensional gel electrophoresis yielded greater than 50 detectable species (Fig. 4C), indicating that Hp synthesizes myriad glycoproteins. The large number of Hp glycoproteins detected is similar in magnitude to the number of glycoproteins identified in large-scale studies of the related species C. jejuni (57, 58) and in more distantly related bacteria, including Mycobacterium tuberculosis (26), Neisseria sp. (20)., and Bacteroides sp. (27). Hp appears to dedicate considerable metabolic energy to glycoprotein synthesis, suggesting glycoproteins play an important role in Hp's physiology.

Fig. 4.

Enrichment of Hp's azide-labeled glycoproteins. A, Targeted isolation of Hp's azide-labeled glycoproteins was facilitated by the Staudinger ligation. First, Hp's azide-labeled glycoproteins were reacted with Phos-FLAG-His₆ to yield glycoproteins containing a tandem affinity tag. Labeled glycoproteins were then enriched by anti-FLAG chromatography followed by Ni²⁺ affinity chromatography. Finally, enriched glycoproteins were trypsinized and analyzed by multidimensional protein identification technology (mudPIT). These experiments were performed along side an azide-free control. B, SDS-PAGE analysis of proteins from Hp treated with Ac₄GlcNAz (Az) or with the azide-free control sugar Ac₄GlcNAc (Ac) before enrichment (input) and after elution from the nickel column (eluent) indicates successful isolation of azide-labeled glycoproteins. Proteins were visualized with zinc stain. C, Two-dimensional gel analysis of Hp's enriched, azide-modified glycoproteins. Proteins were visualized with zinc stain.

Identification of Hp's Azide-Labeled Glycoproteins by Mass Spectrometry Indicates Glycosylation of a Number of Virulence Factors

Motivated by these results, we next sought to identify Hp's glycoproteins by mass spectrometry analysis. Hp's enriched, azide-labeled glycoproteins and a mock-enriched control (which was labeled with Ac₄GlcNAc, reacted with Phos-FLAG-His₆, and enriched via anti-FLAG and Ni-NTA affinity chromatography, as described above) were digested with trypsin, then analyzed by multi-dimensional protein identification technology (mudPIT) (Fig. 4A) (40). Experimental detection of unique peptides enabled the unambiguous identification of 127 proteins in the azide-labeled sample. In contrast, the mock-enriched control that lacked the azide contained only two Hp proteins (HP0721, HP0570), both of which were also present in the azide-labeled sample. These proteins were subtracted from the list of putative glycoproteins identified in the azide-labeled sample, narrowing the pool of candidate glycoproteins to 125 (Table I and Table II). All proteins were identified based on two or more peptides meeting acceptance criteria. Detailed information on all of the proteins identified in the azide-labeled sample and the mock-enriched control, including detected peptides that made assignments possible, can be found in supplemental Table S1. These results demonstrate efficient enrichment and identification of azide-labeled proteins, suggesting this metabolic labeling strategy is a robust method to profile bacterial glycoproteins.

Table I. Putative glycoproteins identified by mudPIT with links to pathogenesis.

Gene	Acc. #	Protein name	Predicted MW (kDa)	Function/sequence features	Link to pathogenesis	Subcellular localizationa	Spect- counts	Uni-pep	Seq-cov%
HP0073	P14916	ureA	27.8	Urease alpha subunit	Enables survival in acid	Cytoplasm, surface	96	4	34.4
HP0072	P69996	ureB	61.6	Urease beta subunit	Enables survival in acid	Cytoplasm, surface	188	7	22.7
HP0527	NP_207323	cag7	230.1	cag pathogenicity island, type IV secretion	Oncogene, gastric cancer	Outer membrane	2	2	1.4
HP1563	P21762	tsaA	23.5	Alkyl hydroperoxide reductase	Resists oxidative damage	Cytoplasm	428	12	65.7
HP0325	O25092	flgH	27.8	Flagellar export	Critical for motility	Outer membrane	4	2	7.2
HP0010	P42383	groEL	61.5	Chaperone	Gastric cancer-associated antigen, immunogen	Cytoplasm, surface	576	18	37.3
HP0011	P0A0R3	groES	13.7	Chaperone	Gastric cancer-associated antigen, immunogen	Cytoplasm, surface	546	7	41.9
HP1161	O25776	fldA	18.5	Flavodoxin	Critical metabolic gene	Unknown	2	2	20.8
HP0653	P52093	pfr	20.3	Nonheme iron-containing ferritin	Crucial for iron acquisition	Cytoplasm	35	3	25.6
HP0389	AAD07454	sodB	25.9	Superoxide dismutase	Resists oxidative damage	Periplasm	10	4	28.9
HP0875	P77872	katA	61.5	Catalase	Combats oxidative stress	Periplasm, surface	91	6	18.0
HP0390	O25151	tagD	19.3	Adhesin thiol peroxidase	Combats oxidative stress	Periplasm	3	3	32.6
HP0243	P43313	napA	17.9	Neutrophil activation	Immunomodulator, biofilm formation	Cytoplasm, surface	9	3	25.5
HP0224	O25011	msrA	43.2	Methionine sulfoxide reductase	Antioxidant, immunogen	Cytoplasm	42	4	18.6
HP0317	AAD07380	babC	85.4	Outer membrane protein	Adhesion, glycan binding	Outer membrane	4	2	5.4
HP1243	AAD08288	babA	83.3	Outer membrane protein	Adhesion, glycan binding	Outer membrane	2	1	2.8
HP1177	AAD08221	hopQ	73.3	Outer membrane protein	Present on disease-related strains	Outer membrane	14	2	5.5
HP1205	P56003	tufB	45.9	Translation elongation factor	Resistance to prolonged acid exposure	Cytoplasm	10	4	17.1
HP1123	O25748	slyD	21.3	Peptidyl-prolyl cis-trans isomerase	Important for urease assembly	Cytoplasm	2	2	16.2
HP1195	P56002	fusA	80.9	Translation elongation factor	Duodenal ulcer-related antigen	Cytoplasm, surface	36	8	21.0
HP0027	P56063	icd	50.1	Isocitrate dehydrogenase	Immunogen, induces humoral immune system	Cytoplasm	21	4	15.6
HP0192	O06913	frdA	84.0	Fumarate reductase	Immunogen, crucial for survival in gastric mucosa	Inner membrane	28	5	13.2
HP0109	P55994	dnaK	70.6	Chaperone and heat shock protein 70	Stress induced surface adhesion, immunogen	Cytoplasm, surface	6	3	8.7
HP0082	AAD07152	tlpC	79.1	Methyl-accepting chemotaxis transducer	Assist colonization of the stomach	Inner membrane	10	3	7.6
HP1019	AAD08063	htrA	50.6	Serine protease, chaperone	Disrupts intracellular adhesion	Periplasm, secreted	5	2	6.6
HP0099	AAD07167	tlpA	78.4	Methyl-accepting chemotaxis protein	Chemotaxis receptor	Outer membrane	3	1	3.2

^a Subcellular localization was based on literature reports or predicted using PSORTb v.3.0.

Table II. Additional putative glycoproteins identified by mudPIT.

Gene	Acc. #	Protein name	Predicted MW (kDa)	Function/sequence features	Subcellular localizationa	Spect- counts	Uni-pep	Seq-cov%
HP0025	AAD07106	omp2	81.6	Outer membrane protein	Outer membrane	5	3	7.9
HP0191	O06914	frdB	29.3	Fumarate reductase beta	Inner membrane	15	4	26.2
HP0599	AAD07662	hylB	51.0	Hemolysin secretion	Outer membrane	31	8	29.5
HP0371	AAD07435	fabE	18.1	Biotin carboxyl carrier	Unknown	35	4	26.7
HP0110	P55970	grpE	23.3	Heat shock protein	Cytoplasm	14	5	29.9
HP0631	AAD07691	hydA	44.6	Quinine reactive hydrogenase	Inner membrane	4	3	9.1
HP1198	O25806	rpoBC	339.5	RNA polymerase	Cytoplasm	9	6	3.6
HP1496	P56078	ctc	20.9	General stress protein	Cytoplasm	6	3	33.7
HP0824	P66928	trxA	12.5	Thioredoxin A	Cytoplasm	20	2	26.8
HP0825	3ISH_B	trxB	35.5	Thioredoxin reductase	Cytoplasm	22	6	33.1
HP1458	AAD08500	—	12.4	Thioredoxin	Cytoplasm	47	2	26.4
HP1325	O25883	fumC	53.6	Fumarase	Cytoplasm	23	5	25.5
HP1555	P55975	tsf	41.6	Translation elongation factor	Cytoplasm	17	3	9.1
HP0266	O25045	pyrC	44.6	Dihydroorotase	Unknown	19	5	25.1
HP1135	AAD08177	atpH	21.3	ATP synthase F1 delta	Membrane	10	2	18.0
HP1134	P55987	atpA	58.1	ATP synthase F1 alpha	Membrane	28	5	13.2
HP1132	P55988	atpD	54.1	ATP synthase F1 beta	Membrane	26	6	22.3
HP1099	AAD08142	eda	23.9	Aldolase	Cytoplasm	5	3	23.2
HP0154	P48285	eno	49.1	Enolase	Cytoplasm	12	4	21.6
HP0512	P94845	glnA	57.4	Glutamate synthase	Cytoplasm	32	4	15.4
HP0779	AAD07828	acnB	97.7	Aconitase	Cytoplasm	72	10	19.9
HP0163	P56074	hemB	38.1	Delta-aminolevulinic acid dehyratase	Cytoplasm	5	3	15.2
HP0974	P56196	pgm	57.7	Phosphoglycerate mutase	Cytoplasm	5	4	18.1
HP0294	O25067	aimE	39.7	Aliphatic amidase	Cytoplasm	31	3	17.9
HP0649	P56149	aspA	54.6	Aspartate ammonia lyase	Cytoplasm	17	5	17.5
HP0865	O25536	dut	16.8	Nucleotidohydrolase	Unknown	3	1	17.5
HP0020	AAD07088	nspC	48.2	Carboxynorpermidine decarboxylase	Unknown	5	4	16.9
HP1540	AAD08580	fbcF	19.2	Ubiquinol cytochrome c oxidase	Inner membrane	18	2	15.9
HP0480	O25225	yihK	69.9	GTP-binding	Inner membrane	10	5	14.1
HP0954	O25608	—	25.4	NAD(P)H reductase	Cytoplasm	3	2	14.0
HP0176	P56109	tsr	35.7	Fructose-bisphosphate aldolase	Cytoplasm	3	2	13.8
HP1110	AAD08154	porA	47.0	Pyruvate ferredoxin oxidoreductase	Unknown	47	4	18.9
HP1111	AAD08155	pfor	36.9	Pyruvate ferredoxin oxidoreductase beta	Unknown	37	3	16.9
HP0632	AAD07692	hydB	67.6	Quinone-reactive Ni-Fe hydrogenase	Inner membrane	7	4	13.5
HP0026	P56062	gltA	50.9	Citrate synthase	Cytoplasm	71	6	13.2
HP0757	AAD07805	—	34.9	Beta-alanine synthase	Cytoplasm	6	2	12.4
HP0485	AAD07551	—	37.7	Catalase-like protein	Periplasm	4	3	12.3
HP0201	AAD07269	plsX	38.4	Fatty acid phospholipid synthesis	Cytoplasm	5	2	12.1
HP0183	P56089	glyA	48.0	Serine hydroxyl-methyltransferase	Cytoplasm	17	3	11.9
HP0690	AAD07742	fadA	43.4	Acetyl coenzyme acetyltransferase	Cytoplasm	5	2	11.9
HP0402	P56145	pheT	89.4	phenylalanyl-tRNA synthase	cytoplasm	20	5	11.7
HP0096	AAD07165	—	36.7	phosphoglycerate dehydrogenase	cytoplasm	6	2	11.7
HP0576	O25300	lepB	35.3	signal peptidase I	inner membrane	11	3	11.1
HP1013	O25657	dapA	34.8	dihydrodipicolinate synthetase	cytoplasm	8	2	10.8
HP0397	AAD07461	serA	60.8	phosphoglycerate dehydrogenase	cytoplasm	12	3	10.7
HP1576	O26096	abc	38.6	ABC transporter	inner membrane	4	2	10.7
HP0006	P56061	panC	32.8	pantoate-beta-alanine ligase	cytoplasm	2	2	10.7
HP0558	NP_207353	fabF	45.8	beta ketoacyl-acyl carrier protein synthase	inner membrane	8	2	10.6
HP0330	O25097	ilvC	38.5	ketol-acid reductoisomerase	cytoplasm	2	2	10.3
HP0672	AAD07733	aspB	45.1	mitochondrial signature protein	cytoplasm	2	2	10.0
HP1103	O25731	glk	38.6	glucokinase	cytoplasm	11	2	9.9
HP0221	AAD07289	—	38.3	nifU-like	cytoplasm	2	2	9.9
HP0604	AAD07669	hemE	40.4	uroporphyrinogen decarboxylase	cytoplasm	3	2	9.8
HP0306	P56115	hemL	49.3	aminomutase	cytoplasm	3	2	9.7
HP1266	AAD08310	nqo3	99.1	NADH-ubiquinone oxidoreductase	unknown	8	4	9.3
HP1366	AAD08410	mboIIR	52.6	type IIS restriction enzyme R	unknown	4	2	8.5
HP0490	AAD07558	—	45.2	putative potassium channel	unknown	2	2	8.5
HP1181	AAD08227	mfs	51.4	multidrug-efflux transporter	inner membrane	4	2	8.4
HP0777	P56106	pyrH	27.4	uridine 5′ monophosphate kinase	cytoplasm	13	2	8.3
HP1422	P56456	ileS	111.3	Isoleucyl-tRNA synthetase	cytoplasm	5	4	8.1
HP0322	AAD07390	—	61.4	poly E-rich protein	cytoplasm	5	3	7.9
HP0382	AAD07451	—	48.5	zinc metallo protease	unknown	3	2	7.9
HP0859	AAD07905	rfaD	39.4	mannoheptose epimerase	cytoplasm	5	2	7.8
HP1100	P56111	—	70.2	6-phophogluconate dehydratase	cytoplasm	55	2	6.4
HP0210	P56116	htpG	74.8	chaperone and heat shock protein	cytoplasm	2	2	7.2
HP0056	AAD07126	—	141.6	pyrroline-5-carboxylate dehydrogenase	cytoplasm	14	5	7.1
HP0728	O25428	tilS	41.7	tRNA(Ile)-lysidine synthetase	unknown	3	2	7.1
HP1045	O25686	acoE	78.8	acetyl-CoA synthetase	cytoplasm	7	3	6.7
HP0169	P56113	prtC	49.9	collagenase	unknown	2	2	6.7
HP1104	AAD08150	cad	40.6	cinnamyl-alcohol dehydrogenase	cytoplasm	7	2	6.6
HP0470	AAD07532	pepF	70.9	oligoendopeptidase F	cytoplasm	5	3	6.4
HP0422	AAD07486	speA	74.0	arginine decarboxylase	periplasm	7	3	6.3
HP0510	P94844	dapB	29.4	dihydrodipicolinate reductase	cytoplasm	4	2	6.3
HP0680	P55982	nrdA	94.7	Ribonucleoside-diphosphate reductase	Cytoplasm	2	2	5.5
HP1430	P56185	—	81.3	Conserved hypothetical ATP-binding protein	Unknown	4	2	5.5
HP1012	AAD08056	pqqE	52.9	Protease	Unknown	2	2	5.3
HP1213	O25812	pnp	80.8	Polynucleotide phosphorylase	Cytoplasm	2	2	4.8
HP1278	P56142	trpB	45.0	Tryptophan synthase beta	Cytoplasm	16	1	4.6
HP1072	P55989	copA	86.1	Copper-transporting ATPase	Inner membrane	3	2	4.3
HP1450	O25989	yidC	65.8	Protein translocase	Inner membrane	6	2	4.3
HP0121	P56070	ppsA	95.8	Phosphoenolpyruvate synthase	Cytoplasm	3	2	4.1
HP0607	NP_207402	acrB	119.6	Acriflavine resistance	Inner membrane	2	2	4.0
HP1402	AAD08445	hsdR	121.7	Type I restriction enzyme	Unknown	2	2	3.4
HP1112	P56468	purB	52.6	Adenylosuccinate lyase	Cytoplasm	5	1	3.4
HP1478	AAD08516	uvrD	81.7	DNA helicase II	Cytoplasm	2	2	3.3
HP1547	P56457	leuS	97.7	Leucyl-tRNA synthase	Cytoplasm	3	2	2.9
HP0791	Q59465	cadA	78.9	Cadmium-transporting ATPase	Inner membrane	2	2	2.9
HP0600	AAD07663	spaB	71.8	Multidrug resistance	Inner membrane	3	1	2.7
HP0696	AAD07747	—	90.7	N-methylhydantoinase	Unknown	3	2	2.6
HP0958		—	31.3	Hypothetical protein	Unknown	2	2	12.3
HP0787		—	48.2	Conserved integral membrane protein	Inner membrane	21	3	9.3
HP0231		—	31.1	Hypothetical protein	Unknown	10	4	22.9
HP0318		—	30.1	Hypothetical protein	Unknown	35	3	18.4
HP0086		—	53.3	Hypothetical protein	Unknown	35	3	9.1
HP0773		—	41.9	Hypothetical protein	Unknown	32	2	8.9
HP0754		—	9.7	Hypothetical protein	Unknown	7	2	34.1
HP0396		—	74.4	Hypothetical protein	Unknown	3	3	8.8
HP1079		—	45.2	Hypothetical protein	Unknown	34	2	6.9
HP1143		—	53.2	Hypothetical protein	Unknown	4	1	5.0

^a Subcellular localization was based on literature reports or predicted using PSORTb v.3.0.

Of the putative glycoproteins identified, 70% have housekeeping functions, 21% are linked to Hp's pathogenesis, and 9% have unknown function. From a basic biology perspective, the identification of a number of putative glycoproteins that have housekeeping functions suggests that glycosylation could play a role in proper folding and function (59), or perhaps that glycosylation provides a selective advantage for survival in the harsh conditions of the gastric tract. From a therapeutic perspective, it is intriguing that 26 of Hp's glycoproteins have known roles in pathogenesis (Table I) that include enabling survival in the host stomach via iron acquisition (pfr) (60), pH resistance (ureA, ureB, tufB) (61), oxidative protection (tsaA, sodB, katA, tagD, msrA) (62, 63), motility (flgH) (64), adhesion (babC, babA, tlpA, dnaK, htrA) (65), eliciting an immune response (napA, msrA, icd, frdA) (66, 67), and causing harm to the host (cag7, groES, groEL, hopQ, fusA) (68, 69). The large percentage of virulence factors in Hp's glycoproteome, relative to the low percentage of virulence factors in Hp's proteome (70, 71), implicates Hp's glycans in pathogenesis. This observation suggests that the known link between protein glycosylation and virulence in Hp, which Logan firmly established with her studies of Hp's flagellin glycosylation (7), may be a widespread phenomenon within this bacterium.

We next considered the subcellular distribution of Hp's putative glycoproteins. Based on literature reports and in silico predictions, candidate glycoproteins appear to be expressed throughout Hp (Fig. 5), including intracellular locations such as the cytoplasm, inner membrane, and periplasm, and extracellular locations such as the outer membrane, cell surface (surface-associated), and the secreted milieu. Moreover, the predicted subcellular distribution of Hp's putative glycoproteins (Fig. 5) appears to reflect the subcellular distribution that we detected experimentally (Fig. 3). Follow-up experiments need to be conducted to establish whether these localization predictions are correct and to confirm that Hp synthesizes glycoproteins throughout the cell, as our data suggest.

Fig. 5.

Predicted subcellular location of candidate glycoproteins. The cellular locations of each identified protein was determined by a combination of literature reports and predictions by PSORTb v.3.0. Secondary surface-associated localization (n = 8) was not included.

We next compared the results of our analyses of Hp's glycoproteins to a recent study by Creuzenet and coworkers. In their study, they employed a periodic acid/hydrazide-based screen to identify nine candidate Hp glycoproteins (15). Five of the putative glycoprotein hits identified here (katA, gltA, tsf, atpA, atpD) were also identified as candidate glycoproteins in that screen, further validating our hit list. Moreover, their analyses indicated the presence of glycoproteins in soluble and membrane fractions of Hp (15), consistent with the subcellular distribution observed here. These findings underscore the value of our targeted strategy to detect and discover bacterial glycoproteins in a manner that is complementary to alternative techniques.

We note the absence of the flagellin proteins FlaA and FlaB, glycoproteins confirmed to be modified with pseudaminic acid, from our list of identified azide-labeled glycoproteins. This result is consistent with our previous studies, which demonstrated that Ac₄GlcNAz treatment does not lead to detectable incorporation of azide into Hp's flagellin proteins (31). Our previously reported data indicate that Ac₄GlcNAz is not converted to the complex sugar pseudaminic acid at detectable levels. Thus, though the studies described here identity a large number of putative Hp glycoproteins, they do not exhaustively identify all Hp glycoproteins.

Mass Spectrometry Analyses Reveal Staudinger Ligation-Glycan Adducts

We sought to characterize the nature of Hp's azide-modified glycans. A sample of Hp's enriched, azide-labeled glycoproteins was analyzed by HPLC-Chip/Q-TOFMS before and after being subjected to beta-elimination to cleave O-linked glycans. For these preliminary experiments, we focused on identifying the simplest Staudinger ligation-glycan adduct that would be formed as a result of metabolic labeling with Ac₄GlcNAz - addition of a single GlcNAz residue or, if the stereochemistry is changed, an N-azidoacetylhexosamine (HexNAz) residue onto a glycosylated protein. Thus, we directed our analysis at the identification of the Staudinger ligation-glycan adduct that would be formed by protein glycosylation with a single HexNAz residue followed by release via beta-elimination - HexNAz-Phos-FLAG-His₆ (1) (Fig. 6). Because this adduct has not been previously characterized by MS, we first synthesized GlcNAz-Phos-FLAG-His₆ and analyzed its chromatographic and mass spectrometric properties using LC-nanoESI and collision-induced dissociation (CID). The nanoESI mass spectrum of the synthetic standard GlcNAz-Phos-FLAG-His₆ (M = 2400.90 Da) is dominated by [M+3H]³⁺, [M+4H]⁴⁺, and [M+5H]⁵⁺ ions (supplemental Fig. S3); additionally, we observed metastable losses of water that were most apparent for the [M+5H]⁵⁺ charge state. The intensities of these peaks were highly dependent on voltages controlling ion transmission, so we attempted to minimize these contributions through careful instrument tuning. CID-MS/MS analysis of the [M+4H]⁴⁺ ion at m/z 601.24 revealed fragmentation rationalized by initial loss of the neutral glycan via pathway (a) to form product ion Ia⁴⁺ (m/z 556.46) (Figs. 6 and 7A), which proceeds, via net loss of CO and CH₂NH, to form the base peak in the spectrum (product ion IIa⁴⁺ at m/z 542.21) (Figs. 6 and 7A; supplemental Table S3). We also detect triply charged versions of product ions at m/z 741.63 and 722.61 (ions Ib³⁺ and IIb³⁺, respectively) (Figs. 6 and 7A), which may be formed by loss of the positively charged glycan via pathway (b) (Fig. 6). This pathway may also be responsible for formation of the glycan-derived product ions that appear in the spectrum at m/z 162.08, m/z 144.07, m/z 126.06, and m/z 96.04 (see proposed structures in Figs. 6 and 7B). We also detected FLAG-His₆-derived product ions. These include abundant immonium ions at m/z 110.07 (His) and m/z 129.10 (Lys), as well as y-type (72) fragments containing the FLAG-His₆ C terminus (see Fig. 7E) and b-type (72) fragments; the latter are denoted by b_n* and contain the glycan-eliminated N terminus (see Fig. 7E) that is attributed to subsequent fragmentations from ion IIa⁴⁺. To determine if the Staudinger ligation-glycan adduct HexNAz-Phos-FLAG-His₆ (1) is released from Hp's enriched, azide-labeled glycoproteins on beta-elimination, a sample was analyzed by HPLC-Chip/Q-TOFMS before and after being subjected to beta-elimination to cleave O-linked glycans. Although analysis of the pre-beta-eliminated sample showed no evidence for product 1, LC-nanoESI analysis of the beta-eliminated sample showed a chromatographic peak appearing at the retention time characteristic of synthetic 1, and producing a mass spectrum with m/z values consistent with the mass of product 1 (M = 2,300.90 Da, mass measurement error < 5 ppm) (supplemental Fig. S4 and supplemental Table S2). Furthermore, when the [M+4H]⁴⁺ ion at m/z 601.24 from putative product 1 was subjected to MS/MS analysis, the measured mass spectrum showed excellent agreement with the synthetic standard (Figs. 7C, 7D; supplemental Table S4). These results suggest that one metabolic fate of Ac₄GlcNAz in Hp is conversion to HexNAz (presumably GlcNAz) and incorporation into Hp's O-linked glycoproteins.

Fig. 6.

Proposed mass spectrometric fragmentation pathways for Staudinger ligation-glycan adducts 1 (HexNAz-Phos-FLAG-His₆) and 2 (HexNAz-Phos-FLAG). CID-MS/MS of nanoESI-generated [M+4H]⁴⁺ (adduct 1) and [M+2H]²⁺ (adduct 2) ions yield fragment ions Ia and IIa, formed via pathway (a), through which loss of the neutral glycan is hypothesized to generate cycylized Ia, which is followed by sequential losses of CO and CH₂NH to generate IIa. Charge-retention on the glycan and subsequent fragmentation via pathway (b) rationalizes formation of glycan-derived putative product ions at m/z 162.08, m/z 144.07, m/z 126.06, and m/z 96.04. Production of y- and b-type fragment ions is attributed to subsequent fragmentation of IIa yielding C- and N-(Phos-containing)-terminus fragments, respectively (see Figs. 7E and 8E).

Fig. 7.

LC-nanoESI-CID-MS/MS spectra of (A, B) synthetic standard GlcNAz-Phos-FLAG-His₆ and (C, D) beta-eliminated glycans from enriched azide-labeled glycoprotein sample. A, CID-MS/MS spectrum of GlcNAz-Phos-FLAG-His₆ synthetic standard 1 showing ions Ia⁴⁺ and Ib³⁺ (see Fig. 6) resulting from glycan loss, and ions IIa⁴⁺ and IIb³⁺ following net loss of CO, NH = CH2 (Fig. 6). As summarized in (E), subsequent fragmentation of ion IIa⁴⁺ is proposed to yield y-type fragment ions that include the C terminus of the FLAG-His₆ peptide and b-type ions, labeled with an asterisk, that include the Phos-modified N terminus. B, Expansion (magnified 5x along the y axis) of the low-mass region from (A) displays a variety of peaks that include glycan-derived fragments with proposed structures represented. Immonium ions (H and K) are also detected in this mass range. C, CID-MS/MS spectra of beta-eliminated glycans from enriched azide-labeled glycoprotein sample showing mass spectral features that agree with those of the synthetic standard (A). D, Expansion (magnified 5x along the y axis) of the low-mass region from (C), showing the production of glycan-derived peaks from the beta-eliminated putative HexNAz-Phos-FLAG-His₆ (1). E, Schematic representation of ions (y- and b-type) derived from further IIa⁴⁺ fragmentation. The asterisk associated with the b-type ions denotes a Phos-modified N terminus. Charge states are labeled when greater than +1. All CID-MS/MS spectra were measured using nitrogen as the collision gas, a 4 Da window for ion isolation, and collision energy of 20 V.

Although the above data indicate the presence of a Staudinger ligation-glycan adduct in an azide-labeled glycoprotein sample, we sought to confirm that these adducts are absent from acetyl-labeled controls. Because of the metastable water losses observed with the Phos-FLAG-His₆ tag and the multiplicity of charge states, we chose to conduct follow-up experiments with the less complex Phos-FLAG tag. We first characterized synthetic GlcNAz-Phos-FLAG (2) (M = 1578.54 Da) by LC-nanoESI and CID. With the basic His₆ chain eliminated, 2 yielded a nanoESI mass spectrum dominated by the lower charge state [M+2H]²⁺ ion at m/z 790.28 (supplemental Fig. S5; supplemental Table S5). As was observed for 1, the CID-MS/MS spectrum for GlcNAz-Phos-FLAG (2) (Fig. 8A and 8B) was dominated by peaks resulting from neutral glycan loss via pathway (a) to form product ions Ia²⁺ and IIa²⁺ (Figs. 6 and 8A); however, glycan-derived fragments (m/z = 96.04, 126.05, 144.07 and 162.08) were weaker, whereas y- and b-type sequence ions (Figs. 8A and 8E) were more abundant in the spectrum of GlcNAz-Phos-FLAG (2), a change that may result from the lower charge state of this precursor.

Fig. 8.

LC-nanoESI-CID-MS/MS spectra of (A, B) synthetic standard GlcNAz-Phos-FLAG (2) and (C, D) beta-eliminated glycans from an azide-labeled, Phos-FLAG-ligated, Hp cell lysate. A, CID-MS/MS spectrum of GlcNAz-Phos-FLAG synthetic standard 2 showing ions Ia²⁺ (see Fig. 6) resulting from glycan loss, and ions IIa²⁺ following net loss of CO, NH = CH2 (Fig. 6). As summarized in (E), subsequent fragmentation of ion IIa²⁺ is proposed to yield y-type fragment ions that include the C terminus of the FLAG peptide and b-type ions, labeled with an asterisk, that include the Phos-modified N terminus. B, Expansion of the low-mass region from (A) displays a variety of peaks that include glycan-derived fragments with proposed structures represented. Immonium ions (K) are also detected in this mass range. C, CID-MS/MS spectra of beta-eliminated glycans from an azide-labeled, Phos-FLAG-ligated, Hp cell lysate showing mass spectral features that agree with those of the synthetic standard (A). D, Expansion of the low-mass region from (C), showing the production of glycan-derived peaks from the beta-eliminated putative HexNAz-Phos-FLAG (2). E, Schematic representation of ions (y- and b-type) derived from further IIa⁴⁺ fragmentation. The asterisk associated with the b-type ions denotes a Phos-modified N terminus. Charge states are labeled when greater than +1. All CID-MS/MS spectra were measured using nitrogen as the collision gas, a 4 Da window for ion isolation, and collision energy of 35 V.

With characterization of the targeted analyte in hand, lysates from Hp supplemented with Ac₄GlcNAz or Ac₄GlcNAc were rinsed extensively to remove residual free sugar, and then subjected to Staudinger ligation with Phos-FLAG. After extensive washing to remove unreacted Phos-FLAG from labeled lysates via ultrafiltration, samples were analyzed by HPLC-Chip/Q-TOFMS before and after being subjected to beta-eliminations to cleave O-linked glycans. LC-nanoESI analysis of the beta-eliminated glycans from the azide-labeled sample revealed a doubly-charged peak at m/z 790.28 that corresponds to the mass of HexNAz-Phos-FLAG (2) (M = 1578.54 Da, mass measurement error < 5 ppm; supplemental Fig. 6E and supplemental Table S2). The CID-MS/MS spectrum of the peak at m/z 790.28 (Fig. 8C and 8D; supplemental Table S6) is consistent with the Staudinger ligation product between GlcNAz and Phos-FLAG - HexNAz-Phos-FLAG (2). This HexNAz-Phos-FLAG adduct is not detectable in the beta-eliminated sample from the corresponding acetyl-labeled control (supplemental Fig. S6C), nor is it present at detectable levels in tagged azide- or acetyl-labeled lysates before beta-elimination (supplemental Fig. S6A and S6B). The exclusive detection of this adduct in beta-eliminated azide-labeled samples indicates that it is derived from glycosylated Hp proteins and that beta-elimination conditions released this glycan adduct. The absence of this adduct in samples before beta-elimination further supports the conclusion that this adduct is derived from Hp glycoprotein conjugates and is released by the beta-elimination reaction. These data provide strong evidence that some Hp glycoproteins are covalently modified with O-linked HexNAz.

N-acetylhexosamine (HexNAc) residues are found in a number of characterized bacterial glycoproteins, including those synthesized by Campylobacter jejuni (73) and Helicobacter pullorum (74). Indeed, H. pullorum has an N-linked protein glycosylation system that makes use of a pentasaccharide with a core HexNAz. Structural studies are underway in our laboratory to assess whether Hp's O-linked HexNAz is elaborated on to form a higher order glycan.

In our experiments, detection of HexNAz-Phos-FLAG-His₆ and HexNAz-Phos-FLAG adducts by HPLC-Chip/Q-TOFMS analysis was greatly aided by the presence of the peptide moiety, which has favorable chromatographic and ionization characteristics. Thus, MOE facilitated detection of Hp's labeled glycans in unanticipated ways. If chromatographic resolution and signal enhancement are generalizable, metabolic glycan labeling coupled to reaction with a readily ionizable tag has the potential to amplify glycan detection in myriad bacteria and overcome challenges associated with the poor resolution and ionizablility of glycans.

ureA and ureB are Biochemically Validated Glycoproteins

Of the putative glycoproteins identified, two of them - urease alpha (ureA) and urease beta (ureB) - are surface-associated colonization and persistance factors (75). We sought to evaluate their glycosylation status, because this has important implications for host-pathogen interactions and the development of novel anti-Hp therapeutics. To assess the glycosylation status of ureA and ureB, these proteins were immunopurified from Hp treated with 1 mm Ac₄GlcNAz or Ac₄GlcNAc, reacted with Phos-FLAG, and then analyzed by Western blot. Probing immunopurified ureA with anti-ureA revealed the presence of a band at ∼27 kDa (Fig. 9A), indicating successful enrichment of ureA. Further, probing with anti-FLAG antibody revealed the presence of azide-dependent signal in ureA (Fig. 9A). Similar analyses of immunopurified ureB indicated successful enrichment of ureB (Fig. 9B) and the presence of azide-dependent signal in ureB (Fig. 9B). These results suggest that ureA and ureB are glycosylated with azide-modified glycans.

Fig. 9.

Secondary screens to assess the glycosylation status of urease alpha (ureA) and urease beta (ureB) subunits. A, Western blot analysis of immunopurified ureA from Hp treated with Ac₄GlcNAz (Az) or Ac₄GlcNAc (Ac). These samples were reacted with Phos-FLAG, electrophoresed, and then probed with anti-ureA or anti-FLAG antibody. B, Western blot analysis of immunopurified ureB from Hp treated with Ac₄GlcNAz (Az) or Ac₄GlcNAc (Ac). These samples were reacted with Phos-FLAG, electrophoresed, and then probed with anti-ureB or anti-FLAG antibody.

Based on our glycan mass spectrometry results, we were curious to see whether urease (composed of alpha and beta subunits) is among the glycoproteins modified with O-linked HexNAz in Hp. To assess this possibility, urease was biochemically purified from Ac₄GlcNAz-labeled Hp. Purified urease (supplemental Fig. S7) was reacted with Phos-FLAG and rinsed extensively to remove residual Phos-FLAG. Hp's labeled urease was analyzed by HPLC-Chip/Q-TOFMS before and after being subjected to beta-elimination to cleave O-linked glycans. Although analysis of the pre-beta-eliminated sample showed no evidence for product 2 (supplemental Fig. S8A), LC-nanoESI analysis of the beta-eliminated sample showed a chromatographic peak appearing at the retention time characteristic of synthetic HexNAz-Phos-FLAG (2) (supplemental Fig. S8B) and producing a mass spectrum (supplemental Fig. S8C) with m/z values consistent with the mass of product 2 (M = 1578.54 Da, mass measurement error < 5 ppm; supplemental Table S2). Furthermore, when the [M+2H]²⁺ ion at m/z 790.28 from putative product 2 was subjected to MS/MS analysis, the measured mass spectrum showed excellent agreement with the synthetic standard (supplemental Figs. S8D and S8E; supplemental Table S7). These results suggest that urease is glycosylated with O-linked HexNAz and have implications for the development of urease-based therapeutic strategies to prevent and treat Hp infection. Taken together, these data validate urease as a glycosylated protein and indicate that MOE is a robust approach to profiling bacterial glycoproteins.

CONCLUDING REMARKS

Glycosylation of Hp's flagellin proteins has been directly linked to Hp's pathogenesis, suggesting that Hp's glycoproteins are potential targets of therapeutic intervention. Despite their potential therapeutic importance, only the two flagellin glycoproteins have been firmly characterized in Hp (7). Mounting evidence from our lab (76) and others (15) suggests protein glycosylation is common in Hp. Employing experimental approaches to selectively study and identify Hp's glycoproteins should facilitate the discovery of new therapeutic targets and shed light on the role of protein glycosylation in Hp's physiology. In this article, we investigated the production of glycosylated proteins in Hp using a combination of metabolic glycan labeling and mass spectrometry analysis. Metabolic labeling of glycans with unnatural sugars and subsequent enrichment has been applied to identify glycoproteins in eukaryotic systems (29, 56, 77). Though metabolic glycan labeling has been used to study bacterial glycolipids (78), to our knowledge this study is the first to demonstrate the applicability of this chemical approach to bacterial glycoproteomics (79).

Metabolic incorporation of azide-labeled sugars, followed by reaction with phosphine probes via Staudinger ligation, facilitates detection, visualization, and enrichment of cellular glycoproteins (29). We began our studies by assessing the subcellular distribution of Hp's azide-labeled glycoproteins to gain insight into their potential functions. Western blot data indicated that glycoproteins are present throughout Hp, including on the cell surface, embedded in both the inner and outer membranes, and within the periplasm and cytoplasm. The observed subcellular distribution suggests that protein glycosylation is abundant, widespread, and critical for Hp's physiology. Broadly, glycans modulate protein function in one of two ways: by providing epitopes for binding and recognition by other proteins (80, 81), and by stabilizing proteins (e.g. folding, solubility, protease susceptibility) (82–84). Hp's intracellular glycans are likely involved in stabilizing proteins, whereas those exposed to the extracellular milieu (surface and secreted glycoproteins) may stabilize proteins as well as modulate interactions with the host.

Based on our results, the subcellular distribution of glycoproteins in Hp appears to be broader than that observed in other bacteria to date. For example, in C. jejuni (85, 86), Neisseria sp. (20, 87), and Bacteroides sp. (27)., general protein glycosylation systems are present within the periplasm and modify proteins that traffic through this compartment (e.g. periplasmic, membrane-bound, and secreted proteins). In Actinobacillus, in contrast, there is a general protein glycosylation system within the cytosol that modifies cytosolic proteins (88). Finally, in Haemophilus influenzae, monosaccharides are added to proteins in the cytoplasm and then transported to the cell envelope (89). Future research will have to be conducted to confirm the localization of our candidate glycoproteins. If the observed, broad distribution is confirmed by follow-up experiments, this result suggests that Hp may have two general protein glycosylation systems (one within the cytosol and one within the periplasm) or an exclusively cytoplasmic glycosylation pathway that modifies proteins before sorting. Based on predictions by the carbohydrate-active enzymes (CAZy) database (90), there are 22 glycosyltransferases within Hp strain 26695, and half of these have unknown functions. Therefore, there are numerous glycosyltransferases within Hp that may be responsible for the synthesis of the observed glycoproteins. Characterizing which of these genes are responsible for carbohydrate assembly pathways is challenging because, in contrast to most bacteria, the genes for Hp carbohydrate biosynthesis pathways (e.g. O-antigen biosynthesis) are spread throughout the chromosome (91). Efforts are underway in our laboratory to identify the glycosyltransferases responsible for protein glycosylation in Hp.

We were excited to observe that a subset of Hp's azide-modified glycoproteins are present on the cell surface, as these surface-exposed glycoproteins and the glycans that modify them are potential therapeutic targets. For example, these glycosylated proteins could provide the basis for a carbohydrate-based vaccine (14). Unusual bacterial monosaccharides, such as di-N-acetylbacillosamine produced by Neisseria sp. (92), can themselves be immunogenic. These glycans, in the context of a glycosylated protein, may serve as particularly effective conjugate vaccines. Alternatively, glycosylation pathways could be targeted with small molecule inhibitors to interrupt interactions between Hp and its host. Finally, our demonstration that azide-labeled glycoproteins are present on the cell surface suggests that azide-covered Hp could be targeted with therapeutic phosphines. Covalent modification with properly designed phosphines could disrupt glycan function or render Hp innocuous within the host (93). Indeed, Kaewsapsak et al. designed phosphine therapeutics conjugated to the immune stimulant 2,4-dinitrophenyl to selectively kill Hp covered with azide-labeled glycans (55).

Staudinger ligation with Phos-FLAG-His₆ allowed for selective and efficient isolation of Hp's azide-labeled glycoproteins. MudPIT analyses of enriched azide-labeled proteins and a mock-enriched control enabled the unambiguous identification of 125 putative glycoproteins. We identified a large number of glycoprotein hits in this study relative to the small number (nine) identified in Creuzenet and coworkers' recent periodic acid/hydrazide-based screen of Hp's glycoproteins (15). We attribute much of this difference to the high incidence of background signal associated with periodic acid/hydrazide chemistry (15), rendering the identification of glycoprotein signal relative to nonglycoprotein noise difficult. In contrast, metabolic glycan labeling has a very high signal-to-noise ratio, enabling the facile detection and selective enrichment of glycoproteins. Thus, metabolic glycan labeling offers a complementary and efficient alternative to existing approaches for bacterial glycoprotein detection and identification.

In our studies, we detected Staudinger ligation-HexNAz adducts in beta-eliminated glycan samples from enriched azide-labeled glycoproteins, azide-labeled Hp, and purified azide-labeled urease. The exclusive presence of these adducts in samples from azide-labeled Hp provides evidence that the Staudinger ligation is exquisitely selective for azide-bearing molecules. Moreover, these results indicate that one metabolic fate of Ac₄GlcNAz in Hp is conversion to HexNAz. The addition of the Phos-FLAG moiety onto the HexNAz epitope greatly enhanced the chromatographic resolution, ionization, and detection of this glycan species. Optimization of the phosphine probe to contain isotopic tags or characteristic fragment ions will further facilitate glycoprotein identification and glycan characterization. Thus, MOE is a robust approach for glycoprotein discovery.

A large number of proteins with known links to colonization, persistence and virulence were identified as glycoprotein hits (Table I), and two of these colonization factors, ureA and ureB, were biochemically validated as glycoproteins in our studies. This finding is consistent both with previous studies indicating that flagellin glycosylation is important to Hp's ability to survive within the host (94), and more broadly, with the known links between protein glycosylation and virulence in a number of medically significant bacterial pathogens (13, 14). Further, this observed link underscores the importance of Hp's glycoproteins as potential therapeutic targets.

The discovery that the urease subunits are glycosylated has potential implications for the development of urease-based therapeutic strategies to prevent and treat Hp infection. Vaccines based on recombinant urease have had some success in preclinical (95, 96) and clinical studies (97, 98). However, these vaccines are based on recombinant proteins produced in Escherichia coli or Salmonella in their nonglycosylated state. New vaccines based on Hp's natively glycosylated urease have the potential to stimulate a robust immune response and thus protect patients more effectively than those based on recombinant urease.

In summary, we were able to use metabolic glycan labeling to enrich and identify 125 putative Hp glycoproteins. Further, we validated the glycosylation status of two of these hits. Finally, we characterized the resulting glycan adducts. These results reveal that glycosylated proteins are abundant in Hp, indicate that Hp's glycoproteins have a wide range of functions, and reveal new links between Hp's pathogenesis and glycosylation. Armed with this information, the stage is set for the development of glycosylation-based therapeutic strategies to eradicate Hp infection. In particular, this work opens the door to phosphine-based therapeutics that target Hp's azide-labeled glycans.

Broadly, this work validates MOE as a viable approach to discover and characterize bacterial glycoproteins. In addition to enabling the discovery of Hp's glycoproteins, MOE has the potential to facilitate the study of glycoproteins in other bacteria. Indeed, reports of metabolic labeling of glycoproteins with azide-modified pseudaminic acid in C. jejuni (54) and with alkyne-bearing fucosylated sugars in Bacteroidales sp. (99) suggest that this glycoproteomic approach should transfer readily to these organisms. The ability to use a general metabolic precursor rather than a specific glycan-binding reagent will facilitate the detection and discovery of glycoproteins in a broad range of bacteria. Thus, MOE will propel bacterial glycoproteomics forward and, in the process, unveil novel therapeutic targets.