N-Glycosylation of Pig Flavin-Containing Monooxygenase Form 1

Chem. Res. Toxicol. 1998, 11, 1145-1153

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N-Glycosylation of Pig Flavin-Containing Monooxygenase Form 1: Determination of the Site of Protein Modification by Mass Spectrometry Katy K. Korsmeyer,† Shengheng Guan,‡ Zi-Cheng Yang,§ Arnold M. Falick,| Daniel M. Ziegler,⊥ and John R. Cashman*,@ Department of Pharmacology and Pharmaceutical Chemistry, University of California, San Francisco, California 94143, National High Field Magnetic Laboratory, 1800 East Paul Dirac Drive, Tallahasee, Florida 32310, Sugen Inc., 351 Galveston Drive, Redwood City, California 94063, Perseptive Biosystems, 871 Dubuque Avenue, South San Francisco, California 94080, Department of Chemistry, Clayton Foundation for Biochemistry, ESB 439, University of Texas, Austin, Texas 78712, and Human BioMolecular Research Institute, 5310 Eastgate Mall, San Diego, California 92121 Received May 26, 1998

By using a combination of biochemical methods (i.e., endoglycosidase H digestion and immunoblot and plant lectin binding studies), it was verified that pig flavin-containing monooxygenase (FMO1) was N-glycosylated. By using mass spectrometry approaches [i.e., peptide mapping, gas chromatography/mass spectrometry, microbore HPLC/electrospray ionization mass spectrometry (LC/ESI/MS), chemical ionization gas chromatography/mass spectrometry (CI/GC/MS), and matrix-assisted laser desorption mass spectrometry (MALDI/ MS)], we were able to confirm that pig FMO1 was N-glycosylated and we were able to identify the site of N-glycosylation. Pig FMO1 contains two putative consensus sites of N-glycosylation. The results showed that pig FMO1 amino acid Asn120 was selectively N-glycosylated. Highly purified pig FMO1 avidly bound concanavalin A and reacted positively for carbohydrates by the periodic acid/Schiff’s base method of analysis. In addition, treatment of pig FMO1 with endo-N-acetylglucosaminidase converted the enzyme to another species with a molecular mass approximately 5000 Da lower than that of the parent protein as determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblot experiments. Peptide mapping of pig FMO1 showed that the protein used in the study was not contaminated with another glycoprotein. MALDI/MS experiments showed that pig FMO1 was present with the expected molecular mass but that higher-molecular mass forms consistent with the presence of N-linked high-mannose oligosaccharide structures were also covalently attached to the enzyme. The presence of N-acetylglucosamine isolated from acid hydrolysates of the N-linked high-mannose oligosaccharide of pig FMO1 was confirmed by high-pH anion exchange HPLC studies and verified by CI/GC/MS studies of derivatized monosaccharide fractions. Further analysis of pig FMO1 proteolytic peptides by LC/ESI/MS showed that the only residue that was N-glycosylated in pig FMO1 was Asn120. Knowledge of the structural aspects of FMO may be useful in understanding the membrane association properties of the enzyme.

Introduction The membrane-associated mammalian flavin-containing monooxygenase (FMO,1 EC 1.14.13.8) catalyzes the NADPH-dependent conversion of nitrogen-, sulfur-, phosphorus-, and other heteroatom-containing drugs, chemicals, and pesticides to their corresponding oxygenated metabolites. Oxygenation of nucleophilic xenobiotics is thought to provide a way for animals and humans to detoxicate chemicals to polar metabolites that are readily * To whom correspondence should be addressed. Telephone: (206) 270-0394. Fax: (206) 270-0385. E-mail: [email protected]. † University of California. ‡ National High Field Magnetic Laboratory. § Sugen Inc. | Perseptive Biosystems. ⊥ University of Texas. @ Human BioMolecular Research Institute. 1 FMO1 represents flavin-containing monooxygenase form 1 (the nomenclature for FMO has been revised; see ref 1).

excreted (1). For example, the pyrolizidine alkaloid plant toxins senecionine, retrorsine, and monocrotaline are efficiently detoxicated by tertiary amine N-oxide formation (2) in a species such as the guinea pig that has a relatively high level of FMO activity and a low level of pyrrole-forming cytochrome P450 (CYP)2 activity (3, 4). N-Oxygenation of another plant alkaloid, (S)-nicotine, probably constitutes a detoxication route in animals and humans as well, shunting the alkaloid through the 2 Abbreviations: CYP, cytochrome P450; HPAE, high-pH anion exchange chromatography; CID, collision-induced dissociation; LC/ESI/ MS, high-performance liquid chromatography/electrospray ionization/ mass spectrometry; HPLC, high-performance liquid chromatography; MALDI/MS, matrix-assisted laser desorption/mass spectrometry; LSIMS, liquid secondary ion mass spectrometry; CI/GC/MS, chemical ionization/gas chromatography/mass spectrometry; BPI, base peak ion; GluC, endoproteinase Glu-C; TRIS, tris(hydroxymethyl)aminomethane; TFA, trifluoroacetic acid; Endo H, endoglycosidase H; HNAT, HEPES/ NaCl/BSA/Tween-20; GlcNAc, N-acetylglucosamine; GlcNH2, glucosamine; GalNH2, galactosamine; Gal, galactose; Man, mannose.

10.1021/tx980117p CCC: $15.00 © 1998 American Chemical Society Published on Web 09/02/1998

1146 Chem. Res. Toxicol., Vol. 11, No. 10, 1998

detoxication pathway leading to the tertiary amine nicotine N-1′-oxide (5). In animals such as pigs, hepatic FMO1 presumably constitutes a major hepatic detoxication catalyst. In human liver, another form of the enzyme, FMO3, apparently is the principal FMO form present (6). However, because of adult human liver enzyme thermal instability, pig FMO1 is the hepatic enzyme that has been most thoroughly studied. Extensive enzyme and physical studies of pig FMO1 have described the kinetic, mechanistic, and product distribution aspects of the enzyme (7). The enzyme is thought to stabilize the 4a-hydroperoxyflavin intermediate that is formed in the presence of molecular oxygen after the flavoprotein has been reduced by NADPH (8, 9). Pig FMO1 is an extremely hydrophobic protein that is sensitive to anionic detergents, has a pH optimum of 8.5, and is rapidly inactivated if placed above 40 °C in the absence of NADPH. Few structural studies of pig FMO1 have been attempted, but preliminary carbohydrate analysis has shown that the enzyme contains complex oligosaccharides linked to high-mannose oligosaccharides (10). The membrane-bound form of the enzyme has been purified to homogeneity, and the properties of the enzyme have been reviewed (6, 7). The primary amino acid sequence of pig FMO1 has been determined by two groups (11, 12). In addition, genes for the human, rabbit, and rat FMO1 orthologues have also been cloned (13). Cloned pig liver (12) and rabbit lung (14) FMO cDNA have been expressed in Escherichia coli. Because both the native and cDNAexpressed pig FMO1 enzyme are active, determination of posttranslational modification of the two enzymes may provide information about the three-dimensional structure of the enzyme. Sequences derived from pig FMO1 cDNA data predict two putative consensus sites of N-glycosylation. However, the nature of posttranslational modifications of pig FMO1 cannot be determined from the cDNA data alone. Gas-phase sequencing of peptides from proteolytic digests has provided some direct amino acid sequence data for rabbit FMO forms (15), and Edman sequencing of rabbit FMOs has shown some microheterogeneity (16). Even though FMO enzymes are often blocked at the N-terminal amino acid that confounds traditional sequence analysis, mass spectrometric analysis of important selected peptide fragments of pig FMO1 has shown that the N terminus was acetylated and verified the existence of FAD- and NADP+-binding domains (17). The membrane topology of pig FMO1 is not known, but studies with truncated forms of rabbit FMO2 have shown that the lipophilic C terminus is not essential for membrane association (14). In the absence of strong evidence for a signal peptide sequence and because a cofactorbinding domain is adjacent to the N terminus, it is unlikely that the N terminus is essential for transmembrane insertion. Presumably, a central portion of pig FMO1 encodes information for the determination of membrane association. Association of the protein in the endoplasmic reticulum membrane could in principle be determined by an identification of the site of N-glycosylation because of the luminal localization of the core glycosylation enzyme system required for N-glycosylation (18). Thus, determination of the site of N-glycosylation could in principle indicate a putative region of Nglycosylation and support the construction of a molecular model of pig FMO1. Aside from the important and

Korsmeyer et al.

interesting structural biological aspects of the multiple forms of FMO, knowledge of the FMO structure and membrane association could be helpful in designing drugs that are not highly efficiently metabolized by FMO. Rapid clearance by the liver is more often than not an unfavorable characteristic for drugs because it can result in poor oral bioavailability, high variability, and short duration of exposure due to a short half-life. Rapid clearance often does not support a drug with once-perday dosing and requires a wide therapeutic index to cope with the likely high drug level variability. In this study, we purified pig FMO1 by a variety of methods and subjected the highly purified preparations to mass spectrometric and other means of physical and biochemical analysis. The pig FMO1 enzyme contains a high-mannose oligosaccharide covalently attached to Asn120. The precise location of N-glycosylation and verification of the structure of the oligosaccharide provide structural information useful for constructing an overall model of pig FMO1.

Experimental Procedures Chemicals. Chemicals used in this study were of the highest purity available and were purchased from the following companies: urea, sodium borohydride, sodium borodeuteride, perchloric acid, and NH4HCO3 from Aldrich (Milwaukee, WI); dithiothreitol, sodium iodoacetate, sodium dodecyl sulfate, and endoproteinase Glu-C from U.S. Biochemicals (San Diego, CA); and trypsin, monosaccharides, water, and acetic acid from Pierce (Rockford, IL). Endoglycosidase H was obtained from Boehringer Mannheim (Groton, CT). Instrumental Analysis. Liquid chromatography/electrospray ionization mass spectrometry (LC/ESI/MS) was carried out on a VG Biotech/Fisons BIOQ mass spectrometer equipped with an electrospray source. Chromatography was performed on an Aquapore 300 reversed-phase microbore column (100 mm × 1.0 mm i.d., Applied Biosystems) that was interfaced by fused silica capillary tubing to the mass spectrometer. The mobile phase was driven by a dual syringe pump (Carlo-Erba Phoenix 20) at a flow rate of 40 µL/min. The mobile phase HPLC program was a gradient from 0.1% trifluoroacetic acid in water to trifluoroacetic acid/water/acetonitrile (0.08:42:58, v/v) with a linear increase at a rate of 0.5%/min. The column effluent was monitored by an Applied Systems 783A variable-wavelength UV detector equipped with a high-sensitivity capillary flow cell (LC Packings) set at 215 nm. Microbore HPLC separations with the water/acetonitrile/trifluoroacetic acid eluent required postcolumn addition of 2-propanol/2-methoxyethanol (1:1) at a flow rate of 5 µL/min via a mixing tee to optimize MS detection. Approximately 5% of the HPLC effluent flowed directly into the mass spectrometer. Generally, the operating voltages for the LC/ESI/MS experiments were as follows: probe tip, 4200 V; counter electrode, 550 V; and sampling orifice, 50 V. The source temperature was held at a constant 60 °C. The mass spectrometer was scanned over a range of m/z 300-2000 at 7 s/scan. Liquid secondary ion mass spectrometry (LSIMS) was carried out on a Kratos MS-50S double-focusing mass spectrometer from Kratos Analytical Instuments (Manchester, U.K.) equipped with a high-field magnet (mass range m/z ) 3000 at an acceleration voltage of 8 kV), a Cs+ liquid secondary ion mass spectrometry source, and a cooled sample introduction probe (19, 20). Tandem mass spectrometry experiments were performed on a Concept II HH four-sector EBEB tandem mass spectrometer fitted with an electrooptical array detector, a Cs+ ion LSIMS source, and a cooled sample probe (21). Precursor ions were generated with a 10 kV Cs+ primary ion beam. The collision energy for collisioninduced dissociaton was 4 kV. The collision gas (He) was used at a pressure sufficient to suppress the precursor ion beam to

FMO1 N-Glycosylation Determined by Mass Spectrometry

Chem. Res. Toxicol., Vol. 11, No. 10, 1998 1147

about 30% of its initial level. The system was controlled and the data were acquired with a DS-90 data system. Data reduction and display were carried out on a Mach 3 data system. Matrix-assisted laser desorption ionization (MALDI) mass spectrometry was done with a Canyon Creek, Texas/Linear Instruments MALDI time-of-flight mass spectrometer (Linear Instruments) operating at a 30 kV accelerating potential in the negative ion mode. Ions emitted from the source were mass selected, and mass spectra were generated from the sum of 2050 laser shots. To a concentrated preparation of pig FMO1 (10 pmol) was added 1 µL of a sinapinic acid matrix solution (50 mM, 1:1 acetonitrile/0.1% TFA in H2O), and the resulting solution was spotted onto a stainless steel sample slide and allowed to evaporate. Arrival time distributions were obtained at the detector after passage through the mass filter. Chemical ionization gas chromatography/mass spectrometry (CI/GC/MS) was carried out with a VG7OS spectrometer fitted with a Varian model 3600 gas chromatograph. A fused DB-1 capillary column (30 m × 0.25 mm i.d., film thickness of 1 µm) was used, and the carrier gas was helium. The linear temperature program was started at 30 °C and increased to 300 °C at 4 °C/min. Ammonia was used as the reactant gas for CI/GC/ MS. Enzyme Purification. FMO1 was purified from pig liver microsomes by two different procedures as previously described (22, 23), and as discussed below, no significant differences were observed for either preparation. Enzyme was prepared and stored at -70 °C in small aliquots (pH 7.6). Enzyme used in the studies had specific activities of 583-1083 nmol of 10-[(N,Ndimethylamino)pentyl]-2-(trifluoromethyl)phenothiazine tertiary amine N-oxide formed min-1 mg of protein-1 (12) or at least 800 nmol of dimethylaniline N-oxide formed min-1 mg of protein-1 (7). Enzyme Assay. The pig FMO1 enzyme activity was measured by determining products by HPLC (12) or by monitoring oxygen consumption polarographically with an oxygen electrode as described previously (23). The protein concentration was determined by the BCA protein assay from Pierce Chemical Co. (Rockford, IL). As described below, both preparations were shown to be extensively glycosylated. Protein Studies. Pig FMO1 was isolated and purified from pig liver microsomes by two methods described above, and each preparation was judged to be homogeneous by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (24). At least six different preparations were used in the following experiments, and no significant differences in the monosaccharide composition or the mass spectral properties of the pig FMO1-derived glycopeptide materials were observed. For peptide mapping studies, highly purified pig liver FMO1 (0.06 mg, 1 nmol) was dissolved in 0.2 mL of denaturing buffer [0.1 M NH4HCO3 (pH 8.2), 8M urea, and 1 mM EDTA]. Dithiothreitol (100 µg, 0.65 µmol) was added, and the sample was purged with an atmosphere of argon and incubated at 37 °C for 1 h. After the disulfides were reduced, sodium iodoacetate (650 mg, 30 nmol) was added and the mixture incubated for 1 h at 37 °C, after which time an additional aliquot of sodium iodoacetate (65.0 mg, 3 nmol) was added and the incubation was continued for 30 min. The reaction was stopped by the addition of 0.2 mL of cold acetone. After sonicaton and vigorous mixing, carboxymethylated pig FMO1 was diluted with a 2-fold volume of 50 mM potassium phosphate buffer (pH 7.8). Endoproteinase Glu-C (5% w/w) was added and the mixture incubated at 37 °C for 5 h followed by an additional aliquot of Glu-C (2% w/w) and the mixture incubated for an additional 12 h at 37 °C. For pig FMO1 digestion by trypsin, the above FMO disulfide reduction and carboxymethylation protocol was followed, 1 µg of trypsin was added to 1 nmol of FMO1, and the sample was incubated at 37 °C for 2 h. For Endo H digests, 1 nmol of pig FMO1 was denatured by heating in the presence of sample buffer [63 mM Tris, 1% SDS, and 0.1 M DTT (pH 6.8)] for 3 min. After the mixture cooled and 6 volumes of potassium phosphate buffer (pH 6) was added, Endo H was added to achieve a final

concentration of 1 milliunit/µL and the mixture incubated for 20 h at 37 °C. Saccharide Analyses. For saccharide analysis, high-pH anion exchange chromatography (HPAE) of acid hydrolysates of pig FMO1 were carried out. Pig FMO1 that was purified by HPLC was used for determination of neutral and amino sugars. The presence of sialic acids was directly determined by evaluating aqueous extracts of acid hydrolysates of highly purified pig FMO1 by GC/MS analysis. Approximately 0.06 mg or 1 nmol of pig FMO1 was placed in (a) 400 µL of 2 M trifluoroacetic acid and the mixture incubated at 100 °C for 3 h (i.e., for neutral sugars), (b) 400 µL of 6 N hydrochloric acid and the mixture incubated at 100 °C for 4 h (i.e., for amino sugars), or (c) 400 µL of 0.1 N hydrochloric acid and the mixture incubated at 80 °C for 4 h (i.e., for sialic acids). The hydrolysate was lyophilized and reconstituted with 200 µL of water, and 150 µL was injected onto a Dionex model BioLC apparatus equipped with a Carbopac AS-6 pellicular anion exchange column, a gradient pump reagent delivery module, and a pulsed amperometric detector. HPLC runs were carried out with a 60 min solvent gradient and a 90 min cycle time under conditions that readily separated monosaccharides (25). For GC/MS analysis of saccharides, samples from Dionex BioLC peaks or hydrolysate samples were directly placed in 0.3 mL Reacti-Vials (Pierce), and a freshly prepared solution of sodium borohydride (or sodium borodeuteride) was added and the mixture allowed to stand overnight at room temperature. At the end of the reaction, acetic acid was added and the entire mixture was dried in vacuo. To the dried alditol were added excess acetic anhydride, acetic acid, and perchloric acid, and the mixture was sonicated and allowed to stand for 8 min at room temperature. Next, 200 µL of water was added and the mixture vigorously mixed, extracted with dichloromethane, and centrifuged to separate the layers. The dichloromethane fraction was reduced in volume and directly injected onto the GC or GC/MS instrument. A reference mixture of n-alkanes (i.e., C6-C36) was injected simultaneously as a marker with each derivatized monosaccharide sample that was analyzed by gas chromatography to provide structure-retention index relationships for derivatized monosaccharides arising from pig FMO1 acid hydrolysates (26). Peptide and Protein Separation and Analysis. Pig FMO1 (i.e., 1-4 nmol) was purified by RP-HPLC on a Rainin model HPX instrument (Emeryville, CA) controlled by a Macintosh SE computer using a Vydac C4 column (4.6 mm × 25 cm) fitted with a C4 precolumn and UV detection at 215 nm. The chromatogram was developed at a flow rate of 1 mL/min by a 90 min linear gradient from 30 to 100% solvent B, where solvent A was 0.1% trifluoroacetic acid in water and solvent B was 0.08% trifluoroacetic acid in acetonitrile. Separation from some minor contaminants was achieved using a 1%/min linear gradient, and pig FMO1 consistently eluted at 57% acetonitrile. The HPLC fractions were individually hand-collected in Eppendorf tubes and lyophilized prior to SDS-PAGE or mass spectrometric analysis. For samples containing peptides or glycopeptides, the RPHPLC chromatograph was recorded on a Vydac C4 column (4.6 mm × 25 cm) at a flow rate of 1%/min by a linear gradient (i.e., from 0 to 60%) of solvent B, where solvent A was 0.1% trifluoroacetic acid in water and solvent B was 0.08% trifluoracetic acid in acetonitrile. Microbore HPLC used a solvent system of 0.1% trifluoracetic acid/water to 42% acetonitrile/ 0.08% trifluoroacetic acid/water at a flow rate of 0.5%/min. A dual syringe pump (Applied Biosystems) was used to deliver the mobile phase at a flow rate of 40 µL/min. Antisera and Immunoblotting. Antibody to pig FMO1 was raised in rabbits as described previously (12). Immunoblots were created according to the method previously described (27). Proteins that were fractionated by SDS-PAGE were transferred electrophoretically to nitrocellulose. After transfer, membranes were placed overnight in blocking buffer that consisted of 1.5% casein, 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic


Korsmeyer et al.

acid (HEPES), 150 mM NaCl, 0.02% BSA, and 0.005% Tween20 (HNAT) at pH 7.6. After three washes, rabbit anti-pig FMO1 was used in a 1:1000 dilution of HNAT for 1 h at room temperature. The second antibody was a goat anti-rabbit IgG conjugated to horseradish peroxidase and was used at a 1:3000 dilution af HNAT and incubated for 1 h. Immunoblotting for concanavalin A binding was done with rabbit anti-concanavalin A antibody diluted 1:1000 for an incubation of 1 h. Both immunoblots that were visualized by color development were created with 4-chloro-1-naphthol in methanol and hydrogen peroxide. For a typical Endo H digestion, 195 µg of pig FMO1 was placed in 50 mM potassium phosphate buffer (pH 6) in the presence of 1 mM dithiothreitol and 50 µg of Endo H. The digestion mixture was incubated at 37 °C for 20 h, and the sample was analyzed for concanavalin A binding and, separately, with rabbit anti-pig FMO1 antibody.

Results Pig FMO1 was isolated and purified to apparent electrophoretic homogeneity by standard procedures. The enzyme was highly purified on the basis that the criteria of SDS-PAGE, immunoreactivity, HPLC, and LC/ESI/ MS analysis showed a single protein. The enzyme was subjected to limited Edman automated sequencing. No useful sequence data were obtained, confirming the previous observation that pig FMO1 was N-terminally acetylated (17). Pig FMO1 was then subjected to extensive physical characterization, including (a) immunoblotting and (b) mass spectrometric studies of N-linked high-mannose oligosaccharides, peptides, and glycopeptides. Immunoblotting. Parallel samples of pig FMO1 were subjected to SDS-PAGE and transferred by electroblotting onto two nitrocellulose membranes. The gel indicated a single protein with an apparent molecular mass of approximately 59 kDa. One of the nitrocellulose membranes was treated with the plant lectin, concanavalin A, and the other was treated with rabbit anti-pig FMO1 antibody. As described previously, the rabbit antipig FMO1 antibody showed clear binding (10). The binding of concanavalin A was also apparent in the other nitrocellulose membrane in the region corresponding to pig FMO1. The results supported the suggestion that high-mannose oligosaccharides were linked to highly purified pig FMO1. These data were consistent with observations that pig FMO1 that fractionated on SDSPAGE stained positive for carbohydrates by the periodic acid/Schiff’s base and the Thymol-sulfate methods (10, 28). In addition, SDS-PAGE fractionation of a sample of pig FMO1 treated with endoglycosidase H (i.e., an enzyme that hydrolyzes N-linked high-mannose oligosaccharides) indicated a decrease in the apparent molecular mass of pig FMO1 of approximately 5 kDa and showed a new protein with an apparent molecular mass of approximately 54 kDa. Pig FMO1 was purified by collecting the immunopositive fraction with a retention time at 57% acetonitrile by repetitive injections on HPLC. The collected fractions were dried to a minimum volume and used immediately or stored at -20 °C for further studies. Under standard conditions (see Experimental Procedures), pig FMO1 eluted with a retention time of 57.2 ( 0.2 min (n ) 8) and ovalbumin (i.e., another glycosylated protein standard) eluted with retention times of 49.4 ( 0.1 and 49.9 ( 0.1 min (n ) 6). Analysis for Amino Sugars. Acid hydrolysis of the HPLC-purified pig FMO1 protein was carried out in

Figure 1. Monosaccharide analysis of acid hydrolysates of pig FMO1 by Dionex HPAE chromatograms with pulsed amperometric detection. Chromatograms of acid hydrolysates were aligned with that of monosaccharide standards (A), a trifluoroacetic acid control sample (B), and a hydrochloric acid control sample (C) that did not contain protein. Pig FMO1 was hydrolyzed in the presence of trifluoroacetic acid (D) that provided evidence for neutral monosaccharides and in the presence of hydrochloric acid that provided evidence for amino sugars (E). The standard monosaccharides included the following: Fuc, fucose; GalNH2, galactosamine; GlcNH2, glucosamine; Glc, glucose; and Man, mannose.

parallel with trifluoroacetic acid (2 M) and hydrochloric acid (6 N) to evaluate the presence of neutral and amino sugars, respectively. Although it was likely that the conditions used for purification of the enzyme hydrolyzed sialic acids, on the basis of the preliminary immunoblot data, we did not anticipate sialic acids being present in the N-linked high-mannose oligosaccharides covalently attached to pig FMO1. Figure 1 shows the Dionex HPLC chromatograms of acid hydrolysates of highly purified pig FMO1 detected with high-pH anion exchange (HPAE) chromatography coupled with pulsed amperometric detection. A number of HPLC peaks were observed, and the most prominent peak coeluted with glucose (probably arising as a contaminant from the sucrose used in the original enzyme purification). Most notable were peaks that coeluted with N-acetylglucosamine and mannose. Acid hydrolysis of HPLC fractions that were collected eluting slightly before or after pig FMO1 showed no evidence of the presence of N-acetylglucosamine or mannose as determined by HPAE chromatography. As shown in Figure 1, control samples containing only HCl or trifluoroacetic acid contained essentially no detectable monosaccharides or only a small amount of glucose, respectively. Pig FMO1 that was extensively purified by HPLC showed an even lower glucose level, and the more



extensively the enzyme preparation was chromatographed, the smaller the amount of glucose observed by HPAE chromatography. As described above, it is likely that glucose is simply a coeluting monosaccharide impurity that arises from the copious amounts of sucrose used in the original enzyme purification. To confirm the chemical identity of the monosaccharides, the HPAE HPLC peaks of the putative monosaccharides were collected, neutralized, dried, reduced with sodium borodeuteride, and acetylated for analysis by gas chromatography/mass spectrometry. The derivatization procedure was carried out in a single reaction vessel, and the loss of the derivatized monosaccharides was thereby minimized (26). Because of the relative abundance and purity of the HPAE fractions, we were only successful in determining the structure of N-acetylglucosamine residues by GC/MS. The derivatized [2H]glucitolamine hexaacetate (expected molecular weight of 435, M + H+) possessed characteristic molecular ions and fragmentation for the expected product. The GC/MS spectrum of the derivatized material [i.e., m/z (relative abundance): 435 (100), 420 (7), 375 (58), 324 (17), 141 (37), 97 (47), 95 (54), 83 (62)] obtained from the extract of the Dionex chromatography peak was essentially identical to the spectrum of authentic derivatized [2H]glucitolamine hexaacetate [i.e., 435 (100), 420 (4), 375 (85), 324 (20), 141 (57), 97 (56), 83 (69)]. In addition, the retention index of the derivatized unknown was essentially identical to that of authentic derivatized glucitolamine as analyzed by gas chromatography (26). Pig FMO1 Peptide Map Developed by Mass Spectrometry. Because we wanted to rule out the possibility that a contaminating protein tightly complexed to pig FMO1 was providing the oligosaccharide moiety observed and because we wanted to verify the pig FMO1 amino acid sequence with the pig FMO1 sequence deduced from the cDNA data, we determined the primary sequence of highly purified pig FMO1 with peptide mapping by mass spectrometry. We also sequenced selected peptides of the protein digest by tandem MS to confirm the primary amino acid sequence deduced from the cDNA data as well as to examine the structure of important peptide digest fragments. Highly purified pig FMO1 was subjected to reduction and carboxymethylation. The carboxymethylated protein was digested with trypsin, endoproteinase Glu-C, or a mixture of both proteolytic enzymes. The proteolytic enzyme digest of pig FMO1 was run out on HPLC as described previously (17), and a total of approximately 90 fractions were collected, evaporated to dryness, and prepared for LSIMS. Each HPLC fraction examined generally contained a number of peptides as judged by the number of molecular ions in the LSIMS spectra. The molecular masses of the peptides in the HPLC fractions were determined by LSIMS and compared with those of the peptides predicted to be present. Some of the fractions required derivatization with hexanol (29) to give useful peptide molecular ions. The measured molecular masses of the peptides from the LSIMS spectra were compared with the peptide molecular masses predicted from the proteolytic cleavage of the protein sequence deduced from the cDNA data. Figure 2 shows the resulting peptide molecular mass map for pig FMO1. Peaks corresponding to the expected molecular masses of approximately 85% of the anticipated peptides were found by LSIMS. The molecular masses of peptides that correponded to extremely lipo-

philic peptides in some cases could not be determined. To confirm the identity of the peptides determined by LSIMS, the amino acid sequence of selected peptides from the molecular ions of the LSIMS experiments was determined by tandem mass spectrometry. The collisioninduced dissociation (CID) spectrum of the protonated peptide ion of selected samples provided amino acid sequence data that were consistent with the sequence deduced from the cDNA data. In every case examined, no evidence for the presence of detectable amounts of nonpig FMO1 peptides was observed, suggesting that no other proteins were present in the highly purified pig FMO1 examined. It is notable that the two possible consensus sequence sites of N-glycosylation located at Asn120 and Asn314 (Figure 3) were determined to be unmodified as shown by LSIMS analysis. However, this is not surprising in view of the large molecular mass and the anticipated low ionization potential of the expected glycopeptide derived from proteolytic cleavage of pig FMO1. Generally, LSIMS does not work well for peptides with a molecular mass of >3000 Da, and the technique does not work particularly well for glycopeptides. These data also suggest that N-glycosylation is probably not complete in the native protein. As described below, we resequenced pig FMO1 proteolytic peptides with LC/ESI/MS, and this technique enabled us to observe the intact glycopeptide. Determination of the Pig FMO1 Molecular Mass. Preliminary negative ion laser desorption mass spectrometry experiments confirmed the expected molecular mass information for pig FMO1 but also showed the presence of higher-molecular mass forms. A typical MALDI mass spectrum of pig FMO1 present in a sinapinic matrix showed a series of peaks centered near a mass of 58 772.3 Da. Of note was the obvious deviation from the peak center (i.e., shoulders) apparent in the mass spectrum that varied from a symmetrical peak shape anticipated for a homogeneous protein. While the molecular mass calculated from the pig FMO1 cDNA data for the protein was similar to the observed mass of 58 772.3 Da (11, 12), the observation of higher-molecular mass species was consistent with the presence of Nlinked high-mannose oligosaccharide structures covalently attached to pig FMO1. The individual glycopeptide linkage was determined, as described below. Pig FMO1 Glycopeptide Linkage. For the determination of the site of glycopeptide linkage, microbore LC/ESI/MS was employed. Peptide maps were generated by reversed-phase HPLC after digestion with trypsin and endoprotease Glu-C. A typical proteolytic digest of pig FMO1 has been published previously (17), and we have found that our more recent digests are more complete and provide a larger number of identifiable peptides by LC/ESI/MS. Endoprotease Glu-C digests of pig FMO1 were analyzed by microbore LC/ESI/MS, and analysis of pig FMO1 peptides by LC/ESI/MS confirmed about 90% of the primary amino acid sequence. Several fractions contained ions whose molecular masses could not be assigned to simple pig FMO1 peptides arising from Glu-C digestion. Because we previously established that no other contaminating proteins were present in the pig FMO1 that was analyzed, we assumed that the nonassignable peptides were glycopeptides. To avoid the task of sequencing every nonassignable proteolytic fragment by tandem mass spectrometry, we searched for the hypothetical high-mannose glycopeptides shown in Fig-


Korsmeyer et al.

Figure 2. Amino acid sequence of pig FMO1. Tryptic (T)- or endoprotease Glu-C (V)-generated peptides observed with LSIMS detection are shown as underlined in the peptide map. Peptides identified by tandem MS analysis of pig FMO1 proteolytic digests are labeled with a bold underline.

Figure 3. Amino acid sequence of pig FMO1 showing two consensus N-glycosylation sites at Asn120 and Asn314. The asterisks indicate the consensus sites of N-glycosylation.

ure 4 by a computer program algorithm, and examined the calculated versus observed m/z values obtained in LC/ ESI/MS experiments. Figure 5 shows the base peak ion (BPI) chromatogram of the Glu-C digest as measured by the electrospray mass spectrometer. The UV chromatogram (215 nm) has been shown previously (17), and because the reconstructed ion chromatogram was dominated by a prominent detergent peak, the chromatograms are not shown. The chromatographic run (acquisition duration of 80 min) and mass range were divided into sectors as typified by the results shown in Figure 5 to better identify the related glycopeptides. The BPI chromatogram of Figure 5, for example, was evaluated for specific ions (i.e., peptide + glycan core + seven or eight mannose residues) by running the data through a computer program algorithm. The ESI/MS analysis showed peaks that corresponded to the predicted masses for the Asn120 glycopeptide that possessed an N-glycan core and a series of seven or eight mannose residues attached. HPLC fractions corresponding to Asn314 glycopeptides containing high-mannose glycans at the predicted masses attached to peptides were not observed. Peaks corre-

sponding to predicted masses for the glycopeptide of Asn120 with more than eight or less that seven mannose residues were also likewise not observed. A schematic structure of the Asn120 glycopeptide containing the oligosaccharide core structure is shown in Figure 4. The calculated m/z values for the Asn120 glycopeptide core with seven mannose residues is 2996.83 Da, and the calculated m/z value for the Asn120 glycopeptide core with eight mannose residues is 3158.97 Da. Figure 5 shows the LC/ESI/MS mass chromatograms of peaks from the glycopeptides. The mass chromatograms indicate the relative abundance of ions having the specified m/z ((1 Da). For example, Figure 6 shows the ESI/MS chromatogram of the peptide eluting from the BPI chromatogram of Figure 5. LC/ESI/MS mass chromatograms of predicted masses for the corresponding highmannose glycopeptides of Asn314 were not observed. After putative glycopeptides were subjected to the computer-driven algorithm, it was apparent that scans near 54.051 min of a typical endoprotease Glu-C digest provided the desired data. Figure 6 clearly showed the expected ESI mass ions (i.e., number of charges equals four) for a glycopeptide with seven mannose residues attached to a peptide arising from proteolytic cleavage of pig FMO1 at Asn120. Likewise, prominent ESI mass ions of 1499.4, 999.9, 632.8, 527.5, and 452.5 were observed in accordance with the anticipated precursor-



Figure 6. Representative electrospray mass spectra of native pig FMO1 peptides generated from a Glu-C digest containing the N-acetylglucosamine core and seven mannose residues as predicted in the analysis of Figure 4 (i.e., m/z 750.6 observed, m/z 750.2 predicted). Scans 491-523 were combined to give the MS spectrum that is shown.

Figure 4. Schematic showing the proposed glycan core of pig FMO1 attached to Asn120 of the eight-amino acid peptide arising from protease digestion (A). The m/z values for the peptide and core structure and seven mannose or eight mannose residues are 2996.83 and 3158.97 Da, respectively. The calculated m/z values for the glycopeptide shown in panel A with an N-acetylglucosamine core and either seven or eight mannose residues is listed (B). The observed values are highlighted with an asterisk.

Figure 5. LC/ESI/MS mass chromatograms of peaks from glycopeptides containing the glycan core and seven mannose residues (main figure) and eight mannose residues (inset). Mass chromatograms indicate the relative abundance of ions having the specified m/z ((1). The base peak ion (BPI) is the relative abundance of the largest peak in each mass spectrum. The UV215 chromatogram (40) and the reconstructed ion chromatogram were dominated by a prominent detergent peak, and the data are not shown.

fragment associations characteristic of the expected glycopeptide of Asn120.

Discussion In the absence of an X-ray structure for FMO, stuctural data can also help to validate a molecular model. In addition, structural information may provide details about how FMO is related to other flavoproteins or other monooxygenases, and these data may provide insight into the physiological function of this important family of enzymes. Analysis of pig FMO1 acid hydrolysates by HPAE chromatography and GC/MS analysis of derivatives

confirmed the presence of N-acetylglucosamine and indicated the presence of mannose (Figure 1). That the glycopeptide observed did not arise from the presence of a minor amount of another contaminating glycoprotein was confirmed by the lack of peptides from any other source after sequencing pig FMO1 by both LSIMS and ESI/MS techniques. In addition to laser desorption mass spectrometry that showed pig FMO1 was comprised of multiple species, ESI/MS of Glu-C peptide digests showed peaks that corresponded to the predicted masses for the Asn120 glycopeptide that possessed an N-glycan core and a series of seven or eight attached mannose residues. Of the two putative consensus sequences for posttranslational N-glycosylation, evidence for modification of only one site was observed. We did not find any evidence that Asn314 was modified by an N-linked high-mannose moiety. The results show that N-glycosylation is specific and not random and may suggest some important structural purpose for the N-glycosylation of pig FMO1. Analysis of tryptic and Glu-C digests of pig FMO1 has provided molecular mass data used to construct molecular mass maps (Figure 2). Appoximately 95% of the primary sequence was confirmed. The N terminus of pig FMO1 is N-acetylated as previously determined by tandem mass spectrometry sequencing of the N-terminal peptide (17). Nevertheless, some amino acid sequence data have been obtained for FMO1 from tryptic digests analyzed by gas-phase sequencing (15, 16, 30), and other work from gas-phase sequence analysis has shown the presence of some amino acid microheterogeneity of FMO1 among enzyme preparations from different species (31). The primary structure of pig FMO1 is schematically illustrated in Scheme 1. To date, the screening of cDNA libraries has provided oligonucleotides approximately 2.2-2.6 kb in length that encode FMO enzymes of approximately 533-535 amino acids (32), but examples of FMOs with 19 (33) or 25 (34) additional C-terminal amino acids have also been observed. Hydropathy profiles show a remarkable similarity even in FMO isoform regions that are only modestly identical (35). Several parts of all FMOs have regions of highly conserved amino acid residues. For example, the highly conserved FADand NADP+-binding domains (i.e., GXGXXG) near deduced amino acid positions 9-14 and between 186 and 196, respectively, apparently are essential for FMO

1152 Chem. Res. Toxicol., Vol. 11, No. 10, 1998 Scheme 1. Schematic Representation of the Pig FMO1 Amino Acid Sequencea

a The shaded region is the lipophilic C terminus. The N terminus is N-acetylated, and amino acids 120 and 314 have the required consensus N-glycosylation sequence Asn-X-Ser/Thr.

function (14). On the basis of enzyme mechanism studies, it is clear that NADPH transfers reducing equivalents to the FAD moiety before molecular oxygen combines with the flavoprotein to form the requisite 4ahydroperoxyflavin necessary for substrate oxygenation. Thus, the NADP- and FAD-binding regions must be proximal to one another. The N terminus of FMO1 does not have an apparent signal peptide sequence, and because the N terminus also has the FAD-binding region, it is likely that this part of the protein does not function as a membrane insertion sequence. The C terminus of pig FMO1 is extremely hydrophobic, and we were unable to obtain complete amino acid sequence information from this region presumably because the peptides were too hydrophobic to be eluted from the HPLC column or insoluble after it was hydrolyzed and possibly lost prior to HPLC analysis. We conclude that membrane association is not a passive event dictated simply by hydrophobic interactions of C-terminal amino acids (6). We anticipate that membrane association is signaled by an internal sequence (i.e., selected residues between amino acids 15 and 230). With the localization of the site of N-glycosylation at Asn120 reported herein, we can begin to construct a more detailed picture of the enzyme. The construction of an FMO1 model is also aided by other data because other regions of FMO possess some remarkable homology to other well-characterized esterases (16). Thus, the consensus sequence containing the histidine residue of the catalytic triad of carboxylesterases and the highly conserved Ser194 of FMOs are among the amino acids of highly conserved peptide sequences of esterases. Further, the highly conserved Asp225 of FMOs is contained in an amino acid sequence quite similar to one present in thioesterases and rabbit liver microsomal esterases (16). Systematic analysis of the distance between the glycosylation site and the transmembrane hydrophobic segment suggests that the glycosylation site should be separated from the membrane by at least 13 amino acid residues (36), although shorter spans have been observed. Twelve amino acids to the N-terminal side of the N-glycosylation site is a region with a number of charged amino acids and is thus unlikely to be a transmembrane segment. However, it is likely that membrane association ocurrs between amino acid residues Cys30 and Leu90 because, presumably, the NADPand FAD-binding domains must be on the same side of the membrane. To the C-terminal side of Asn120, there is a highly conserved region with extremely hydrophobic amino acids (i.e., Ser137-Thr151). This is a putative transmembrane region that fulfills all of the criteria for membrane association. To test these postulates, further experiments will be required. It is possible that like CYP(arom) (37), the majority of FMO is located on the cytoplasmic side of the endoplas-

Korsmeyer et al.

mic membrane. Retention of FMO1 in the endoplasmic reticulum thus represents an unusual mechanism compared with the two other types of endoplasmic reticulum retention mechanisms that include retention by the KDEL region (38) or by a short cytoplasmic segment (39). In this regard, it is notable that rabbit FMO2 has been shown to be tightly associated with the KDEL-containing calcium binding protein, calreticulum (40). Not all FMO enzymes possess the same requirement for N-glycosylation because the number and position of putative Nglycosylation sites vary within the family of five mammalian FMOs currently known (35). In addition, highly active FMO enzymes have been expressed in E. coli, and bacteria do not possess the ability to enzymatically N-glycosylate proteins. While N-glycosylation of FMO is not likely to grossly alter enzyme function, this work has provided insight into the structure of FMO. The prediction is that N-glycosylation will be at a location distal to the substrate binding channel and cofactor binding regions.

Acknowledgment. A generous gift of the anti-guinea pig FMO supplied by Drs. K. Oguri and H. Yamada (Kyushi University, Fukuoda, Japan) is gratefully acknowledged. The authors are grateful to Dr. Reid Townsend (Dionex Corp.) for his help with the HPAEPAD. We thank Linear Instruments for providing the MALDI spectrum. This research was supported by grants from the National Institutes of Health (GM36426 to J.R.C.) and grants to the UCSF Bioorganic Biomedical Mass Spectrometry Resource (A. L. Burlingame, Director), supported by NIH Division of Research Grant RR01614 and NSF Grant DIR 8700766.

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TX980117P

N-Glycosylation of Pig Flavin-Containing Monooxygenase Form 1

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