Mucins and their O-Glycans from human bronchial epithelial cell cultures

Jessica M. Holmén ,1,* Niclas G. Karlsson,2,* Lubna H. Abdullah,3,* Scott H. Randell,3,4 John K. Sheehan,3,5 Gunnar C. Hansson,1 and C. William Davis3,4

1Department of Medical Biochemistry, Göteborgs universitet, 405 30 Gothenburg, Sweden; 2Proteome Systems, Locked Bag 2073, Sydney, New South Wales 1670, Australia; and 3Cystic Fibrosis/Pulmonary Research and Treatment Center, 4Department of Cell and Molecular Physiology, and 5Department of Biochemistry and Biophysics, University of North Carolina, Chapel Hill, North Carolina 27599

Submitted 24 March 2004 ; accepted in final form 9 June 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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A longstanding question in obstructive airway disease is whether observed changes in mucin composition and/or posttranslational glycosylation are due to genetic or to environmental factors. We tested whether the mucins secreted by second-passage primary human bronchial epithelial cell cultures derived from noncystic fibrosis (CF) or CF patients have intrinsically different specific mucin compositions, and whether these mucins are glycosylated differently. Both CF and non-CF cultures produced MUC5B, predominantly, as judged by quantitative agarose gel Western blots with mucin-specific antibodies: MUC5B was present at ~10-fold higher levels than MUC5AC, consistent with our previous mRNA studies (Bernacki SH, Nelson AL, Abdullah L, Sheehan JK, Harris A, William DC, and Randell SH. Am J Respir Cell Mol Biol 20: 595–604, 1999). O-linked oligosaccharides released from purified non-CF and CF mucins and studied by HPLC mass spectrometry had highly variable glycan structures, and there were no observable differences between the two groups. Hence, there were no differences in either the specific mucins or their O-glycans that correlated with the CF phenotype under the noninfected/noninflammatory conditions of cell culture. We conclude that the differences observed in the mucins sampled directly from patients are most likely due to environmental factors relating to infection and/or inflammation.

cystic fibrosis; obstructive airway disease; mucus overproduction


SINCE THE DEVELOPMENT 6 to 8 years ago of a primary human bronchial epithelial (HBE) cell culture system yielding a well-differentiated mucociliary phenotype (21, 33, 34), our understanding of airway biology and disease has been advanced considerably, particularly the pathogenesis of cystic fibrosis (CF) (27). Despite their utility, however, there are aspects of HBE cell cultures that are poorly understood, particularly with regard to mucin production and glycosylation, and in relation to disease.

A hallmark of obstructive airway disease is mucus overproduction, which in CF contributes to reduced mucociliary clearance and increased susceptibility to lung infection. The mucociliary clearance defect in CF is now believed to be more complex, however, and to result from a failure in cystic fibrosis transmembrane conductance regulator (CFTR)-mediated Cl secretion and a related increase in Na+ absorption, the combined effects of which deplete airway surface liquid to the extent that the mucus gel impacts onto cilia and interferes physically with ciliary function (27, 33, 39). Despite its longer history, the basis of CF-related mucin hypersecretion and molecular alterations is not as well understood as the consequences of ion and fluid transport abnormalities. In principle, the mechanism of mucin secretion from goblet cells in the superficial epithelium appears to be normal in CF (30), and mucus overproduction may be driven by inflammatory processes associated with infection that lead to a hyper- and metaplastic expansion of mucin-producing secretory cells (12, 44).

Mucins are massive glycoconjugates, generally >80% carbohydrate by weight, which form a heterogeneous family comprising monomeric mucins that may be secreted or tethered to cell surfaces and polymeric mucins that are secreted and form the scaffolding of mucus gels (35). Of the four known human polymeric mucins, MUC2, MUC5AC, MUC5B, and MUC6, MUC5AC and MUC5B are normally expressed in the lung, where they are secreted from goblet/mucous cells in the airways, the superficial epithelium, and submucosal glands, respectively (13, 42). Under pathological conditions as in CF, the intestinal mucin, MUC2, is also expressed in small amounts in the airways (14, 26, 31), and MUC5B expression expands to include the surface epithelium (13).

Mucin glycans are connected by O-linkages to Ser or Thr, which are clustered in large glycosylated "mucin domains." Although not known in detail, these O-glycans exhibit a remarkable heterogeneity and diversity, in part, because of blood group variability due to the ABO and Lewis blood group systems (24). One controversy that has attracted attention, before and after the discovery of CFTR in 1989, is whether there are direct, disease-related alterations in glycosylation. Several such alterations have been described in well-purified sputum samples as well as in the glycocalyx of cell lines (29, 46). The glycosylation alterations described vary but generally include increased sulfation, fucosylation, and/or sialylation, as well as decreased sialylation. The most consistent observation, increased sulfation of mucins, has also been observed in the glycocalyx of human airway epithelial cells grown in culture (10) and in mucins harvested from such cells grown in xenografts (52). Decreased sialylation of glycocalyx components has been observed in CF airway cells (17), whereas increased sialylation was described for airway mucins (15). These studies are inherently complicated, however, and the results to date have not allowed a clear discrimination between differences being due to primary or secondary effects of a dysfunctional CFTR.

We investigated the suitability of HBE cultures in approaching problems of this type, by harvesting and quantifying the mucins secreted from well-differentiated, primary HBE cell cultures derived from non-CF and CF lungs and by analyzing their oligosaccharides. Our goals were to apply newly available mucin-specific reagents to quantify the MUC5AC and MUC5B mucins secreted from HBE cultures, to analyze and quantify the oligosaccharides on these molecules for composition and structure using liquid chromatography-mass spectrometry (LC-MS) and tandem mass spectrometry (MS-MS), and to test whether there are readily detectible differences between the mucins harvested from non-CF and CF cultures using these techniques.


    MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
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Cell Culture and Harvesting of Mucins

Cell culture. Bronchial epithelial cells were harvested and cultured as previously described (5, 41), in triplicate, from six non-CF and six CF lungs (Table 1), under an institutional review board-approved protocol. Three individuals of each condition had blood type O; after quantitation, mucins collected from the cultures derived from these individuals were used in glycosylation studies. Lungs from the other individuals were selected without regard to blood type. Primary cells were cultured on type I and III collagen-coated (Cohesion, Palo Alto, CA) plastic plates and were cryopreserved at 75–90% confluence. When needed, these passage 1 cells were thawed, resuspended, and expanded on collagen-coated plastic dishes, and the resulting passage 2 cells were used to seed Transwell-Col membranes (24 mm, Corning Costar, Cambridge, MA). The cultures were maintained in air-liquid interface (ALI) medium (5, 21, 41) in BioCoat Deep-Well cluster plates (BD Biosciences, San Jose, CA). Beginning at confluence, 3–5 days after seeding, the cultures were maintained at an ALI and fed, from the bottom only, at 3-day intervals. As a point of reference, the day of confluence for each culture was taken as day 0.


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Table 1. Donor characteristics

 
Collection of mucins. Mucus was collected from HBE cell culture surfaces at 3-day intervals, from days 2 to 35 as indicated in RESULTS. For each collection, 0.5 ml of PBS was applied gently to the luminal surface of each culture, and the cultures were placed back in the incubator for 15 min. Holding each culture at a 45° angle, we collected the surface liquid by pipette and reapplied it to the upper edge of the culture, allowing the liquid to flow slowly down over the culture surface. This procedure was repeated twice, and then the liquid was collected and pipetted into a 2-ml screw-cap vial. Two more 0.5-ml PBS washes/collections were then made using the same procedure and added to the vial, which was then sealed and frozen (–20°C).

Based on preliminary assessments of mucin content in the collected samples, a subset consisting of samples collected on days 2, 5, 14, 26, and 35 postconfluence was chosen for detection of MUC5AC and MUC5B by agarose Western blotting. From the quantitative data derived from these blots (see Fig. 3), all of the samples derived from each of the six individuals with the blood type O (3 non-CF, 3 CF) between days 14 and 26 postconfluence, inclusive, were pooled and subjected to further analyses.



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Fig. 3. MUC5AC (A) and MUC5B (B) contents in the surface liquid of non-CF and CF HBE cell cultures. Each point represents the mean of triplicate HBE cell cultures derived from non-CF (filled symbols) and CF (open symbols) lungs sampled on the indicated days following confluence. For clarity, the means ± SE were offset from individual data points by 1 day, and individual overlapping symbols were displaced to the right. *By 2-way analysis of variance, MUC5AC and MUC5B levels at days 14 and 26 were elevated over those observed on day 5 postconfluence (P < 0.001); however, there were no significant differences in the levels of MUC5AC and MUC5B between the non-CF and CF samples (P > 0.05; n = 6). The mucins at days 2 and 35 were not included in the analysis due to smaller sample sizes (n = 3).

 
Quantitative Agarose and Western Blotting

Sample and standard mucin preparations. At room temperature, the vials containing HBE cell culture secretions were thawed, the samples from the triplicate cultures for each collection were combined, and 100-µl volumes were dialyzed (Mini Dialysis Unit, Pierce, Rockford, IL) against 6 M urea buffer containing 0.1 M Tris acetate, 5 mM EDTA, pH 8.0. Subsamples of the dialyzed material were diluted quantitatively and prepared for electrophoresis. To each 100 µl of sample and standard, 10 µl of 10x DTT sample buffer [50% glycerol, 1.0 M Tris acetate, 50 mM EDTA, 45 mM DTT, 1.0% (wt/vol) SDS, 0.01% bromphenol blue, pH 8.0] were added, and the mixtures were heated at 100°C for 15 min.

Mucin standards in 6 M urea buffer were prepared from stock solutions of known concentration using the same protocol. As described previously (26), the MUC5AC and MUC5B standard mucins were isolated from HT-29 cells and human saliva, respectively, and their concentrations were determined by refractive index. A series of five dilutions of the appropriate standard was included on each gel, 1.25–20 ng/well for MUC5AC and 62.5–2,000 ng/well for MUC5B.

Electrophoresis and vacuum transfer. Forty microliters of samples and standards were loaded into the wells of 15 x 15 cm x 1.5 mm thick, 1.0% agarose gels. The gels were electrophoresed for 2.5–3.0 h in TAE buffer (40 mM Tris acetate, 1 mM EDTA, pH 8.0) at a constant 80 V. After electrophoresis, the gels were washed in SSC buffer (0.6 M NaCl, 60 mM Na citrate, pH 7.0), and the resolved molecules were transferred to nitrocellulose (0.45 µm) in the same buffer for 1.5 h at a negative pressure of 45 mbar using a VacuGene XL vacuum-blotting apparatus (Amersham Biosciences, Piscataway, NJ).

Mucin detection and analysis. The nitrocellulose membranes were washed with PBST, blocked with 5% milk, and probed with mucin-specific antibodies (26, 50) to MUC5AC (MAN-5ACI) or to MUC5B (MAN-5BI). After being washed in PBST, the blots were developed with an alkaline phosphatase-conjugated secondary antibody (Jackson ImmunoResearch, West Grove, PA), using nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate as the substrate.

The developed membranes were digitized in monochrome at 1,200 x 1,200 dpi x 12 bit (depth) using a UMAX Powerlook III flatbed scanner (Dallas, TX). The integrated staining intensities of each MUC5AC- or MUC5B-positive band in the resulting TIFF images were determined with MetaMorph image processing software (Universal Imaging, Downington, PA) using box-shaped cursors (Fig. 1). Background intensities were determined using the same cursors positioned in a nonstained portion of each lane, below the positive band. Calibration curves (see Fig. 1) were constructed from background-subtracted intensities from each mucin dilution series by nonweighed, least-squares regression analysis. Differences between mean values in the final data set were analyzed statistically by two-way ANOVA.



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Fig. 1. MUC5AC and MUC5B agarose Western blots of mucin standards and resulting standard curves. Left and right: image of typical Western blot (top) and standard curves derived from its quantitation (bottom). The standards, stored in 6 M urea buffer, were reduced and electrophoresed on 1% agarose gels along with samples of non-cystic fibrosis (CF) and CF human bronchial epithelial (HBE) cell culture surface liquid. After vacuum transfer to nitrocellulose, the blots were probed with MUC5AC- and MUC5B-specific antibodies and quantitated as described in MATERIALS AND METHODS. Means ± SE of nonnormalized data from 3 standard curves are shown. Boxes surrounding selected regions illustrate the type of cursors used to quantify the staining (see MATERIALS AND METHODS).

 
Mucin Purification and Amino Acid Analysis

The materials collected from the cultures derived from blood type O individuals (3 non-CF, 3 CF) were pooled, and the mucins were purified by dual CsCl gradient centrifugation following previously published procedures (2, 48). Briefly, the material was prepared for density gradient centrifugation by adding guanidinium Cl to a concentration of 4 M and CsCl (both from Fisher Scientific, Pittsburgh, PA) to an initial density of 1.40 g/ml. The solution was centrifuged for 3 days in a Beckman L7-55 ultracentrifuge equipped with a 50.2 Ti rotor at 36,000 rpm and 15°C. Each tube was divided into 16 fractions (2.5 ml) that were weighed for density and assessed for DNA (OD260) and sialic acid (11). Sialic acid-positive fractions were pooled and dialyzed against 0.2 M GuHCl in PBS overnight at 4°C; the density of the solution was adjusted to 1.50 g/ml with CsCl, and the material was centrifuged a second time, as above, to separate mucins from contaminating DNA. The sialic acid-containing fractions were pooled and dialyzed against deionized water.

Amino acid analysis was performed on purified mucins as described previously (24), using an Alpha Plus amino acid analyzer (Pharmacia Biotech, Uppsala, Sweden).

Monosaccharide Analysis of Mucin Oligosaccharides

By use of the previously described procedures (23), monosaccharide analysis of the purified mucins was performed on nonreleased oligosaccharides, i.e., while still attached to the protein backbone. Additionally, oligosaccharides were analyzed following their release from the mucins (~250 µg) by reductive {beta}-elimination (1 M sodium borohydride) under alkaline conditions (0.05 M potassium hydroxide) (25). Collected oligosaccharides were desalted and separated into two subfractions containing neutral and acidic oligosaccharides, respectively (25). The separation was performed on a DEAE-Sephadex A-25 column, and the neutral fraction was eluted with dry methanol, while the acidic species were eluted with pyridinium acetate. The monosaccharides of the released neutral and acidic oligosaccharide alditols were analyzed, as above (23).

Mass Spectrometric Analysis of Oligosaccharides

Oligosaccharides were released from isolated mucin subunits (~50 µg) using 50 mM sodium hydroxide and 0.50 M sodium borohydride at 50°C for 16 h, followed by desalting. Approximately 5 µg of each mucin sample were analyzed by negative ion LC-MS as described (47). Separation was performed using a constant 10 mM ammonium bicarbonate mobile phase with a linear acetonitrile gradient, 0–30% in 30 min, on a 5-µm particle Hypercarb column (150 x 0.32 mm; Thermo-Hypersil, Runcorn, UK). The detector was an LCQ Deca ion trap mass spectrometer (Thermo-Finnigan, San Jose, CA). As a measurement for the abundance of individual oligosaccharide components, the total intensities of each full scan of identified pseudomolecular ions and their isotopes, eluting as single peaks in the LC-MS trace, were integrated. Relative intensities were calculated from the total amount of structure presented. Partial oligosaccharide sequence information was obtained by automatically switching between LC-MS and LC-MS/MS. MS/MS spectra were manually interpreted, and the GlycosuiteDB oligosaccharide database (www.glycosuite.com) was used to evaluate predicted oligosaccharide epitopes in relation to those previously described in respiratory mucins.

Analysis of Mucin Electrostatic Charge by SDS-PAGE

Approximately 25-µg aliquots of the purified mucins from the non-CF and CF samples specified in RESULTS were reduced, subjected to SDS-PAGE, blotted onto polyvinylidene difluoride (PVDF) membrane, and acidic oligosaccharides were detected by Alcian blue (47). The stained area was cut from the membrane, and the CF and control lanes were divided into four equal pieces (see Fig. 6). The number of sialic acid and sulfate groups on the oligosaccharides released from these pieces was determined by LC-MS (47). The mass spectrometric intensities of the ions for individual oligosaccharide components in the chromatogram were used to estimate the average charge of the oligosaccharides in each area of the gel. Separate lanes in the same gel were transferred to a PVDF membrane and used to detect MUC5B by Western blotting with the PANH2 (37) antibody. To allow access by the antibody, the materials in these lanes were deglycosylated with trifluoromethanesulfonic acid (10 ml) and toluene (600 µl) at –20°C for 5 h, followed by thorough washes with methanol and water. The blot was blocked using 0.1% Tween and 0.5% BSA in PBS at 4°C for 16 h, incubated with the primary antibody (1 h), secondary antibody (horseradish peroxidase-conjugated sheep anti-mouse; Silenus, Melbourne, Australia) diluted 1/500 for 1 h, and developed for 30 min in 1 mg/ml chloronaphthol in 1:2 methanol/water (24 ml) with 30 µl of hydrogen peroxide added.



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Fig. 6. Agarose-polyacrylamide gel electrophoresis and charge analysis of MUC5B from N3 and CF1 cultures. Duplicate lanes containing N3 and CF1 purified mucins were separated on an agarose-polyacrylamide composite gel and blotted as described in MATERIALS AND METHODS. Left: 1 pair of lanes was stained with Alcian blue to reveal charged oligosaccharides. The stained area was cut from the membrane, divided as indicated, and the released O-linked oligosaccharides were analyzed for charge by LC-MS. The average charge from the recorded mass spectra is given to the left (N3) and right (CF1) of the indicated bands. Right: other lanes were stained with the anti-MUC5B monoclonal antibody PANH2 (37) followed by horseradish peroxidase-conjugated anti-mouse antibody and detected with chloronaphthol.

 

    RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
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 DISCUSSION
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MUC5AC and MUC5B in HBE Cell Culture Surface Liquids

Previously, we found that all four known polymeric mucin genes are detected by RT-PCR in well-differentiated HBE cell cultures, but only MUC5AC, MUC5B, and MUC6 expression was upregulated in a retinoic acid-sensitive manner, and only MUC5B was expressed sufficiently strongly as to be detectable by Northern blot analysis (5). Both the MUC5AC and MUC5B mucins were readily detected by Western blotting in the surface liquid of HBE cell cultures derived from non-CF and CF lungs. Figure 2 shows examples of agarose Western blots for both mucins from each condition. MUC5AC resolved as a single compact band with a mobility in 1% agarose gels approximately equal to that for the HT-29 cell-derived standard. MUC5B from both non-CF and CF HBE cell cultures also resolved as a single, compact band, in contrast to the two glycoforms visible in the immunoblots of human saliva-derived standards electrophoresed on the same gels (Fig. 1) (26). The single band visible in the agarose Western blots indicates that a single glycoform of MUC5B is expressed in HBE cell cultures, in contrast to the two glycoforms that are commonly observed from mucus collected from the airways and saliva (26). Interestingly, the mobility of MUC5B from HBE cultures appeared to be very uniform, approximately midway between the two glycoforms of the MUC5B standard, in contrast to the more variable mobility observed in the material derived from airway samples. To illustrate the sensitivity of the method in detecting differences in net charge, Fig. 2, inset, shows the effects on MUC5B mobility of digesting terminal sialic acid residues from HBE mucins with neuraminidase. The motilities of MUC5AC were homogeneous, and like MUC5B, there were no obvious differences in band position on the agarose gels between the mucins from different individuals or between non-CF and CF lungs (data not shown). In ongoing experiments on material harvested from more than 12 HBE non-CF and CF cultures and subjected to analysis by mass spectrometry, we failed to detect a single peptide fragment derived from either MUC2 or MUC6. In contrast, several different peptide fragments from MUC5AC and MUC5B are commonly detected in every sample (data not shown).



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Fig. 2. Sample agarose Western blots for MUC5AC and MUC5B secreted from non-CF and CF HBE cell cultures. Mucins harvested on days 14 and 26 postconfluence from culture surfaces are shown at 2 or 3 dilutions. The samples were solubilized in GuHCl, then electrophoresed, blotted, and probed in the same way as the mucin standards (Fig. 1). Note that MUC5AC and MUC5B from both non-CF and CF HBE cell cultures resolved as compact, single bands on the agarose gels. Inset: purified HBE mucins from other non-CF cultures before (C) and after (N) digestion for 30 min with neuraminidase to illustrate the effects of removing sialic acid on MUC5B mobility.

 
MUC5AC and MUC5B secretion from non-CF and CF HBE cell cultures developed with similar time courses (Fig. 3): the quantities increased from essentially undetectable levels at 1 day postconfluence to apparent steady-state levels from days 14 through 26. Thereafter, MUC5AC levels remained at this plateau level through day 35, whereas MUC5B levels declined to variable degrees. For both mucins, the amounts collected from the cultures every 3 days were normally distributed and varied over about a 10-fold range between individuals. The mean steady-state levels of MUC5AC sampled from both non-CF and CF HBE cell cultures were ~30 µg/culture, whereas the MUC5B levels were ~10 times higher. MUC5B levels appeared to be nominally higher in CF than in non-CF HBE cell cultures at days 14 (281 ± 78 vs. 413 ± 72 µg/culture) and 26 (264 ± 72 vs. 320 ± 68 µg/culture); however, these apparent differences were not statistically significant.

Chemical Composition of HBE Mucins

The secreted mucins were purified from the secretions from HBE cell cultures derived from three non-CF (N1-N3) and three CF patients (CF1-CF3), all blood group type O, by two rounds of CsCl density gradient ultracentrifugation. The purified mucins were analyzed first for amino acid and monosaccharide composition. The results revealed that 84–90% of the total mucin mass was made up by oligosaccharides, a common characteristic of mucins (Table 2). Both the total amounts of oligosaccharides observed and the relative distributions of monosaccharides were similar for the CF and non-CF patients and were typical of those normally associated with secreted mucins. The relative distribution of amino acids also showed a similar composition for the CF and non-CF patients (Table 3) with high percentages of the amino acids, Ser (8–12%), Thr (15–22%), and Pro (8–12%). The overall amino acid composition was similar for the six mucin samples and resembles the composition of the MUC5B and MUC5AC mucins (Table 3) (16, 19). The content of Ser was higher in the mucin samples than expected, but the results are still generally compatible with MUC5B being the predominant mucin (see Fig. 3).


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Table 2. Chemical composition of human epithelial cell mucins

 

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Table 3. Relative amino acid composition of the purified human epithelial cell mucins MUC5B and MUC5AC

 
O-glycans of the HBE Mucins

The O-linked oligosaccharides were released from the mucins and separated into two subfractions containing neutral and acidic compounds by ion-exchange chromatography (25). The GalNAc residues at the reducing ends were converted to GalNAcol by sodium borohydride during the release, and the amounts of the derivatized monosaccharide were used to express the molar amounts of Fuc, GlcNAc, GalNAc, and Gal oligosaccharides in the neutral and acidic fractions relative to the GalNAcol (Table 4). No major differences were observed in comparing the relative oligosaccharide compositions of the non-CF and CF samples, although individual differences were evident. The glycosylation of N1 is clearly different from the other non-CF and CF patients in the higher relative amounts of neutral glycans and longer acidic glycans compared with the other mucin samples.


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Table 4. Relative distribution of monosaccharides in the oligosaccharides released from purified mucins of non-CF and CF HBE cultures

 
To obtain additional information on glycosylation, the O-linked oligosaccharides from the purified mucins were released and analyzed by LC-MS. The total ion chromatograms from the three non-CF and CF mucin samples are presented in Fig. 4. The chromatograms reveal a complex mixture of oligosaccharides, with only limited similarities within the two groups. Interestingly, two of the CF samples, CF1 and CF3, were most similar to the N1 sample. The composition and structure of each of the detected oligosaccharides were determined from their molecular mass and tandem mass spectra according to published principles of fragmentation (47). The results are gathered in Fig. 5 and the identified components are indicated in Fig. 4. Fragmentation by MS/MS of the oligosaccharides was consistent with the compositions assigned and could in many cases be used for detailed structural assignment (Table 5). The structural analyses revealed very complex mixtures as have been observed previously in respiratory mucins (6, 7, 51). More than 60 oligosaccharide components were found above the threshold of 10% of the intensity of the major component in any of the samples. Components were identified by retention time on the graphitized carbon column and molecular mass, allowing elucidation of both compositional and often isomeric information. Oligosaccharide structures were also found below that threshold but were often of high molecular mass in heterogeneous mixtures, not resolving as single peaks in the chromatogram. Many of the major components (Table 5) were successfully sequenced from the MS/MS. Mass spectrometry typically provides sequence information, but linkage information is possible only in some instances. The nature of isomeric monosaccharide residues has to be deduced from monosaccharide compositional data (see Tables 2 and 4). If previously identified oligosaccharides of respiratory mucins are taken into consideration (Glycosuite DB), the presence of core 1 [Gal{beta}1–3GalNAcol], core 2 [Gal{beta}1–3(GlcNAc{beta}1–6)GalNAcol], core 3 [GlcNAc{beta}1–3GalNAcol], and core 4 [GlcNAc{beta}1–3(GlcNAc{beta}1–6)GalNAcol] is suggested in the current study. Blood group type oligosaccharides having blood group H-type sequences [Fuc{alpha}1–2Gal{beta}1-], as well as Lewis-type sequences, were found. The high proportion of type 2 chains [Gal{beta}1–4GlcNAc{beta}1-] found [depicted from specific cross ring cleavages in MS/MS (47)] indicates that the Lewis-type structures were predominantly Lewis x [Gal{beta}1–4(Fuc{alpha}1–3)GlcNAc{beta}1-] and Lewis y [Fuc{alpha}1–2Gal{beta}1–4(Fuc{alpha}1–3)GlcNAc{beta}1-], rather than Lewis a [Gal{beta}1–3(Fuc{alpha}1–4)GlcNAc{beta}1-] and Lewis b [Fuc{alpha}1–2Gal{beta}1–3(Fuc{alpha}1–4)GlcNAc{beta}1-]. Mono- and disialylated components of both three- and six-linked sialic acid were also found together with sulfated compounds.



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Fig. 4. Total ion chromatograms from the HPLC-MS analyses of released oligosaccharides from the mucins of 3 non-CF (N1-N3) and 3 CF (CF1-CF3) HBE cell cultures. The released oligosaccharides were separated on a Hypercarb column and detected with a LCQ Deca ion trap mass spectrometer. Peak designations refer to the type and size of the identified oligosaccharides, e.g., N4.1 means a neutral oligosaccharide with 4 sugar residues and Si3.1 a sialylated oligosaccharide with 3 sugar residues. The number following the decimal point enumerates oligosaccharides with the same number of sugar residues, e.g., Si3.1 and Si3.2.

 


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Fig. 5. Histogram of relative distribution of oligosaccharides in the combined non-CF and CF HBE mucins as obtained from HPLC-MS analyses. Filled bars are CF and open bars are non-CF samples. Each bar is the relative percentage of each compound given with its standard deviation. Each set of bars is due to 1 oligosaccharide as marked below these. The relative composition is based on the total ion count divided by the molecular mass. Only oligosaccharides >10% in any chromatogram are presented. The code identifying the oligosaccharides in a peak is presented in Fig. 4, and the composition and detailed structure of each structure are detailed in Table 5.

 

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Table 5. Proposed structures as based on MS/MS of O-glycans released from HBE

 
The approach using LC-MS for detecting mucin oligosaccharides was found to be efficient for the analysis of small- and medium-sized oligosaccharides. This indicates that the coverage of neutral oligosaccharides was better than for acidic species, because neutral oligosaccharides were shown to have a shorter average length (Table 4). Preliminary data from the oligosaccharides below the 10% threshold in the LC-MS trace (not reported in Fig. 5) indicated that these were larger and mainly sialylated (loss of sialic acid in the MS/MS) or sulfated (loss of sulfate in the MS/MS). As the predicted average length especially of the acidic glycans was longer, the HBE mucins were separated by composite agarose-polyacrylamide gel electrophoresis and the oligosaccharides released from different mucin subpopulations (Fig. 6). It has previously been shown that MUC5B will separate into different subpopulations by electrophoresis, depending on charge of the attached oligosaccharides (26), where the more highly charged mucins have higher mobilities. The mucins from CF and non-CF HBE mucins, however, had uniform electrophoretic mobilities, indicating a similar charge distribution of the oligosaccharides attached to the mucins (Fig. 6). Extensive LC-MS analysis of the oligosaccharides was also carried out where the average number of negative charges (sialic acid and sulfate) per oligosaccharide based on mass spectrometric intensities of the oligosaccharides released was calculated. The results showed the expected charge-dependent electrophoretic separation of the mucins, but the non-CF and CF samples exhibited similar charge gradients (from 1.0 to 1.3 charges). The monosaccharide analysis (Table 4) indicated that additional, longer acidic oligosaccharides were present in the HBE mucins than the oligosaccharide structures indicated by MS-MS (Table 5). This difference is due, in part, to an underestimation of the number of oligosaccharide chains as calculated based on GalNAcol in Table 4. The acidic fractions, especially, contain some glycopeptides with unreleased sugar chains, as indicated by the relative amount of GalNAc. Any significant differences in the charge distribution of these oligosaccharides between CF and non-CF mucins, however, would have been revealed as different patterns of electrophoretic mobility.

Comparison of the oligosaccharides found in the non-CF and CF mucins revealed no specific qualitative differences. To further explore potential differences, the relative amount of each individual oligosaccharide was analyzed by combining the information from three non-CF and three CF samples (Fig. 5). The results illustrate what could be observed in the initial chromatograms, a large variation between individuals, as seen by the large standard deviation on many compounds. No specific differences that could be explained by specific up- or downregulation of individual glycosyltransferases due to nonfunctional CFTR were found. As discussed above, some of the larger more negatively charged oligosaccharides could not be analyzed, but the similar distribution of charged oligosaccharides on the mucins as illustrated in Fig. 6 argues against specific CF-related differences also among this type of glycans.

The results of the analyses of the O-linked glycans suggest a large variation between different individuals but no clear difference that could be attributed to the CF and non-CF genotype.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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The complexities of the respiratory tract require simplified experimental models to allow a detailed understanding of its function in health and its pathology in disease. Cell culture models are very attractive and the HBE cell cultures have made important contributions to the understanding of the pathogenesis in CF (21, 33). The present studies were performed on passage 2 cultures, which reliably undergo mucociliary differentiation, to minimize artifacts associated with the passaging of primary cells, yet allow the cultures from CF cultures more time under conditions free of infection and inflammation than would be the case with cultures from native or passage 1 cells.

Mucins Produced by HBE Cultures

The mucins produced by the HBE cell cultures had typical amino acid and carbohydrate compositions, i.e., more than 84% of the mucin mass comprised sugars, and there were relatively high amounts of the amino acids, Ser, Thr, and Pro. These mucins were shown by immunological methods to contain both MUC5B and MUC5AC, with ~10-fold more MUC5B (Fig. 3). The predicted amino acid compositions for MUC5B and MUC5AC are similar to one another and to the compositions observed for the mucins harvested from both non-CF and CF cultures (Tables 2 and 3). One exception, however, was that the quantities determined for Ser, from both sets of cultures, were slightly higher than expected (15–17 residues vs. the 11–13 predicted). A possible explanation for this discrepancy is the copurification of other mucins with MUC5AC and MUC5B during the CsCl gradient centrifugation procedure. By mass spectrometry, we have not observed other polymeric mucins in the materials harvested from HBE cell cultures; however, we routinely detect peptide fragments from multiple tethered mucins (data not shown). Comparing the relative amounts of MUC5AC and MUC5B in the six non-CF and CF mucins by Western blotting, which included the three blood group O cultures in each condition, revealed no major differences. Hence, the similarities in MUC5AC and MUC5B distributions, and in the mucin amino acid and carbohydrate compositions, suggest that the mucins produced by the non-CF and CF HBE cell cultures are similar.

MUC5B mucin is normally expressed in the glands of the lungs (13, 42), but it is also the predominant gel-forming mucin expressed in HBE cell cultures (Fig. 3). This pattern is not what is expected for cells derived from airway superficial epithelium but is consistent with a previous report (5). There are three conceivable explanations for the dominance of MUC5B instead of MUC5AC. First, the development of the HBE cell cultures, in vitro, may recapitulate the development of the airways, in vivo. In human development, MUC5B appears early in the airways, followed by ciliation, then by MUC5AC expression. As the airways mature and submucosal glands develop, MUC5B expression migrates into the glands that represent its major site of expression in the normal adult human lung. In this light, the decline in MUC5B levels between days 24 and 35 may reflect analogous changes occurring in culture. Second, the HBE cell cultures may represent not the superficial epithelium of the airways, but instead the ciliated ducts. In the epithelium lining gland ducts, MUC5B is generally expressed (42), and in the HBE cell cultures ciliated, duct-like cysts commonly form within the epithelium as the cultures age, as though they are attempting to form glands. Third, the culture conditions used to support the growth and differentiation of HBE cell cultures may mimic inflammatory conditions that cause an upregulation of MUC5B expression. This possibility is supported by the observation that MUC5B mucin is expressed in the superficial epithelium of inflamed lungs (13). The present data do not allow a distinction among these three possibilities. That the relative abundance of MUC5AC and MUC5B was similar between non-CF and CF HBE cultures, however, supports the notion that the species of mucin produced is independent of CFTR.

Mucin Oligosaccharides

The mucin oligosaccharides were released and analyzed for length and composition. The mucins in all the samples analyzed carried approximately equal number of neutral and acidic saccharides. The acidic saccharides were considerably longer, as suggested from both the compositional analyses and the mass spectrometry studies (Table 4). Preliminary mass spectrometry analysis of these larger acidic oligosaccharides suggests that they are sulfated, rather than sialylated, a suggestion supported by observation of long sulfated oligosaccharides in human tracheal mucin glycoproteins (45). There were no apparent differences, however, in saccharide lengths or composition between the mucins analyzed from non-CF and CF HBE cultures.

When the O-linked glycans on the three blood group O cultures from the non-CF and CF donors were compared in detail, the oligosaccharides revealed a complex pattern that varied between the individuals. Furthermore, the variation in individual oligosaccharides was larger than that observed between the non-CF and CF groups. The N1 and N2 cultures from non-CF donors formed the two extremes of observed variability, between which the CF cultures were distributed. The monosaccharide compositional analysis suggests that additional oligosaccharides are present in the samples that deserve thorough structural characterization, especially considering the unresolved question about the presence of large sulfated structures in connection with CF. Even though the analysis in Table 4 may overestimate the actual length of the oligosaccharides, the mass spectrometry studies suggest the presence of larger oligosaccharides that are extremely heterogeneous, not only in size but also in isomeric complexity, precluding more detailed characterization due to the low amount of each individual species.

An important factor to consider when analyzing mucin glycosylation is natural individual variation. Although the cultures analyzed were of blood group O origin, there is a possibility that the difference could be due to another blood group (i.e., Lewis or Secretor status) or other glycosyltransferases. We conclude that the present study does not support intrinsic abnormalities in the glycosylation of polymeric mucins in CF airways. This is based on both the monosaccharide compositional analyses and mass spectrometric results, acknowledging that the global glycomic approach used here might overlook minor differences among high-molecular-mass acidic oligosaccharides. The present results do not support any pronounced CF-specific up- or downregulation of sulfo- or glycosyltransferases.

Glycosylation alterations in CF are controversial. Specific alterations leading to decreased sialylation and increased fucosylation have been proposed (43, 46), based primarily on glycoproteins other than polymeric mucins that comprise the glycocalyx of various cell lines. A hypothesis linking CFTR to glycosylation posited that dysfunctions in CFTR cause increases in Golgi pH, with the result that glycosyltransferase activities are altered (4). In support of this hypothesis, it has been shown that neutralization of Golgi pH using a weak base caused a redistribution of glycosyltransferases within the Golgi, as well as a decrease in O-glycan chain lengths of the mucins secreted from LS 174T cells (3). Other studies, however, showed that Golgi pH is not altered significantly in cells expressing dysfunctional CFTR (9, 20) and that CFTR has no effect on glycosyltransferase activities (7). Thus, although glycoproteins other than mucins may be altered in CF through other mechanisms, it is clear that direct effects of CFTR on cellular systems can only occur in CFTR-expressing cells. The available evidence suggests a lack of CFTR in airway and intestinal goblet cells. CFTR was undetectable in goblet-like SPOC1 cell lysates by Western blotting (1), and it was not detected in human colonic goblet cells by immunostaining (32), whereas it was detected readily in nonsecretory cells in both cases. Similarly, CFTR was positively detected in non-CF or CF human airway ciliated cells by immunostaining, but the goblet and mucous cells in the superficial epithelium and submucosal glands, respectively, were negative (Kreda SM and Boucher RC, personal communication). This apparent absence of CFTR in airway goblet and mucous cells is consistent with the lack of detectable differences in the polymeric mucins expressed and their glycosylation observed in this study. In ciliated cells and submucosal gland serous cells in the airways, and in the mucin-secreting epithelial cells of the gallbladder and pancreatic ducts, however, dysfunctional CFTR may lead to alterations in glycoproteins of their respective glycocalyx and/or secreted glycoproteins because these cells normally express CFTR (18, 28, 40). It is also conceivable that these cells could respond to injury or inflammation with CFTR-dependent changes in their glycosylated proteins.

For polymeric mucins, Morelle et al. (36) suggested that differences in glycosylation observed in sputum samples from CF and chronic bronchitis patients are due to infection and inflammation rather than dysfunctional CFTR. A relationship was observed between the presence of Pseudomonas aeruginosa and the expression levels of sialyl-Lewis x. Hence, this study is in line with an emerging understanding of glycosylation being a dynamic process modulated by infection and inflammation of mucosal surfaces. We showed that a parasitic infection (Nippostrongylus brasiliensis) in mouse small intestine gives rise to a transient induction of the blood group H glycosyltransferase, Fuc{alpha}1–2 transferase (named Fut2; Ref. 22). Interestingly, we also observed an induction of the Fut2 enzyme in the small intestine of mice lacking a functional CFTR (49). This enzyme, however, is present in the large intestine of both wild-type and CF mice, an organ always more heavily colonized by bacteria (49), and in the small intestine it has been previously shown to be induced on bacterial colonization of germ-free mice (8). Hence, both the gnotobiotic and CF mouse models support the notion of specific induction of a glycosyltransferase in response infection/inflammation (8, 49).

In summary, HBE cells from non-CF and CF lungs cultured under noninflammatory conditions produced MUC5B as the dominant mucin. Extensive compositional analysis showed that there were no differences between the non-CF and CF groups in the amounts of MUC5B and MUC5AC produced or in their glycosylation. In general, the variabilities in glycosylation observed were larger between individuals than between the two groups. Given the lack of CFTR expression in airway mucin-secreting cells, we therefore conclude that a direct relationship between CFTR and the species of polymeric mucin produced or in their glycosylation is highly unlikely. Furthermore, at the sensitivity of our methods, we also failed to detect any indirect relationship between CFTR dysfunction and glycosylation of polymeric mucins in cultured HBE cells. These observations therefore support the notion that changes in mucin composition observed in vivo in patients with CF or other obstructive lung diseases are related primarily to infection and inflammation.


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Support for this work was provided by Swedish Research Council Grant 07461, IngaLill and Arne Lundberg's Foundation (Sweden), National Heart, Lung, and Blood Institute Grant P50-HL-60280, and by the Cystic Fibrosis Foundation of North America.


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Address for reprint requests and other correspondence: C. W. Davis, 6009 Thurston-Bowles, Univ. of North Carolina, Chapel Hill, NC 27599-7248 (E-mail: cwdavis{at}med.unc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

* J. M. Holmén, N. G. Karlsson, and L. H. Abdullah made equal contributions to this study. Back


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