Processing of N-linked carbohydrate chains in a patient with glucosidase I deficiency (CDG type IIb)

Christof Völker2, Claudine M. De Praeter3, Birgit Hardt2, Willi Breuer2, Burga Kalz-Füller2, Rudy N. Van Coster4 and Ernst Bause1,2

2 Institut für Physiologische Chemie, Universität Bonn, Bonn, Germany; 3 Department of Pediatrics, Division of Neonatal Intensive Care, Gent University Hospital, Gent; and 4 Department of Pediatrics, Division of Neurology and Metabolic Diseases, Gent University Hospital, Gent

Received on February 18, 2001; revised on March 15, 2002; accepted on March 18, 2002


    Abstract
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Recently, we reported a novel congenital disorder of glycosylation (CDG-IIb) caused by severe deficiency of the glucosidase I. The enzyme cleaves the {alpha}1,2-glucose residue from the asparagine-linked Glc3-Man9-GlcNAc2 precursor, which is crucial for oligosaccharide maturation. The patient suffering from this disease was compound-heterozygous for two mutations in the glucosidase I gene, a T->C transition in the paternal allele and a G->C transition in the maternal allele. This gives rise in the glucosidase I polypeptide to the substitution of Arg486 by Thr and Phe652 by Leu, respectively. Kinetic studies using detergent extracts from cultured fibroblasts showed that the glucosidase I activity in the patient’s cells was < 1% of the control level, with intermediate values in the parental cells. No significant differences in the activities of other processing enzymes, including oligosaccharyltransferase, glucosidase II, and Man9-mannosidase, were observed. By contrast, the patient’s fibroblasts displayed a two- to threefold higher endo-{alpha}1,2-mannosidase activity, associated with an increased level of enzyme-specific mRNA-transcripts. This points to the lack of glucosidase I activity being compensated for, to some extent, by increase in the activity of the pathway involving endo-{alpha}1,2-mannosidase; this would also explain the marked urinary excretion of Glc3-Man. Comparative analysis of [3H]mannose-labeled N-glycoproteins showed that, despite the dramatically reduced glucosidase I activity, the bulk of the N-linked carbohydrate chains (>80%) in the patient’s fibroblasts appeared to have been processed correctly, with only ~16% of the N-glycans being arrested at the Glc3-Man9–7-GlcNAc2 stage. These structural and enzymatic data provide a reasonable basis for the observation that the sialotransferrin pattern, which frequently depends on the type of glycosylation disorder, appears to be normal in the patient.

The human glucosidase I gene contains four exons separated by three introns with exon-4 encoding for the large 64-kDa catalytic domain of the enzyme. The two base mutations giving rise to substitution of Arg486 by Thr and Phe652 by Leu both reside in exon-4, consistent with their deleterious effect on enzyme activity. Incorporation of either mutation into wild-type glucosidase I resulted in the overexpression of enzyme mutants in COS 1 cells displaying no measurable catalytic activity. The Phe652Leu but not the Arg486Thr protein mutant showed a weak binding to a glucosidase I–specific affinity resin, indicating that the two amino acids affect polypeptide folding and active site formation differently.

Key words: CDG type II/genomic DNA/glucosidase I deficiency/glycoprotein processing/sialotransferrin


    Introduction
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Asparagine-linked oligosaccharides have been implicated as playing an important role in many biological processes, such as cell growth, cell development, and communication, as well as for protein stability and the control of protein folding (Varki, 1993Go; Parodi, 2000Go). The biosynthetic pathway includes preassembly of Dol-PP-GlcNAc2-Man9-Glc3, en bloc transfer of the activated tetradecasaccharide onto specific asparagine residues, and subsequent remodeling of the protein-bound precursor to give the mature structure (Kornfeld and Kornfeld, 1985Go). Processing begins in the endoplasmic reticulum (ER) with removal of the distal {alpha}1,2- and the two inner {alpha}1,3-linked glucose residues, catalyzed by glucosidase I and glucosidase II, respectively. The resulting Man9-GlcNAc2 structure is then acted on by different ER- and Golgi-resident {alpha}1,2-mannosidases, which remove up to four mannoses, thereby generating different high-mannose intermediates. After transfer of a GlcNAc residue onto the {alpha}1,2-mannose-depleted Man5-GlcNAc2 core, the outer {alpha}1,3/{alpha}1,6-mannosyl branch is excised by {alpha}-mannosidase II; this is followed by conversion of the GlcNAc-Man3-GlcNAc2 hybrid to complex type N-glycans involving a variety of Golgi-located glycosyltransferases.

Given the very complex nature of the N-glycosylation pathway, it is not surprising that genetic defects resulting in aberrant N-glycan chain structures have a dramatic impact on many intra- and extracellular functions. Several congenital disorders of glycosylation (CDGs; previously known as carbohydrate-deficient glycoprotein syndromes) have been identified that can be attributed to individual enzyme defects in the biosynthesis of N-glycoproteins (Marquardt and Freeze, 2001Go; Schachter, 2001Go). The CDGs are multisystem diseases, often associated with delayed development and neurological dysfunction up to and including severe mental retardation, as well as with dysfunction of organs or the endocrine and coagulation system. A recent definition classifies the CDGs into two groups: type I CDGs comprise defects for enzymes catalyzing formation of dolichol-PP-activated GlcNAc2-Man9-Glc3 and its transfer by oligosaccharyltransferase (OST) on to protein, whereas type II CDGs include defects for enzymes involved in the processing of the asparagine-linked oligosaccharide precursor to its mature structure (Participants of the First International Workshop on CDGS, 2000).

We have previously reported on a novel type of CDG (type IIb) in a neonate that was shown to be caused by a deficiency of processing glucosidase I (De Praeter et al., 2000Go). The clinical course of the disease was progressive and characterized by hepatomegaly, hypoventilation, feeding problems, seizures, and fatal outcome at 74 days after birth. The patient was compound-heterozygous for two mutations in the glucosidase I gene with a G->C transition in the maternal allele and a T->C transition in the paternal allele, resulting in the substitution of arginine-486 by threonine and phenylalanine-652 by leucine, respectively. Each substitution gave rise to an inactive enzyme protein. In this article, we study the functional and cellular consequences associated with this glucosidase I deficiency in cultured fibroblasts of the patient, including potential effects on other processing enzymes in the glycosylation pathway and on the structure of N-linked glycans.


    Results
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Structural analysis of high-mannose oligosaccharides synthesized in fibroblasts
Removal of the glucose residues from the protein-bound Glc3-Man9-GlcNAc2 precursor is an essential step in N-linked oligosaccharide processing (Kornfeld and Kornfeld, 1985Go). To study the accumulation of Glc3-containing glycan intermediates caused through lack of glucosidase I activity, fibroblast cells from the patient, both parents, and a control were cultured in the presence of [3H]-mannose, followed by cell lysis and isolation of [3H]glycoproteins after extraction with chloroform/methanol/water (3:2:1, v/v/v). Radiolabeled N-glycoproteins were then treated with either endo H, which releases N-linked high-mannose oligosaccharides, or with PNGase F, which removes most types of N-linked glycan. Irrespective of the source of the fibroblasts used, we found that endo H released ~50% of the radioactively labeled carbohydrate, compared to almost 100% for PNGase F (data not shown). This observation indicates that > 50% of the N-linked oligosaccharides must have been processed beyond the high-mannose stage, yielding complex type sugars, not only in the parental and control cells but also in those from the patient.

High-performance liquid chromatography (HPLC) analysis of the endo H–susceptible [3H]glycan fraction obtained from control fibroblasts indicated a range of high-mannose oligosaccharides from Man5-GlcNAc to Glc1-Man9-GlcNAc, with the Man9–6-GlcNAc structures being in the majority (Figure 1A). The relative amount of Man9-GlcNAc, Man8-GlcNAc, Man7-GlcNAc, and Man6-GlcNAc was estimated to be 23%, 27%, 19%, and 22%, respectively, with only 6% of Man5-GlcNAc and 3% of Glc1-Man9-GlcNAc, based on the total number of mannose residues (Figure 1E). An identical pattern of high-mannose intermediates was found after endo H treatment of the [3H]glycoprotein fraction isolated from father’s (Figure 1B and 1E) and mother’s fibroblast cells (Figure 1C and 1E). By contrast, the [3H]oligosaccharide pattern derived from N-glycoproteins synthesized by the patient’s fibroblasts differed, in particular, by containing two larger species (Figure 1D; Pk-2 and Pk-3) as well as an increased proportion of an intermediate that was eluted from the column in the position of Glc1-Man9-GlcNAc (Pk-1). Apart from these larger compounds, however, ~66% of the high-mannose glycans were converted to Man9–5-GlcNAc. Furthermore, the relative ratio of these intermediates to one another was similar to that seen in control cells and fibroblast cells from both parents (Figure 1E).



View larger version (44K):
[in this window]
[in a new window]
 
Fig. 1. Analysis of the endo H–released [3H]oligosaccharides. Fibroblast cells from a control (A), the father (B), the mother (C), and the patient (D) were cultured for 24 h in the presence of [3H]mannose. Radiolabeled glycoproteins were then isolated by chloroform/methanol/water extraction, followed by cleavage with endo H and analysis of the released [3H]oligosaccharides by HPLC. M5, M9, and G3M9 denote elution positions of [14C]Man5-GlcNAc, [14C]Man9-GlcNAc, and [14C]Glc3Man9-GlcNAc, which were added as internal standards for each HPLC run. Peaks occurring only in the patient’s [3H]glycans are numbered and referred to in the text as Pk-1, Pk-2, and Pk-3. The bar chart (E) shows the relative proportion of [3H]labeled high-mannose intermediates in A (open), B (light gray), C (dark gray), and D (black), corrected for the total number of [3H]mannoses.

 
To determine the structure of Pk-1, Pk-2, and Pk-3, aliquots of the corresponding HPLC fractions were incubated with either endo-{alpha}1,2-mannosidase or glucosidase I/glucosidase II and the cleavage products analyzed by HPLC. As shown in Figure 2, Pk-3 was degraded specifically by endo-{alpha}1,2-mannosidase to Glc3-Man and Man8-GlcNAc (panel A), whereas glucosidase I/glucosidase II treatment yielded Man9-GlcNAc as the major cleavage product (panel B), consistent with Pk-3 being Glc3-Man9-GlcNAc. The minor amount of Glc1-Man9-GlcNAc still present (panel B), may be due to incomplete deglucosylation. Incubation of Pk-2 with endo-{alpha}1,2-mannosidase or glucosidase I/glucosidase II gave either Glc3-Man and Man7-GlcNAc or Man8-GlcNAc as cleavage products, respectively, showing that Pk-2 is Glc3-Man8-GlcNAc rather than Glc2-Man9-GlcNAc (panels C and D). Endo-{alpha}1,2-mannosidase cleavage of Pk-1, on the other hand, resulted in the formation of Glc1-Man, Glc3-Man, Man6-GlcNAc, and Man8-GlcNAc as the main products (not shown). Thus the Pk-1 fraction, unlike Pk-2 and Pk-3, is heterogeneous but contains Glc3-Man7-GlcNAc and Glc1-Man9-GlcNAc as major components. The [3H]oligosaccharide eluting in position M9 (see Figure 1D) was not susceptible to either endo-{alpha}1,2-mannosidase or glucosidase I/II, indicating that it is Man9-GlcNAc rather than a glucose-substituted, partially demannosylated intermediate. Considered together these data show that, in contrast to the high-mannose fraction isolated from control and parental fibroblasts, approximately 33% of the high-mannose [3H]glycans synthesized in patient’s fibroblasts still contain the Glc3 unit, attached to either Man9-GlcNAc or to a partially demannosylated Man7 and Man8 core.



View larger version (25K):
[in this window]
[in a new window]
 
Fig. 2. Structural characterization of the Pk-2 and Pk-3 glycans in the high-mannose fraction isolated from the patient’s glycoproteins. Aliquots of purified Pk-3 and Pk-2 (see Figure 1D) were incubated with either endo-{alpha}1,2-mannosidase (A and C) or with glucosidase I and II (B and D) as described in Materials and methods. Cleavage products were then separated by HPLC. M7M9 denote [3H]Man7–9-GlcNAc, G3M Glc3-[3H]Man, G1M9 Glc-[3H]Man9-GlcNAc, and G1M8 Glc-[3H]Man8-GlcNAc. Elution of [14C]labeled oligosaccharide standards is marked by triangles; the elution position for undigested Pk-2 and Pk-3 is indicated.

 
Activity of endo-{alpha}1,2-mannosidase and other processing enzymes
To analyze whether the lack of glucosidase I activity may interfere with the expression of other processing enzymes in the N-glycosylation pathway, we measured the activity of OST, glucosidase I, glucosidase II, Man9-mannosidase, and endo-{alpha}1,2-mannosidase in cultured fibroblast cells of the patient, comparing these with fibroblast cells derived from the parents and a healthy individual. The results of the measurements are summarized in Figure 3. As shown in panel A, incubation of detergent extracts from control fibroblasts with [14C]Glc3-Man9-GlcNAc2 resulted in a rapid and time-dependent release of [14C]glucose by glucosidase I; however, substrate degradation was found to be reduced by approximately twofold in cell extracts from either the father or the mother. This observation is consistent with previous data showing that both parents are heterozygous for the glucosidase I defect (De Praeter et al., 2000Go). Under identical assay conditions, no substrate hydrolysis by glucosidase I was measurable in detergent extracts of cultured fibroblasts from the patient. On extending the incubation time to 5 h, approximately 2% free [14C]glucose was detectable in the assay mixture. Assuming that this 2% originated from [14C]Glc3-Man9-GlcNAc2 degradation by glucosidase I, it can be estimated that the residual glucosidase I activity in fibroblast cells of the patient must be significantly lower than 1% of the control value.



View larger version (23K):
[in this window]
[in a new window]
 
Fig. 3. Catalytic activity of processing enzymes in cultured fibroblasts. Aliquots of detergent extracts prepared from cultured fibroblasts of the patient (closed circles), the father (triangles), the mother (plus signs), and control cell lines (open circles, open squares) were incubated in the presence of [14C]labeled oligosaccharides. After given times, reaction were stopped by the addition of acetic acid and substrate hydrolysis by glucosidase I (A), glucosidase II (C), Man9-mannosidase (D), and endo-{alpha}1,2-mannosidase (E and F) determined as detailed in Materials and methods. Oligosaccharyltransferase activity (B) was measured by using N-benzoyl-Asn-Gly-Thr-NHCH3 as acceptor peptide and Dol-PP-[3H]GlcNAc2 as the glycosyl donor (Bause et al., 1995Go). Each point represents the average of duplicate measurements; within each set of measurements the rates are normalized to the same protein content, but this is not true for comparisons between different sets of measurements.

 
In contrast to the substantial differences for glucosidase I, comparable activities were measured for OST (Figure 3B), glucosidase II (panel C), and Man9-mannosidase (panel D), indicating that these processing activities are not affected by the glucosidase I defect. Interestingly, however, the activity of endo-{alpha}1,2-mannosidase, known to degrade [14C]Glc31-Man9-GlcNAc2 to [14C]Glc3–1-Man and Man8-GlcNAc2 specifically (Lubas and Spiro, 1988Go), was increased by two- to threefold in the patient’s fibroblasts compared to control cells, with intermediate cleavage rates for both parents. The time course of [14C]Glc3-Man9-GlcNAc2 degradation by endo-{alpha}1,2-mannosidase is shown for two independent experiments in panels E and F (note the different time scales). The deviation from linearity of [14C]Glc3-Man9-GlcNAc2 degradation, seen at longer incubation times (60 h, panel E), is probably due to both substrate depletion and instability of endo-{alpha}1,2-mannosidase activity in the detergent extracts. Based on initial cleavage rates (E and F), the relative endo-{alpha}1,2-mannosidase activity was 1.0, ~1.8, and ~2.5 in control, parental, and patient’s fibroblasts, respectively. This suggests that the glucosidase I/II-mediated deglucosylation pathway, which is not functional in the patient’s fibroblasts, can be compensated for (at least partially) by increased expression of endo-{alpha}1,2-mannosidase activity, consistent with and explaining the massive accumulation of Glc3-Man in the patient’s urine (De Praeter et al., 2000Go). The apparent discrepancy between the lack of change in endo-{alpha}1,2-mannosidase activity reported by us previously (De Praeter et al., 2000Go) and the results reported herein is explained simply by our using more 14C-labeled substrate and more cell protein for the current measurements. Although small differences were seen in the original results, these were below the limits of reliability.

Immunoblot analysis of glucosidase I in fibroblasts
Aliquots of detergent extracts prepared from cultured fibroblasts of the parents, the patient, and a healthy individual were subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE), followed by immunoblotting using a polyclonal antibody against glucosidase I for detection. The results in Figure 4 show that the anti–glucosidase I antibody stained a 95-kDa protein, identical in molecular mass to that of human glucosidase I overexpressed in COS 1 cells (panel A, lane 1) with a similar staining intensity for control (lane 2) and for maternal fibroblast cells (lane 4), whereas the immunoreaction was markedly reduced in cell homogenates from fibroblasts of the patient and the father (lanes 3 and 5). This suggests that the inactive form of glucosidase I, encoded for by the paternal allele and containing the F652L mutation, appears to be less stable than the R486T mutant protein encoded for by the maternal allele. This difference in antibody response is unlikely to be caused by unequal protein loading of the gel as shown by the observation that under identical loading conditions immunostaining of the glucosidase II-{alpha} subunit was similar in all cell samples (panel B, lanes 2–5) (Hentges and Bause, 1997Go).



View larger version (27K):
[in this window]
[in a new window]
 
Fig. 4. Immunoblot analysis of glucosidase I and glucosidase II in fibroblast cells. Detergent extracts prepared from control cells (lane 2), fibroblast cells of the patient (lane 3), the mother (lane 4), and the father (lane 5) were subjected to SDS–PAGE, followed by electrophoretic transfer of the proteins onto nitrocellulose. Immunostaining was carried out using an affinity purified antibody raised against either glucosidase I (A) or glucosidase II (B). Lane 1, glucosidase I overexpressed in COS 1 cells. The identity of the cross-reacting material in (A), which appears to be fibroblast-specific, is at present unknown.

 
RT-PCR analysis of glucosidase I- and endo-{alpha}1,2-mannosidase-specific mRNA transcripts
The amounts of glucosidase I- and endo-{alpha}1,2-mannosidase-specific mRNA transcripts were determined by reverse transcription polymerase chain reaction (RT-PCR) in fibroblasts of the patient and a healthy individual to analyze whether the transcription level may be altered by the expression of inactive glucosidase I. As an internal control, RT-PCR was carried out for glyceraldehyde phosphate dehydrogenase (GAPDH) transcripts (for primers used, see Materials and methods). The PCR products were separated by gel electrophoresis and stained with ethidium bromide. The results are summarized in Figure 5 and show that enzyme-specific cDNA fragments are amplified in a cycle-dependent manner. Quantification of the PCR products by Southern blotting using radioactively labeled cDNA probes pointed to a similar level of glucosidase I-specific cDNA/mRNA in control and patient’s fibroblasts, whereas it appeared that the amount of endo-{alpha}1,2-mannosidase-specific transcripts was slightly higher (1.5- to 2.5-fold) in the patient’s fibroblasts relative to the amount of the GAPDH-specific PCR products in control and patient’s cells. This observation would be in line with endo-{alpha}1,2-mannosidase activity being two- to threefold higher in the patient.



View larger version (32K):
[in this window]
[in a new window]
 
Fig. 5. Analysis of glucosidase I- and endo-{alpha}1,2-mannosidase-specific transcripts by RT-PCR. Total RNA isolated from fibroblast cells of the patient and a control was reverse-transcribed into cDNA and aliqots of the cDNA subjected to PCR using primer combinations specific for either glucosidase I, endo-{alpha}1,2-mannosidase or GAPDH. The PCR products obtained after given reaction cycles were separated by gel electrophoresis and visualized by ethidium bromide staining.

 
Characterization of glucosidase I mutants after expression in COS 1 cells
The open reading frame (ORF) in the glucosidase I–specific cDNA encoded for a polypeptide of 836 amino acids corresponding to a molecular mass of ~92 kDa (Kalz et al., 1995Go). To study potential effects on structural and functional parameters caused by the R486T and F652L mutations, corresponding base exchanges were introduced into the cDNA of the human wild-type enzyme by site-directed mutagenesis. COS 1 cells were then transfected with the cDNAs subcloned into pSV.SPORT 1 (pSV-GIWT, pSV-GIR486T, and pSV-GIF652L) and the overexpressed proteins characterized by immunoblotting and measuring their catalytic activity. The results of a typical immunoblot analysis is shown in Figure 6A. The anti–glucosidase I antibody stained a ~95-kDa protein specifically in detergent extracts from COS 1 cells overexpressing either wild-type glucosidase I (lane 2), the R486T (lane 3), or the F652L mutant (lane 4), confirming the structures of the vector constructs. Under the loading conditions used, no immunostaining of the endogenous enzyme was detectable when COS 1 cells were transfected with pSV.SPORT1 (lane 1, control).



View larger version (26K):
[in this window]
[in a new window]
 
Fig. 6. The glucosidase I R486T and F652L mutants are overexpressed in COS 1 cells as inactive enzyme proteins. COS 1 cells transfected with either pSV-SPORT 1 (control), pSV-GIWT, pSV-GIR486T, or pSV-GIF652L were cultured for 48 h and the overexpressed enzyme proteins analyzed by immunoblotting (A) and by measuring their catalytic activity using [14C]Glc3-Man9-GlcNAc2 as substrate (B). (A) Lane 1, control; lane 2, wild-type glucosidase I; lane 3, R486T-mutant; lane 4, F652L-mutant. (B) (open circles), control; (closed circles), wild-type glucosidase I; (squares), R486T-mutant; (triangles), F652L-mutant. Each point represents the average of duplicate measurements.

 
As shown in Figure 6B, COS 1 cells overexpressing wild-type glucosidase I exhibited an approximately 15-fold higher glucosidase I activity compared with control cells. Essentially no increase of [14C]Glc3-Man9-GlcNAc2 hydrolysis over control values was detectable after transfection with either the R486T- or the F652L-specific vector constructs, although both protein mutants were overexpressed efficiently (Figure 6A and 6B). Thus, substitution in the glucosidase I polypeptide of either arginine-486 by threonine or phenylalanine-652 by leucine both lead to a catalytically inactive glucosidase I protein, consistent with the failure to detect enzyme activity in fibroblast cells of the patient.

Binding of glucosidase I mutants to CP-dNM-Sepharose
The two inactive R486T and F652L protein mutants were tested for their ability to bind to a glucosidase I–specific affinity resin; this resin contained N-5-carboxypentyl-1-deoxynorjirimycin covalently attached to AH-Sepharose (CP-dNM-Sepharose). CP-dNM-Sepharose has previously been shown to bind glucosidase I specifically, allowing efficient purification of the enzyme from calf and pig liver (Hettkamp et al., 1984Go; Bause et al., 1989Go). The binding experiments were carried out batchwise by adding the affinity resin to the detergent extracts prepared from transfected COS 1 cells. After 12 h, the affinity resin was separated by centrifugation and bound enzyme protein eluted with 100 mM dNM in binding buffer. Aliquots of the detergent extracts before and after affinity binding, as well as the dNM eluate, were then analyzed by immunoblotting.

The results, summarized in Figure 7, show that wild-type glucosidase I as well as the inactive R486T and F652L enzyme mutants were overexpressed efficiently (panel A, lanes 1–3). After CP-dNM-Sepharose treatment the supernatant was found to be depleted significantly in wild-type glucosidase I (panel B, lane 1), whereas the amounts of both the R486T and F652L proteins were not severely reduced (lanes 2 and 3). This shows that the catalytically active and inactive enzyme species must differ substantially in their affinity for the CP-dNM-Sepharose. As expected, the wild-type enzyme could be eluted from the affinity resin by 1-dNM (panel C, lane 1), whereas the F652L protein was essentially not detectable in the dNM eluate fraction (panel C, lane 3). By contrast, a specific ~95-kDa protein band was observed on the immunoblot in case of the R486T mutant, indicating that this protein does have some affinity for CP-dNM-Sepharose (panel C, lane 2). Thus the R486T mutation interfers less severely with active site structure than does the F652L mutation.



View larger version (48K):
[in this window]
[in a new window]
 
Fig. 7. Wild-type glucosidase I and glucosidase I mutants show a different affinity for CP-dNM-Sepharose. COS 1 cells transfected with pSV-SPORT 1, pSV-GIWT, pSV-GIR486T, or pSV-GIF652L were cultured for 48 h and the cells solubilized in 200 mM sodium phophate buffer, pH 6.5, containing 1% Triton X-100 and 10 µM phenylmethanesulfonylchloride. After addition of CP-dNM-Sepharose the suspensions were stirred for 12 h, followed by separation of the affinity resin and elution of bound enzyme proteins with 100 mM dNM. Aliquots of detergents extracts before (A) and after (B) affinity binding, as well as aliquots of the dNM eluate (C), were subjected to SDS–PAGE and immunoblotting using an anti–glucosidase I antibody. Lane 1, wild-type glucosidase I; lane 2, R486T-mutant; lane 3, F652L-mutant.

 
Exon–intron structure of the human glucosidase I gene and position of the point mutations
The patient’s glucosidase I defect is caused by a T->C transition in the paternal allele and a G->C transition in the maternal allele (De Praeter et al., 2000Go). We analyzed the exon–intron structure of the human glucosidase I gene to assign the two base mutations to specific peptide domains. A human placental genomic DNA library in EMBL3 was screened by hybridization using glucosidase I–derived cDNA fragments. A specific {lambda} clone containing a ~15-kbp insert was isolated, from which appropriate subfragments were generated by restriction cleavage, followed by subcloning into pUC BM20. Based on overlapping sequences in the three independent clones gPBN1, gPEE1, and gPBH1, a 4.4-kbp genomic DNA could be reconstructed representing the complete cDNA sequence for glucosidase I (Figure 8). Comparative analysis of these sequences showed that the cDNA sequence was distributed among four exons, with exon-1 covering the 5' untranslated region as well as part of the ORF encoding for the N-terminal cytosolic polypeptide and the transmembrane domain; exon-2 and exon-3, on the other hand, encode rather short polypeptides comprising 75 and 65 amino acid residues, respectively; the bulk of the luminal polypeptide (~64 kDa) assumed to be responsible for the catalytic activity of the enzyme, is encoded for by exon-4. Thus the structural organization of the glucosidase I gene shows a relationship to the functional domains of the enzyme (Khan et al., 1999Go).



View larger version (16K):
[in this window]
[in a new window]
 
Fig. 8. Schematic representation of the human glucosidase I gene. Coding sequences are shown as black boxes, 5'- and 3'-untranslated regions of the glucosidase I cDNA as gray boxes. Exon I encodes for the cytosolic N-terminus and the transmembrane domain, exon IV encodes for the ~64-kDa catalytic domain. Arg-486 and Phe-652 denote the amino acid substitutions caused by the G->C transition in the maternal allele and the T->C transition in the paternal allele, respectively.

 
The two mutations in the cDNA/mRNA of the glucosidase I from the patient that give rise to the expression of inactive enzyme proteins are both located in exon-4, pointing up the catalytic significance of the ~64-kDa domain. Comparison with other {alpha}-glucosidases revealed a substantial degree of homology within the ~64-kDa polypeptide sequence ranging from > 84% for rat and mouse glucosidase I to 23% for glucosidase I from yeast and 36–37% from A. thaliana and C. elegans, suggesting a degree of evolutionary conservation. The R486T mutation was located within a highly conserved stretch of ~35 amino acids, which was identical in rat and mouse glucosidase I and 57% to 69% homologous in the other three enzymes. The 35-residue peptide always contained arginine-486, indicating a potential role for this sequence. The 26-amino-acid region adjacent to the F652L mutation was identical in rat and mouse glucosidase I, but the enzymes from yeast, C. elegans, and A. thaliana showed a significantly lower degree of homology (25% to 42%). It is remarkable, however, that phenylalanine-652 was never replaced by an amino acid other than tyrosine, indicating that an aromatic amino acid is essential in this particular position.


    Discussion
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
This article describes the results of structural analysis of N-linked oligosaccharides synthesized in fibroblasts from a patient with glucosidase I deficiency, as well as the biochemical and genetic characterization of two amino acid substitutions, previously identified to be the molecular cause for the glucosidase I defect (De Praeter et al., 2000Go). Differential analysis of [3H]mannose-labeled N-glycoproteins isolated from cultured fibroblasts showed that endo H released approximately 50% and PNGase F almost 100% of the N-linked [3H]glycans, for the patient as well as for both parents and a control. We found that the relative ratio of Man9–5-GlcNAc intermediates in the endo H–susceptible fraction was similar, differing only for the patient by the occurence of ~33% Glc3-Man9–7-GlcNAc. These observations indicate that > 80% of N-glycoproteins synthesized in the patient’s fibroblasts are correctly processed, containing carbohydrate chains similar to those found in control and parental fibroblasts. Because removal of Glc3 from Glc3-Man9-GlcNAc2 is essential for precursor maturation and because we were unable to detect glucosidase I activity in patient’s cell extracts, deglucosylation of the oligosaccharide precursor must use an alternative pathway involving Golgi-located endo-{alpha}1,2-mannosidase. This is supported by (1) the patient excreting large amounts of a urinary Glc3-Man tetrasaccharide and (2) by the activity of endo-{alpha}1,2-mannosidase, as well as the level of enzyme-specific mRNA transcripts, increasing two- to threefold in the patient’s fibroblasts.

The observed increase in endo-{alpha}1,2-mannosidase activity indicates that the glucosidase I- and endo-{alpha}1,2-mannosidase-mediated pathways are not operating independently. It seems rather that their individual processing capacity may be up- or down-regulated depending on the particular situation within the cell. In contrast to the observed differences for endo-{alpha}1,2-mannosidase, neither the activity of OST nor the activities of other processing enzymes downstream from the glucosidase I block (including glucosidase II and Golgi-Man9-mannosidase), were affected in the patient’s fibroblasts. These data, together with the structural information provided by the endo H/PNGase F digestion experiments, lend support to the supposition that, independently of the glucosidase I deficiency, relatively "normal" N-glycosylation by OST, as well as further remodeling of the Glc3-Man9-GlcNAc2 precursor to cell-specific glycan structures, should take place in the patient’s fibroblasts as well as in control or parental cells; this will occur as long as the glucose residues are removed by endo-{alpha}1,2-mannosidase. It is not surprising, therefore, that the serum sialotransferrin pattern determined by IEF was found to be normal in the patient.

The presence in the endo H–susceptible fraction of approximately 16% N-linked glycans containing Glc3 indicates on the other hand that endo-{alpha}1,2-mannosidase is not capable of fully compensating for the glucosidase I defect. Several mechanisms could be used to explain this observation: (1) the processing capacity of endo-{alpha}1,2-mannosidase is (despite a two- to threefold increase) still too low to allow complete precursor degradation (see incubation times in Figure 3D and F); (2) the N-glycoproteins that still contain Glc3 are not recognized and accepted as substrates; and (3) the N-glycoprotein fraction is retained in the ER and thus is not accessible to endo-{alpha}1,2-mannosidase, which is located in the Golgi (Lubas and Spiro, 1987Go; Zuber et al., 2000Go). This spatial restriction obviously does not hold for the bulk of N-glycoproteins in the patient’s fibroblasts, which have acquired correctly processed N-glycan chains. Transport of Glc3-Man9-GlcNAc2-containing N-glycoproteins to the Golgi and removal of the Glc3-unit in the Golgi would imply that this N-glycoprotein fraction is not subject to the ER-resident quality control system (Ellgaard and Helenius, 2001Go). This suggests that the proportion of cellular N-glycoproteins, processed by the ER system, is smaller than generally expected. It should be noted, however, that these conclusions are derived from structural and enzymatic data determined in fibroblast cells and may not necessarily reflect the situation in other cell types. Recent studies using various cell lines showed that utilization of the endo-{alpha}1,2-mannosidase-mediated deglucosylation route was cell type–specific with catalytic activity differing over a surprisingly wide range (Karaivanova et al., 1998Go; Dong et al., 2000Go).

The endo-{alpha}1,2-mannosidase-catalyzed deglucosylation pathway fails to explain the occurence of Man9-GlcNAc2 in the endo H–susceptible glycan fraction from the patient, suggesting that this intermediate may be generated by the concerted action of glucosidase I and II. This seems rather unlikely, however, because we were unable to detect glucosidase I activity in the patient’s cell extracts. Also, neither the R486T nor the F652L glucosidase I protein mutant were found to display measurable glucosidase I activity when overexpressed in COS 1 cells. Thus deglucosylation of Glc3-Man9-GlcNAc2 or formation of Man9-GlcNAc2 may occur by an as-yet-unknown mechanism. A likely explanation is that OST transfers not only natural Glc3-Man9-GlcNAc2 but also nonglucosylated or Glc2–1-containing Man9-GlcNAc2 structures, as observed in lower eukaryotes and mammalian cells (Parodi, 2000Go; Romero and Herscovics, 1986Go). Also, it is possible that Man8-GlcNAc2 generated by endo-{alpha}1,2-mannosidase may be subject to remannosylation as part of a quality control mechanism for protein folding in the Golgi. This would be comparable to the de-/reglucosylation system in the ER but involving {alpha}1,2-specific mannosidases and mannosyltransferases as central components (Ellgaard and Helenius, 2001Go).

Glucosidase I is an ER-resident type II membrane protein containing a short cytosolic tail, a highly hydrophobic transmembrane sequence and a large catalytic domain directed towards the lumen. The two base mutations in the glucosidase I gene resulting in the expression of inactive enzyme proteins are located within exon-4, encoding for the catalytic domain of the enzyme. Both the substitution of Arg486 by Thr and of Phe652 by Leu affect positions which are highly conserved in the glucosidase I polypeptide, pointing to the functional significance of these regions for catalysis. Several reasons can be put forward to explain loss of catalytic activity: (1) both amino acid substititions located in the ~64-kDa domain of the glucosidase I protein, interfere with polypeptide folding; or (2) arginine-486 and/or phenylalanine-652 are directly involved in substrate binding and/or catalysis. The observation that the glucosidase I mutant carrying the Arg486-Thr substitution maintained some affinity for CP-dNM-Sepharose indicates that substrate binding is less affected for this mutation than for the Phe652-Leu mutation. Although a direct involvement of Arg486 and Phe652 in substrate binding and/or catalysis cannot be excluded by these data, it appears more likely that failure of the glucosidase I chain to adopt its native conformation may be the key for the loss of activity in the mutants. The weak binding of the Arg486-Thr glucosidase I mutant to the affinity resin shows, on the other hand, that this protein is still able to recognize the affinity ligand. It may also have residual catalytic activity responsible for the occurrence of Man9-GlcNAc2 in the high-mannose oligosaccharide fraction, although we have been unable to reliably measure any activity given the high endogenous level for glucosidase I in COS 1 cells.

Our data show that the endo-{alpha}1,2-mannosidase-dependent deglucosylation pathway in the patient is able to bypass the glucosidase I defect to a considerable extent. The observation that ~16% of N-linked glycans appear as Glc3 intermediates indicates, however, that sequential deglucosylation by glucosidase I and II in the ER is essential for the oligosaccharide maturation of certain N-glycoproteins. This highlights the functional significance of the glucosidase I enzyme for normal development and life.


    Materials and methods
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Materials
Materials and chemicals were obtained from the following sources: synthetic oligonucleotides, Dulbecco’s modified Eagle medium, pSV.SPORT1 (Life Technologies); restriction endonucleases, Taq DNA polymerase (MBI Fermentas); U.S.E., mutagenesis kit (Amersham Bioscience); genomic DNA libary from human placenta in EMBL3 (Clontech); pUC-BM 20 vector, rapid ligation kit (Boehringer Mannheim/Roche); DNA gel extraction kit, plasmid isolation kit, NucleosilR-NH2-column (Macherey and Nagel); Omniscript reverse transcriptase (Qiagen); platinum Taq polymerase (Invitrogen); nitrocellulose membranes (Schleicher & Schuell); D-[2-3H]mannose, specific activity 26 Ci/mmol (Hartmann Analytic); PNGase F, endo H (New England Biolabs); and COS 1 cells (Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH). All other chemicals and compounds used were of analytical-grade purity.

Cell culture and COS 1 cell transfection
Fibroblast cells and COS 1 cells were grown at 37°C and 5% CO2 as monolayer cultures in Dulbecco’s modified Eagle medium supplemented with 10% fetal bovine serum, 50 U/ml penicillin, and 100 µg/ml streptomycin. Transfection of COS 1 cells with the glucosidase I–specific vector constructs was carried out using the DEAE-dextran/chloroquine method (Ausubel et al., 1987Go).

[3H]Mannose labeling and structural analysis of N-linked [3H]glycans
Fibroblast cells were grown on 100-mm culture dishes. After the cells had reached ~60% confluency, 1 mCi [3H]mannose (specific activity 26 Ci/mmol) was added to the medium. The cells were cultured for a further 24 h, harvested by scraping, and washed several times with phosphate buffered saline to remove excess [3H]mannose. The cell pellets were then taken up in chloroform/methanol/water (3/2/1 by volume) to precipitate the [3H]labeled glycoproteins, followed by centrifugation for 5 min at 1000 x g (Wessel and Flügge, 1984Go). The precipitate at the chloroform/water interface containing the [3H]glycoprotein fraction was removed and after reextraction solubilized by heating in 2% SDS/water at 80°C for 10 min. Aliquots of the solution were then diluted with 50 mM citrate buffer, pH 5.5, and with 50 mM sodium phosphate buffer, pH 7.5, followed by incubation with endo H and PNGase F, respectively (Bieberich and Bause, 1995Go). After 12 h at 25°C, the reaction mixtures were made biphasic by the addition of chloroform/methanol/water (3/2/1 by volume). Released [3H]oligosaccharides, which partitioned into the aqueous upper phase, were measured by scintillation counting. The [3H]labeled high-mannose intermediates obtained after endo H cleavage were separated by HPLC on a NucleosilR-NH2-column as previously described (Schweden et al., 1986Go; Bause et al., 1992Go).

For structural characterization of the Pk-3, Pk-2, and Pk-1 glycans present in the endo H–susceptible [3H]oligosaccharide fraction from the patient, aliquots of the purified compounds were incubated either with detergent extracts prepared from COS 1 cells overexpressing glucosidase I and II or with detergent extracts prepared from pig liver microsomes (endo-{alpha}1,2-mannosidase). Incubations were carried out in the presence of either 2 mM 1-deoxymannojirimycin (glucosidase I/II assay) or in the presence of 2 mM 1-dNM and 1 mM 1-deoxymannojirimycin (endo-{alpha}1,2-mannosidase assay) to prevent nonspecific substrate degradation (Hettkamp et al., 1984Go; Bause and Burbach, 1996Go).

Enzyme assays
Cultured fibroblasts were taken up in 50 mM sodium phosphate, pH 6.8, containing 1% Thesit. After disruption with ultrasound, the suspensions were kept on ice for 30 min and then centrifuged at 5000 x g for 10 min. The activity of {alpha}-glycosidases was measured by incubating 10–40-µl aliquots of the resulting supernatants with [14C]labeled oligosaccharide (see methods discussed later). Reactions were run at 37°C and stopped at given times by the addition of an equal volume of acetic acid, followed by separation of the cleavage products by paper chromatography using acetic acid/methanol/water (Hettkamp et al., 1984Go; Bause and Burbach, 1996Go). The activities of glucosidase I, glucosidase II, and Man9-mannosidase were measured using 500 cpm of [14C]Glc3-Man9-GlcNAc2, [14C]Glc1-Man9-GlcNAc2, and [14C]Man5-GlcNAc2, respectively; the {alpha}-mannosidase assay was supplemented with 1 mM CaCl2 (Schweden and Bause, 1989Go). Endo-{alpha}1,2-mannosidase activity was determined using 2000 cpm of [14C]Glc3-Man9-GlcNAc2 in the presence of 1 mM ethylenediamine tetra-acetic acid and 2 mM 1-dNM to inhibit nonspecific degradation of [14C]Glc3-Man9-GlcNAc2 by glucosidase I and II (Bause and Burbach, 1996Go).

OST activity was measured using fibroblast cells suspended in 50 mM Tris–HCl buffer, pH 7.5, containing 500 mM sodium acetate, 10 mM MnCl2, 1 M sucrose, and 0.8% Triton-X100. After centrifugation, 40-µl aliquots of the detergent extract were added to 60 µl lysis buffer (see procedure previously dicussed) containing 2000 cpm of Dol-PP-[14C]GlcNAc2 and 1.7 mM acceptor tripeptide (N-benzoyl-Asn-Gly-Thr-NHCH3). Reactions were carried out at 25°C and stopped at given times by the addition of 500 µl methanol. Isolation and quantification of [14C]glycopeptides were done as described previously (Bause et al., 1995Go).

Binding of glucosidase I mutants to CP-dNM-Sepharose
COS 1 cells grown to ~60% confluency were transfected with 15 µg pSV-GIwt, pSV-GIR486T, and pSV-GIF652L vector DNA using the DEAE-Dextran/chloroquine method. Forty-eight hours after transfection cells were harvested, washed with phosphate buffered saline, and solubilized in 500 µl lysis buffer containing 200 mM sodium phosphate, pH 6.5, 1% Triton-X100, and 10 µM phenylmethanesulfonylfluoride. The detergent extracts were centrifuged at 5.000 gav for 5 min and the resulting supernatants incubated with 30 µl of CP-dNM-Sepharose previously equilibrated in lysis buffer (Hettkamp et al., 1984Go). After stirring the suspension for 12 h at 4°C, the affinity resin was removed by centrifugation and washed several times with 10 ml lysis buffer. Bound glucosidase I protein was then eluted batchwise by treatment of the affinity resin for 3 h at 4°C with 200 µl lysis buffer containing 100 mM 1-dNM. Aliquots of the detergent extracts before and after affinity binding, as well as of the dNM eluate, were then analyzed by SDS–PAGE and immunoblotting.

RT-PCR
Total RNA isolated from cultured fibroblasts using the acidic phenol procedure (Chomczynski and Sacchi, 1987Go), was incubated with DNase I, followed by extraction with phenol/chloroform (1:1 by volume), chloroform, and precipitation by addition of iso-propanol. The RNA was taken up in water previously treated with diethylpyrocarbonate and its quality checked by gel electrophoresis. First-strand cDNA was synthesized from 1 µg of RNA by using the Omniscript reverse transcriptase. Aliquots of the incubation mixture were then subjected to PCR amplification using the platinum Taq polymerase. Amplified cDNA fragments were separated by gel electrophoresis and either visualized by staining with ethidium bromide or quantitated by hybridization with radiolabeled cDNA probes after blotting onto nylon membranes. The following sense/antisense primer combinations were used for cDNA synthesis and PCR amplification: glucosidase I, ggagagtgactgtagagc/ggtccagaagacattgtag; endo-{alpha}1,2-mannosidase, gtgggcccaggatacatagatac/ttatgaaacaggcagctggcgatc; glyceraldehyde-3-phosphate-dehydrogenase, caagaccccttcattgacctc/taagcaggtggtggtgcagga.

Screening for human genomic clones
A human placenta genomic DNA library (in EMBL3) was screened for by plaque hybridization with a 728-bp cDNA fragment, previously synthesized by PCR amplification using the glucosidase I–specific full-length cDNA (Kalz et al., 1995Go). One glucosidase I–specific lambda clone was isolated containing a ~15-kb insert that was processed further by digestion with either BamHI and NcoI, BamHI and HindIII, or EcoRI. Resulting restriction fragments were cloned into pUC-BM 20 and glucosidase I–specific subclones screened for using colony hybridization, followed by sequencing.

Vector construction and generation of glucosidase I mutants
The full-length cDNA encoding human glucosidase I was subcloned into the mammalian expression vector pSV.SPORT1 using common restriction sites (Kalz et al., 1995Go). The recombinant vector (pSV-GIWT) contained the complete 2505-bp ORF of glucosidase I as well as 196 bp and 130 bp from the noncoding 3'- and 5'-sequence, respectively. Using pSV-GIWT as the template, vector constructs containing either the G->C mutation at position 1446 of the ORF (pSV-GIR486T) or the T->C mutation at position 1974 (pSV-GIF652L) were generated by mutagenesis following the Amersham Bioscience manual. Target mutagenic primers of 167 bp and 139 bp length for mutant DNA synthesis were generated by PCR amplification using the sense/antisense primer combinations ctggattgggacggagcagatactg/ctctagcatatgggctacag and gagctaggagtccttgcagactttg/gactgacatagccaagagc, respectively. Base exchanges (shown in boldface type) were verified by sequencing.

General methods
Library screening, subcloning, PCR amplification, and other standard techniques were performed as described elsewhere (Ausubel et al., 1987Go; Sambrook et al., 1989Go; White, 1993Go). SDS–PAGE, western blotting, and immunoblotting were carried out as detailed by Laemmli (1970)Go, Blake et al. (1984)Go, and Schweden and Bause (1989)Go. [14C]Glc3–1-Man9-GlcNAc2, [14C]Man5-GlcNAc2, Dol-PP-[14C]GlcNAc2, and N-benzoyl-Asn-Gly-Thr-NHCH3 were synthesized as described by Hettkamp et al. (1982)Go, Schweden et al. (1986)Go, and Bause et al. (1995)Go. Polyclonal antibodies against glucosidase I and glucosidase II were prepared as detailed by Bause et al. (1989)Go and Hentges and Bause (1997)Go. Synthesis of CP-dNM and preparation of CP-dNM-Sepharose followed the procedures described by Hettkamp et al. (1984)Go.


    Acknowledgments
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
This work was supported by the Deutsche Forschungsgemeinschaft (Sonderforschungsbereich 284). The authors are indebted to R.A. Klein (Universität Bonn) for critical reading of the manuscript.


    Abbreviations
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
CDG, congenital disorder of glycosylation; CP-dNM, N-5-carboxypentyl-1-deoxynorjirimycin; ER, endoplasmic reticulum; GAPDH, glyceraldehyde phosphate dehydrogenase; HPLC, High-performance liquid chromatography; ORF, open reading frame; OST, oligosaccharyltransferase; RT-PCR, reverse transcription polymerase chain reaction; SDS–PAGE, sodium dodecyl sulfate–polyacrylamide gel electrophoresis.


    Footnotes
 
1 To whom correspondence should be addressed; E-mail: bause@institut.physiochem.uni-bonn.de Back


    References
 Top
 Abstract
 Introduction
 Results
 Discussion
 Materials and methods
 Acknowledgments
 Abbreviations
 References
 
Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (1987) Current protocols in molecular biology. John Wiley and Sons, New York.

Bause, E. and Burbach, M. (1996) Purification and enzymatic properties of endo-{alpha}1, 2-mannosidase from pig liver involved in oligosaccharide processing. Biol. Chem., 377, 639–646.[ISI][Medline]

Bause, E., Breuer, W., and Peters, S. (1995) Investigation of the active site of oligosaccharyltransferase from pig liver using synthetic tripeptides as tools. Biochem. J., 312, 979–985.[ISI][Medline]

Bause, E., Breuer, W., Schweden, J., Roeser, R., and Geyer, R. (1992) Effect of substrate structure on the activity of Man9-mannosidase from pig liver involved in N-linked oligosaccharide processing. Eur. J. Biochem., 208, 451–457.[Abstract]

Bause, E., Schweden, J., Gross, A., and Orthen, B. (1989) Purification and characterization of trimming glucosidase I from pig liver. Eur. J. Biochem., 183, 661–669.[Abstract]

Bieberich, E. and Bause, E. (1995) Man9-mannosidase from human kidney is expressed in COS cells as a Golgi-resident type II transmembrane N-glycoprotein. Eur. J. Biochem., 233, 644–649.[Abstract]

Blake, M.S., Johnson, K.H., Russel-Jones, G.J., and Gotschlich, E.C. (1984) A rapid, sensitive method for detection of alkaline phosphatase-conjugated anti-antibody on western blots. Anal. Biochem., 136, 175–179.[ISI][Medline]

Chomczynski, P. and Sacchi, N. (1987) Single-step method of RNA isolation by acid guanidium thiocyanate-phenol-chloroform extraction. Anal. Biochem., 162, 156–159.[CrossRef][ISI][Medline]

De Praeter, C., Gerwig, G.J., Bause, E., Nuytinck, L.K., Vliegenthart, J.F.G., Breuer, W., Kamerling, J.P., Espeel, M.F., Martin, J.J.R., De Paepe, A.M., and others. (2000) A novel disorder in a newborn caused by defective biosynthesis of N-linked oligosaccharides due to glucosidase I deficiency. Am. J. Hum. Genet., 66, 1744–1756.[CrossRef][ISI][Medline]

Dong, Z., Zuber, C., Spiro, M.J., Spiro, R.G., and Roth, J. (2000) Immunohistochemical evaluation of endomannosidase distribution in rat tissues: evidence for cell type–specific expression. Histochem. Cell Biol., 114, 461–467.[ISI][Medline]

Ellgaard, L. and Helenius, A. (2001) ER quality control: towards an understanding at the molecular level. Curr. Opin. Cell Biol., 13, 431–437.[CrossRef][ISI][Medline]

Hentges, A. and Bause, E. (1997) Affinity purification and characterization of glucosidase II from pig liver. Biol. Chem., 378, 1031–1038.[ISI][Medline]

Hettkamp, H., Bause, E., and Legler, G. (1982) Inhibition by nojirimycin and 1-deoxynojirimycin of microsomal glucosidases from calf liver acting on the glycoprotein oligosaccharides Glc1–3-Man9-GlcNAc2. Biosci. Reports, 2, 899–906.[ISI][Medline]

Hettkamp, H., Legler, G., and Bause, E. (1984) Purification by affinity chromatography of glucosidase I, an endoplasmic reticulum hydrolase involved in the processing of asparagine-linked oligosaccharides. Eur. J. Biochem., 142, 85–90.[Abstract]

Kalz, B., Bieberich, E., and Bause, E. (1995) Cloning and expression of glucosidase I from human hippocampus. Eur. J. Biochem., 231, 344–351.[Abstract]

Karaivanova, V.K., Luan, P., and Spiro, R.G. (1998) Processing of viral envelope glycoprotein by the endomannosidase pathway: evaluation of host specificity. Glycobiology, 8, 725–730.[Abstract/Free Full Text]

Khan, F.A., Varma, G.M., and Vijay, I.K. (1999) Genomic organization and promoter activity of glucosidase I gene. Glycobiology, 9, 797–806.[Abstract/Free Full Text]

Kornfeld, R. and Kornfeld, R. (1985) Assembly of asparagine-linked oligosaccharides. Annu. Rev. Biochem., 54, 631–664.[CrossRef][ISI][Medline]

Laemmli, U.K. (1970) Cleavage of structural proteins during assembly of the head of bacteriophage T4. Nature, 227, 680–685.[ISI][Medline]

Lubas, W.A. and Spiro, R.G. (1987) Golgi endo-{alpha}-D-mannosidase from rat liver, a novel N-linked carbohydrate unit processing enzyme. J. Biol. Chem., 262, 3775–3781.[Abstract/Free Full Text]

Lubas, W.A. and Spiro, R.G. (1988) Evaluation of the role of rat liver Golgi endo-{alpha}-D-mannosidase in processing of N-linked oligosaccharide. J. Biol. Chem., 263, 3990–3998.[Abstract/Free Full Text]

Marquardt, T. and Freeze, H. (2001) Congenital disorders of glycosylation: glycosylation defects in man and biological models for their study. Biol. Chem., 382, 161–177.[ISI][Medline]

Parodi, A.J. (2000) Protein glucosylation and its role in protein folding. Annu. Rev. Biochem., 69, 69–93.[CrossRef][ISI][Medline]

Participants of the First International Workshop on CGDS (2000) Carbohydrate-deficient glycoprotein syndromes become congenital disorders of glycosylation: an updated nomenclature for CDG. Glycobiology, 10, iii–iv.[Medline]

Romero, P.A. and Herscovics, A. (1986) Transfer of nonglucosylated oligosaccharide from lipid to protein in a mammalian cell. J. Biol. Chem., 261, 15936–15940.[Abstract/Free Full Text]

Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989) Molecular cloning: a laboratory manual. Cold Spring Habor Laboratory Press, New York.

Schachter, H. (2001) Congenital disorders involving defective N-glycosylation of proteins. Cell Mol. Life Sci., 58, 1085–1104.[CrossRef]

Schweden, J. and Bause, E. (1989) Characterization of trimming Man9-mannosidase from pig liver: purification of a catalytically active fragment and evidence for the transmembrane nature of the intact 65 kDa enzyme. Biochem. J., 264, 347–355.[ISI][Medline]

Schweden, J., Legler, G., and Bause, E. (1986) Purification and characterization of a neutral processing mannosidase from calf liver acting on (Man)9(GlcNAc)2 oligosaccharides. Eur. J. Biochem., 157, 563–570.[Abstract]

Varki, A. (1993) Biological roles of oligosaccharides: all of the theories are correct. Glycobiology, 3, 97–130.[Abstract]

Wessel, D. and Flügge, U.I. (1984) A method for the quantitative recovery of protein in dilute solution in the presence of detergents and lipids. Anal. Biochem., 138, 141–143.[ISI][Medline]

White, B.A. (1993) PCR-protocols—current methods and applications. Human Press, Ottawa, Canada.

Zuber, C., Spiro, M.J., Guhl, B., Spiro, R.G., and Roth, J. (2000) Golgi apparatus immunolocalization of endomannosidase suggests post-endoplasmic reticulum glucose trimming: implication for quality control. Mol. Biol. Cell, 11, 4227–4240.[Abstract/Free Full Text]