(Received for publication, August 12, 1996, and in revised form, December 2, 1996)
From the Department of Biochemistry, Carbohydrate-deficient glycoprotein (CDG)
syndrome type I is a congenital disorder that involves the
underglycosylation of N-glycosylated
glycoproteins (Yamashita, K., Ideo, H., Ohkura, T.,
Fukushima, K., Yuasa, I., Ohno, K., and Takeshita, K. (1993) J. Biol. Chem. 268, 5783-5789). In an
effort to further elucidate the biochemical basis of CDG syndrome type
I in our patients, we investigated the defect in the multi-step pathway
for biosynthesis of lipid-linked oligosaccharides (LLO) by the
metabolic labeling method using [3H]glucosamine,
[3H]mannose, and [3H]mevalonate. The LLO
levels in synchronized cultures of fibroblasts from these patients were
severalfold lower than those in control fibroblasts in the S phase, and
the oligosaccharides released from LLO showed the same structural
composition, Glc1~3·Man9·GlcNAc·GlcNAc, in the case of both the patients and controls. The amount of
[3H]mannose incorporated into mannose 6-phosphate,
mannose 1-phosphate, and GDP-mannose was greater in fibroblasts from
these patients than in the control fibroblasts in the G1
period, although the ratios of these acidic mannose derivatives as
indicated by the relative levels of radioactivity were the same for the
two types of fibroblasts. Furthermore, upon metabolic labeling with
[3H]mevalonate, the level of
[3H]dehydrodolichol in fibroblasts from these patients
increased in the S phase, and the levels of [3H]dolichol
and [3H]dolichol-PP oligosaccharides concomitantly
decreased, although the chain length distribution of the respective
dolichols and dehydrodolichols was the same in the two types of
fibroblasts. These results indicate that the conversion of
dehydrodolichol to dolichol is partially defective in our patients and
that the resulting loss of dolichol leads directly to
underglycosylation.
It has recently been determined that carbohydrate-deficient
glycoprotein (CDG)1 syndrome type I is an
autosomal recessive endoplasmic reticulum disorder (1-3). In 1980, Jaeken et al. (4) first reported this syndrome as
encountered in 2-year-old twin girls with neuroimpairment, cerebellar
atrophy, hepatomegaly, abnormal subcutaneous fat deposition, and
skeletal abnormalities. Both patients had a number of biochemical abnormalities mainly affecting glycoproteins. Later, it was reported that sialic acid-deficient serum and cerebrospinal fluid transferrins are features of this newly recognized genetic syndrome (5), and the
finding of pathologic transferrin heterogeneity led to the
demonstration of a partial deficiency of the
sialyl-N-acetyllactosamine group in serum transferrins (6,
7). More recently, it was elucidated that CDG syndrome type I is a
partial asparagine-N-linked sugar chain transfer deficiency
in endoplasmic reticulum (8-10). However, the fundamental defect
remains unclear, and at least two models, defective mannose uptake (11)
and defective conversion of mannose 6-phosphate to mannose 1-phosphate
(12), have been suggested. In preliminary experiments by the metabolic
labeling method using [3H]mannose, fibroblasts from our
patients with CDG syndrome type I did not accumulate any smaller
sized [3H]mannose-labeled oligosaccharide moieties in LLO
as compared with those from patients in the previously reported
families (11, 13, 14), although most of the clinical features and the
clinical biochemical features in our patients (15) are the same as
those of other groups' families (1, 2). In order to elucidate the
primary defect responsible for CDG syndrome type I in our patients, the
biosynthetic pathway from mevalonate to
Glc3·Man9·GlcNAc2·PP·dolichol in the fibroblasts was investigated by metabolic labeling with [3H]glucosamine, [3H]mannose, and
[3H]mevalonate.
Synchronized cultures of the CDG syndrome type I and control
fibroblasts were used for the metabolic labeling studies, because we
found, using synchronized cultures of rat 3Y1 cells, that the biosynthesis of LLO sharply increases in the S phase and decreases in
the G1, G2, and M periods, concomitantly with
dehydrodolichyl diphosphate synthase
activity.2
In this paper we report that the biosynthesis of LLO sharply
increases in the S phase, and the LLO levels in CDG syndrome type I
fibroblasts are severalfold lower than those in control fibroblasts.
The oligosaccharides released from LLO in both types of fibroblasts
have the same structure,
Glc1-3·Man9·GlcNAc2, which may
result in random failure of glycosylation of the available Asn-X-(Ser/Thr) sites. Furthermore, since the biosynthetic
pathway for dolichol is proposed to be mevalonate [2-3H]Mannose (17.6 Ci/mmol), [1,6-3H]glucosamine·HCl (54.1 Ci/mmol),
GDP-[14C]mannose (286 mCi/mmol), and
[5-3H]mevalonate triethylammonium salt (40 Ci/mmol) were
purchased from DuPont NEN. [6-3H]Thymidine (26.0 Ci/mmol)
was from Amersham Life Sciences, Inc. Diethylaminoethyl cellulose
(DE52) was from Whatman Biosystems Ltd. The Sep-Pak Vac C18 cartridges
and µPorasil silica column (3.9 × 150 mm) were purchased from
Waters. A reverse-phase column, Chemcopak Nucleosil 7C18 column
(4.6 × 250 mm), was from Chemco Scientific Co., Ltd. (Osaka,
Japan). Acid phosphatase from potato was from Sigma. Dehydrodolichol
and dolichol were generous gifts from Kuraray Co. (Japan). Bio-Gel P-4
(extra fine) and AG-50W-X2 (100-200 mesh) were purchased from Bio-Rad.
Concanavalin A-Sepharose was from Pharmacia Biotech Inc. Yeast
hexokinase was obtained from Oriental Yeast Co., Ltd. (Tokyo, Japan).
Methyl- The clinical features of three
patients, A.H., Y.M., and M.M., with CDG syndrome type I, whose
established fibroblast lines were used in this study (15), can be
summarized as follows. The major abnormalities displayed by these
patients were mental retardation, hepatopathy, cerebellar
hypoplasia/atrophy, cerebellar ataxia, polyneuropathy, growth
retardation, and dysmorphic features. The hepatopathy appeared in early
infancy, the cerebellar hypoplasia was congenital, the atrophy
progressed for several years after birth, the polyneuropathy appeared
in childhood, and the growth retardation became apparent in late
childhood. During infancy, internal strabismus was noticed. The
biochemical description is as follows: the pI values of various serum
glycoproteins were higher than those in normal controls. Both the
activities and antigen levels of plasma antithrombin III, protein C,
and protein S were decreased to nearly half normal levels; however, the
activities of lysosomal Primary fibroblast
cultures, established from skin punch biopsies, were grown in
Dulbecco's modified Eagle's medium (Life Technologies, Inc.)
supplemented with 10% fetal bovine serum and serially passaged by
trypsinization. The number of population doublings was calculated as
described previously (18). The culture age was calculated as the
population doubling level (PDL) starting from the first passage
in vitro. All of the experiments were performed at PDL
5-15. After PDL 20, reproducible results could not be obtained. Moreover, the ages of the patients, when the cultured fibroblasts were
established, influenced the data. In this study, cells from three
patients with CDG syndrome type I (A.H., 13 months old; Y.M., 8 months
old; M.M., 9 years old), and cells from age-matched controls were
used.
Synchronized cultures were obtained by the density-arrested culture
method (19). In brief, cells were inoculated into 90-mm plastic dishes
and then cultured to confluence. The confluent cells were refed with
fresh medium and then cultured for an additional 3 days. Then, the
density-arrested cells were stimulated by replating in fresh medium at
a density of 1 × 106 cells per 90-mm dish. At regular
intervals, [3H]thymidine (1 µCi/ml) was added to the
medium, and the cells were labeled for 2 or 3 h. After the cells
had been lysed in 5% sodium dodecyl sulfate, 5% trichloroacetic acid,
the [3H]thymidine-labeled insoluble macromolecules
(i.e. DNA) were trapped on a glass filter (Whatman GF/C).
The size of the cell population entering the S phase was determined
based on the synchronous increase in [3H]thymidine
incorporation. The size of the cell population entering the M period
was determined by counting mitotic cells among total cells.
After metabolic labeling, the
medium containing [3H]glucosamine was immediately
replaced with 4 ml of methanol, 5 mM Tris-HCl buffer (pH
8.0) containing 5 mM EDTA (1:1) at 4 °C; the cells were
scraped off, and 2 ml of chloroform was added, followed by mixing for 2 min. Since the LLO synthesis reaction is very rapid, such immediate
treatment is indispensable for obtaining reproducible results. After
centrifugation, the upper phase and interfacial phases were mixed with
2 ml of chloroform, and the combined lower phases were pooled as the
lipid fraction (chloroform/methanol (2:1, v/v)) (20). The
chloroform/methanol (2:1, v/v) fraction should contain components
ranging from GlcNAc- to
Man2GlcNAc2-pyrophosphate-dolichols. The upper
phase and interfacial phases were mixed with 2 ml of methanol and then
centrifuged, and the precipitate was washed with 6 ml of 50% methanol.
Tritium-labeled LLO were twice extracted from the precipitate with 5 ml
of chloroform/methanol/water (10:10:3, v/v) (CMW) (20).
Oligosaccharides were released from the LLO and lipid fractions by
hydrolysis with 0.5 ml of n-propyl alcohol, 1 ml of 0.01 N HCl at 100 °C for 25 min (20), passed through an AG-3
(OH [3H]Mannose 6-phosphate was prepared as
follows. A 100-µl reaction mixture consisting of 0.1 M
Tris-HCl (pH 7.4), 10 mM MgCl2, 10 mM ATP, 0.1 mCi of [3H]mannose (17.6 Ci/nmol), and 1 unit of yeast hexokinase was incubated at 37 °C for
5 h (22). After boiling for 2 min, the tritium-labeled mannose
6-phosphate was purified by paper electrophoresis at pH 5.4. Yield,
95%. Phosphomannomutase activity was measured as follows. A 50-µl
reaction mixture consisting of 50 mM Tris-HCl (pH 7.4), 2 mM MgCl2, 5 × 104 dpm (25 nM) [3H]mannose 6-phosphate, 1 mM
glucose 1,6-bisphosphate, and cell homogenate (5-10 µg of protein)
of subconfluent CDG syndrome type I or control fibroblasts was
incubated at 37 °C for 30 min. After boiling for 2 min, the
tritium-labeled mannose 1,6-bisphosphate was separated from
[3H]mannose 6-phosphate by paper electrophoresis at pH
5.4. The radioactivity of [3H]mannose 1,6-bisphosphate,
which was diluted with approximately 104 times more glucose
1,6-bisphosphate, was measured as phosphomannomutase activity, because
glucose 1,6-bisphosphate can substitute for mannose 1,6-bisphosphate as
a catalyst (23) and 1/104 of the [3H]mannose
1,6-bisphosphate synthesized should be converted to [3H]mannose 1-phosphate.
Synchronized cultures of
fibroblasts (1 × 106 cells per dish) were
pulse-labeled with 0.25 mCi of [3H]mannose in
glucose-free medium at the respective indicated times for 20 min,
because the rate of incorporation of [3H]mannose into LLO
was constant for at least 20 min. After pulse labeling, the medium in
the dishes was immediately replaced with 7 ml of chilled 50% methanol
in 10 mM Tris-HCl buffer (pH 8.2) containing 1 mM EDTA, and then the cells were scraped off. After 3000 dpm of GDP-[14C]mannose had been added to the tubes as an
internal standard, the mixtures were boiled for 2 min, sonicated five
times for 5-s periods, and then centrifuged. The supernatants were
dried, and the residues were subjected to paper chromatography with
95% methanol, 1 M ammonium acetate (pH 7.5) (5:2, v/v) as
the developing solvent (24). Mannose-monophosphates and GDP-mannose
were extracted and subjected to paper electrophoresis (pH 5.4) at 70 V/cm for 40 min. GDP-[3H]- or [14C]mannose
(Rf, 0.22), and
[3H]mannose-monophosphates (Rf, 0.58)
were separated by paper chromatography with 1-butanol/acetic acid/water
(3:3:2, v/v) as the developing solvent. The radioactive GDP-mannose was further adsorbed to a concanavalin A-Sepharose column equilibrated with
20 mM acetate buffer (pH 3.5) and eluted with 0.1 M methyl- Synchronized cultures of CDG
syndrome type I and control fibroblasts were obtained by the
density-arrested culture method. The density-arrested cells were
replated in fresh medium at the density of 1 × 106
cells per 90-mm dish, and five dishes were used at each point. Synchronization was checked by measuring the incorporation of [3H]thymidine into DNA and by determining the frequency
of mitotic cells among total cells of monolayers in the mitotic phase.
The synchronized cultures were incubated with
[3H]mevalonate (25 µCi/ml) for 27, 30, 36, 42, and
48 h. After the cells had been washed with phosphate-buffered
saline and scraped, they were centrifuged. The packed cells were
homogenized and extracted three times with 10 volumes of 1-butanol
saturated with water (27). The 1-butanol extracts were washed with
water saturated with 1-butanol, dried, and then applied to a DE52
column (bed volume, 1 ml), which had been equilibrated with
1-butanol/methanol (9:1). The pass-through fractions were mixed with
0.1 µg each of dehydrodolichol and dolichol as internal standards and
then dried. The samples were dissolved in methanol and then applied to
a Sep-Pak C18 column (Waters). After washing with methanol, the column
was eluted with n- hexane, and the eluate was analyzed by
HPLC. HPLC was performed using a µPorasil silica column (3.9 × 150 mm) with 3% diethyl ether, 0.2% acetic acid in hexane as the
eluent at a flow rate of 0.6 ml/min (28). Internal standards were
detected at 210 nm, and 0.6-ml fractions were collected. Dehydrodolichol and dolichol are clearly separated by this silica column chromatography, although these polyprenols are not separated according to their chain lengths. The chain lengths of dolichol and
dehydrodolichol were determined on a Chemcopak Nucleosil 7C18 reverse-phase column (4.6 × 250 mm), with isopropyl
alcohol/methanol/hexane/H2O (240:120:40:9) as the eluent at
a flow rate of 0.5 ml/min (29).
LLO from cells metabolically labeled
with [3H]mevalonate was extracted as described under
"Extraction of LLO Metabolically Labeled with
[3H]Glucosamine," and [3H]polyprenols
were released from LLO by hydrolysis with 0.5 ml of n-propyl
alcohol, 1 ml of 0.01 N HCl at 100 °C for 25 min, and
then digested with potato acid phosphatase. The released
[3H]polyprenols were extracted with n-hexane
and then analyzed on a µPorasil silica column (3.9 × 150 mm)
with 3% diethyl ether, 0.2% acetic acid in hexane as the eluent at a
flow rate of 0.6 ml/min.
Dehydrodolichol
diphosphate or dolichol diphosphate was digested with potato acid
phosphatase (30) as follows. A 50-µl reaction mixture consisting of 1 M sodium acetate buffer (pH 5.0), 0.1% Triton X-100, 0.2 mg of acid phosphatase, 40% methanol, and 20% 1-butanol was incubated
at 37 °C overnight.
The defect in the pathway of LLO synthesis
responsible for CDG syndrome type I was investigated by metabolic
labeling with [3H]glucosamine, which is a precursor in
LLO synthesis. Fibroblasts with similar population doubling levels
(5-15) and similar proliferation ability from the patients and control
subjects were synchronized by the density-arrested culture method and
then metabolically pulse-labeled with [3H]glucosamine
(125 µCi/ml) or [3H]thymidine (3 µCi/ml) at 3-h
intervals from 14 to 48 h after replating. In repeated
experiments, reproducible results were obtained only for cultures with
tight synchronization (cultures in which 50-80% of the mitotic cells
were in the mitotic period) at the earlier passage levels (less than 15 PDLs). [3H]Thymidine-labeled DNA was acid-precipitated,
and [3H]glucosaminelabeled LLO were extracted with
CMW as described under "Experimental Procedures." The time course
of incorporation of [3H]thymidine into DNA showed that
the DNA synthesis activity in the synchronized CDG syndrome type I and
control fibroblasts was the same (Fig. 1, dashed
lines), and the time of maximum DNA synthesis was 27-30 h after
replating. Since it takes only several minutes to complete the
conversion of dolichol to
Glc3·Man9·GlcNAc2·PP·Dol (31), labeling for 3 h was considered to be long enough to achieve the steady state level of LLO. The amount of radioactivity incorporated into LLO in the control fibroblasts increased in the S phase (Fig. 1,
When oligosaccharides synthesized at the
respective times were released from
[3H]glucosamine-labeled LLO by mild acid hydrolysis and
examined by Bio-Gel P-4 (extra fine) column chromatography (see
"Experimental Procedures"), the sizes of the oligosaccharides
derived from CDG syndrome type I and control fibroblasts were the same,
and over 90% displayed the structure
Glc1-3·Man9·GlcNAc2 (Fig.
2A), which can be transferred en
bloc to nascent polypeptide chains to initiate
N-glycosylation. No smaller oligosaccharides ranging from
GlcNAc to Man2GlcNAc2 were detected (Fig.
2B). These results indicate that the various enzymes
responsible for the conversion of GlcNAc·P·P-dolichol
to
Glc3·Man9· GlcNAc·GlcNAc·P·P-dolichol are functioning as normal, and some earlier step in the LLO
synthetic pathway may be partially deficient in our CDG syndrome
type I patients. Results similar to those shown in Fig. 2 were obtained upon metabolically pulse labeling with [3H]mannose, and
no smaller sized oligosaccharides were released from LLO (data not
shown). These phenomena are clearly different from findings
previously reported for other groups' patients. Panneerselvam and
Freeze (11), and Krasnewich et al. (14) previously reported
that the initial velocities of [3H]mannose uptake were
2-3-fold less in CDG syndrome type I cells compared with controls
(11), and the size of the lipid-linked oligosaccharide precursor is
much smaller than in controls (11, 13, 14). These results suggest the
possibility that there could be multiple causes of CDG syndrome
type I.
Our findings indicate that CDG syndrome type I in our
patients is caused by a partial deficiency of an enzyme in the earliest steps of LLO biosynthesis. However, Van Schaftingen and Jaeken (12)
recently proposed that this syndrome is caused by a phosphomannomutase deficiency. Thus, we examined whether the fibroblasts in our patients, A.H., Y.M., and M.M., display decreased phosphomannomutase activity, as
seen in the case of CDG syndrome type I patients of European families.
Phosphomannomutase (Enz) mediates the following reactions: mannose
6-phosphate + Enz-P In order to
determine whether the biosynthetic pathway from mannose to GDP-mannose
in vivo is deficient, metabolic labeling with
[3H]mannose was performed using synchronized cultures of
both types of fibroblasts. [3H]Mannose incorporated into
fibroblasts should be sequentially metabolized as follows (Equation 1).
Institute for Chemical Reaction Science,
isopentenyl
diphosphate
geranyl diphosphate
farnesyl diphosphate
dehydrodolichyl diphosphate
dehydrodolichyl monophosphate
dehydrodolichol
dolichol (16, 17), the biosynthetic pathway for
dolichol in relation to LLO in CDG syndrome type I fibroblasts was also
investigated by means of metabolic labeling with
[3H]mevalonate.
Chemicals and Enzymes
-mannoside, mannose 1-phosphate, glucose 1,6-bisphosphate,
and dolichyl phosphate were from Sigma. All other chemicals were of
analytical grade.
-hexosaminidases and
-L-fucosidase in the sera of these patients were
severalfold higher than those in normal controls. These clinical
features and clinical biochemical features in our patients (3, 15) are
the same as those of other reported groups (1, 2), except one of the
characteristic features designated as "fat pads on the buttocks"
was not noticed in our patients.
form) column, and then analyzed by chromatography
using a Bio-Gel P-4 (extra fine) column (2 × 120 cm) (21). The
amount of LLO was calculated based on the amount of radioactivity
corresponding to oligosaccharides ranging from
Glc3·Man9·GlcNAc2 to
Man3·GlcNAc2. Radioactivity was measured with
a Beckman liquid scintillation counter, model LS6000LL.
-mannoside in 10 mM Tris-HCl (pH
7.4) (25). Recovery, ~80%. Quantitative analysis of mannose
1-phosphate and mannose 6-phosphate was performed by hydrolysis of the
mannose-monophosphates with 0.02 N HCl at 100 °C for 30 min, since mannose 1-phosphate is acid-labile and converted to free
mannose, and mannose 6-phosphate is resistant (26).
Cell Cycle-dependent Incorporation of
[3H]Glucosamine into Lipid-linked
Oligosaccharides
), since many membrane-bound glycoproteins are synthesized in the
course of cell proliferation. Although the radioactivity incorporated
into LLO in synchronized cultures of CDG syndrome type I fibroblasts
also increased in the S phase, the amount was 2-fold lower than that in
control fibroblasts in the S phase (Fig. 1,
-
). In contrast, the
amounts of labeled LLO in the G1 and G2 periods
were equally low in fibroblasts from the patients and controls. These
results indicate that the amount of LLO synthesized in the S phase in
CDG syndrome type I fibroblasts is smaller than that in control
fibroblasts, and an insufficient amount of LLO could result in random
failure of glycosylation of the available Asn-X-Ser/Thr
sites. The equivalence of labeling in the G1 and G2 periods shows clearly that the difference in S phase
between normal and CDG syndrome type I fibroblasts is not merely due to a different mannose pool size as described by other groups (11, 13,
14).
Fig. 1.
Cell cycle-dependent
incorporation of [3H]glucosamine into LLO in CDG syndrome
type I and control fibroblasts. Synchronized cultures of patient
A.H. and age-matched control fibroblasts (1 × 106
cells per dish) with 10 PDL were pulse-labeled with
[3H]glucosamine or [3H]thymidine for 3-h
intervals. - - -, [3H]thymidine incorporated into DNA;
, [3H]glucosamine incorporated into LLO.
indicates
control and
indicates patient A.H. Patients Y.M. and M.M. showed
similar results (data not shown).
[View Larger Version of this Image (21K GIF file)]
Fig. 2.
Maturation of
[3H]glucosamine-labeled oligosaccharides in LLO in Fig.
1. Arrows at the top indicate the elution
positions of glucose oligomers (numbers indicate the glucose
units). Black triangles indicate the elution positions of
authentic
Glc3·Man9·GlcNAc· GlcNAc
(a), Glc2·Man9·GlcNAc·GlcNAc
(b), Man5·GlcNAc·GlcNAc (c), Man·GlcNAc·GlcNAc (d), GlcNAc·GlcNAc (e),
and GlcNAc (f). The elution patterns on Bio-Gel P-4
(extra fine) column chromatography of
[3H]glucosamine-labeled oligosaccharides released from
the LLO (A) and the chloroform/methanol (2:1, v/v) fraction
(B) (3-h pulse labeling, 27-30 h after replating) on mild
acid hydrolysis (0.01 N HCl:n-propyl
alcohol = 2:1, at 100 °C for 25 min). Solid line, control; dotted line, patient A.H. The same elution patterns
were observed for LLO at the other respective intervals, and similar results were obtained using fibroblasts from patients Y.M. and M.M.
(data not shown).
[View Larger Version of this Image (24K GIF file)]
Enz + mannose 1,6-bisphosphate
Enz-P + mannose 1-phosphate. The linearity of the rate of product (mannose
1,6-bisphosphate) formation was maintained until approximately 80% of
the substrates had been consumed in our enzyme assay system. Because an
excess amount of glucose 1,6-bisphosphate at concentrations up to 1 mM did not inhibit the phosphomannomutase activity, but acted as a catalyst of this enzyme, we measured the amount of the
synthesized radioactive intermediate, i.e.
[3H]mannose 1,6-bisphosphate, which was diluted with at
least a 104-fold excess amount of glucose 1,6-bisphosphate,
as the phosphomannomutase activity. The mean value of
phosphomannomutase activity in fibroblasts from five control subjects
was 6.02 ± 1.05 pmol/min/mg protein and in the fibroblasts from
our three patients the value was 2.71 pmol/min/mg protein (A.H.), 1.08 pmol/min/mg protein (M.M.), and 1.50 pmol/min/mg protein (Y.M.). The
phosphomannomutase activity in CDG syndrome type I fibroblasts was
18-45% that in the control fibroblasts. Similar data were obtained
for CDG syndrome type I fibroblasts in the study by Jaeken et
al. (41).3 When fibroblasts that had
been frozen and thawed were used as the enzyme source, the
phosphomannomutase activity in control fibroblasts remained rather
constant, whereas the fibroblasts from our patients decreased to
several percent that in the controls. These results indicate that the
phosphomannomutase in the fibroblasts of these patients is more easily
inactivated than that in control fibroblasts. If the conversion of
mannose 6-phosphate to mannose 1-phosphate is deficient, CDG syndrome
type I fibroblasts metabolically labeled with
[3H]glucosamine should accumulate 3H-labeled
GlcNAc
1
4GlcNAc-PP-dolichol in the lipid fraction. However, this
compound could not be detected in either the control or CDG syndrome
type I fibroblasts (Fig. 2B, dotted line). These results
showed that decreased phosphomannomutase activity is not a cause of CDG
syndrome type I, at least in our patients. But there still remains the
possibility that the conversion of mannose 6-phosphate to mannose
1-phosphate is mediated by phosphoglucomutase.
(Eq. 1)
Therefore, we measured the radioactivity of
[3H]mannose 6-phosphate, [3H]mannose
1-phosphate and GDP-[3H]mannose as described under
"Experimental Procedures." After synchronization and stimulation,
CDG syndrome type I and control fibroblasts (1 × 106
cells per dish) were pulse-labeled with [3H]mannose (0.25 mCi/ml) in glucose-free medium for 20 min at the indicated times; the
labeling medium in the dishes was immediately replaced with chilled
50% methanol in 10 mM Tris-HCl buffer (pH 8.2), containing
1 mM EDTA, and then the cells were scraped off and boiled
for 2 min. Such immediate inactivation is indispensable for obtaining
reproducible results, since these acidic [3H]mannose
derivatives are immediately transferred to other compounds.
The levels of [3H]mannose 6-phosphate,
[3H]mannose 1-phosphate, and
GDP-[3H]mannose in synchronized control fibroblasts were
constant throughout from the G1 to the S phase (Fig. 3,
B--D, ). In contrast, the levels of these acidic [3H]mannose derivatives in CDG
syndrome type I fibroblasts varied depending on the cell cycle and were
constantly higher than in controls in the G1 period (Fig.
3, B-D,
). The ratios of
[3H]mannose 6-phosphate to [3H]mannose
1-phosphate in the CDG syndrome type I fibroblasts (Fig. 3E,
) and control fibroblasts (Fig. 3E,
) were equal, and
the equilibrium was shifted toward mannose 6-phosphate rather than mannose 1-phosphate in the ratio of approximately 4:1. A similar equilibrium has been reported for the phosphomannomutase of rabbit brain (23) and that of plant (32). These results suggest that even
though phosphomannomutase activity in vitro is decreased, in
intact cells this enzyme activity is sufficient for a normal rate of
conversion of mannose 6-phosphate to mannose 1-phosphate in CDG
syndrome type I. The higher levels of GDP-[3H]mannose in
CDG syndrome type I fibroblasts may indicate that the synthesized
GDP-mannose is not effectively used as a substrate by
dolichylphosphomannose synthase and LLO:
- and
-mannosyltransferases involved in the biosynthesis of LLO. In fact,
when [3H]mannose-labeled LLO obtained from the
precipitates by extraction with CMW was analyzed, the levels in CDG
syndrome type I and control fibroblasts were highly increased in the S
phase; however, the levels in fibroblasts from patient A.H. were
severalfold lower than the levels in control fibroblasts, and the
smaller sized lipid-linked oligosaccharide precursor was not detected,
similar to the data in Fig. 2 (data not shown). These results also
support the view that the site of the defect in biosynthesis of LLO in these patients is a step earlier than the synthesis of
GlcNAc-PP-polyprenol.
Incorporation of [3H]Mevalonate into Polyprenols
In order to examine this possibility directly, we
incubated synchronized CDG syndrome type I fibroblasts and control
fibroblasts, from three persons in each instance (similar population
doubling levels, 5-15, 1 × 106 cells per dish), with
[3H]mevalonate (25 µCi/ml) for 30 h after
inoculation (i.e. in the S phase). After incubation, lipidic
molecules were extracted with saturated 1-butanol as described under
"Experimental Procedures," and the evaporated extracts were first
fractionated on a DE52 column, which had been equilibrated in
1-butanol/methanol (9:1), to separate neutral lipids from anionic
lipids. The DE52 pass-through neutral lipids were further purified on a
Sep-Pak C18 column. Most of the radioactivity was found in ubiquinone,
cholesterol, cholesteryl ester, and polyprenols. The polyprenols
derived from [3H]mevalonate were pooled and analyzed by
HPLC on a silica column, which can easily separate dolichol and
dehydrodolichol, but not according to chain length. Although total
radioactivity incorporated into polyprenols in CDG syndrome type I
fibroblasts and control fibroblasts was not meaningfully different, the
compositions of [3H]polyprenols in the two types of
fibroblasts in the S phase were clearly different from each other. As
shown in Fig. 4, A-F, the relative ratio of [3H]dehydrodolichol (completely
unsaturated polyisoprenoid) versus [3H]dolichol (the terminal isoprene unit being saturated)
derived from the three lots of CDG syndrome type I fibroblasts were
1.0, 1.5, and 5.1. In contrast, dolichols were dominant in the three lots of control fibroblasts, and the ratios of dehydrodolichol versus dolichol were 0.2, 0.1, and 0.3. These results
indicated that the CDG syndrome type I fibroblasts partially lack the
enzymatic activity, which reduces the terminal isoprene unit of
dehydrodolichol.
The level of DE52-bound acidic polyprenols was below one-tenth that of neutral polyprenols, and all of the acidic polyprenols were converted to neutral polyprenols by phosphatase digestion; however, further analysis could not be performed.
Incorporation of [3H]Mevalonate into LLOBecause it has been reported that the oligosaccharides of dehydrodolichyl pyrophosphate oligosaccharides are not efficiently transferred to nascent polypeptides (33), polyprenols of LLO were analyzed by metabolically labeling with [3H]mevalonate. After metabolically labeling with [3H]mevalonate under the same conditions as described above, dehydrodolichol, dolichol phosphate, dolichol phosphate monosaccharides, and dolichol pyrophosphate mono- and di-saccharides were removed by extraction with chloroform/methanol (2:1), and 3H-LLO were sequentially extracted with chloroform/methanol/water (10:10:3) from the residue. The 3H-LLO derived from CDG syndrome type I and control fibroblasts were hydrolyzed with n-propyl alcohol, 0.01 N HCl (1:2) at 100 °C for 25 min and then digested with potato acidic phosphatase. The released [3H]polyprenols were extracted with n-hexane, and then analyzed on a silica gel column. All of the polyprenols derived from both the CDG syndrome type I fibroblasts and control fibroblasts were exclusively dolichols (Fig. 4, G and H). These results indicated that dolichol is preferentially utilized in vivo for the glycosylation reaction in human fibroblasts.
The Chain Length Distributions of Polyprenols in CDG Syndrome Type I and Control FibroblastsIn order to elucidate whether the chain
length distributions of dehydrodolichol and dolichol in CDG syndrome
type I and control fibroblasts are different, dehydrodolichol and
dolichol from one patient (Fig. 4D) and one control (Fig.
4A), respectively, were analyzed on a reverse-phase column,
which easily separates polyisoprenoids of different chain lengths. As
shown in Fig. 5, the chain length distribution of
dehydrodolichols and dolichols in fibroblasts from the patient was
equal to that in the control in every preparation from 17 to 20 isoprene units, with the major labeled species corresponding to 18 and
19 isoprene units. These results suggest that the amount of
dehydrodolichyl diphosphate synthase activity is equal in CDG syndrome
type I and control fibroblasts.
Reduction of Dehydrodolichol to Dolichol
We reported in the
preceding part that the biosynthesis of LLO is dependent on the cell
cycle and that a lower level of LLO synthesis in CDG syndrome type I
fibroblasts can be clearly observed in the S phase. In order to
elucidate whether the slower conversion of dehydrodolichol to dolichol
in the fibroblasts from these patients depends on the cell cycle,
synchronized CDG syndrome type I (patient A.H.) and control fibroblasts
(similar population doubling levels, 11-15; 1 × 106
cells per dish) were continuously labeled with
[3H]mevalonate (25 µCi/ml) for 27, 30, 36, 42, and
48 h. After the respective incubation times, polyisoprenoids were
extracted from the dishes and analyzed by HPLC on a silica column.
Although radioactivity incorporated into dehydrodolichol and dolichol
of CDG syndrome type I and control fibroblasts continuously increased
with incubation time in the presence of [3H]mevalonate,
as shown in Fig. 6, B and C, the
amounts of radioactivity incorporated into dehydrodolichol (Fig.
6B) and dolichol (Fig. 6C) in the fibroblasts
from the patients and the controls were different. The ratio of
[3H]dolichol (patient/control) (Fig. 6C,
-
) was constantly lower than 1.0, and the ratio of
[3H]dehydrodolichol (patient/control) (Fig.
6B,
-
) was constantly higher than 1.0. Also, the
levels of [3H]dehydrodolichol were extremely elevated in
the S phase as compared with those in the G2 and M periods.
These results indicate that the dehydrodolichol reductase in CDG
syndrome type I fibroblasts is partially deficient, and the effectively
synthesized dehydrodolichol might be accumulated in the S phase.
Furthermore, the amount of radioactivity incorporated into the dolichol
moiety of LLO in CDG syndrome type I fibroblasts was severalfold lower
than that in control fibroblasts (Fig. 6D). These results
are in accordance with the ratio of
[3H]glucosamine-labeled LLO in patient and control
fibroblasts (see Fig. 1).
Metabolic labeling with [3H]glucosamine or
[3H]mannose showed that the levels of LLO in synchronized
CDG syndrome type I fibroblasts were severalfold lower than those in
control fibroblasts, although the sizes of oligosaccharides released
from these LLO showed the same composition corresponding to
Glc1-3·Man9·GlcNAc2 in the two
types of fibroblasts. In contrast, the amount of
[3H]mannose incorporated into mannose-6-P, mannose-1-P,
and GDP-mannose in CDG syndrome type I fibroblasts was greater than
that in control fibroblasts, and the ratio of mannose-6-P to
mannose-1-P was lower than that in control fibroblasts. These results
indicate that even though phosphomannomutase activity in
vitro is decreased, in intact cells this activity is sufficient
for a normal rate of conversion of mannose 6-phosphate to mannose
1-phosphate, and the sequentially synthesized GDP-mannose was not
effectively used for LLO synthesis in our patients with CDG syndrome
type I. As seen upon [3H]mevalonate labeling of
synchronized skin fibroblasts from CDG syndrome type I patients and
controls, [3H]dehydrodolichol was accumulated in the S
phase in the fibroblasts from these patients, and the amounts of
[3H]dolichol and [3H]LLO were lower than
those in control fibroblasts, although the chain length distributions
of the dolichol and dehydrodolichol were equal in every preparation. On
the basis of these results, the behavior (, ) and possible
primary defect site (-//) in the LLO biosynthetic pathway
responsible for CDG syndrome type I in our patients are summarized in
Fig. 7.
Considering that dehydrodolichyl diphosphate synthase activity increases in the S phase and decreases in the G1, G2, and M periods,2 substantial quantities of dehydrodolichol should be synthesized in the S phase. At the same time, because a lot of membrane-bound glycoproteins are also synthesized in the S phase,2 an abnormality of dolichol synthesis could be clearly detected in the S phase.
The biosynthesis of LLO is dependent on the cell cycle, and the cellular level of LLO is at least 10 times higher in the S phase than in the G1 and G2 periods (see Fig. 1). Also, the level of LLO gradually decreases with an increase in PDL (data not shown). Accordingly, in order to obtain reproducible results, it was important that cells with similar population doubling levels and with similar proliferative potential should be tightly synchronized for metabolic studies. Furthermore, the medium used for metabolic labeling with [3H]glucosamine or [3H]mannose was immediately replaced with 50% methanol in buffer containing EDTA, and the harvested cells were then treated with chloroform or boiled at 100 °C for 2 min. Since metabolically labeled LLO, mannose 1-phosphate, mannose 6-phosphate, and GDP-mannose, which are intermediates of N-linked oligosaccharide synthesis, are immediately transferred to the corresponding polypeptides within several minutes, such immediate inactivation of enzymes in the metabolically labeled fibroblasts was also important for obtaining reproducible results.
It is known that several mutants of yeast that are devoid of
certain enzymes required for the complete formation of LLO
accumulate small-sized LLO, as follows (34): alg 1 (1
4-mannosyltransferase deficiency), alg 2 (
1
3-mannosyltransferase deficiency), alg 3 (dolichylphosphomannose synthase deficiency?),
alg 5 (dolichylphosphoglucose synthase deficiency), and
alg 6 (
1
3-glucose transferase deficiency) accumulate
GlcNAc2·PP·dolichol,
Man2·GlcNAc2·PP·dolichol,
Man5·GlcNAc2·PP·dolichol, Man9·GlcNAc2·PP·dolichol, and
Man9·GlcNAc2·PP·dolichol,
respectively. Because no accumulation of small-sized LLO was
observed in our CDG syndrome type I patients, it appears that none of
the glycosyltransferases responsible for LLO biosynthesis was
missing.
It has been reported that Chinese hamster ovary cells of the Lec 9 recessive complementation group contain a lower amount of dolichol, and mutants biosynthesize underglycosylated glycoproteins (29, 35) similar to those described in our CDG syndrome type I cases (8). One of the mutants, F2A8 cells accumulated 10 times more polyprenols than the wild type, 99% of the polyprenols being dehydrodolichol and the polyprenols of LLO being exclusively dehydrodolichol (36), showing that F2A8 cells completely lack dehydrodolichol reductase activity. On the other hand, the conversion of dehydrodolichol to dolichol in our CDG syndrome type I patients was not blocked, but was slower, since the accumulation of dehydrodolichol was not so high.
When the biosynthesis of dolichol is partially deficient, not only glycoproteins containing N-linked sugar chains but also glycosylphosphatidylinositol (GPI)-anchored glycoproteins should be partially underglycosylated, because dolichylphosphomannose is used as a substrate for glycan formation of the GPI anchor (37). In preliminary experiments, normal serum alkaline phosphatases were found to be exclusively linked to glycans derived from the GPI anchor; however, a portion of the alkaline phosphatases in the sera of patients with CDG syndrome type I were devoid of the glycan portion derived from the GPI anchor.3 These results also support the view that a partial deficiency of dehydrodolichol reduction is the primary cause of CDG syndrome type I in our patients.
Two models have already been suggested for the fundamental defect in CDG syndrome type I as follows: Panneerselvam and Freeze (11) proposed defective mannose uptake and Van Schaftingen and Jaeken (12) proposed a phosphomannomutase deficiency. In our patients, we suggest that the abnormal mannose metabolism in CDG syndrome type I is not the actual problem but that conversion of dehydrodolichol to dolichol is partially defective. The resulting loss of dolichol may lead directly to underglycosylation. Judging from the results so far described, there might be multiple causes of CDG syndrome type I.
It has been observed that the abundance of various glycoproteins decreases in CDG syndrome type I. For example, 1) partially underglycosylated glycoproteins tend to be precipitable (9), resulting in their immediate clearance from the bloodstream. 2) Intracellular lysosomal enzymes in CDG syndrome type I decrease, as in the case of I-cell disease, because a trafficking signal for lysosomes might be absent. 3) mRNA expression of decorin is reduced in CDG syndrome type I fibroblasts, which might be due to the decrease in cytokine receptors on the cell surface (38). The decrease in phosphomannomutase in our patients might be due to a mechanism similar to that in the case of other glycoproteins so far described. Such decreases in various enzymes and glycoproteins in CDG syndrome type I should be taken into account when we search for the defect in this disease.
Yasugi et al. (39) previously proposed that CDG syndrome type I is not due to a deficiency of the enzyme responsible for the biosynthesis of dolichyl phosphate, because total dolichols, including free dolichol and esterified dolichol, in skin fibroblasts and sera from patients with CDG syndrome type I and normal controls were the same. However, because it was previously reported regarding the intracellular distribution of dolichols in pig liver that 76.7% of the free dolichol was located in the mitochondria, while 52.7% of the esterified dolichol was associated with a "cell debris plus nuclei" fraction, and only 10% of the free dolichol was located in the supernatant including microsomes (40), most free and esterified dolichol in sera and skin fibroblasts should not represent the dolichol and dolichyl phosphate content of the rough endoplasmic reticulum involved in the biosynthetic pathway for LLO and GPI-anchored glycans. [3H]Dolichol in LLO incorporated upon metabolic labeling with [3H]mevalonate might rather reflect the dolichol content involved in the biosynthesis of N-linked sugar chains.
Because dehydrodolichol reductase activity in cultured cells has not yet been measured, we are now improving the microassay system for this enzyme activity, and we intend to use it to compare CDG syndrome type I and control fibroblasts in the near future.