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Studies of mannose metabolism and effects of long-term mannose ingestion in the mouse
Biochimica et Biophysica Acta 1528 (2001) 116^126
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Studies of mannose metabolism and e¡ects of long-term mannose ingestion in the mouse Joseph A. Davis, Hudson H. Freeze * Glycobiology Program, The Burnham Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037, USA Received 9 May 2001 ; received in revised form 3 July 2001; accepted 3 July 2001
Abstract Dietary mannose is used to treat glycosylation deficient patients with mutations in phosphomannose isomerase (PMI), but there is little information on mannose metabolism in model systems. We chose the mouse as a vertebrate model. Intravenous injection of [2-3 H]mannose shows rapid equilibration with the extravascular pool and clearance t1=2 of 28 min with 95% of the label catabolized via glycolysis in 6 2 h. Labeled glycoproteins appear in the plasma after 30 min and increase over 3 h. Various organs incorporate [2-3 H]mannose into glycoproteins with similar kinetics, indicating direct transport and utilization. Liver and intestine incorporate most of the label (75%), and the majority of the liver-derived proteins eventually appear in plasma. [2-3 H]Mannose-labeled liver and intestine organ cultures secrete the majority of their labeled proteins. We also studied the long-term effects of mannose supplementation in the drinking water. It did not cause bloating, diarrhea, abnormal behavior, weight gain or loss, or increase in hemoglobin glycation. Organ weights, histology, litter size, and growth of pups were normal. Water intake of mice given 20% mannose in their water was reduced to half compared to other groups. Mannose in blood increased up to 9-fold (from 100 to 900 WM) and mannose in milk up to 7-fold (from 75 to 500 WM). [2-3 H]Mannose clearance, organ distribution, and uptake kinetics and hexose content of glycoproteins in organs were similar in mannose-supplemented and non-supplemented mice. Mannose supplements had little effect on the specific activity of phosphomannomutase (Man-6-PHMan-1-P) in different organs, but specific activity of PMI in brain, intestine, muscle, heart and lung gradually increased 6 2-fold with increasing mannose intake. Thus, long-term mannose supplementation does not appear to have adverse effects on mannose metabolism and mice safely tolerate increased mannose with no apparent ill effects. ß 2001 Elsevier Science B.V. All rights reserved. Keywords : Mannose; Metabolism; Transporter; Mouse; Phosphomannose isomerase ; Phosphomannomutase
1. Introduction Mannose is used for N- and O-glycosylation, C-mannosylation and GPI anchor synthesis [1] (Fig. 1). The donor GDP-Man is synthesized by the pathway Man-6PCMan-1-PCGDP-Man. Man-6-P can be formed by two pathways: (1) from Fru-6-P using phosphomannose isomerase (PMI), and (2) ATP-dependent phosphorylation of mannose imported into the cell via a mannose-speci¢c transporter [2]. Studies in human hepatoma and ¢broblast cell lines suggest that under physiological conditions, transported mannose is e¤ciently used for glycosylation even though the concentration of glucose in normal blood is 100-fold higher than mannose [2,3]. Recent studies in humans suggest that direct utilization of mannose may be
* Corresponding author. Fax: +1-858-713-6281. E-mail address : [email protected] (H.H. Freeze).
more important than previously appreciated [4]. Several studies have shown that mannose is e¡ectively absorbed from the gastrointestinal tract [4^6] and evidence for mannose-speci¢c transporters has been found in kidney, intestine and in many cell lines [2,7^11]. Patients with congenital disorder of glycosylation, CDG 1b, caused by a de¢ciency in PMI can be successfully treated by oral mannose therapy [5,12,13], but the long-term e¡ects of mannose are unknown. Mammals contain 35^120 WM mannose in their blood that could be used for glycoprotein synthesis [14]. Metabolic studies of mannose in an intact animal model have only been done in the rat [14]. Here, we extend this study to a murine model. Further, we also examine the e¡ects of mannose supplements on health and mannose metabolism, since mouse models of CDG-1b carrying a null allele would probably depend on mannose for survival. The results show that mannose supplements raise steady state mannose levels in the blood, but do not have any obvious detrimental e¡ects on the mice. These results are impor-
0304-4165 / 01 / $ ^ see front matter ß 2001 Elsevier Science B.V. All rights reserved. PII: S 0 3 0 4 - 4 1 6 5 ( 0 1 ) 0 0 1 8 3 - 0
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tant since some PMI-de¢cient humans may require mannose therapy for prolonged periods. 2. Materials and methods ATP, Phosphomannose isomerase (P5153), hexokinase (H5625), glucose-6-phosphate dehydrogenase (G8404), phosphoglucose isomerase (P5381), amyloglucosidase (A7255), NADP (N 0505), mannose-1-phosphate, mannose-6-phosphate, rabbit liver glycogen, trichloroacetic acid (TCA), glycated hemoglobin (Hb A1 C) kit (Cat. no. 441-B), glutamate oxaloacetate transaminase (GOT) and glutamate pyruvate transaminase (GPT) kit (Cat. no. 505OP), bu¡er salts and all ¢ne chemicals were obtained from Sigma, St. Louis, MO. D-Mannose was obtained from Ho¡man International, Calgary, AB, Canada. Hank's balanced salt solution was from Irvine Scienti¢c, Santa Clara, CA. [2-3 H]Mannose (15 Ci/mmol) and [1-14 C]mannose (53 mCi/mmol) were obtained from American Radiolabeled Chemicals, St. Louis, MO. Age-matched female and male Sprague^Dawley mice (C57 Bl/6) were obtained from Harlan, San Diego, CA. Before the experiment, the mice were kept in individual cages with controlled light (lights on 07:00^19:00 h), relative humidity (55 þ 5%) and temperature (22 þ 1³C). 2.1. [2-3 H]Mannose injection and blood sampling Mice were anesthetized with avertin (0.015 ml/g body weight) prior to injection or blood collection. In each experiment, two mice were injected between 09:00 and 10:00 h through the tail vein with 50 WCi of [2-3 H]mannose in pre-¢ltered (0.2 WM, Puradisc, 25 AS, Whatmann) PBS, pH 7.2. For mannose clearance studies, 50 Wl of blood was collected through the ocular vein into heparin-treated (100 units/ml) capillary tubes at 3, 15, 30, 45, 60, 120, 180 and 240 min post-injection. The samples were diluted 10 volumes with cold PBS, pH 7.2 containing protease inhibitors (Boehringer Mannheim, Germany, Cat. no. 1697498) and centrifuged at 6000Ug for 10 min. All measurements were performed on the supernatant. Data are the mean of two mice, and the variation between multiple experiments was not greater than 10% for any determination. Intracellular conversion of [2-3 H]mannoseCC fructose-6-phosphate+3 H2 O and [2-3 H]mannose in samples were determined as described previously [14]. 3 H2 O was expressed as the percentage of total radioactivity. The actual mannose concentration in the plasma was estimated spectrophotometrically [15]. Glutamate oxaloacetate transaminase (GOT) and glutamate pyruvate transaminase (GPT) were assayed in plasma of control and mannose-fed mice as per the supplier's protocol. HbA1 C was quantitated in 50 Wl of heparinized whole blood of control and mannose-fed mice as per the protocol given by the supplier.
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2.2. [2-3 H]Mannose metabolism To measure kinetics of mannose incorporation into different organs, 50 WCi [2-3 H]mannose was injected in 0.15 ml of pre-¢ltered PBS (pH 7.2) through the tail vein and the mice were killed by whole body perfusion at 15, 30, 60, 120, 180, 240 and 360 min as described previously [14]. Heparinized blood samples were collected at 3 min in addition to the indicated time points. Calculations of the amount of label incorporated into each organ were normalized to 3 H in the blood at 3 min. After the indicated time, the major organs were removed and processed immediately or frozen at 320³C or ¢xed in 4% paraformaldehyde in PBS (pH 7.2) overnight for histological examination. In the case of mannose supplemented mice, the organs were removed at 240 min. Mouse organs were homogenized in 50 mM Tris^HCl, pH 7.2 (1 g wet tissue/10 ml bu¡er containing protease inhibitors). Glycoprotein-associated [2-3 H]mannose was determined by adding 1.0 ml of 10% TCA to 200 Wl of the homogenate or 100 Wl of the diluted plasma or serum. The mixture was vortexed, incubated on ice for 60 min and the precipitated protein was collected by centrifugation at 12 000Ug for 10 min. The pellet was washed twice with 10% TCA and twice with cold acetone to remove the free [3 H]mannose, dolichol-phosphate mannose and other lipid intermediates. The ¢nal pellet was resuspended in 200 Wl water, sonicated and transferred to a scintillation vial and counted for radioactivity. 2.3. Organ culture Secretion of [2-3 H]mannose-labeled glycoproteins was monitored in 2^4 mm organ slices by incubating in KMEM containing 5 mM glucose, 100 WM mannose, 50 U each of penicillin, streptomycin and 100 mM glutamine and 20 WCi of [2-3 H]mannose with or without 7 WCi of [14 C]mannose at 37³C for 3 h in a six-well culture plate. After removing labeled medium, slices were washed four times with ice-cold PBS, resuspended in 500 Wl PBS and sonicated for 30 s.Five hundred Wl of the medium and 100 Wl of the sonicate were TCA precipitated, acetone-washed and the pellet scintillation counted for radioactivity. The counts were normalized to protein content. 2.4. Mannose supplementation The mice were divided into ¢ve groups of three females each; male mice were maintained individually. All were fed standard rat chow ad lib. Total mannose in the food was estimated at 6 0.2% based on the manufacturer's composition reports, but the bioavailability is unknown. Assuming food intake of 5 g/mouse per day, basal level of mannose ingestion is 6 10 mg/day. Mice were given ad libitum drinking water containing either 0%, 1%, 3%, 10% or 20% (w/v) mannose, which was changed thrice a
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week for 5 months. During this period, females were impregnated twice. The pups were weaned at 21 days and maintained on the same mannose dose as their mother for 9 days. The water consumption and body weight were recorded throughout the study. Both mother and pups were monitored for ill health such as bloating, diarrhea and abnormal weight gain or loss. Behavior of these animals were also monitored as described [16]. 2.5. Mannose levels in blood and milk Plasma was separated from 200 Wl heparinized blood samples collected from adult mice at various times and from 30-day-old pups about 9 days after weaning. Milk samples were collected from lactating females in each group. Mannose concentration in plasma was determined as described [15] and adapted to estimate mannose in milk. Brie£y, 50 Wl whole milk was treated with 10 Wl 10% TCA, vortexed, and incubated on ice for 60 min followed by centrifugation at 10 000Ug for 5 min. The supernatant (60 Wl) was quantitatively collected and passed through 0.5 ml C-18 spin column to remove the lipids and washed twice with 60 Wl water. All the eluates were pooled and the mannose concentration was determined as described. Recoveries were estimated at 90^95% by adding 50^100 WM mannose to milk prior to analysis. 2.6. Glycogen estimation
50 Wg protein, 5 mM MgCl2 , 0.25 mM NADP, and 1 U/ml each of PGI and G6PDH. PMM was measured as described with 10 WM glucose 1,6-bisphosphate as cofactor [19]. 3. Results 3.1. Fate of [2-3 H]mannose in the blood Both male and female mice have 100 WM þ 20 WM mannose in their blood (data not shown). Kinetics of mannose metabolism are shown in Fig. 2. Within 3 min of injecting [2-3 H]mannose into the blood, 90% of the total label equilibrates with the extravascular pool, and the label remaining in the vasculature is exclusively in [2-3 H]mannose. The total radioactivity in the blood decreases by 35% over the next hour and then remains steady over the next 2^3 h. 95% of the label in the blood is converted to 3 H2 O within 2 h. This indicates rapid intracellular conversion of [23 H]mannose to fructose-6-phosphate+3 H2 O and equilibration with non-labeled H2 O in the rest of the body [14]. About 30 min after injecting [2-3 H]mannose, labeled TCAprecipitable material begins to appear in the blood and continues to accumulate up to 3 h. This increase probably results from glycoproteins secreted by the liver (Fig. 2A). The t1=2 for [2-3 H]mannose clearance is 28 þ 2 min. (Fig. 2B).
Glycogen was estimated in liver by measuring the released glucose [17]. 200 Wl liver homogenate was evaporated to dryness and reconstituted in 200 Wl 0.2 M sodium acetate bu¡er (pH 4.8) containing 2 U amyloglucosidase for 2 h at 40³C. The released glucose in 20 Wl digest was measured by coupling glucose to NADPH production by adding 1 U/ml each of glucokinase/glucose-6-phosphate dehydrogenase. Similar treatment of rabbit liver glycogen control gave about 90^95% glucose. The total glycogen was normalized to protein. 2.7. Quantitation of glycoprotein-associated hexoses Hexose content of glycoproteins in tissue homogenates was determined as described [18] except that 10% TCA was used to precipitate the protein. The pellets were washed three times in 10% TCA and the hexose quantitated. 2.8. PMI and PMM assays Phosphomannose isomerase (PMI) and phosphomannomutase (PMM) activities were measured in 100 000Ug supernatant of freshly homogenized tissues in 50 mM HEPES (pH 7.1) containing protease inhibitors. PMI was measured as described earlier [18] except that the assay was carried out in 50 mM HEPES (pH 7.1) containing
Fig. 1. Mannose metabolism in mammalian cells. Mannose (Man) in glycoconjugates can be derived directly from mannose imported through the Man transport (O), or derived from the glycolytic pathway using imported glucose (Glc) (R) or internal glycogen stores. Man and Glc are both phosphorylated by hexokinase (HK) to Man-6-P and Glc-6-P, respectively. Phosphoglucose isomerase (PGI) converts Glc-6-P to Fructose-6-P (Fru-6-P). Phosphomannose isomerase (PMI) interconverts Fru6-P and Man-6-P in a reaction that causes irreversible loss of 3 HOH from [2-3 H]Man-6-P. This reaction allows either mannose or glucose to contribute to glycolysis or to glycoconjugate synthesis. Man-6-P is converted into Man-1-P by phosphomannomutase (PMM) and then to GDP-Man via GDP-Man pyrophosphorylase for incorporation into glycoconjugates.
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3.2. Incorporation of [2-3 H]mannose into glycoproteins of various organs To determine the kinetics of [2-3 H]mannose incorporation into glycoproteins of various organs, we administered [2-3 H]mannose intravenously and killed mice at various times. Following thorough perfusion, the organs were removed, homogenized and proteins precipitated with TCA. Preliminary experiments were also carried out to account for the lipid intermediates precipitated along with TCA. Acetone removes all the lipid intermediates. TCA precipitable radioactivity is due to only glycoproteins in the homogenate (data not shown). Results of triplicate determinations in two separate experiments are normalized for the amount of label in the blood at 3 min post-injection. Fig. 3 shows the changes in speci¢c activity of various organs. The kinetics are similar for all organs, indicating that all take up [2-3 H]mannose from the blood, but the extent of incorporation varies. Fig. 3A shows high speci¢c activity in liver, intestine, spleen and plasma. The similar level and kinetic pattern of labeled plasma glycoproteins seen in both Figs. 2 and 3 indicate that mannose metabolism is highly reproducible. Incorporation into the liver is high at 60 min and then decreases as the amount of labeled plasma glycoprotein increases, consistent with glycoprotein secretion by the liver. The luminal contents from the intestine also contain labeled glycoproteins, which increase with time (data not shown). We are not able to correlate this increase directly with intestinal secretion, since some of the label may also come from gastric and pancreatic secretions in addition to the intestine [20]. Fig. 3B shows the speci¢c activity of [2-3 H]mannose incorporated into kidney, lung and brain glycoproteins. Kidney and lung incorporation is about half that of liver and intestine. Fig. 3C shows that muscle and heart incorporate 10^15fold less label than liver and intestine. Fig. 4 shows the distribution of [2-3 H]mannose into newly synthesized glycoproteins of various organs and plasma at 30, 60, 180 and 360 min, respectively. Results are expressed as a percentage of the total precipitable label (84 000^160 000 dpm) which is de¢ned as 100% at each time point. At 30 min, liver and intestine account for 75% of the total with very little in plasma (2%). Labeled liver glycoprotein decreases with time while plasma increases to 44% of the total at 360 min. About half of the total TCA-precipitable label seen at 30 min is lost by 360 min. This probably results from a combination of initial oligosaccharide processing, and subsequent glycoprotein turnover. 3.3. Secretion of [2-3 H]radiolabeled glycoproteins in organ culture Liver and intestine slices incubated with [2-3 H]mannose for 3 h in vitro secreted 80% and 69%, respectively, of their total labeled glycoproteins into the medium (data
Fig. 2. Metabolic fate of [2-3 H]mannose injected into mice. [2-3 H]Mannose was injected through the tail vein of anesthetized mice and blood samples drawn at indicated time points. (A) 3 H2 O (a), [2-3 H]mannose (F), total 3 H (E), labeled [3 H]GP (b). (B) Linear regression analysis of mannose clearance (t1=2 = 28 min). The values are representative of three mice.
not shown). Radiolabeled glycoproteins were not detected in the medium of brain, kidney and testis slices. 3.4. E¡ects of long-term mannose supplementation Four-week-old mice were placed on drinking water containing 0%, 1%, 3%, 10% or 20% (w/v) mannose for 5 months to assess the physiological and potential pathological e¡ects of long-term mannose intake. Water consumption in various groups was similar (4.5^6.0 ml/day), except for mice placed on 20% mannose, where consumption was about half (2^2.5 ml/day) that of all others (Fig. 5, upper panel). This reduction is presumably due to the bitter taste of the L-anomer of mannose. Even though mice drinking 20% mannose water showed reduced water consumption, their average total intake of mannose per day was higher than those given 10% mannose (Fig. 5, middle panel). Weight gain was similar in all groups (Fig. 5, lower panel). There were no apparent ill e¡ects of mannose during the study such as bloating or diarrhea. Behaviors such as mode of drinking and eating, climbing on the bars, grooming, social behavior, and locomotion were normal in both the control and the mannose-fed mice (data not shown). Since mannose is about ¢ve times as reactive as glucose
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nose supplements did not appear to increase glycogen accumulation in liver as shown by PAS stain of histological sections. Further, glycoprotein-bound hexose contents in the organs did not increase in the mannose-fed groups (data not shown). Liver enzymes in plasma such as GOT and GPT were normal in both control and mannose-fed mice (normal range: GOT, 40^50 IU/l ; GPT, 15^25 IU/l) except 10% mannose-fed mice where GPT increased slightly, but was still within the normal range. Thus, there was no evidence of liver abnormalities. During 5 months of mannose supplements, female mice were impregnated twice. Litter sizes (7^9 pups) and all pups were normal at birth. The dams continued on mannose-supplemented water during nursing. Pups grew at a normal rate, were healthy, and all survived until weaning at 21 days. After that, the pups were placed on mannose supplements for 9 days along with the mother. 3.5. Mannose supplements increase mannose in blood and milk Mannose concentration in plasma of pregnant female
Fig. 3. Kinetics of [2-3 H]mannose incorporation into glycoproteins of various mouse organs. Mice were injected 50 WCi of [2-3 H]mannose and killed at indicated time points, and the radiolabel incorporated into various organs and plasma measured as described in Section 2. (A) Liver (7), intestine (F), spleen (O) and plasma (a); (B) kidney (R), lung (E) and brain (b); (C) muscle (U) and heart (8). Some of the error bars are omitted for clarity. The values show mean þ S.D. of two mice and triplicate determinations. Results expressed as dpm/mg protein and dpm/Wl plasma.
in non-enzymatic protein glycation [21], long-term mannose supplements could lead to diabetic-like complications. To assess protein glycation, we measured hemoglobin A1 C (HbA1 C) in mice after 3 months. All groups showed nearly the same levels (2.9^3.1%) regardless of mannose supplements, suggesting that glycated proteins do not accumulate in the blood in this animal. All organ weights and their histological appearance were normal in all groups. Although mannose can be converted into glycogen in rabbits [22] and presumably other animals, man-
Fig. 4. Distribution of 3 H into glycoproteins of various mouse organs. Mice were injected 50 WCi of [2-3 H]mannose and the total label incorporated into all glycoproteins in plasma and various organs were measured by TCA precipitation at indicated time points, shaded in the pie chart as follows: liver (black), brain (dashed), intestine (horizontal bars), spleen (white dots on black), muscle (check), kidney (oblique), heart (dark gray), lung (light gray) and plasma (white). The numbers in the ¢gure represent the percentage distribution. Total dpm is indicated. The values are based on two mice.
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mice, 30-day-old pups, and milk samples from nursing dams was determined (Fig. 6). Mannose level in the blood gradually elevated with increasing mannose in the water, reaching 900 WM with 20% mannose supplements compared to 100 WM in control. A similar mannose concentration pro¢le was also observed in males and non-pregnant females (data not shown). Blood mannose levels in 30-day-old pups also increased steadily and reached 900 WM with 20% mannose water. In the milk samples, mannose rose from 60 WM in normal mice up to s 500 WM in mice given 20% mannose water.
Fig. 6. Mannose levels in mouse plasma and milk. Blood and milk samples were collected from mannose-supplemented mice as described in Section 2. Mannose levels in the blood of dams, pups and in milk rose with increasing mannose in water. Blood samples were collected from pups after 21 days' weaning and maintained on the same dose as their mother's for 9 days. Milk samples were collected during lactation. Values represent mean þ S.D. of triplicate assays.
3.6. Mannose metabolism in mannose-supplemented mice Fig. 5. Water and mannose consumption and weight gain of mice. Four-week-old mice were given water containing (w/v) 0% (white bars), 1% (oblique bars), 3% (hatched bars), 10% (light-gray bars) and 20% (dark-gray bars) mannose for 5 months. The water and mannose consumption and body weight of the mice were monitored every 3 days for 4 months. Upper panel: water consumption ml/day per mouse. Middle panel: mannose consumed mg/day per mouse; (*) calculated basal level of mannose ingestion is 6 10 mg/day. Lower panel: percent di¡erential weight gain of each group at the indicated intervals. Data shown are mean þ S.D. of three mice per group.
To determine whether long-term mannose supplementation a¡ected overall mannose metabolism, all groups were given plain water for 16 h before labeling with [2-3 H]mannose. Blood mannose returns to normal (100 WM) within this time in all groups (data not shown). This purging was essential because high endogenous mannose in the circulation would compete with the injected [2-3 H]mannose. Each group was i.v. injected with 50 WCi [2-3 H]-
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8, mannose supplements had no major and consistent effect on the distribution of [2-3 H]mannose into glycoproteins of di¡erent organs or plasma. We also determined the speci¢c activity of [2-3 H]mannose incorporated into glycoproteins by di¡erent organs of control and mannose-fed groups after 5 months on mannose (Fig. 9). Two mice of each group were killed 4 h after mannose injection and the labeled glycoproteins were determined. As seen previously, liver, intestine and spleen had a high speci¢c activity ( s 200 dpm/mg protein) followed by lung, kidney and brain. Muscle and heart had a very low speci¢c activity. In 3% mannose-fed group, the speci¢c activity of 3 H mannose incorporated into glycoproteins increased slightly in most organs, but this was
Fig. 7. E¡ect of mannose supplements on the metabolism of [2-3 H]mannose. After 5 months of mannose feeding, mannose was removed for 16 h, and the mice were i.v. injected with 50 WCi [2-3 H]mannose and blood samples drawn as indicated: 0% mannose (a), 1% mannose (E), 3% mannose (b), 20% mannose (O). (A) Mannose clearance in di¡erent groups. (B) Kinetics of [3 H]glycoprotein accumulation in the plasma. For clarity, data from 10% mannose-supplemented mice were omitted since it showed a similar pro¢le to 20% mannose. Data shown are mean þ S.D. of two mice per group.
mannose and analyzed for its clearance, catabolism, and incorporation into glycoproteins as described above. t1=2 was 28 þ 2 min (Fig. 7A) and 95% of the label is metabolized via PMI with release of 3 H2 O in all groups. It is also possible that high mannose supplements might a¡ect the labeling kinetics of serum glycoproteins. However, there was no di¡erence between the control and mannose-fed groups in either parameter (Fig. 7B). We also measured the relative distribution of [2-3 H]mannose incorporated into di¡erent organs. Two agematched 5-month mannose-fed and control mice were injected 50 WCi [2-3 H]mannose and killed after 4 h as described in Section 2. Total TCA precipitable material in homogenates of di¡erent organs were measured and expressed as percentage of total precipitable label (118 000^ 168 000 dpm), which is de¢ned as 100%. As shown in Fig.
Fig. 8. Distribution of [3 H]mannose-labeled glycoproteins from various mouse organs. Mice were fed mannose supplemented water for 5 months. After removal of mannose for 16 h, mice were injected 50 WCi [2-3 H]mannose and the total label incorporated into glycoproteins in plasma and various organs were measured after 4 h, shaded in the pie chart as follows : liver (black), brain (dashed), intestine (horizontal bars), spleen (di¡use gray), muscle (check), kidney (oblique), heart (dark gray), lung (light gray) and plasma (white) ; (A) 0%, (B) 1%, (C) 3%, (D) 10%, (E) 20% mannose. The numbers in the ¢gures represent percent distribution of total dpm which varied from 118 000 to 168 000. The values are based on two mice per group.
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followed by a substantial decrease in the 10% and 20% mannose-fed groups. 3.7. E¡ects of mannose supplements on PMM and PMI activities Mannose is catabolized through glycolysis via PMI or delivered to glycan synthesis starting with PMM (Fig. 1). Mannose supplements could conceivably alter the levels of these enzymes since they are involved in early metabolism. The speci¢c activities of PMI were measured in various organs from each group of mice (Fig. 10). PMM activities were high in liver in all groups tested. There was a slight increase ( 6 2-fold) in the PMM speci¢c activity in liver, intestine, kidney and heart in the 10% mannose-fed group followed by a decrease in the 20% mannose-fed group (data not shown). However, PMI speci¢c activity increased gradually in brain, intestine, muscle, heart and lung between 1% and 10% mannose-fed and decreased substantially at 20%. 4. Discussion Most studies of mannose metabolism [4,23,24] have been done in humans, since mannose is an e¡ective therapy for congenital disorder of glycosylation Type Ib (PMI-de¢ciency) [5,12,13]. Details of mannose metabolism and e¡ects of long-term mannose feeding in humans are unknown. Therefore, an animal model is desirable to examine the potential pathological e¡ects. Previous studies in a few patients showed that mannose increases glycated Hb A1c which could lead to diabetic-like complications
Fig. 9. Speci¢c activity of [2-3 H]mannose incorporated into glycoproteins of various mouse organs. Mice were allowed to drink mannosesupplemented water for 5 months, then placed on plain water for 16 h and injected 50 WCi [2-3 H]mannose as described in Section 2. Mannosefed groups: 0% (white bars), 1% (oblique bars), 3% (hatched bars), 10% (light-gray bars) and 20% (dark-gray bars). After 4 h, mice were killed and the label incorporated into various organs were measured. The values represent mean þ S.D. of two mice per group and triplicate experiments.
Fig. 10. PMI activity in organs of female mice fed mannose supplemented water. Two mice from each group were killed, organs removed, homogenized and assayed for the enzyme activity. Mannose-fed groups: 0% (white bars), 1% (oblique bars), 3% (hatched bars), 10% (light-gray bars) and 20% (dark-gray bars). The values represent mean þ S.D. of duplicate assays of two mice per group in two separate experiments.
[6,12,18]. This is probably caused by the greater glycation potential of mannose compared to glucose [21]. Several studies show that high mannose concentration ( s 2 mM) is teratogenic in rat embryos [25^27]. The developmental abnormalities result from intracellular energy depletion because the amount of PMI activity is insu¤cient to direct mannose into the glycolytic pathway. PMI insu¤ciency also explains why honeybees rapidly die when given only mannose [25,28]. Together, these studies prompted cautionary notes on the potential danger of mannose ingestion during pregnancy [29,30]. On the other hand, oral mannose supplements have been suggested as therapy for urinary tract infections since Escherichia coli binds to the epithelial cells through a mannose-sensitive lectin-type interaction [31]. Mannose also in£uences binding of other bacteria to the large and small intestinal epithelial cells [32,33]. An animal model system to study mannose is important and the murine model is currently the best choice, since mice with a null MPI allele might serve as a model of CDG-Ib. The ¢rst step is to assess normal mannose metabolism and the e¡ects of elevated mannose on normal mice. Previous studies in both rats and humans indicated that mannose is cleared from the blood with a t1=2 of about 25^ 30 min. In rats, the liver and intestine were the major consumers of mannose [4,14,23]. Similar results are seen in the mouse model. As with the rat, i.v. injected [2-3 H]mannose rapidly equilibrates with the extravascular pool, and at 3 min post-injection, all of the radioactivity in the blood is found exclusively in mannose. It is rapidly imported into cells of all organs where the great majority (95%) enters the glycolytic pathway. This conclusion is based on the fact that metabolism of [2-3 H]mannose-6-P leads to either glycan biosynthesis or to glycolysis. The former pathway begins with PMM (Man-6-PCMan-1-P) and the latter with PMI, which produces non-labeled fruc-
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tose-6-P and 3 H2 O that rapidly di¡uses throughout the body. Thus, measuring the production of 3 H2 O should re£ect the proportion of mannose metabolized through glycolysis. On the other hand it might be argued that 3 H2 O overestimates this amount. Since the equilibrium constant is near unity, [2-3 H]mannose-6-P could lose its radiolabel in a ¢rst encounter with PMI and then, acting in the reverse direction, PMI would reconvert it again to non-labeled mannose-6-P and used for glycoprotein biosynthesis. To test this, we also measured glycoprotein synthesis by labeling with [14 C]mannose, which does not lose its label when converted into fructose-6-P. Preferential incorporation of [14 C]- over [2-3 H]mannose would indicate that [2-3 H]mannose labeling underestimates protein glycosylation. Several experiments using mouse organ cultures or established cell lines showed no preference for [14 C]mannose incorporation. In fact, there was a slight preference for incorporation of [2-3 H]mannose (data not shown). This is likely due to an isotope e¡ect of 3 H vs. 1 H at the 2-position [34], making it a less e¤cient substrate for PMI. All organs appear to incorporate injected mannose into protein with very similar kinetics, indicating that all take up mannose directly from the circulation ; however, the extent of incorporation into glycoprotein varies (Fig. 3). The substantial di¡erences probably re£ect di¡erences in rate of glycoprotein synthesis and/or the amount of mannose transported into the cells. Clearly, liver and intestine incorporate the most radioactivity, together accounting for 75% of the total incorporated into glycoprotein (Fig. 4). Most of the glycoproteins initially synthesized in the liver eventually appear in the plasma (Figs. 2^4), and organ cultures of both liver and intestine labeled with mannose also secrete most of their glycoproteins. The high glycosylation demands of these organs probably explain why much of the pathology seen in CDG patients involves one or both of these systems and their secreted products. The substantial decrease in TCA precipitable label seen in liver and intestine beyond 60 min mostly results from glycoprotein secretion. Some loss of radioactivity may also result from processing of high mannose type oligosaccharides to complex types of N-linked chains which are present in most of the serum glycoproteins. Other organs that do not secrete substantial amounts of glycoproteins show a smaller proportionate decrease of label beyond 60 min (Fig. 3). Mannose-supplemented water given to mice for 5 months did not show any adverse or pathological e¡ects on growth, behavior, organ size or weight, accumulation of glycated hemoglobin, liver glycogen, serum transaminases or the histological appearance of any major organs or tissues. Humans drinking mannose sometimes experience bloating and osmotic diarrhea [4,18,23], but this did not occur in mice. Those receiving 20% mannose drank only half as much water as the control or the other groups. Nevertheless, their average daily mannose intake
was still the highest (Fig. 5, middle panel). The reduced water consumption in the 20% group is probably due to the bitter taste to humans of the L-anomer of mannose in solution. The absence of increased HbA1 C in the mannose mice may be due to more rapid rate of blood iron turnover, which is about 3^4 times faster than in humans [35]. Oral mannose intake leads to a dose-dependent increase in mannose levels in the blood of males, females, and recently weaned pups. Mannose also increases the level in the milk from nursing dams. These increases may be important to accommodate the glycosylation needs of mice carrying an MPI null allele, which is now under study in our laboratory. Our previous work in rats shows that [2-3 H]mannose can cross the placental barrier and label the fetus [14]. We also wanted to determine if hyper-physiological amounts of mannose led to changes in the clearance rate, fate, organ distribution, or rate of glycoprotein secretion into the blood. None of these parameters changed in mice given mannose for 5 months and placed on plain water for 16 h prior to labeling with mannose. We noticed that the speci¢c activity of the glycoproteins made in some organs (liver, intestine, spleen and lung) appeared to decrease by as much as 3^4-fold in mice given 10% and 20% mannose in their water. This might re£ect a change in the size of the precursor pools in these organs. Consistent with this ¢nding, rats given 15% mannose (w/w) in their chow increased the size of the man-6-P pools in liver by 3^5-fold which would to lower the speci¢c activity of injected radiolabeled mannose used for glycoprotein synthesis [36^ 38]. The speci¢c activity of PMM in any organ did not vary s 2-fold at any concentration of mannose compared to mice given no mannose. The one exception was the intestine of mice placed on 10% mannose, which was considerably higher than the rest. PMI, the other enzyme likely to be a¡ected by mannose ingestion showed gradually increasing speci¢c activities ( 6 2-fold) with increasing mannose, and then a decrease in most organs in mice given 20%. It is di¤cult to assess the signi¢cance of these changes, but both are relatively small. There are no studies on the regulation of either PMM or PMI activity in mice, but rats show a similar distribution of speci¢c activities of both enzymes in these organs [14]. In summary, hyper-physiological amounts of mannose do not seem to have adverse a¡ects on activity, physiology, reproduction, or lead to obvious macroscopic or histological pathology. We cannot rule out that some changes may occur which are either below our detection limits or do not occur within ¢ve months on mannose. Further, the mannose metabolism in mice also corroborates our earlier ¢ndings in rats. The mice in this study received up to an equivalent of 20 g/kg per day which is more than 25 times the typical dose (0.6^0.8 g/kg per day) given to CDG-1b patients. These initial studies in mice suggest that mannose consumption in humans will probably not have adverse e¡ects. Mannose added to the
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drinking water can increase the mannose concentration in blood and milk. It remains to be seen whether this amount of mannose is su¤cient to ful¢ll the glycosylation demands of mice homozygous for the null allele of MPI, which should be totally dependent on exogenous mannose for survival. Acknowledgements This work was supported by the National Institutes of Health R01-GM55695 to H.H.F. We are indebted to Dr. Nissi Varki for the pathological assessment of the mice, the members of the Animal Care Facility for the scrupulous attention to the care of the mice and Antonio Melgoza for assistance in the surgical procedures. We thank Dr. Vandana Sharma for her advice with the in vitro experiments, and Dr. Geetha Srikrishna for ongoing advice and insight throughout this project.
References [1] H.H. Freeze, Metabolism of sugars and sugar nucleotides, in: Ernst, Hart, Sinay (Eds.), Saccharides in Chemistry and Biology: A Comprehensive Handbook, Vol. 3, Wiley-VCH, Weinheim, 2000, pp. 3^ 18. [2] K. Panneerselvam, H.H. Freeze, Mannose corrects altered N-glycosylation in carbohydrate-de¢cient glycoprotein syndrome ¢broblasts, J. Clin. Invest. 97 (1996) 1478^1487. [3] K. Panneerselvam, J.R. Etchison, H.H. Freeze, Human ¢broblasts prefer mannose over glucose as a source of mannose for N-glycosylation, J. Biol. Chem. 272 (1997) 23123^23129. [4] G. Alton, S. Kjaergarrd, J.R. Etchison, F. Skovby, H.H. Freeze, Oral ingestion of mannose elevates blood mannose levels : a ¢rst step toward a potential therapy for carbohydrate de¢cient glycoprotein syndrome type I, Biochem. Mol. Med. 60 (1997) 127^133. [5] R. Niehues, M. Hasilik, G. Alton, C. Korner, M. Schiebe-Sukumar, H.G. Koch, K.P. Zimmer, R. Wu, E. Harms, K. Reiter, K. von Figura, H.H. Freeze, H.K. Harms, T. Marquardt, Carbohydrate de¢cient glycoprotein syndrome Type 1b. Phosphomannose isomerase de¢ciency and mannose therapy, J. Clin. Invest. 101 (1998) 1414^ 1420. [6] H.H. Freeze, H.K. Harms, T. Marquardt, Low-tech mannose therapy for protein glycosylation de¢ciencies, in: Aubery (Ed.), Glycans in Cell Interaction and Recognition, Gordon and Breach Publishing Group, London, 2001, pp. 169^192. [7] M. Silverman, M.A. Aganon, F.P. Chinard, Speci¢city of monosaccharide transport in dog kidney, Am. J. Physiol. 218 (1970) 743^750. [8] J. Pritchard, G.W. Booz, A. Kleinzeller, Renal sugar transport in the winter £ounder. VI. Reabsorption of D-mannose, Am. J. Physiol. 242 (1982) F415^F422. [9] E. Ogier-Denis, G. Trugman, C. Sapin, M. Aubery, P. Codogno, Dual e¡ect of 1-deoxymannojirimycin on the mannose uptake and on the N-glycan processing of the human colon cancer cell line HT29, J. Biol. Chem 265 (1990) 5366^5369. [10] E. Ogier-Denis, A. Blais, J.J. Houri, T. Voisin, G. Trugman, P. Codogno, The emergence of a basolateral 1-deoxymannojirimycinsensitive mannose carrier is a function of intestinal epithelial cell di¡erentiation. Evidence for a new inhibitory e¡ect of 1-deoxymanojirimycin on facilitative mannose transport, J. Biol. Chem. 269 (1994) 4285^4290.
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[11] T. Blasco, J.J. Aramayona, A.I. Alcalde, N. Halaihel, M. Sarasa, V. Sorribas, Expression and molecular characterization of rat renal Dmannose transport in xenopus oocytes, J. Membr. Biol. 178 (2000) 127^135. [12] P.D. Lonlay, M. Cuer, S. Vuillaumier-Barrot, G. Beaune, P. Castelnau, M. Kretz, G. Durand, J.M. Saudubray, N. Seta, Hyperinsulinemic hypoglycemia as a presenting sign in phosphomannose isomerase de¢ciency: a new manifestation of carbohydrate-de¢cient glycoprotein syndrome treatable by mannose, J. Pediatr. 135 (1999) 379^383. [13] D. Babovic-Vuksanovic, M.C. Patterson, W.F. Schwenk, J.F. O'Brien, J. Vockley, H.H. Freeze, D.P. Mehta, V.V. Michels, Severe hypoglycemia as a presenting symptom of carbohydrate-de¢cient glycoprotein syndrome, J. Pediatr. 135 (1999) 775^781. [14] G. Alton, M. Hasilik, R. Niehues, K. Panneerselvam, J.R. Etchison, F. Fana, H.H. Freeze, Direct utilization of mannose for mammalian glycoprotein synthesis, Glycobiology 8 (1998) 285^295. [15] J.R. Etchison, H.H. Freeze, Enzymatic assay of D-mannose in serum, Clin. Chem. 43 (1997) 533^538. [16] P. Martin, P. Bateson (Eds.), Measuring Behaviour, An Introductory Guide, second edition, Cambridge University Press, Cambridge, UK, 1993, pp. 125^160. [17] M. Gomez-Lechon, X. Ponsoda, J.V. Castell, A microassay for measuring glycogen in 96-well-cultured cells, Anal. Biochem. 236 (1996) 296^301. [18] V. Westphal, S. Kjaergaard, J.A. Davis, S.M. Peterson, F. Skovby, H.H. Freeze, Genetic and metabolic analysis of the ¢rst adult with congenital disorder of glycosylation type 1b: long-term outcome and e¡ects of mannose supplementation, Mol. Genet. Metab. 73 (2001) 77^85. [19] E. Van Schaftingen, J. Jaeken, Phosphomannomutase de¢ciency is a cause of carbohydrate de¢cient glycoprotein syndrome type 1, FEBS Lett. 377 (1995) 318^320. [20] A.C. Guyton (Ed.), Human Physiology and Metabolism of Disease, Fifth edition, Saunders, Philadelphia, 1992, pp. 474^499. [21] H. Bunn, P.J. Higgins, Reactions of monosaccharides with proteins: possible evolutionary signi¢cance, Science 213 (1981) 222^224. [22] V. Harding, T.F. Nicholson, A.R. Armstrong, Cutaneous blood-sugar curves after the administration of fructose, mannose and xylose, J. Biol. Chem. 128 (1933) 2035^2042. [23] F.C. Wood, G.F. Cahill, Mannose utilization in man, J. Clin. Invest. 42 (1963) 1300^1312. [24] S. Kjaergaard, B. Kristiansson, H. Stibler, H.H. Freeze, M. Schwartz, T. Martinsson, F. Skovby, Failure of short-term mannose therapy of patients with carbohydrate-de¢cient glycoprotein syndrome type 1A, Acta Paediatr. 87 (1998) 884^888. [25] N. Freinkel, N.J. Lewis, S. Akazawa, S.I. Roth, L. Gorman, The honeybee syndrome-implications of the teratogenicity of mannose in rat-embryo culture, New Engl. J. Med. 310 (1984) 223^230. [26] T. Buchanan, N. Freinkel, N.J. Lewis, B.E. Metzger, S. Akazawa, Fuel-mediated teratogenesis. Use of D-mannose to modify organogenesis in rat embryo in vivo, J. Clin. Invest. 75 (1985) 1927^1934. [27] D. Moore, M. Stanisstreet, F. Beck, C.A. Clark, The e¡ects of mannose on rat embryos grown in vitro, Life Sci. 41 (1987) 1885^1893. [28] S.A. Saunders, R.W. Gracy, K.D. Schnackerz, E.A. Noltmann, Are honeybees de¢cient in phosphomannose isomerase?, Science 164 (1969) 858^859. [29] M.J. Adams, Honeybee syndrome, glycolysis, and birth defects (continued), New Engl. J. Med. 311 (1984) 860. [30] P.A. Lazo, Honeybee syndrome, glycolysis, and birth defects, New Engl. J. Med. 310 (1979) 1535^1536. [31] S. Akazawa, B.E. Metzger, N. Freinkel, Relationships between glucose and mannose during late gestation in normal pregnancy and pregnancy complicated by diabetes mellitus: Concurrent concentrations in maternal plasma and amniotic £uid, J. Clin. Endocrinol. Metab. 62 (1986) 984^989. [32] J. Pak, Y. Pu, Z.T. Zhang, D.L. Hasty, X.R. Wu, Tamm-Horsfall protein binds to type 1-¢mbriated E. coli and prevents the E. coli
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from binding to uroplakin Ia and Ib receptors, J. Biol. Chem. 276 (2001) 9924^9930. [33] L. Golderman, E. Rubinstein, Salmonella and shigella adherence to the intestine of mice, Isr. J. Med. Sci. 18 (1982) 1032^1036. [34] B.A. Oyofo, J.R. Deloach, D.E. Corrier, J.O. Norman, R.L. Ziprin, H.H. Mollenhauer, Prevention of Salmonella typhimurium colonization of broilers with D-mannose, Poult. Sci. 68 (1989) 1357^1360. [35] G. Shearer, J.R. Jones, D.H. Kohl, The consequences of the isotope e¡ect on proline dehydrogenation rates estimated by the tritium loss method, Anal. Biochem. 203 (1992) 191^200.
[36] E. Cotchin, F.J.C. Roe, Haematology of rats and mice, in: E. Cotchin, F.J.C. Roe (Eds.), Pathology of Laboratory Rats and Mice, Blackwell Scienti¢c Publications, Philadelphia, 1967, pp. 501^536. [37] N. Asikin, R.E. Koeppe, Mannose-6-p and mannose-1-p in rat brain, kidney and liver, Biochem. Biophys. Res. Commun. 89 (1979) 279^ 285. [38] S. Manochioping, Mannose-6-p Concentrations in Hepatocytes and Perfused Rat Livers, Ph.D., Oklahoma State University, Stillwater, 1981.
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Studies of mannose metabolism and e¡ects of long-term mannose ingestion in the mouse Joseph A. Davis, Hudson H. Freeze * Glycobiology Program, The Burnham Institute, 10901 North Torrey Pines Road, La Jolla, CA 92037, USA Received 9 May 2001 ; received in revised form 3 July 2001; accepted 3 July 2001
Abstract Dietary mannose is used to treat glycosylation deficient patients with mutations in phosphomannose isomerase (PMI), but there is little information on mannose metabolism in model systems. We chose the mouse as a vertebrate model. Intravenous injection of [2-3 H]mannose shows rapid equilibration with the extravascular pool and clearance t1=2 of 28 min with 95% of the label catabolized via glycolysis in 6 2 h. Labeled glycoproteins appear in the plasma after 30 min and increase over 3 h. Various organs incorporate [2-3 H]mannose into glycoproteins with similar kinetics, indicating direct transport and utilization. Liver and intestine incorporate most of the label (75%), and the majority of the liver-derived proteins eventually appear in plasma. [2-3 H]Mannose-labeled liver and intestine organ cultures secrete the majority of their labeled proteins. We also studied the long-term effects of mannose supplementation in the drinking water. It did not cause bloating, diarrhea, abnormal behavior, weight gain or loss, or increase in hemoglobin glycation. Organ weights, histology, litter size, and growth of pups were normal. Water intake of mice given 20% mannose in their water was reduced to half compared to other groups. Mannose in blood increased up to 9-fold (from 100 to 900 WM) and mannose in milk up to 7-fold (from 75 to 500 WM). [2-3 H]Mannose clearance, organ distribution, and uptake kinetics and hexose content of glycoproteins in organs were similar in mannose-supplemented and non-supplemented mice. Mannose supplements had little effect on the specific activity of phosphomannomutase (Man-6-PHMan-1-P) in different organs, but specific activity of PMI in brain, intestine, muscle, heart and lung gradually increased 6 2-fold with increasing mannose intake. Thus, long-term mannose supplementation does not appear to have adverse effects on mannose metabolism and mice safely tolerate increased mannose with no apparent ill effects. ß 2001 Elsevier Science B.V. All rights reserved. Keywords : Mannose; Metabolism; Transporter; Mouse; Phosphomannose isomerase ; Phosphomannomutase
1. Introduction Mannose is used for N- and O-glycosylation, C-mannosylation and GPI anchor synthesis [1] (Fig. 1). The donor GDP-Man is synthesized by the pathway Man-6PCMan-1-PCGDP-Man. Man-6-P can be formed by two pathways: (1) from Fru-6-P using phosphomannose isomerase (PMI), and (2) ATP-dependent phosphorylation of mannose imported into the cell via a mannose-speci¢c transporter [2]. Studies in human hepatoma and ¢broblast cell lines suggest that under physiological conditions, transported mannose is e¤ciently used for glycosylation even though the concentration of glucose in normal blood is 100-fold higher than mannose [2,3]. Recent studies in humans suggest that direct utilization of mannose may be
* Corresponding author. Fax: +1-858-713-6281. E-mail address : [email protected] (H.H. Freeze).
more important than previously appreciated [4]. Several studies have shown that mannose is e¡ectively absorbed from the gastrointestinal tract [4^6] and evidence for mannose-speci¢c transporters has been found in kidney, intestine and in many cell lines [2,7^11]. Patients with congenital disorder of glycosylation, CDG 1b, caused by a de¢ciency in PMI can be successfully treated by oral mannose therapy [5,12,13], but the long-term e¡ects of mannose are unknown. Mammals contain 35^120 WM mannose in their blood that could be used for glycoprotein synthesis [14]. Metabolic studies of mannose in an intact animal model have only been done in the rat [14]. Here, we extend this study to a murine model. Further, we also examine the e¡ects of mannose supplements on health and mannose metabolism, since mouse models of CDG-1b carrying a null allele would probably depend on mannose for survival. The results show that mannose supplements raise steady state mannose levels in the blood, but do not have any obvious detrimental e¡ects on the mice. These results are impor-
0304-4165 / 01 / $ ^ see front matter ß 2001 Elsevier Science B.V. All rights reserved. PII: S 0 3 0 4 - 4 1 6 5 ( 0 1 ) 0 0 1 8 3 - 0
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tant since some PMI-de¢cient humans may require mannose therapy for prolonged periods. 2. Materials and methods ATP, Phosphomannose isomerase (P5153), hexokinase (H5625), glucose-6-phosphate dehydrogenase (G8404), phosphoglucose isomerase (P5381), amyloglucosidase (A7255), NADP (N 0505), mannose-1-phosphate, mannose-6-phosphate, rabbit liver glycogen, trichloroacetic acid (TCA), glycated hemoglobin (Hb A1 C) kit (Cat. no. 441-B), glutamate oxaloacetate transaminase (GOT) and glutamate pyruvate transaminase (GPT) kit (Cat. no. 505OP), bu¡er salts and all ¢ne chemicals were obtained from Sigma, St. Louis, MO. D-Mannose was obtained from Ho¡man International, Calgary, AB, Canada. Hank's balanced salt solution was from Irvine Scienti¢c, Santa Clara, CA. [2-3 H]Mannose (15 Ci/mmol) and [1-14 C]mannose (53 mCi/mmol) were obtained from American Radiolabeled Chemicals, St. Louis, MO. Age-matched female and male Sprague^Dawley mice (C57 Bl/6) were obtained from Harlan, San Diego, CA. Before the experiment, the mice were kept in individual cages with controlled light (lights on 07:00^19:00 h), relative humidity (55 þ 5%) and temperature (22 þ 1³C). 2.1. [2-3 H]Mannose injection and blood sampling Mice were anesthetized with avertin (0.015 ml/g body weight) prior to injection or blood collection. In each experiment, two mice were injected between 09:00 and 10:00 h through the tail vein with 50 WCi of [2-3 H]mannose in pre-¢ltered (0.2 WM, Puradisc, 25 AS, Whatmann) PBS, pH 7.2. For mannose clearance studies, 50 Wl of blood was collected through the ocular vein into heparin-treated (100 units/ml) capillary tubes at 3, 15, 30, 45, 60, 120, 180 and 240 min post-injection. The samples were diluted 10 volumes with cold PBS, pH 7.2 containing protease inhibitors (Boehringer Mannheim, Germany, Cat. no. 1697498) and centrifuged at 6000Ug for 10 min. All measurements were performed on the supernatant. Data are the mean of two mice, and the variation between multiple experiments was not greater than 10% for any determination. Intracellular conversion of [2-3 H]mannoseCC fructose-6-phosphate+3 H2 O and [2-3 H]mannose in samples were determined as described previously [14]. 3 H2 O was expressed as the percentage of total radioactivity. The actual mannose concentration in the plasma was estimated spectrophotometrically [15]. Glutamate oxaloacetate transaminase (GOT) and glutamate pyruvate transaminase (GPT) were assayed in plasma of control and mannose-fed mice as per the supplier's protocol. HbA1 C was quantitated in 50 Wl of heparinized whole blood of control and mannose-fed mice as per the protocol given by the supplier.
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2.2. [2-3 H]Mannose metabolism To measure kinetics of mannose incorporation into different organs, 50 WCi [2-3 H]mannose was injected in 0.15 ml of pre-¢ltered PBS (pH 7.2) through the tail vein and the mice were killed by whole body perfusion at 15, 30, 60, 120, 180, 240 and 360 min as described previously [14]. Heparinized blood samples were collected at 3 min in addition to the indicated time points. Calculations of the amount of label incorporated into each organ were normalized to 3 H in the blood at 3 min. After the indicated time, the major organs were removed and processed immediately or frozen at 320³C or ¢xed in 4% paraformaldehyde in PBS (pH 7.2) overnight for histological examination. In the case of mannose supplemented mice, the organs were removed at 240 min. Mouse organs were homogenized in 50 mM Tris^HCl, pH 7.2 (1 g wet tissue/10 ml bu¡er containing protease inhibitors). Glycoprotein-associated [2-3 H]mannose was determined by adding 1.0 ml of 10% TCA to 200 Wl of the homogenate or 100 Wl of the diluted plasma or serum. The mixture was vortexed, incubated on ice for 60 min and the precipitated protein was collected by centrifugation at 12 000Ug for 10 min. The pellet was washed twice with 10% TCA and twice with cold acetone to remove the free [3 H]mannose, dolichol-phosphate mannose and other lipid intermediates. The ¢nal pellet was resuspended in 200 Wl water, sonicated and transferred to a scintillation vial and counted for radioactivity. 2.3. Organ culture Secretion of [2-3 H]mannose-labeled glycoproteins was monitored in 2^4 mm organ slices by incubating in KMEM containing 5 mM glucose, 100 WM mannose, 50 U each of penicillin, streptomycin and 100 mM glutamine and 20 WCi of [2-3 H]mannose with or without 7 WCi of [14 C]mannose at 37³C for 3 h in a six-well culture plate. After removing labeled medium, slices were washed four times with ice-cold PBS, resuspended in 500 Wl PBS and sonicated for 30 s.Five hundred Wl of the medium and 100 Wl of the sonicate were TCA precipitated, acetone-washed and the pellet scintillation counted for radioactivity. The counts were normalized to protein content. 2.4. Mannose supplementation The mice were divided into ¢ve groups of three females each; male mice were maintained individually. All were fed standard rat chow ad lib. Total mannose in the food was estimated at 6 0.2% based on the manufacturer's composition reports, but the bioavailability is unknown. Assuming food intake of 5 g/mouse per day, basal level of mannose ingestion is 6 10 mg/day. Mice were given ad libitum drinking water containing either 0%, 1%, 3%, 10% or 20% (w/v) mannose, which was changed thrice a
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week for 5 months. During this period, females were impregnated twice. The pups were weaned at 21 days and maintained on the same mannose dose as their mother for 9 days. The water consumption and body weight were recorded throughout the study. Both mother and pups were monitored for ill health such as bloating, diarrhea and abnormal weight gain or loss. Behavior of these animals were also monitored as described [16]. 2.5. Mannose levels in blood and milk Plasma was separated from 200 Wl heparinized blood samples collected from adult mice at various times and from 30-day-old pups about 9 days after weaning. Milk samples were collected from lactating females in each group. Mannose concentration in plasma was determined as described [15] and adapted to estimate mannose in milk. Brie£y, 50 Wl whole milk was treated with 10 Wl 10% TCA, vortexed, and incubated on ice for 60 min followed by centrifugation at 10 000Ug for 5 min. The supernatant (60 Wl) was quantitatively collected and passed through 0.5 ml C-18 spin column to remove the lipids and washed twice with 60 Wl water. All the eluates were pooled and the mannose concentration was determined as described. Recoveries were estimated at 90^95% by adding 50^100 WM mannose to milk prior to analysis. 2.6. Glycogen estimation
50 Wg protein, 5 mM MgCl2 , 0.25 mM NADP, and 1 U/ml each of PGI and G6PDH. PMM was measured as described with 10 WM glucose 1,6-bisphosphate as cofactor [19]. 3. Results 3.1. Fate of [2-3 H]mannose in the blood Both male and female mice have 100 WM þ 20 WM mannose in their blood (data not shown). Kinetics of mannose metabolism are shown in Fig. 2. Within 3 min of injecting [2-3 H]mannose into the blood, 90% of the total label equilibrates with the extravascular pool, and the label remaining in the vasculature is exclusively in [2-3 H]mannose. The total radioactivity in the blood decreases by 35% over the next hour and then remains steady over the next 2^3 h. 95% of the label in the blood is converted to 3 H2 O within 2 h. This indicates rapid intracellular conversion of [23 H]mannose to fructose-6-phosphate+3 H2 O and equilibration with non-labeled H2 O in the rest of the body [14]. About 30 min after injecting [2-3 H]mannose, labeled TCAprecipitable material begins to appear in the blood and continues to accumulate up to 3 h. This increase probably results from glycoproteins secreted by the liver (Fig. 2A). The t1=2 for [2-3 H]mannose clearance is 28 þ 2 min. (Fig. 2B).
Glycogen was estimated in liver by measuring the released glucose [17]. 200 Wl liver homogenate was evaporated to dryness and reconstituted in 200 Wl 0.2 M sodium acetate bu¡er (pH 4.8) containing 2 U amyloglucosidase for 2 h at 40³C. The released glucose in 20 Wl digest was measured by coupling glucose to NADPH production by adding 1 U/ml each of glucokinase/glucose-6-phosphate dehydrogenase. Similar treatment of rabbit liver glycogen control gave about 90^95% glucose. The total glycogen was normalized to protein. 2.7. Quantitation of glycoprotein-associated hexoses Hexose content of glycoproteins in tissue homogenates was determined as described [18] except that 10% TCA was used to precipitate the protein. The pellets were washed three times in 10% TCA and the hexose quantitated. 2.8. PMI and PMM assays Phosphomannose isomerase (PMI) and phosphomannomutase (PMM) activities were measured in 100 000Ug supernatant of freshly homogenized tissues in 50 mM HEPES (pH 7.1) containing protease inhibitors. PMI was measured as described earlier [18] except that the assay was carried out in 50 mM HEPES (pH 7.1) containing
Fig. 1. Mannose metabolism in mammalian cells. Mannose (Man) in glycoconjugates can be derived directly from mannose imported through the Man transport (O), or derived from the glycolytic pathway using imported glucose (Glc) (R) or internal glycogen stores. Man and Glc are both phosphorylated by hexokinase (HK) to Man-6-P and Glc-6-P, respectively. Phosphoglucose isomerase (PGI) converts Glc-6-P to Fructose-6-P (Fru-6-P). Phosphomannose isomerase (PMI) interconverts Fru6-P and Man-6-P in a reaction that causes irreversible loss of 3 HOH from [2-3 H]Man-6-P. This reaction allows either mannose or glucose to contribute to glycolysis or to glycoconjugate synthesis. Man-6-P is converted into Man-1-P by phosphomannomutase (PMM) and then to GDP-Man via GDP-Man pyrophosphorylase for incorporation into glycoconjugates.
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3.2. Incorporation of [2-3 H]mannose into glycoproteins of various organs To determine the kinetics of [2-3 H]mannose incorporation into glycoproteins of various organs, we administered [2-3 H]mannose intravenously and killed mice at various times. Following thorough perfusion, the organs were removed, homogenized and proteins precipitated with TCA. Preliminary experiments were also carried out to account for the lipid intermediates precipitated along with TCA. Acetone removes all the lipid intermediates. TCA precipitable radioactivity is due to only glycoproteins in the homogenate (data not shown). Results of triplicate determinations in two separate experiments are normalized for the amount of label in the blood at 3 min post-injection. Fig. 3 shows the changes in speci¢c activity of various organs. The kinetics are similar for all organs, indicating that all take up [2-3 H]mannose from the blood, but the extent of incorporation varies. Fig. 3A shows high speci¢c activity in liver, intestine, spleen and plasma. The similar level and kinetic pattern of labeled plasma glycoproteins seen in both Figs. 2 and 3 indicate that mannose metabolism is highly reproducible. Incorporation into the liver is high at 60 min and then decreases as the amount of labeled plasma glycoprotein increases, consistent with glycoprotein secretion by the liver. The luminal contents from the intestine also contain labeled glycoproteins, which increase with time (data not shown). We are not able to correlate this increase directly with intestinal secretion, since some of the label may also come from gastric and pancreatic secretions in addition to the intestine [20]. Fig. 3B shows the speci¢c activity of [2-3 H]mannose incorporated into kidney, lung and brain glycoproteins. Kidney and lung incorporation is about half that of liver and intestine. Fig. 3C shows that muscle and heart incorporate 10^15fold less label than liver and intestine. Fig. 4 shows the distribution of [2-3 H]mannose into newly synthesized glycoproteins of various organs and plasma at 30, 60, 180 and 360 min, respectively. Results are expressed as a percentage of the total precipitable label (84 000^160 000 dpm) which is de¢ned as 100% at each time point. At 30 min, liver and intestine account for 75% of the total with very little in plasma (2%). Labeled liver glycoprotein decreases with time while plasma increases to 44% of the total at 360 min. About half of the total TCA-precipitable label seen at 30 min is lost by 360 min. This probably results from a combination of initial oligosaccharide processing, and subsequent glycoprotein turnover. 3.3. Secretion of [2-3 H]radiolabeled glycoproteins in organ culture Liver and intestine slices incubated with [2-3 H]mannose for 3 h in vitro secreted 80% and 69%, respectively, of their total labeled glycoproteins into the medium (data
Fig. 2. Metabolic fate of [2-3 H]mannose injected into mice. [2-3 H]Mannose was injected through the tail vein of anesthetized mice and blood samples drawn at indicated time points. (A) 3 H2 O (a), [2-3 H]mannose (F), total 3 H (E), labeled [3 H]GP (b). (B) Linear regression analysis of mannose clearance (t1=2 = 28 min). The values are representative of three mice.
not shown). Radiolabeled glycoproteins were not detected in the medium of brain, kidney and testis slices. 3.4. E¡ects of long-term mannose supplementation Four-week-old mice were placed on drinking water containing 0%, 1%, 3%, 10% or 20% (w/v) mannose for 5 months to assess the physiological and potential pathological e¡ects of long-term mannose intake. Water consumption in various groups was similar (4.5^6.0 ml/day), except for mice placed on 20% mannose, where consumption was about half (2^2.5 ml/day) that of all others (Fig. 5, upper panel). This reduction is presumably due to the bitter taste of the L-anomer of mannose. Even though mice drinking 20% mannose water showed reduced water consumption, their average total intake of mannose per day was higher than those given 10% mannose (Fig. 5, middle panel). Weight gain was similar in all groups (Fig. 5, lower panel). There were no apparent ill e¡ects of mannose during the study such as bloating or diarrhea. Behaviors such as mode of drinking and eating, climbing on the bars, grooming, social behavior, and locomotion were normal in both the control and the mannose-fed mice (data not shown). Since mannose is about ¢ve times as reactive as glucose
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nose supplements did not appear to increase glycogen accumulation in liver as shown by PAS stain of histological sections. Further, glycoprotein-bound hexose contents in the organs did not increase in the mannose-fed groups (data not shown). Liver enzymes in plasma such as GOT and GPT were normal in both control and mannose-fed mice (normal range: GOT, 40^50 IU/l ; GPT, 15^25 IU/l) except 10% mannose-fed mice where GPT increased slightly, but was still within the normal range. Thus, there was no evidence of liver abnormalities. During 5 months of mannose supplements, female mice were impregnated twice. Litter sizes (7^9 pups) and all pups were normal at birth. The dams continued on mannose-supplemented water during nursing. Pups grew at a normal rate, were healthy, and all survived until weaning at 21 days. After that, the pups were placed on mannose supplements for 9 days along with the mother. 3.5. Mannose supplements increase mannose in blood and milk Mannose concentration in plasma of pregnant female
Fig. 3. Kinetics of [2-3 H]mannose incorporation into glycoproteins of various mouse organs. Mice were injected 50 WCi of [2-3 H]mannose and killed at indicated time points, and the radiolabel incorporated into various organs and plasma measured as described in Section 2. (A) Liver (7), intestine (F), spleen (O) and plasma (a); (B) kidney (R), lung (E) and brain (b); (C) muscle (U) and heart (8). Some of the error bars are omitted for clarity. The values show mean þ S.D. of two mice and triplicate determinations. Results expressed as dpm/mg protein and dpm/Wl plasma.
in non-enzymatic protein glycation [21], long-term mannose supplements could lead to diabetic-like complications. To assess protein glycation, we measured hemoglobin A1 C (HbA1 C) in mice after 3 months. All groups showed nearly the same levels (2.9^3.1%) regardless of mannose supplements, suggesting that glycated proteins do not accumulate in the blood in this animal. All organ weights and their histological appearance were normal in all groups. Although mannose can be converted into glycogen in rabbits [22] and presumably other animals, man-
Fig. 4. Distribution of 3 H into glycoproteins of various mouse organs. Mice were injected 50 WCi of [2-3 H]mannose and the total label incorporated into all glycoproteins in plasma and various organs were measured by TCA precipitation at indicated time points, shaded in the pie chart as follows: liver (black), brain (dashed), intestine (horizontal bars), spleen (white dots on black), muscle (check), kidney (oblique), heart (dark gray), lung (light gray) and plasma (white). The numbers in the ¢gure represent the percentage distribution. Total dpm is indicated. The values are based on two mice.
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mice, 30-day-old pups, and milk samples from nursing dams was determined (Fig. 6). Mannose level in the blood gradually elevated with increasing mannose in the water, reaching 900 WM with 20% mannose supplements compared to 100 WM in control. A similar mannose concentration pro¢le was also observed in males and non-pregnant females (data not shown). Blood mannose levels in 30-day-old pups also increased steadily and reached 900 WM with 20% mannose water. In the milk samples, mannose rose from 60 WM in normal mice up to s 500 WM in mice given 20% mannose water.
Fig. 6. Mannose levels in mouse plasma and milk. Blood and milk samples were collected from mannose-supplemented mice as described in Section 2. Mannose levels in the blood of dams, pups and in milk rose with increasing mannose in water. Blood samples were collected from pups after 21 days' weaning and maintained on the same dose as their mother's for 9 days. Milk samples were collected during lactation. Values represent mean þ S.D. of triplicate assays.
3.6. Mannose metabolism in mannose-supplemented mice Fig. 5. Water and mannose consumption and weight gain of mice. Four-week-old mice were given water containing (w/v) 0% (white bars), 1% (oblique bars), 3% (hatched bars), 10% (light-gray bars) and 20% (dark-gray bars) mannose for 5 months. The water and mannose consumption and body weight of the mice were monitored every 3 days for 4 months. Upper panel: water consumption ml/day per mouse. Middle panel: mannose consumed mg/day per mouse; (*) calculated basal level of mannose ingestion is 6 10 mg/day. Lower panel: percent di¡erential weight gain of each group at the indicated intervals. Data shown are mean þ S.D. of three mice per group.
To determine whether long-term mannose supplementation a¡ected overall mannose metabolism, all groups were given plain water for 16 h before labeling with [2-3 H]mannose. Blood mannose returns to normal (100 WM) within this time in all groups (data not shown). This purging was essential because high endogenous mannose in the circulation would compete with the injected [2-3 H]mannose. Each group was i.v. injected with 50 WCi [2-3 H]-
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8, mannose supplements had no major and consistent effect on the distribution of [2-3 H]mannose into glycoproteins of di¡erent organs or plasma. We also determined the speci¢c activity of [2-3 H]mannose incorporated into glycoproteins by di¡erent organs of control and mannose-fed groups after 5 months on mannose (Fig. 9). Two mice of each group were killed 4 h after mannose injection and the labeled glycoproteins were determined. As seen previously, liver, intestine and spleen had a high speci¢c activity ( s 200 dpm/mg protein) followed by lung, kidney and brain. Muscle and heart had a very low speci¢c activity. In 3% mannose-fed group, the speci¢c activity of 3 H mannose incorporated into glycoproteins increased slightly in most organs, but this was
Fig. 7. E¡ect of mannose supplements on the metabolism of [2-3 H]mannose. After 5 months of mannose feeding, mannose was removed for 16 h, and the mice were i.v. injected with 50 WCi [2-3 H]mannose and blood samples drawn as indicated: 0% mannose (a), 1% mannose (E), 3% mannose (b), 20% mannose (O). (A) Mannose clearance in di¡erent groups. (B) Kinetics of [3 H]glycoprotein accumulation in the plasma. For clarity, data from 10% mannose-supplemented mice were omitted since it showed a similar pro¢le to 20% mannose. Data shown are mean þ S.D. of two mice per group.
mannose and analyzed for its clearance, catabolism, and incorporation into glycoproteins as described above. t1=2 was 28 þ 2 min (Fig. 7A) and 95% of the label is metabolized via PMI with release of 3 H2 O in all groups. It is also possible that high mannose supplements might a¡ect the labeling kinetics of serum glycoproteins. However, there was no di¡erence between the control and mannose-fed groups in either parameter (Fig. 7B). We also measured the relative distribution of [2-3 H]mannose incorporated into di¡erent organs. Two agematched 5-month mannose-fed and control mice were injected 50 WCi [2-3 H]mannose and killed after 4 h as described in Section 2. Total TCA precipitable material in homogenates of di¡erent organs were measured and expressed as percentage of total precipitable label (118 000^ 168 000 dpm), which is de¢ned as 100%. As shown in Fig.
Fig. 8. Distribution of [3 H]mannose-labeled glycoproteins from various mouse organs. Mice were fed mannose supplemented water for 5 months. After removal of mannose for 16 h, mice were injected 50 WCi [2-3 H]mannose and the total label incorporated into glycoproteins in plasma and various organs were measured after 4 h, shaded in the pie chart as follows : liver (black), brain (dashed), intestine (horizontal bars), spleen (di¡use gray), muscle (check), kidney (oblique), heart (dark gray), lung (light gray) and plasma (white) ; (A) 0%, (B) 1%, (C) 3%, (D) 10%, (E) 20% mannose. The numbers in the ¢gures represent percent distribution of total dpm which varied from 118 000 to 168 000. The values are based on two mice per group.
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followed by a substantial decrease in the 10% and 20% mannose-fed groups. 3.7. E¡ects of mannose supplements on PMM and PMI activities Mannose is catabolized through glycolysis via PMI or delivered to glycan synthesis starting with PMM (Fig. 1). Mannose supplements could conceivably alter the levels of these enzymes since they are involved in early metabolism. The speci¢c activities of PMI were measured in various organs from each group of mice (Fig. 10). PMM activities were high in liver in all groups tested. There was a slight increase ( 6 2-fold) in the PMM speci¢c activity in liver, intestine, kidney and heart in the 10% mannose-fed group followed by a decrease in the 20% mannose-fed group (data not shown). However, PMI speci¢c activity increased gradually in brain, intestine, muscle, heart and lung between 1% and 10% mannose-fed and decreased substantially at 20%. 4. Discussion Most studies of mannose metabolism [4,23,24] have been done in humans, since mannose is an e¡ective therapy for congenital disorder of glycosylation Type Ib (PMI-de¢ciency) [5,12,13]. Details of mannose metabolism and e¡ects of long-term mannose feeding in humans are unknown. Therefore, an animal model is desirable to examine the potential pathological e¡ects. Previous studies in a few patients showed that mannose increases glycated Hb A1c which could lead to diabetic-like complications
Fig. 9. Speci¢c activity of [2-3 H]mannose incorporated into glycoproteins of various mouse organs. Mice were allowed to drink mannosesupplemented water for 5 months, then placed on plain water for 16 h and injected 50 WCi [2-3 H]mannose as described in Section 2. Mannosefed groups: 0% (white bars), 1% (oblique bars), 3% (hatched bars), 10% (light-gray bars) and 20% (dark-gray bars). After 4 h, mice were killed and the label incorporated into various organs were measured. The values represent mean þ S.D. of two mice per group and triplicate experiments.
Fig. 10. PMI activity in organs of female mice fed mannose supplemented water. Two mice from each group were killed, organs removed, homogenized and assayed for the enzyme activity. Mannose-fed groups: 0% (white bars), 1% (oblique bars), 3% (hatched bars), 10% (light-gray bars) and 20% (dark-gray bars). The values represent mean þ S.D. of duplicate assays of two mice per group in two separate experiments.
[6,12,18]. This is probably caused by the greater glycation potential of mannose compared to glucose [21]. Several studies show that high mannose concentration ( s 2 mM) is teratogenic in rat embryos [25^27]. The developmental abnormalities result from intracellular energy depletion because the amount of PMI activity is insu¤cient to direct mannose into the glycolytic pathway. PMI insu¤ciency also explains why honeybees rapidly die when given only mannose [25,28]. Together, these studies prompted cautionary notes on the potential danger of mannose ingestion during pregnancy [29,30]. On the other hand, oral mannose supplements have been suggested as therapy for urinary tract infections since Escherichia coli binds to the epithelial cells through a mannose-sensitive lectin-type interaction [31]. Mannose also in£uences binding of other bacteria to the large and small intestinal epithelial cells [32,33]. An animal model system to study mannose is important and the murine model is currently the best choice, since mice with a null MPI allele might serve as a model of CDG-Ib. The ¢rst step is to assess normal mannose metabolism and the e¡ects of elevated mannose on normal mice. Previous studies in both rats and humans indicated that mannose is cleared from the blood with a t1=2 of about 25^ 30 min. In rats, the liver and intestine were the major consumers of mannose [4,14,23]. Similar results are seen in the mouse model. As with the rat, i.v. injected [2-3 H]mannose rapidly equilibrates with the extravascular pool, and at 3 min post-injection, all of the radioactivity in the blood is found exclusively in mannose. It is rapidly imported into cells of all organs where the great majority (95%) enters the glycolytic pathway. This conclusion is based on the fact that metabolism of [2-3 H]mannose-6-P leads to either glycan biosynthesis or to glycolysis. The former pathway begins with PMM (Man-6-PCMan-1-P) and the latter with PMI, which produces non-labeled fruc-
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tose-6-P and 3 H2 O that rapidly di¡uses throughout the body. Thus, measuring the production of 3 H2 O should re£ect the proportion of mannose metabolized through glycolysis. On the other hand it might be argued that 3 H2 O overestimates this amount. Since the equilibrium constant is near unity, [2-3 H]mannose-6-P could lose its radiolabel in a ¢rst encounter with PMI and then, acting in the reverse direction, PMI would reconvert it again to non-labeled mannose-6-P and used for glycoprotein biosynthesis. To test this, we also measured glycoprotein synthesis by labeling with [14 C]mannose, which does not lose its label when converted into fructose-6-P. Preferential incorporation of [14 C]- over [2-3 H]mannose would indicate that [2-3 H]mannose labeling underestimates protein glycosylation. Several experiments using mouse organ cultures or established cell lines showed no preference for [14 C]mannose incorporation. In fact, there was a slight preference for incorporation of [2-3 H]mannose (data not shown). This is likely due to an isotope e¡ect of 3 H vs. 1 H at the 2-position [34], making it a less e¤cient substrate for PMI. All organs appear to incorporate injected mannose into protein with very similar kinetics, indicating that all take up mannose directly from the circulation ; however, the extent of incorporation into glycoprotein varies (Fig. 3). The substantial di¡erences probably re£ect di¡erences in rate of glycoprotein synthesis and/or the amount of mannose transported into the cells. Clearly, liver and intestine incorporate the most radioactivity, together accounting for 75% of the total incorporated into glycoprotein (Fig. 4). Most of the glycoproteins initially synthesized in the liver eventually appear in the plasma (Figs. 2^4), and organ cultures of both liver and intestine labeled with mannose also secrete most of their glycoproteins. The high glycosylation demands of these organs probably explain why much of the pathology seen in CDG patients involves one or both of these systems and their secreted products. The substantial decrease in TCA precipitable label seen in liver and intestine beyond 60 min mostly results from glycoprotein secretion. Some loss of radioactivity may also result from processing of high mannose type oligosaccharides to complex types of N-linked chains which are present in most of the serum glycoproteins. Other organs that do not secrete substantial amounts of glycoproteins show a smaller proportionate decrease of label beyond 60 min (Fig. 3). Mannose-supplemented water given to mice for 5 months did not show any adverse or pathological e¡ects on growth, behavior, organ size or weight, accumulation of glycated hemoglobin, liver glycogen, serum transaminases or the histological appearance of any major organs or tissues. Humans drinking mannose sometimes experience bloating and osmotic diarrhea [4,18,23], but this did not occur in mice. Those receiving 20% mannose drank only half as much water as the control or the other groups. Nevertheless, their average daily mannose intake
was still the highest (Fig. 5, middle panel). The reduced water consumption in the 20% group is probably due to the bitter taste to humans of the L-anomer of mannose in solution. The absence of increased HbA1 C in the mannose mice may be due to more rapid rate of blood iron turnover, which is about 3^4 times faster than in humans [35]. Oral mannose intake leads to a dose-dependent increase in mannose levels in the blood of males, females, and recently weaned pups. Mannose also increases the level in the milk from nursing dams. These increases may be important to accommodate the glycosylation needs of mice carrying an MPI null allele, which is now under study in our laboratory. Our previous work in rats shows that [2-3 H]mannose can cross the placental barrier and label the fetus [14]. We also wanted to determine if hyper-physiological amounts of mannose led to changes in the clearance rate, fate, organ distribution, or rate of glycoprotein secretion into the blood. None of these parameters changed in mice given mannose for 5 months and placed on plain water for 16 h prior to labeling with mannose. We noticed that the speci¢c activity of the glycoproteins made in some organs (liver, intestine, spleen and lung) appeared to decrease by as much as 3^4-fold in mice given 10% and 20% mannose in their water. This might re£ect a change in the size of the precursor pools in these organs. Consistent with this ¢nding, rats given 15% mannose (w/w) in their chow increased the size of the man-6-P pools in liver by 3^5-fold which would to lower the speci¢c activity of injected radiolabeled mannose used for glycoprotein synthesis [36^ 38]. The speci¢c activity of PMM in any organ did not vary s 2-fold at any concentration of mannose compared to mice given no mannose. The one exception was the intestine of mice placed on 10% mannose, which was considerably higher than the rest. PMI, the other enzyme likely to be a¡ected by mannose ingestion showed gradually increasing speci¢c activities ( 6 2-fold) with increasing mannose, and then a decrease in most organs in mice given 20%. It is di¤cult to assess the signi¢cance of these changes, but both are relatively small. There are no studies on the regulation of either PMM or PMI activity in mice, but rats show a similar distribution of speci¢c activities of both enzymes in these organs [14]. In summary, hyper-physiological amounts of mannose do not seem to have adverse a¡ects on activity, physiology, reproduction, or lead to obvious macroscopic or histological pathology. We cannot rule out that some changes may occur which are either below our detection limits or do not occur within ¢ve months on mannose. Further, the mannose metabolism in mice also corroborates our earlier ¢ndings in rats. The mice in this study received up to an equivalent of 20 g/kg per day which is more than 25 times the typical dose (0.6^0.8 g/kg per day) given to CDG-1b patients. These initial studies in mice suggest that mannose consumption in humans will probably not have adverse e¡ects. Mannose added to the
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drinking water can increase the mannose concentration in blood and milk. It remains to be seen whether this amount of mannose is su¤cient to ful¢ll the glycosylation demands of mice homozygous for the null allele of MPI, which should be totally dependent on exogenous mannose for survival. Acknowledgements This work was supported by the National Institutes of Health R01-GM55695 to H.H.F. We are indebted to Dr. Nissi Varki for the pathological assessment of the mice, the members of the Animal Care Facility for the scrupulous attention to the care of the mice and Antonio Melgoza for assistance in the surgical procedures. We thank Dr. Vandana Sharma for her advice with the in vitro experiments, and Dr. Geetha Srikrishna for ongoing advice and insight throughout this project.
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