Abstract

O-Linked fucose is an unusual carbohydrate modification in which fucose is linked directly to the hydroxyl groups of serines or threonines. It has been found on the epidermal growth factor-like modules of several secreted proteins involved in blood coagulation and fibrinolysis. We have recently reported the existence of an elongated form of O-linked fucose in Chinese hamster ovary cells consisting of a glucose linked to the 3′-hydroxyl of fucose (Glcβ1,3Fuc-O-Ser/Thr). This structure is highly unusual for two reasons. First, in mammalian systems fucose is usually a terminal modification of Nand O-linked oligosaccharides. Here the fucose is internal. Secondly, terminal β-linked glucose is extremely rare on mammalian glycoconjugates. Thus, the Glcβ1,3Fuc structure is a very unique mammalian carbohydrate structure. Here we report the identification and initial characterization of a novel enzyme activity capable of forming this unique linkage: UDP-glucose: O-linked fucose β1,3 glucosyltransferase. The enzyme utilizes UDP-glucose as the high energy donor and transfers glucose to α-linked fucose residues. The activity is linearly dependent on time, enzyme, and substrate concentrations and is enhanced in the presence of manganese ions. Activity is present in extracts of cultured cells from a variety of species (hamster, human, mouse, rat, chicken) and is enriched in brain and spleen of a normal adult rat. Thus, while this glycosyltransferase appears to be widespread in biology, it forms a very unique linkage, and it represents the first mammalian enzyme identified capable of elongating fucose.

Introduction

The posttranslational modification known as O-linked fucose consists of the sugar L-fucose directly attached to protein through the hydroxyl group of either serine or threonine (Harris and Spellman, 1993). This unique form of glycosylation was discovered in the past decade on a small number of mammalian serum proteins including urokinase (Kentzer et al., 1990), tissue-type plasminogen activator (Harris et al., 1991), and clotting factors VII (Bjoern et al., 1991), IX (Nishimura et al., 1992), and XII (Harris et al., 1992). Comparison of the glycosylation sites from these proteins resulted in a putative consensus sequence for O-linked fucose modification: C-X-X-G-G-S/T-C, where X is any amino acid and S/T is the modified residue (Harris et al., 1993). The consensus site is found within an epidermal growth factor-like (EGF) module of these proteins between the second and third conserved cysteines. Although the actual number of proteins identified with the O-linked fucose modification is small, a database search reveals that numerous secreted and cell-surface proteins from nematodes to humans contain the consensus sequence for the addition of O-linked fucose (see Table I and Harris and Spellman, 1993; Harris et al., 1993).

The initial reports of O-linked fucose described the modification as a simple monosaccharide. Subsequently, human clotting factor IX was shown to contain a tetrasaccharide form of O-linked fucose with the structure: NeuAcα2,6 Galβ1,4 GlcNAcβ1,3 Fucα1-O-Ser (Harris et al., 1993). This finding was unusual because fucose on other oligosaccharides in mammalian systems had always been seen as a terminal sugar. Factor IX represents the only protein currently identified with an elongated form of O-linked fucose.

Recently, we reported that O-linked fucose can be elongated with a β1,3-linked glucose residue on proteins from Chinese hamster ovary (CHO) cells (Moloney et al., 1997). We also demonstrated that elongation of O-linked fucose is a proteinspecific phenomenon as it occurs only on a subset of O-linked fucose-modified proteins. The discovery of multiple elongated forms of O-linked fucose led us to suggest the presence of a previously unrecognized glycosylation pathway in mammalian cells (Moloney et al., 1997). In this novel pathway, O-linked fucose on specific proteins may be elongated by either a β-linked GlcNAc (as on factor IX) or a β-linked glucose. Elongation by either sugar would be mutually exclusive because they would occupy the same hydroxyl of O-linked fucose. Thus, we hypothesized that specific signals may be encoded within a polypeptide chain to direct the elongation of O-linked fucose.

The biosynthesis of these carbohydrate structures within the O-linked fucose glycosylation pathway requires several novel glycosyltransferases. The enzyme responsible for the addition of fucose to the consensus sequence, GDP-fucose: polypeptide fucosyltransferase, was recently purified and characterized (Wang et al., 1996; Wang and Spellman, 1998). Here we report the identification of an enzyme activity which is specific for adding β-linked glucose onto O-linked fucose. This study includes the development of an enzymatic assay for the O-linked fucose β1,3 glucosyltransferase and initial characterization of this activity. The results presented suggest that the glucose β1,3 fucose modification is widespread and may have a very specific role in biology. The identification and characterization of enzymes within this unusual class of glycosyltransferases of the O-linked fucose pathway is an important step in the process that will lead us to a greater understanding of these unique carbohydrates.

Results

Development of an enzymatic assay for the O-linked fucose β1,3 glucosyltransferase

All glycosyltransferase reactions require a source of active enzyme, a high-energy sugar donor substrate, and a suitable acceptor substrate. We identified the glucosylfucose protein modification in CHO cells, and therefore decided to use CHO whole-cell lysates as the source of the enzyme. For the high-energy sugar donor, the sugar nucleotide UDP-glucose was used. Since no specific proteins modified with glucosylfucose have yet been identified, we tried using a low molecular weight compound, para-nitrophenyl (pNp) -α-L-fucose, as a suitable acceptor substrate. Although we predict that the O-linked fucose β1,3 glucosyltransferase will recognize specific signals encoded within the proteins it modifies, low molecular weight acceptors have been used in the past to initially identify other proteinspecific glycosyltransferases (e.g., the lysosomal enzyme GlcNAc-1-phosphotransferase (Lang et al., 1984) and the glycoprotein hormone-specific GalNAc transferase (Smith and Baenziger, 1988)). In addition, low molecular weight acceptors are frequently utilized to demonstrate activity in recombinant glycosyltransferases identified by homology searches but for which endogenous substrates are unknown (Amado et al., 1998). There are many advantages of using a low molecular weight compound as a substrate. These compounds are small and can often fit into an active site when conformational requirements are unknown. In the case of pNp-fucose, it is mildly hydrophobic and will adhere to a reverse-phase column. It fluoresces and can be monitored by UV absorbance. Hydrolysis of the substrate by a competing enzyme can be easily detected due to the fact that the solution turns yellow as p-nitrophenol is produced. While the in vivo protein targets of this enzyme are not yet known, pNp-fucose proved to work as an acceptor substrate.

Fig. 1.

Fucose-specific glucosyltransferase activity is linearly dependent on concentration of substrates and CHO cell protein. Enzyme activity was measured as described in Materials and methods. (A) UDP-Glucose concentration was varied with pNp-fucose (4 mM) and CHO cell protein held constant. (B) pNp-α-fucose concentration was varied with UDP-glucose (3 µM) and CHO cell protein held constant. (C) CHO cell protein was varied with pNp-fucose (2.8 mM) and UDP-glucose (3 µM) held constant.

Fig. 1.

Fucose-specific glucosyltransferase activity is linearly dependent on concentration of substrates and CHO cell protein. Enzyme activity was measured as described in Materials and methods. (A) UDP-Glucose concentration was varied with pNp-fucose (4 mM) and CHO cell protein held constant. (B) pNp-α-fucose concentration was varied with UDP-glucose (3 µM) and CHO cell protein held constant. (C) CHO cell protein was varied with pNp-fucose (2.8 mM) and UDP-glucose (3 µM) held constant.

After incubating CHO cell lysate with pNp-fucose and UDP-[3H]-glucose, the products were separated from the unincorporated label using a C18 cartridge. Fucose-specific incorporation of [3H]-glucose was determined by comparing radiolabel in the presence or absence of pNp-fucose. Initially, we examined the effects of varying the amount of enzyme and substrate concentrations on enzyme activity. As shown in Figure 1, activity for the enzyme was linearly dependent on the presence of both the acceptor substrate, pNp-fucose, and the donor substrate, UDPglucose. Activity was also linearly dependent on the amount of CHO cell lysate used as the source of enzyme. The Km of the enzyme for UDP-glucose under these conditions was ∼4 µM. We could not saturate activity using the substrate, pNp-fucose, even at 5 mM concentration.

Characterization of the product from the enzymatic assay

To demonstrate that the product produced in the enzymatic assay was glucose β-linked to the 3′-hydroxyl of fucose, the product was analyzed by high pH anion-exchange chromatography (HPAEC). The radiolabeled sugar component was released from the p-nitrophenyl group using mild acid hydrolysis, reduced with sodium borohydride, and subjected to HPAEC. As shown in Figure 2, the product comigrated exclusively with the glucose-β1,3-fucitol standard under conditions known to separate all of the positional isomers of β-glucosylfucitol (Moloney et al., 1997). This is identical to the disaccharide product which we previously released from CHO cell proteins by β-elimination (Moloney et al., 1997).

Optimization of time, temperature, pH, and cofactor requirements

We next sought to optimize the conditions of our assay. First, we demonstrated that activity was linear for up to 2 h under the conditions of our assay (Figure 3A). Not surprisingly, the optimal temperature was 37°C (Figure 3B) with little to no activity at 0°C. In addition, the enzyme displayed activity over a broad pH range with an optimum at pH 7.5 (Figure 3C). Because metal cations often have been reported to affect glycosyltransferases (Beyer et al., 1981), we wanted to know how cations affect the fucose-specific glucosyltransferase activity. Consistent with many ER/Golgi glycosyltransferases (Beyer et al., 1981), activity was enhanced in the presence of manganese (Figure 4A). There is some residual activity even when no cations are added to the reaction, but the fact that EDTA inhibits this activity suggests that either low levels of cations are present in the buffers or some cations are tightly bound to the enzyme. Manganese had an optimal concentration of 10 mM (Figure 4B). At higher concentrations (50 mM) activity decreased. Calcium and magnesium had no effect on activity, while both cobalt and zinc, interestingly, inhibited activity.

Fig. 2.

Enzyme activity yields glucose β-linked to the 3′-hydroxyl of fucose. The radiolabeled product from the assay was subjected to HPAEC after mild acid hydrolysis and reduction as described in Materials and methods. The product comigrated with the expected glucose β1,3 fucitol sugar standard. Migration positions for the following standards are indicated: 1, fucitol; 2, Glcβ1,2fucitol; 3, Glcβ1,4fucitol; 4, Glcβ1,3fucitol; and 5, glucose.

Fig. 2.

Enzyme activity yields glucose β-linked to the 3′-hydroxyl of fucose. The radiolabeled product from the assay was subjected to HPAEC after mild acid hydrolysis and reduction as described in Materials and methods. The product comigrated with the expected glucose β1,3 fucitol sugar standard. Migration positions for the following standards are indicated: 1, fucitol; 2, Glcβ1,2fucitol; 3, Glcβ1,4fucitol; 4, Glcβ1,3fucitol; and 5, glucose.

Substrate specificity of the β1,3 glucosyltransferase

To address the substrate specificity of the enzyme, we examined a variety of compounds to see which the enzyme prefers. As shown in Figure 5, the enzyme has a requirement for α-linked fucose. β-linked fucose substrates did not work as acceptors. This is consistent with the fact that fucose is found exclusively in the α-anomer in mammals, including O-linked fucose. In addition, these data demonstrate that not any pNp-sugar will act as a substrate for the activity described. The enzyme very specifically modified fucose only in α-linkage. Interestingly, the thioester α-linked fucose was not as good a substrate as the hydroxy α-linked fucose. Also, methylumbelliferyl-fucose did not work as well as pNp-fucose. We also tested free fucose as a competitive inhibitor of pNp-fucose in our assay and found that at up to 10 times the concentration of pNp-fucose, free fucose did not reduce the glucosylation of pNp-fucose (data not shown). When we tested free fucose directly as a suitable substrate for the glucosyltransferase using an anion-exchange column to separate the reactants from products, we found that ∼20 times more free fucose was required to get an equivalent amount of activity using pNp-fucose (data not shown). Taken together, these data indicate that the enzyme not only recognizes the sugar, but that the anomeric linkage and the aglycone component of the substrate play an important role as well.

As mentioned earlier, O-linked fucose has been reported to be elongated on the 3′-hydroxyl with a β-linked glucose and with a β-linked GlcNAc (Harris and Spellman, 1993). It is possible that the activity we detected is the result of a single promiscuous enzyme which can accept both UDP-Glc and UDP-GlcNAc as sugar nucleotide donor substrates. Thus, we examined for fucose-specific GlcNAc transferase activity by replacing UDP-[3H]-Glc with UDP-[3H]-GlcNAc in our assay. In addition, we tested whether UDP-[3H]-Gal would serve as a donor. Neither UDP-GlcNAc nor UDP-Gal served as a donor substrate under the conditions of our assay (data not shown). Interestingly, a small amount of product was formed when UDP-[3H]-Gal was used as a donor. However, product analysis revealed that all of the radiolabel transferred to pNp-fucose under these conditions was in fact [3H]-glucose, presumably the result of an epimerase present in the cell extract. These data indicate that the enzyme which we identified is specific for UDP-Glc and is distinct from the as yet unidentified O-linked fucose β1,3 N-acetylglucosaminyltransferase.

Fig. 3.

Optimization of time, temperature, and pH of the β1,3 glucosyltransferase activity. Fucose-specific glucosyltransferase activity was measured as described in Materials and methods as a function of time (A), temperature (B), and pH (C).

Fig. 3.

Optimization of time, temperature, and pH of the β1,3 glucosyltransferase activity. Fucose-specific glucosyltransferase activity was measured as described in Materials and methods as a function of time (A), temperature (B), and pH (C).

Fig. 4.

β1,3 glucosyltransferase activity is dependent on the presence of manganese. (A) Activity was measured as described in Materials and methods in the presence of EDTA or various divalent cations (5 mM). In the “None” sample, cations were not added to the assay. (B) The amount of MnCl2 was varied to determine the optimal concentration for activity.

Fig. 4.

β1,3 glucosyltransferase activity is dependent on the presence of manganese. (A) Activity was measured as described in Materials and methods in the presence of EDTA or various divalent cations (5 mM). In the “None” sample, cations were not added to the assay. (B) The amount of MnCl2 was varied to determine the optimal concentration for activity.

Cellular distribution of the β1,3 glucosyltransferase

We next wanted to examine the distribution of this enzymatic activity in a variety of cell-lines and tissues. Using the assay we developed, we tested extracts of several tissue-culture cell-lines from different species for activity. As shown in Figure 6A, we detected activity in human, hamster, rat, mouse, and chicken cell-lines. There were, however, substantial differences in activity between the various cell-lines. While HT29 (human colon carcinoma) had a moderate level of activity, HeLa (human cervical carcinoma), K562 (human erythroleukemia), and Jurkat (human T-cell leukemia) cells had very little activity comparatively. Surprisingly, the chicken macrophage cell-line, HD11, showed a relatively high amount of activity. Taken together, these data indicate that this enzyme activity is widespread in biology, but levels of activity vary greatly from one cell-type to another.

Fig. 5.

Initial analysis of the substrate specificity of the O-linked fucose β1,3 glucosyltransferase. α- vs. β-linked fucose. Different α- and β-linked fucose compounds (2 mM) were tested in comparison to pNp-α-fucose (2 mM) as substrates for the fucose-specific glucosyltransferase. The assay was performed as described for pNp-fucose.

Fig. 5.

Initial analysis of the substrate specificity of the O-linked fucose β1,3 glucosyltransferase. α- vs. β-linked fucose. Different α- and β-linked fucose compounds (2 mM) were tested in comparison to pNp-α-fucose (2 mM) as substrates for the fucose-specific glucosyltransferase. The assay was performed as described for pNp-fucose.

We also wanted to determine if there were tissue-specific differences in enzymatic activity. Determination of which tissues have the most activity may give clues about the role the modification plays in biology and will reveal the best starting materials for the future purification of the enzyme. We made extracts of normal adult rat tissues using a polytron homogenizer. As shown in Figure 6B, activity was found in all tissues examined but was specifically enriched in spleen and brain. Interestingly, the kidney preparation contained a strong fucosidase activity not present in the other tissues which hydrolyzed the substrate, pNp-α-fucose, giving the appearance of little or no glucosyltransferase activity. We were able to inhibit this fucosidase activity through the addition of 25 mM free fucose to the reaction. As mentioned above, free fucose at this concentration does not interfere in the assay. This allowed measurement of glucosyltransferase activity in kidney extracts.

Cellular fractionation of the β1,3 glucosyltransferase

In an initial attempt to localize the glucosyltransferase activity within cells, we prepared microsomes from CHO cells. To our surprise, nearly all of the activity was recovered in the high-speed centrifugation supernatant and very little activity was found in the membrane-associated pellet (Table II). To ensure that our preparation was faithful, we examined the membrane association of a known membrane-bound Golgi glycosyltransferase: β1,4 galactosyltransferase (Roth, 1997). Nearly all of the β1,4 galactosyltransferase activity was found as expected in the pellet. These data indicate that the glucosyltransferase may be a soluble enzyme. This finding is surprising because all known Golgi glycosyltransferases to date contain a membrane anchor. Interestingly, when the polypeptide fucosyltransferase was identified, the majority of its activity was also found in the high-speed centrifugation supernatant (Wang et al., 1996). It may be that the glucosyltransferase, as proposed for the fucosyltransferase (Wang et al., 1996), is highly susceptible to proteolysis which may cleave the transmembrane “stem” releasing active enzyme as a soluble protein. Apparently, the β1,4 galactosyltransferase was not susceptible to proteolysis as it remained associated with the membrane fraction. We find it intriguing that the polypeptide fucosyltransferase and the O-linked fucose β1,3 glucosyltransferase both fractionated in this unusual fashion (as soluble enzymes) which may be because they belong to a unique family of enzymes.

Discussion

In this report we have established an enzymatic assay for an α-fucose-specific β1,3 glucosyltransferase which is likely the enzyme responsible for addition of a glucose in β-linkage to the 3′-hydroxyl of O-linked fucose. The linkage formed results in a very unique mammalian carbohydrate structure. After initial characterization of activity, we were able to demonstrate that this activity could be found in a variety of tissues and across species, implying that this is a biologically important modification. Interestingly, the highest activity was found in brain, suggesting that the modification may play a role in normal neuronal functions or development.

This report marks the first identification of an activity for a mammalian enzyme which is capable of elongating the sugar fucose. We showed that the enzyme requires α-linked fucose, as found in O-linked fucose. β-linked fucose was an ineffective substrate for the enzyme. The nature of the aglycone portion of the substrate had large effects on enzyme activity, and free fucose was a poor substrate relative to pNp-fucose. These data suggested to us that the enzyme not only recognizes the fucose to which a glucose will be added, but also how and to what the fucose is attached. Perhaps the presence of the mildly hydrophobic p-nitrophenyl group next to the sugar aids in binding to the active site mimicking fucose found attached directly to protein. Thus, unlike some glycosyltransferases (like β1,4 galactosyltransferase) which primarily recognize the sugar being modified, a significant portion of substrate recognition by the β1,3 glucosyltransferase includes the aglycone component of the molecule. These results are consistent with our previous finding of specific protein species bearing O-linked fucose elongated by a β-linked glucose (Moloney et al., 1997). Since O-linked fucose is also known to be elongated with a β1,3 linked GlcNAc (Harris and Spellman, 1993), it was important to determine whether the activity described here could synthesize both Glcβ1,3Fuc and GlcNAcβ1,3Fuc. Based on the fact that we could not detect any transfer of GlcNAc from UDP-GlcNAc to pNp-fucose, we conclude that there must exist a distinct, as yet undescribed, enzyme responsible for the synthesis of the GlcNAcβ1,3Fuc structure.

The unusual nature of the product formed strongly supports our conclusion that the activity we have characterized is that of the O-linked fucose glucosyltransferase. As discussed above, elongation of fucose in mammalian systems has only been observed on O-linked fucose saccharides (Moloney et al., 1997). In addition, glucose is a relatively rare sugar within oligosaccharide modifications. The main involvement of glucose in post-translational modifications is the glucose cap on N-linked carbohydrates (Cummings, 1992; Hebert et al., 1995). This is a transient modification involved in protein-folding quality control where three glucoses are attached in α-linkage to mannose on an N-glycan. The only other glucose modifications are found in glycosphingolipids, O-linked glucose, and on collagen and glycogenin. In glycosphingolipids and O-linked glucose, the glucose residue forms the core reducing end sugar in β-linkage which is elongated by other sugars, such that the glucose is not exposed (Harris and Spellman, 1993; Kundu, 1992). Collagen contains α-linked glucose residues attached to the 2′ hydroxyl of galactose (Smith et al., 1983). Glycogenin, a protein involved in the initiation of glycogen synthesis which is primarily found in liver and muscle, contains glucosyltransferase activity where it adds glucose in α-linkage to itself at tyrosine 194 (Pitcher et al., 1988; Smythe et al., 1988). The glucose β1,3 fucose structure is therefore a very unique modification on mammalian proteins. The fact that the enzyme described here can form this linkage strongly argues that this enzyme is involved in O-linked fucose elongation in vivo.

In several cases in the past, unusual modifications have been shown to perform specific functions in biological systems (Varki, 1993). For example, mannose-6-phosphate is involved in targeting lysosomal enzymes to lysosomes (Dahms et al., 1989), and sialyl Lewis x structures are involved in recruiting leukocytes to sites of inflammation (McEver et al., 1995). It will be interesting to examine whether the glucose β1,3 fucose structure also plays such a specific function in biology.

Because O-linked fucose modifications have been found only on secreted proteins, the modifications are presumed to occur within the secretory pathway. We would predict the β1,3 glucosyltransferase to be localized to the Golgi compartment based on the fact that the O-linked fucose addition to protein is expected to occur within the Golgi. The polypeptide fucosyltransferase utilizes GDP-fucose (Wang et al., 1996), and GDP-fucose transporters are absent in the ER (Hirschberg and Snider, 1987). GDP-fucose and UDP-glucose transporters have been found localized to the Golgi compartments (Hirschberg and Snider, 1987). The fact that the majority of the O-linked fucose β1,3 glucosyltransferase fractionated as a soluble enzyme suggests that either it underwent proteolysis or it represents a novel class of Golgi glycosyltransferases that do not contain a membrane anchor. In the latter case, an enzyme of this class might localize to the Golgi by association with an anchored protein or only become active upon entering the Golgi where it would interact with substrates. Resolution of these possibilities will occur when the enzyme is purified and cloned.

O-linked fucose modifications have been found exclusively on EGF modules. Interestingly, EGF modules are often involved in protein-protein interactions. The Notch receptor protein, which contains thirty-six tandem EGF modules and multiple sites for O-linked fucose addition, is known to bind its cell-surface ligands, Delta, Serrate, or Jagged, through the EGF modules of both interacting proteins (Rebay et al., 1991). Delta, Serrate, and Jagged also contain several EGF modules with multiple sites for the O-linked fucose modification. Thus, it is interesting to speculate that the glycosylation of the EGF modules within these proteins may influence receptor-ligand interactions. We have recently shown that the Notch protein is modified with O-linked fucose saccharides (Moloney and Haltiwanger, 1998). We are currently examining which forms of O-linked fucose are found on specific EGF modules in the protein. It will be interesting to see if Notch, which is known to play a key role in neuronal development (Fleming et al., 1997), is modified by the O-linked fucose β1,3 glucosyltransferase.

Although the functions of glucosylfucose and other O-linked fucose modifications are still unknown, we hope that by identifying the enzymes responsible for the synthesis of these unusual structures we will gain a better understanding of the modifications and develop the tools necessary for determining their biological significance. We hope that through the purification of the glucosyltransferase, we will be able to identify proteins that can interact with and are specifically modified by this enzyme. We would also like to determine the specific signals that the enzyme recognizes. Determining the signals for the glucosylation of O-linked fucose will allow us to design proteins, such as tissue-type plasminogen activator (Harris et al., 1991) which normally bears the monosaccharide O-linked fucose, to carry the disaccharide, glucosylfucose, to display an altered characteristic. We are currently focusing our efforts on the purification of this novel activity.

Materials and methods

Materials

The strain of CHO cells used in this study was Lec1 cells developed by Dr. Pamela Stanley (Stanley and Siminovitch, 1977). Lec1, NRK52E, NRK49F, NIH3T3, HT29, HeLa, Jurkat (clone E6-1), and K562 cells were from American Type Culture Collection (Rockville, MD). Lec1 cells were grown in EMEM media supplemented with 10% fetal bovine serum. NRK, NIH3T3, HT29, and HeLa cells were grown in DMEM media supplemented with 10% fetal bovine serum. Jurkat and K562 cells were grown in RPMI 1640 media supplemented with 10% fetal bovine serum. HD11 cells (Beug et al., 1979) were a gift from Dr. Michael Hayman (SUNY Stony Brook). UDP-[6-3H]-Glucose (60 Ci/mmol) and UDP-[6-3H]-galactose (15 Ci/mmol) were from American Radiolabeled Chemicals (St. Louis, MO). UDP-[6-3H]-N-acetylglucosamine (25.8 Ci/mmol) was from NEN Life Science Products (Boston, MA). pNp-α-Lfucose, pNp-β-L-fucose, mUMB-α-L-fucose, mUMB-β-L-fucose, pNp-β-D-GlcNAc, pNp-α-thio-L-fucose, and pNp-β-thio-Lfucose were from Sigma (St. Louis, MO). Free L-fucose was from Calbiochem (San Diego, CA). The β-linked glucosylfucose standards (Glcβ1,2Fuc, Glcβ1,3Fuc, and Glcβ1,4Fuc) were synthesized, characterized, and generously provided by Dr. Khushi Matta (Roswell Park Memorial Institute, Buffalo, NY) (Moloney et al., 1997). Alditol sugar standards were prepared by reduction of the corresponding sugar with sodium borohydride as described previously (Haltiwanger et al., 1990). Sep-Pak cartridges (C18) were obtained from Waters Associates (Milford, MA). Tissue-culture reagents were obtained from Life Technologies (Gaithersburg, MD). Protease inhibitor cocktails I and II were prepared as described previously (Holt and Hart, 1986). All other reagents were of the highest quality available.

Preparation of cell lysates

Tissue culture cells were grown in 100 mm dishes, collected by using either a cell-scraper (adherent cells) or centrifugation (suspension cells), and washed three times with Tris-buffered saline (10 mM Tris-HCl, pH 7.5, 0.15 M NaCl). Cell pellets (∼106 cells per ml lysis buffer) were lysed on ice in Tris-buffered saline with 1% (w/v) Nonidet P-40 and protease inhibitor cocktails I and II. Lysates were clarified by centrifugation (12,000 × g, 4 min), aliquoted, frozen in liquid nitrogen, and stored at -80°C until needed.

Preparation of microsomes

Microsomes were prepared essentially as described previously (Abmayr and Workman, 1990). Briefly, cells were collected and washed three times with Tris-buffered saline and then suspended in hypotonic buffer (10 mM HEPES pH 7.9, 1.5 mM MgCl2, 10 mM KCl and protease inhibitor cocktails I and II) five times the cell pellet volume. Immediately, the cells were centrifuged 1000 × g for 5 min. The supernatant was discarded, and the cell pellet was resuspended in three times the original cell pellet volume with hypotonic buffer and allowed to swell on ice for 10 min. The cells were transferred to a glass Dounce homogenizer and homogenized with 10 up and down strokes using a type B pestle. Cell lysis was monitored using trypan blue. The lysed cells were centrifuged at 3300 × g for 15 min at 4°C to pellet nuclei and unbroken cells. To the supernatant, 0.11 volume of a 10× dilution buffer (0.3 M HEPES, pH 7.9, 1.4 M KCl, 0.03 M MgCl2) was added and mixed thoroughly. The sample was then subjected to ultracentrifugation at 100,000 × g for 1 h at 4°C. The pellet (microsomal membrane fraction) was resuspended in Tris-buffered saline containing 1% Nonidet P-40 with protease inhibitor cocktails I and II. The pellet fraction and the supernatant were aliquoted, frozen in liquid nitrogen, and stored at -80°C until needed.

Preparation of rat tissue extracts

Tissues from an adult female rat were washed with cold Tris-buffered saline, and homogenized on ice using a polytron homogenizer (setting 4, ∼2 min) with 1 ml of Tris-buffered saline with protease inhibitor cocktails I and II for every gram of tissue. Mitochondria, nuclei, cellular debris, and connective tissues were removed by centrifugation (10,000 × g, 10 min, 4°C). The postmitochondrial supernatant was subjected to ultracentrifugation (100,000 × g, 1 h, 4°C). The supernatants were aliquoted, frozen in liquid nitrogen, and stored at -80°C until needed. Assays of tissue extracts were performed using high-speed supernatants since these had lower amounts of competing activities than whole-cell lysates. We have also shown that the majority of the O-linked fucose β1,3 glucosyltransferase activity is present in the high-speed supernatant (see Table II and data not shown).

Fucose-specific glucosyltransferase assay

All assays were performed as follows unless indicated. The reaction mixture (final volume = 100 µl) contained 50 mM HEPES, pH 7.5, 5–10 mM MnCl2, 2.97 µM UDP-[6-3H]-glucose (3.33 Ci/mmol), 2–5 mM pNp-α-L-fucose and 50–100 µg of cell protein. The reaction was initiated by the addition of cell protein and continued for 1 h at 37°C. The reaction was terminated by the addition of 900 µl of 50 mM EDTA, pH 8, followed by 5 min heating at 70°C. After cooling, the sample was loaded onto a C18 cartridge (Sep-Pak, Waters) equilibrated in water. The column was washed with 15 ml of water, and the pNp-sugar was eluted with 5 ml of 50% methanol. Incorporation of [3H]-Glucose into pNp-fucose was determined by scintillation counting of the eluant. All samples were assayed with and without pNp-fucose to determine pNp-fucose-specific incorporation of [3H]-glucose.

Product analysis by high pH anion-exchange chromatography (HPAEC)

Radiolabeled product in the eluant from the glucosyltransferase assay was dried by evaporation in a Speed-Vac (Savant). The sample was then subjected to mild acid hydrolysis by the addition of 0.1 M HCl for 30 min at 75°C, which released the sugar product from the p-nitrophenyl group without cleavage of the glycosidic bond. The sample was again dried by evaporation using a Speed-Vac, resuspended in 0.1 M NaOH with 1 M NaBH4, and incubated at 37°C for 18 h to reduce the sugar product. The sample was neutralized with acetic acid and desalted using Dowex-50 (H-form) equilibrated in water. Borate was removed by repeated evaporation in the presence of methanol. The sugar product was subjected to HPAEC using a Dionex DX300 HPLC system equipped with pulsed amperometric detection (PAD-2 cell). The sample was chromatographed using a CarboPac MA-1 column (Dionex Corp.) as described previously (Moloney et al., 1997).

Galactosyltransferase assay

The galactosyltransferase assay was performed as follows. The reaction mixture (final volume = 100 µl) contained 50 mM HEPES pH 7.4, 5 mM MnCl2, 0.67 µM UDP-[6-3H]-galactose (15 Ci/mmol), 2.8 mM pNp-β-GlcNAc, 0.9% Triton X-100, and 50-100 µg of CHO cell microsomal preparation supernatant or pellet protein. The reaction was initiated by the addition of CHO cell protein and continued for 1 h at 37°C. The rest of the assay was performed exactly as the fucose-specific glucosyltransferase assay.

Acknowledgments

We thank Mr. Scott Busby, Mr. Sean Li, Ms. Li Shao, and Dr. James Trimmer for critical reading and helpful discussions. We also thank Ms. Elizabeth Hahn and Dr. Michael Hayman for providing the HD11 cells. This work was supported by National Institutes of Health Grant GM 48666 and by a grant from Neose Technologies, Inc.

Abbreviations

    Abbreviations
  • EGF, epidermal growth factor-like
  • CHO, Chinese hamster ovary
  • fuc, L-fucose
  • Glc, D-glucose
  • GlcNAc,N-acetyl-D-glucosamine
  • Gal, D-galactose
  • GalNAc,N-acetyl-D-galactosamine
  • NeuAc,N-acetyl-neuraminic acid
  • pNp,para-nitrophenyl
  • pNpfuc,para-nitrophenyl-α-L-fucose
  • mUMB, methylumbelliferyl
  • HPAEC, high pH anion-exchange chromatography

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