2Laboratoire de glycobiologie structurale et fonctionnelle, Unité Mixte de Recherche du CNRS no. 8576, Université des Sciences et Technologies de Lille I, 59655 Villeneuve dAscq cedex, France, and 3Station de Pathologie Comparée INRA/Unité Mixte de Recherche du CNRS no. 5087/ Université de Montpellier II, 30380 Saint-Christol lez Alès, France.
Received on January 17, 2001; revised on March 20, 2001; accepted on March 21, 2001.
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Abstract |
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Key words: baculovirus/insect cells/recombinant glycoproteins/sialic acid/trans-sialidase
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Introduction |
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However, it is now well known that N- and O-glycosylation are highly cell typedependent modifications, and it has been extensively reviewed that insect cells glycosylation substantially differs from the mammalian-type glycosylation (März et al., 1995). As summarized by Altmann et al. (1999)
, this constitutes a serious barrier for therapeutic use of baculovirus-expressed glycoproteins. Typically, the early events in glycosylation pathways are conserved from insects to mammals, and the divergences reside in the terminal reactions. The O-glycans produced by insect cells are generally reduced to a single GalNAc (Tn antigen) or the disaccharide Galß1,3GalNAc (T antigen) (Thomsen et al., 1990
; Grabenhorst et al., 1993
; Sugiyama et al., 1993
; Lopez et al., 1999
). The N-glycosylation potential has been investigated more, both on endogenous and baculovirus-expressed glycoproteins. Most studies showed that the major structures are truncated N-glycans, highly fucosylated (Grabenhorst et al., 1993
; Manneberg et al., 1994
; Lopez et al., 1997
). These short species are explained by a trimming reaction catalyzed by a Golgi-located ß-N-acetylglucosaminidase (Altmann et al., 1995
; Wagner et al., 1996a
; Marchal et al., 1999
). However, in some cell lines showing low levels of this enzyme, such as Ea4 from Estigmene acrea or in High Five cells (TN-5B1-4) derived from Trichoplusia ni, the glycosylation capacities can be extended to N-glycans containing low amounts of terminal galactose (Ogonah et al., 1996
; Hsu et al., 1997
; Hooker et al., 1999
; Rudd et al., 2000
). The presence of sialic acids (reviewed by Marchal et al., 2001
) has been reported in some cases; however, it must be recognized that the sialylation in insect cells occurs with extreme low frequency and seems to be restricted to particular glycoproteins (see, for example, Davidson et al., 1990
) or to be tissue-specific. Sialic acids were detected during the development of Drosophila embryos (Roth et al., 1992
) and, more recently, in larval tissues of Galleria mellonella (Lepidoptera) (Karaçali et al., 1997
, 1999) and of the cicada Philaenus spumarius (Malykh et al., 1999
). Thus, it can be speculated that complex N-glycosylation might be essential at some developmental stages in insects and that it is repressed at other stages. Accordingly, Karaçali et al. (1999)
observed a reduction of 16 to 1 of sialic acids levels from larvae to adults.
When looking at the enzyme levels, several authors have shown that cultured insect cells lack terminal transferases like ß1,4-galactosyl- and sialyltransferases (Lopez et al., 1999; Hooker et al., 1999
). Furthermore, Hooker et al. (1999)
were unable to detect any cytidine monophosphate N-acetylneuraminic acid (CMP-Neu5Ac) among the nucleotide pool of uninfected cells from Spodoptera frugiperda (Sf9 and Sf21) and E. acrea (Ea4) cells, as well as in baculovirus-infected Sf21 cells.
Accordingly, insect cells are a promising model for engineering the glycosylation pathways (reviewed by Jarvis et al., 1998). The first attempts to correct the glycan structures by adding mammalian glycosyltransferases were promising: the overexpression of human N-acetylglucosaminyltransferase I (GNTI) was found to lead to a fourfold increase in terminal GlcNAc residues on a baculovirus-expressed fowl plague hemagglutinin (Wagner et al., 1996b
). Similarly, the expression of bovine ß1,4-galactosyltransferase under the control of a viral immediate early promoter, either by the way of a baculovirus expression vector (Jarvis and Finn, 1996
) or in stably transfected cells, enabled the cells to galactosylate the N-glycans on a viral glycoprotein (Hollister et al., 1998
) or on recombinant human transferrin (Ailor et al., 2000
). The elongation of N-glycans can be achieved because insect cells contain the proper sugar donors, namely, uridine diphospho-N-acetylglucosamine (UDP-GlcNAc) and uridine diphosphogalactose (UDP-Gal). However, the availability of the sialic acid donor CMP-Neu5Ac is not likely to be sufficient in cultured lepidopteran cells (Hooker et al., 1999
), therefore, engineering the sialylation in these cells will probably require the addition of more than one gene.
The protozoan parasite Trypanosoma cruzi, the causing agent of Chagas disease, expresses a developmentally regulated trans-sialidase (TS) (reviewed by Schenkman et al., 1994). This enzyme allows the parasite, which does not synthesize sialic acids, to sialylate mucin-like molecules on its surface, at the expense of glycoconjugates from the host. This sialylation is thought to have a role in adhesion of T. cruzi trypomastigotes to red blood cells, in the invasion (Schenkman et al., 1991
), and in preventing recognition by the immune system (Pereira-Chioccola and Schenkman, 1999
). The enzyme has the unique property to transfer
2,3-linked sialic acids linked to ß-galactosides from various donors to an acceptor ß-galactoside (Scudder et al., 1993
).
In this study, we developed an approach for sialylation in the baculovirus-insect cells system using a membrane-bound TS. As a first approach, T. cruzi TS was expressed under the control of the strong baculovirus promoter p10. We report the expression of a chimeric protein consisting in the catalytic domain of T. cruzi TS fused to the C-terminally located transmembrane domain of the major envelope glycoprotein gp67 (also called gp64) of the baculovirus Autographa californica multicapsid nuclear polyhedrosis virus (AcMNPV). The recombinant enzyme was found to be active and membrane-bound, although partially soluble, and was able to sialylate ß-galactosides using fetuin or sialyllactose as sialic acid donor.
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Results |
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Another hypothesis to explain this soluble TS activity could be its incorporation into vesicular membranes or exosomes (Trams et al., 1981). To determine if this could be the case, the clarified infection medium was submitted to ultracentrifugations at increasing forces, and the TS activity remaining in the supernatant was assayed (Figure 6A). We observed that even after a 15 h centrifugation at 100,000 x g, a condition where all potential vesicles should be sedimented, 60% of the TS activity remained in the supernatant. Similarly, after filtration through a 0.22-µm filter, which should retain the vesicular material, 90% of the TS activity was recovered in the filtrates (Figure 6B). Taken together, these results strongly indicate that the TS activity in the infection medium was mainly present as a soluble protein and not integrated into exosomes. Because this protein possesses the hydrophobic tail from gp67, it could be able to form stable aggregates. To test this possibility, the proteins contained in the clarified infection medium were resolved on a nondenaturing PAGE and the TS was detected by western blot. Figure 6C shows that the immunoreactive material could penetrate into the gel, remaining in the upper part. This result shows that the TS present in the clarified infection medium was not associated to vesicles but rather was present as aggregates of very high molecular weight.
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In a first experiment, Sf9 cells were infected with AcP10TS, and 1 day after infection the medium was replaced by a fresh medium containing 3 mg/ml fetuin and 0.2 mg/ml lactose. Figure 7A shows the paper chromatography analysis of the medium after a 24-h incubation (i.e., 2 days p.i.). More than 30% of the lactose was converted to sialyllactose, that is, approximately 600 µg of sialyllactose was produced in 24 h by 5 x 106 infected cells.
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Discussion |
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This study was aimed to explore another possibility for the sialylation in the baculovirus system using T. cruzi S. This enzyme, which has a unique catalytic activity, has been extensively studied (Schenkman et al., 1994), and several groups are concerned with the exploitation of this activity in vitro (Tomlinson et al., 1992
; Ito and Paulson, 1993
).
Due to the surface expression of this enzyme, T. cruzi is able to sialylate mucin-like acceptors using circulating or cellular donors. Our idea was to mimic this property of the parasite in insect cells. To achieve this purpose, the catalytic domain of the enzyme was fused to the transmembrane domain of gp67. The envelope fusion protein gp67 or gp64 has been shown to drive the viral budding (Oomens and Blissard, 1999) and to be the main insect cell receptor (Hefferon et al., 1999
). An envelope fusion domain and a trimerization domain have been defined in the gp67 ectodomain, and a number of mutations in the ectodomain result in viruses with reduced infectivity (Monsma and Blissard, 1995
). In contrast, the deletion of the C-terminal cytoplasmic tail did not significantly modify the infectivity or the virions production (Oomens and Blissard, 1999
), suggesting that this sequence does not play any essential role in the virion assembly. Therefore, it was postulated in this study that the addition of only the transmembrane sequence and the cytoplasmic tail, that is, the last 30 amino acids of AcMNPV gp67 sequence (Whitford et al., 1989
), would direct the chimeric protein to be integrated in the plasma membrane and not specifically in the envelope of the virus. Our strategy was different from the baculovirus display strategy (Boublik et al., 1995
; Grabherr et al., 1997
; Ernst et al., 2000
), which takes advantage of the specific integration of gp67 into the viral envelope to express proteins on the surface of budded virions. However, because the envelope of budded virions is known to derive from the cytoplasmic membrane of infected cells, it was expected that the TS construction would also be present in the viral envelope. In fact, after sedimentation of the viral particles through a sucrose gradient, we were able to detect TS activity in the virus-containing fractions.
When the chimeric TS was overexpressed under the control of the p10 promoter in Sf9 cells, the TS activity recovered was mainly associated with the cells and in the infection medium. Because in the flow cytofluorimetry experiments the expression of the TS was comparable for intact and permeabilized AcP10TS-infected cells, we concluded that the majority of the cellular TS was bound to the plasma membrane. Only a minor part seemed to be associated with virus budding.
It was more surprising to observe that an equal amount of TS activity was found on the plasma membrane and as soluble material in the infection medium, even after high-speed centrifugations in the conditions used to pellet the viruses. It was first thought that the protein was shed from the cell surface by a proteolytic cleavage resulting in an active, truncated form of the enzyme. But after SDSPAGE and western blot analysis we could not detect any difference in the size of the protein found in cell lysates and in the infection supernatants. Therefore, another possible explanation for this soluble activity could be the incorporation of the TS, through the hydrophobic segment from gp67, into vesicles exfoliated from the cellular membrane. This exfoliation process was first observed by Trams et al. (1981) who proposed the name of exosomes for these vesicles. However, taking their experimental conditions (ultracentrifugation and ultrafiltration), we were unable to characterize the association of soluble TS activity with exosomes. Furthermore, an electrophoresis under native conditions showed that the TS could penetrate into the gel and confirmed that the enzyme was not linked to vesicular membranes. Our interpretation is that the TS synthesized in the endoplasmic reticulum follows the secretion pathway, because both cellular and soluble forms are N-glycosylated. Due to the overexpression driven by the p10 promoter, a part of the glycoprotein remains plasma membraneassociated, while an other part is released as aggregates. Indeed, the time-course analysis shows that the TS associated to the membranes rapidly reaches a plateau, whereas the released activity continuously increases during the infection. The plasma membranebound activity was found to represent one third to one half of the total cellular activity as assayed in the lysates, and this appears not to be in agreement with the flow cytofluorimetry analyses, which indicated that the majority of the TS was bound to the plasma membrane, because no difference in the antibody fixation was observed for permeabilized cells as compared with intact cells. One possible explanation is that a part of the intracellular TS activity is soluble and hence is lost after permeabilization. But this discrepancy may also reflect a difference in activity or in substrate accessibility between the solubilized and membrane-bound forms of the enzyme.
More important is the observation that the expression of a TS in the baculovirus system enables the insect cells to sialylate exogenous galactosylated acceptors, such as lactose and asialo-N-glycans. But further efforts should concern the optimization of the model. The first point to consider is that in this study the TS was overexpressed under the control of a strong late promoter. Because this could affect the synthesis of a glycoprotein of interest, it could be useful to express the TS under the control of a moderate and early promoter (for review, see Jarvis et al., 1990). The second point is that we were only concerned to sialylate exogenous galactosylated acceptors, because Sf9 cells do not produce significant amounts of galactose-terminated glycoproteins. As discussed above, the expression of TS in insect cells engineered to extend the glycosylation pathway up to the galactose seems to be a key point. Future studies should concern the coexpression of a ß-galactosyltransferase in combination with the TS, together with a glycoprotein of interest. Alternatively, it could be interesting to express the TS in insect cell lines that are known to synthesize higher levels of galactosylated glycoproteins, such as Ea4 or High Five cells.
Another essential improvement concerns the efficiency of the TS reaction and the optimization of the sialylation levels. The choice of the most appropriate sialic acid donor seems to be crucial. Several requirements have to be adressed. First, the donor will have to allow an easy purification of the glycoprotein of interest. For example, a variety of low molecular weight or insolubilized sialic acid donors can be substrates for the TS. A good donor should also be cost-effective. Finally, the quality of the donor and the ratio donor versus acceptor have to be tested for optimal sialylation levels.
Though a major limitation for sialylation of recombinant glycoproteins using TS is that it transfers only 2,3-linked sialic acids, the main advantage is that a variety of acceptor motifs, including O- and N-glycans, can be sialylated. This is generally not the case with sialyltransferases, as these enzymes have a very high specificity toward the acceptor (for a review, see Harduin-Lepers et al., 1995
). Moreover, the strategy, which uses in vitro sialylation of the purified glycoprotein by a sialyltransferase, is constrained by the cost of CMP-NeuAc, which is the exclusive donor for this family of enzymes.
Although the recent study of Hollister and Jarvis (2001) reports that the coexpression of a ß1,4-galactosyltransferase and an 2,6-sialyltransferase in a stably transformed cell line derived from Sf9 allows the sialylation of a recombinant glycoprotein, the way by which CMP-NeuAc is synthesized remains unclear. In addition, the sialylation levels reached in this system presumably are limited by the CMP-NeuAc availability. Therefore, the trans-sialidase appears to be a potential alternative for the production of
2,3-sialylated glycoproteins in the baculovirus-insect cells system.
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Materials and methods |
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The viruses were propagated in Sf9 cells, and all procedures were done essentially as described by Summers and Smith (1987). For infection, cells were inoculated with a viral suspension at multiplicities of infection (MOIs) of 510 PFU/cell. After a 1-h adsorption at room temperature, the viral inoculum was removed and fresh culture medium was added. Infected cells were further incubated at 28°C for various times. The viral titers were determined by plaque assay.
Construction of the recombinant baculovirus
The cDNA encoding a 6-His tagged truncated form of T. cruzi trans-sialidase (GenBank accession number L26499), cloned in the EcoRI site of the pTrcHisA vector (Invitrogen, The Netherlands) was a kind gift of Dr A.C.C. Frasch (see Buscaglia et al., 1998). This plasmid served as a template for polymerase chain reaction (PCR) to generate modified ends of the TS sequence (Figure 1), using primers from Eurogentec (Belgium) and the VENT polymerase (New England Biolabs, Beverly, MA). Oligonucleotides TST1 (5'-CGA-TTC-TAG-ACT-GGC-ACC-CGG-ATC-GAG-CCG-AGT-TGA-3') and TST2 (5'-CAC-TGT-GGG-ATC-CAC-CAC-ACG-AGA-AAC-AGA-3') were designed for generation of a modified 5' end containing an additional XbaI site and deleted of the 6-His tag. The amplified fragment (307 bp) was excised from the agarose gel and cloned into a pBlueScript plasmid for sequencing using the T7 polymerase kit from Pharmacia (Uppsala, Sweden) and d[35S]ATP from Amersham (UK). The obtained plasmid was then digested by XbaI and BamHI. The digestion fragment was reinserted into pTrcHisATS to generate pTrcHisATSt. Oligonucleotides TSQ1 (5'-GGG-TAA-GAG-GTA-CCA-CGT-CGT-TCT-3') and TSQ2 (5'-CTG-AGA-ATT-CCT-AGG-GCA-CTC-GTG-TCG-CTG-CTG-CTG-TC-3') were used for generation of a modified 3' end containing an additional AvrII site and deleted of the stop codon. The amplified 315-bp-long fragment was excised from the gel and cloned into a pUC19 plasmid for sequencing. The KpnIEcoRI fragment was then reinserted into pTrcHisATSt to generate pTrcHisATStq.
A degenerated sequence encoding the C-terminal transmembrane domain from the baculovirus envelope glycoprotein gp67 (Whitford et al., 1989, GenBank accession number M25420) and containing unique restriction sites at both ends was constructed using a set of 10 overlapping oligonucleotides. The dephosphorylated oligonucleotides were denatured by heating, mixed, and allowed to renature overnight. The resulting fragment (135 bp long) was cloned into a HpaI/SacI-cut pUC vector carrying the signal sequence from the ecdysteroid glucosyltransferase gene of the baculovirus AcMNPV (O'Reilly and Miller, 1989
, GenBank accession number M22619). Several independent clones obtained after ligation and transformation of Escherichia coli were sequenced, and one clone was found to contain the right sequence.
The modified TS sequence was excised by XbaI/AvrII digestion and inserted between the signal peptide and the transmembrane sequence. The final construction was then subcloned into the transfer vector p119, designed for insertion into the p10 locus of the baculovirus.
Sf9 cells were cotransfected by lipofection (Felgner and Ringold, 1989) using DOTAP (Boehringer Mannheim, Germany) with p119-TS and genomic DNA of the modified baculovirus AcSLP10. AcSLP10 is derived from wild-type AcMNPV and possesses only one strong promoter, p10, with the polyhedrin coding sequence inserted downstream of this promoter (Chaabihi et al., 1993
). Thus, it has an occlusion bodypositive phenotype, whereas the recombinant virus is occlusion bodynegative. The screening and purification of the recombinant baculovirus AcP10TS were carried out as described by Summers and Smith (1987)
.
Production of the antibodies
E. coli strain DH5 bacteria were transformed with pTrcHisATS, cultured in 200 ml LB medium with 50 µg/ml; ampicillin and induced by 1 mM isopropyl ß-D-thiogalactoside (Sigma) when the optical density reached 0.6. After a 4-h induction at 30°C, cells were harvested, washed in a cold Tris buffer (pH 7.5) with 1 mM phenylmethylsulfonyl fluoride (PMSF) and were lysed using a French press at a pressure of 10,000 psi. After centrifugation and SDSPAGE analysis, the pellet was found to contain the TS. The pellet was resuspended in denaturing buffer (urea 8 M, NaH2PO4 100 mM, Tris 10 mM, pH 8, PMSF 1 mM). Then the 6-His-tagged TS was batch-purified using an Ni-NTA-agarose resin (Qiagen GmbH, Germany) as described in the manufacturers protocol. Each fraction was analyzed by SDSPAGE; the fractions containing the pure TS were pooled, dialyzed against water, and lyophilized. An approximate yield of 800 µg was obtained and used for immunization of a rabbit by three successive injections.
The antiserum was collected 8 weeks after the first injection, and the antibodies were precipitated with ammonium sulfate and resuspended in a half volume of phosphate buffered saline (PBS, pH 7.2) and dialyzed. The specificity of the antibodies was evaluated by western blot analysis (not shown).
TS assays
For TS assays, we used a protocol in which the sialic acid donor was fetuin (rich in 2,3-linked sialic acids) and the acceptor was [14C]-labeled lactose (Amersham, UK). Each assay contained 1 mM lactose (0.4 µCi) and 20 mg/ml fetuin (Sigma, Neu5Ac content approximately 5%) in a final volume of 100 µl in PBS pH 7.2, with or without 0.5% Triton X100. The incubations were performed at 28°C for 1 h, then the macromolecules were precipitated by the addition of 900 µl cold ethanol. Following centrifugation, the supernatants were dried under nitrogen flow and resuspended in 70% ethanol, and the labeled compounds were separated by paper electrophoresis as described by Leguizamon et al. (1994)
. Sialyllactose and lactose could also be separated by descending paper chromatography in the following solvent: pyridine/ethyl acetate/acetic acid/water 5/5/1/3 (by volume).
For analysis of the TS activity in cell lysates, the infected cells were harvested, washed with cold PBS pH 7.2, and lysed in the lysis buffer (50mM TrisHCl, pH 7.5, 1% Triton X100) and the TS assay was performed as described above.
For analysis of the membrane-bound activity, the infected cells were harvested, washed with cold PBS pH 7.2, and resuspended in PBS. The assay was carried out as described above with this cell suspension (without Triton X100). The cell viability was estimated by Trypan blue exclusion to be around 90%.
For analysis of the soluble activity, the infection medium was clarified by centrifugation 20 min at 50,000 x g in a SW41TI rotor at 4°C. The clarified infection medium was then centrifuged at various speeds and for various times in a SW41TI rotor at 4°C, and the supernatants were assayed for TS activity as above. The clarified infection medium was also filtered through 0.45- and 0.22-µm filters (Pall Gelman Sciences, Ann Arbor, MI) and the filtrates were assayed for TS activity.
When asialo-1-AGP was used as sialic acid acceptor, the desialylation was carried out by incubation of human
1-AGP (Sigma) with 3 M acetic acid at 80°C for 3 h, and the desialylation was checked by gas-liquid chromatography. Pure sialyl-
2,3-lactose was the kind gift of Dr. Gérard Strecker.
Flow cytofluorimetry analysis
Sf9 cells grown in EXTRA 1x medium were infected with the recombinant baculovirus AcP10TS or with the control baculovirus AcSLP10. The cells were harvested at 48 h p.i. and washed three times in cold PBS, then incubated with the antibody or with the preimmune serum diluted in PBS containing 0.1% bovine serum albumin (BSA) for 1 h in ice. Cells were washed then incubated 1 h with the secondary antibody: fluorescein conjugated anti-rabbit (Sigma) diluted in PBSBSA. Then the cells were analyzed in a FACScalibur (Beckton Dickinson, Sunnyvale, CA).
Permeabilization of the cells prior to incubation with the antibodies was performed by incubating the cells for 2 h in ice-cold 70% ethanol at a temperature of 20°C. Then the cells were washed three times with cold PBS and incubated with the antibodies as described above.
Preparation of viral particles and sedimentation on sucrose gradient
Sf9 cells cultured in EXTRA 1x medium were infected with AcP10TS in a 75-cm2 T-flask. Three days after infection, the medium was collected and the viral particles were pelleted by ultracentrifugation at 50,000 x g for 20 min at 4°C (according to Loisel et al., 1997). The supernatant was saved and assayed for TS activity. The pellet was resuspended in cold PBS and centrifuged at low speed to eliminate any cellular debris. This supernatant was then pelleted by another ultracentrifugation at 50,000 x g for 20 min at 4°C. The pellet was resuspended in 1.2 ml of TE buffer (Tris 10 mM, EDTA 1 mM, pH 7.4) containing 1 mM PMSF (Sigma), assayed for TS activity, and layered onto a linear continuous sucrose gradient (2556% w/v in TE buffer). The gradient was centrifuged at 100,000 x g for 90 min in a SW41TI rotor at 4°C. Fractions were collected from the bottom to the top of the gradient, dialyzed against water, and assayed for TS activity as described. An aliquot of each fraction was analyzed by SDSPAGE, and western blot analysis was performed using a mouse monoclonal antibody against gp67, the major envelope glycoprotein of the baculovirus. This anti-gp67 antibody (AcV5, Hohmann and Faulkner, 1983
; Monsma and Blissard, 1995
) was a kind gift of Dr. G.W. Blissard. The secondary antibody was a goat anti-mouse antibody conjugated to horseradish peroxydase (Sigma). The immunoreactive bands were detected by chemiluminescence.
Western blot analysis of the recombinant TS
To allow an easier detection, Sf9 cells adapted to a serum-free medium, EXTRA SB5 supplemented with cholesterol, were used for western blot analysis. Sf9 cells adapted to this medium were infected with AcP10TS or with the control AcSLP10 at MOIs of 10. At 3 days p.i., the cells were harvested and lysed in the lysis buffer (TrisHCl, pH 7.5, 1% Triton X100) and assayed for TS activity. The infection supernatants were clarified by centrifugation at 50,000 x g for 20 min at 4°C and concentrated five times using a Microsep unit (Pall Gelman Sciences). PNGase F digestions were carried out according to the manufacturers protocol (New England Biolabs). The samples were first denatured 10 min at 100°C in denaturing buffer (0.5% SDS, 1% ß-mercaptoethanol), then the incubations were carried out for 1 h at 37°C with 15 mIU PNGase F in the incubation buffer (sodium phosphate 0.05 M, pH 7.5) with 1% NP40. Control samples were denatured similarly and treated with the buffer only. Finally, the samples were placed in Laemmli buffer, boiled, and ran on denaturing gel electrophoresis (Laemmli, 1970). Proteins were electrotransferred to nitrocellulose sheets (Schleicher & Schuell, Germany), and visualized by Ponceau red staining (Sigma). The nitrocellulose membranes were blocked in 2% gelatin (Sigma) in Tris buffered saline (TBS) buffer (TrisHCl 15mM, pH 8, NaCl 140 mM, 0.05% Tween 20). The proteins were detected by using the polyclonal anti-TS antibody as the primary antibody (1/10,000 in TBS buffer) and a goat anti-rabbit IgG conjugated to horseradish peroxidase (Dako, Glostrup, Denmark, 1/1000 in TBS buffer). The immunoreactive bands were detected by chemiluminescence using an ECL kit from Pharmacia (Uppsala, Sweden).
Native gel electrophoresis
For nondenaturing gels, the samples were placed in a sample buffer (187 mM TrisHCl, pH 8.8, 1% sucrose, 0.005% bromophenol blue) and ran in a 515% gradient gel without SDS in the electrophoresis buffer (TrisHCl, 0.025 M, glycine 0.2 M, pH 8.5). The western blot was performed as above.
Lectin blotting
After SDSPAGE and electrotransfer as described above, the nitrocellulose sheets were blocked in polyvinylpyrrolidone (2% in TBS buffer), then incubated with digoxigenin-conjugated MAA (specific for 2,3-linked sialic acids); blocked again in blocking reagent; incubated with Fabanti-digoxigenin fragments, conjugated to alkaline phosphatase, and finally revealed with nitroblue tetrazolium chloride-5-bromo-4-chloro-3-indolyl-phosphate (all reagents from Boehringer Mannheim). Control samples were treated before SDSPAGE with 50 mU/ml C. perfringens sialidase (Sigma) in 100 µl of incubation buffer (sodium citrate 50 mM, pH 6, NaCl 0.9%, CaCl2 0.1%) for 1 h at 37°C.
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Acknowledgments |
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Abbreviations |
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Footnotes |
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References |
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