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. 2012 Dec;84(21):24–30. doi: 10.1016/j.phytochem.2012.08.023

Myrosinases TGG1 and TGG2 from Arabidopsis thaliana contain exclusively oligomannosidic N-glycans

Eva Liebminger a, Josephine Grass b, Jakub Jez a, Laura Neumann b, Friedrich Altmann b, Richard Strasser a,⁎
PMCID: PMC3494833  PMID: 23009876

Graphical abstract

All N-glycosylation sites of endogenous thioglucoside glucohydrolases TGG1 and TGG2 from Arabidopsis leaves are occupied by oligomannosidic N-glycan structures, which are generated by class I α-mannosidases.

graphic file with name fx1.jpg

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Highlights

► TGG1 and TGG2 are glycosylated with oligomannosidic N-glycans. ► TGG1 N-glycans are processed by class I alpha-mannosidases. ► TGG1 transcript levels are higher in leaves of mns1 mns2 mns3 plants.

Keywords: Arabidopsis thaliana, Post-translational modification, N-Glycosylation, Glycan, Glycoprotein, Mannosidase, Thioglucoside glucohydrolase

Abstract

In all eukaryotes N-glycosylation is the most prevalent protein modification of secretory and membrane proteins. Although the N-glycosylation capacity and the individual steps of the N-glycan processing pathway have been well studied in the model plant Arabidopsis thaliana, little attention has been paid to the characterization of the glycosylation status of individual proteins. We report here the structural analysis of all N-glycans present on the endogenous thioglucoside glucohydrolases (myrosinases) TGG1 and TGG2 from A. thaliana. All nine glycosylation sites of TGG1 and all four glycosylation sites of TGG2 are occupied by oligomannosidic structures with Man5GlcNAc2 as the major glycoform. Analysis of the oligomannosidic isomers from wild-type plants and mannose trimming deficient mutants by liquid chromatography with porous graphitic carbon and mass spectrometry revealed that the N-glycans from both myrosinases are processed by Golgi-located α-mannosidases.

1. Introduction

Arabidopsis thaliana is the preferred organism for the investigation of the glycan structure–function relationship in plants. By analysis of mutants with defects in various maturation steps it has become clear that N-glycan processing is essential for plant development (Boisson et al., 2001; Burn et al., 2002; Gillmor et al., 2002; Liebminger et al., 2009) and for abiotic as well as biotic stress reactions (Häweker et al., 2010; Kang et al., 2008; Lu et al., 2009). N-glycosylation mutants displaying a severe developmental defect have also drastically altered cell walls and many proteins involved in cell wall synthesis or remodelling are glycoproteins (Kang et al., 2008; Liebminger et al., 2009; Zhang et al., 2011). Moreover, plasma membrane located receptor kinases essential for plant growth (e.g. BRI1) or pathogen perception (e.g. EFR, FLS2) are heavily glycosylated and contain numerous N-glycosylation sites in their extracellular ligand binding domains (Gómez-Gómez and Boller, 2000; Li and Chory, 1997; Zipfel et al., 2006). In spite of these important findings, the analysis of the glycosylation site occupancy and oligosaccharide structures present on individual A. thaliana glycoproteins have been largely ignored in the past. The glycosylation status of endogenous A. thaliana proteins has mainly been determined by shifts in mobility on SDS–PAGE upon enzymatic or chemical deglycosylation and consequently the extent and heterogeneity of the attached oligosaccharide structures are unknown. Apart from the glycan analysis of recombinant proteins expressed in different A. thaliana organs (e.g. Schähs et al., 2007; Van Droogenbroeck et al., 2007) structural information of N-glycans is derived mainly from studies performed by enzymatic release of the oligosaccharides from total protein extracts (Henquet et al., 2008; Kajiura et al., 2010b; Rayon et al., 1999; Rendić et al., 2007; Strasser et al., 2004). The first comprehensive analysis of N-glycosylation sites present in the A. thaliana proteome has been reported recently (Zielinska et al., 2012) and in one study glycopeptides from the putative cell wall proteome have been identified by MALDI-TOF MS (Zhang et al., 2011).

Here, we performed for the first time a detailed characterization of the N-glycan structures present on two endogenous glycoproteins isolated from A. thaliana leaves. The oligosaccharide structures present on the myrosinases THIOGLUCOSIDE GLUCOHYDROLASE1 (TGG1) and TGG2 were determined by mass spectrometry of glycopeptides and chromatographic comparison by porous graphitic carbon-liquid chromatography–electrospray ionization mass spectrometry (PGC-LC–ESI-MS). TGG1 and TGG2 belong to the glycoside hydrolase superfamily and produce toxic compounds against pathogens and insects by catalyzing the hydrolysis of glucosinolates (Barth and Jander, 2006; Husebye et al., 2002). The two myrosinases display an organ-specific expression pattern and accumulate mainly in rosette leaves, flowers and siliques, but not in roots (Ueda et al., 2006). In previous studies it has been shown that TGG1 and TGG2 have redundant functions in terms of glucosinolate degradation and hormone signalling (Barth and Jander, 2006; Islam et al., 2009). Both enzymes are glycosylated, but the number of glycans, the type of glycans and the specific role of N-glycosylation for their biological function are unknown (Ueda et al., 2006; Zhou et al., 2012).

2. Results and discussion

2.1. TGG1 and TGG2 accumulate abnormally in mns mutants

In a previous study we have identified mns mutants with defects in processing of oligomannosidic N-glycans (Liebminger et al., 2009). The mns mutants display altered root formation and the triple mutant (mns1 mns2 mns3) which completely lacks class I α-mannosidase activity, exhibits reduced leaf growth and rosette diameter. To identify potential candidate glycoproteins whose functions are altered in mns mutants compared to wild-type plants, we analyzed total protein extracts from leaves by SDS–PAGE and Coomassie Brilliant Blue staining. Two aberrant bands of ca. 75 and 65 kDa were detectable in the mutants (Fig. 1A). The two protein bands were excised from the gel, tryptic digested and analyzed by ESI-MS. Peptides from the 75 kDa protein band could be assigned to TGG1 and peptides from the 65 kDa band to TGG2 (data not shown). Immunoblot analysis with antibodies specific for TGG1 and TGG2 confirmed that both the mobility and the abundance of the two myrosinases are altered in mns mutants compared to control plants (Fig. 1B and C). In particular, the mns1 mns2 mns3 triple mutant displayed a pronounced shift and an almost 3-fold increase in signal intensity for TGG1 and TGG2. Differences in abundance of TGG1 have also been described for mutants with defects in the assembly of N-glycans (Farid et al., 2011; Koiwa et al., 2003; Zhang et al., 2008, 2009) suggesting that N-glycosylation of myrosinases could affect their stability.

Fig. 1.

Fig. 1

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TGG1 and TGG2 abundance and mobility are altered in mns mutants. (A) Coomassie Brilliant Blue staining of protein extracts from wild-type, mns3 single (3), mns1 mns2 double (12) and mns1 mns2 mns3 triple (123) mutants. (B) Immunoblot analysis using anti-TGG1 and (C) anti-TGG2 antibodies. Antibodies against UGPase were used as a control.

2.2. TGG1 and TGG2 are glycosylated with oligomannosidic N-glycans

To assess the degree of N-glycan processing protein extracts from A. thaliana wild-type leaves were subjected to endoglycosidase H (Endo H) and Peptide: N-glycosidase F (PNGase F) digestion followed by immunoblotting with TGG1 and TGG2 specific antibodies. For both myrosinases shifts in mobility were visible upon enzymatic deglycosylation and the Endo H and PNGase F digested bands showed co-migration, which is indicative for the presence of predominantly oligomannosidic N-glycans (Fig. 2A and B). However, plant complex N-glycans frequently contain α1,3-fucose residues linked to the innermost GlcNAc, which make them insensitive to PNGase F digestion (Tretter et al., 1991). Therefore we performed endoglycosidase treatments of TGG1 and TGG2 in the fut11 fut12 line, which generates complex N-glycans lacking core α1,3-fucose (Strasser et al., 2004). Deglycosylated TGG1 and TGG2 from fut11 fut12 were indistinguishable from deglycosylated wild-type forms strongly indicating the absence of any α1,3-fucosylated complex N-glycans (Fig. 2C and D).

Fig. 2.

Fig. 2

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TGG1 and TGG2 are sensitive to endoglycosidase digestion. Protein extracts of wild-type (A and B) and fut11 fut12 (C and D) were digested with Endo H and PNGase F, subjected to SDS–PAGE and analyzed by immunoblotting with TGG1 (A and C) and TGG2 (B and D) specific antibodies.

Next we investigated the glycan site occupancy and glycan structures of TGG1 and TGG2. Protein extracts from A. thaliana leaves were separated by SDS–PAGE, bands corresponding to TGG1 and TGG2 were excised from the gel, digested by proteases and analyzed by mass spectrometry. For TGG1 nine glycopeptides were identified showing that all potential N-glycosylation sites are subjected to glycosylation (Figs. 3 and S1). The TGG1 glycopeptides displayed masses matching with different oligomannosidic structures, but lack paucimannosidic or complex N-glycans. Man5GlcNAc2 was the main oligosaccharide found on the majority of TGG1 glycopeptides. In addition, Man6GlcNAc2, Man7GlcNAc2 and Man8GlcNAc2 were also present on several glycosylation sites while on Asn108 the predominant peak corresponds to Man9GlcNAc2 indicating that this glycopeptide is not accessible for further N-glycan trimming. The same N-glycan structures were observed when TGG1 was purified by Concanavalin A Sepharose from protein extracts followed by LC–ESI-MS analysis (data not shown).

Fig. 3.

Fig. 3

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TGG1 glycopeptides contain exclusively oligomannosidic N-glycans. The spectra of all nine glycopeptides (GP 1-GP 9) derived from endogenous A. thaliana TGG1 are shown. Peaks corresponding to N-glycan structures are labelled. Sodium adducts are indicated (#). Spectra were obtained by LC–ESI-MS analysis of trypsin/GluC double digests.

For TGG2, four glycopeptides could be identified revealing that TGG2 is fully glycosylated. The assigned glycan profile of TGG2 is very similar to TGG1 showing only oligomannosidic N-glycans with Man5GlcNAc2 as predominant peak (Fig. S2).

2.3. TGG1 from mns mutants display incompletely processed oligomannosidic N-glycans

The observed shifts in mobility on protein gels suggest that both myrosinases are aberrantly glycosylated in the mannose trimming deficient mns mutants. Consequently, we analyzed corresponding glycopeptides of TGG1 from different mns mutants to figure out if the myrosinase is indeed processed by MNS proteins. Consistent with the previously described glycosylation defect (Liebminger et al., 2009) TGG1 N-glycans from mns3 plants displayed increased levels of Man6GlcNAc2 and Man7GlcNAc2 structures. The spectra derived from mns1 mns2 double and mns1 mns2 mns3 triple knockouts contained almost exclusively Man8GlcNAc2 and Man9GlcNAc2 N-glycans, respectively (Figs. 4 and S3). These data show clearly that TGG1 N-glycans are processed by MNS3 and MNS1/MNS2.

Fig. 4.

Fig. 4

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TGG1 N-glycans are trimmed by class I α-mannosidases. LC–ESI-MS analysis of TGG1 glycopeptide 1 (GP 1) from wild-type, mns3, mns1 mns2 and mns1 mns2 mns3 mutants. Sodium adducts are indicated (#).

Since oligomannosidic N-glycans very often represent a mixture of different isomers we subjected released N-glycans from TGG1 to PGC-LC–ESI-MS analysis, which allows the separation and identification of oligomannosidic isomers (Pabst et al., 2012). PGC-LC analysis confirmed the presence of Man5GlcNAc2 to Man9GlcNAc2 oligosaccharide structures. The Man5GlcNAc2 isomer (M6M3)M was the most abundant glycan on TGG1 extracted from wild-type plants (Fig. 5). Accordingly, we could assign the predominant Man6GlcNAc2 structure to (M6M2-3)M in mns3 as well as the Man8GlcNAc2 to (M2-6M3)M2-2 in mns1 mns2 and the Man9GlcNAc2 to (M2-6M2-3)M2-2 in mns1 mns2 mns3 (Table 1). These findings are in good agreement with current models of N-glycan processing in the early secretory pathway of A. thaliana (Liebminger et al., 2009).

Fig. 5.

Fig. 5

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Determination of the predominant N-glycan isomers on TGG1. Selected ion chromatograms (SIC) of oligomannosidic N-glycans from TGG1 are shown. TGG1 was extracted from leaves of wild-type plants and reduced N-glycans were separated by PGC-LC–ESI-MS. The assignment of structural isomers was based on reference glycans as described in detail previously (Pabst et al., 2012).

Table 1.

N-Glycan isomers of TGG1.

N-Glycan structure Proglycan code Relative abundance on individual glycosylation sites (%)
Wild-type mns3 mns1 mns2 mns1 mns2 mns3
Man5 Man5.1 (M6M3)M 56.8 15.8 – –



Man6 Man6.1 (M6M3)M2 15.3 – – –
Man6.10 (M6M2-3)M – 51.5 – –



Man7 Man7.1 (M6M3)M2-2 7.1 – 9.5 –
Man7.2 (M2-6M3)M2 3.8 – 20.8 –
Man7.7 (M6M2-3)M2 – 10.5 – –



Man8 Man8.1/8.4 (M2-6M3)M2-2/(M2-6M2-3)M2 8.7 9.7 57.6 22.8
Man8.2 (M6M2-3)M2-2 – 2.0 – 3.1



Man9 Man9.1 (M2-6M2-3)M2-2 8.3 10.5 12.1 74.1
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The Proglycan code for oligomannosidic glycans has been described in detail recently (Pabst et al., 2012).

2.4. TGG1 transcript levels are higher in leaves of mns1 mns2 mns3 plants

The most prominent difference in expression levels were found for TGG1 in the mns1 mns2 mns3 triple mutant (Fig. 1). It has been proposed that TGG1 is cell type-specific expressed in specialized myrosin cells and is also highly abundant in guard cells of leaves (Husebye et al., 2002; Ueda et al., 2006; Zhao et al., 2008). To monitor if the elevated TGG1 levels are caused by an increased number of guard cells we compared the stomata in leaves from mns1 mns2 mns3 to wild-type plants (Fig. 6A). Neither the number nor the morphology of stomata was affected in the mutant.

Fig. 6.

Fig. 6

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Phenotypic analysis of wild-type and mutant plants. (A) Images of the epidermis of mature rosette leaves. Lower panel: staining of stomata from Col-0 and mns1 mns2 mns3 (mns123) with FM4-64 dye. Bar = 50 μm. (B) Quantitative real time PCR (qPCR) analysis of TGG1 transcript levels in wild-type and mutant plants. (C) Phenotypes of 3-week-old wild type and mutant plants grown on soil under long day conditions. (D) 10-day-old seedlings grown on 1× Murashige and Skoog medium supplemented with 2% sucrose. Bar = 5 mm.

Next we addressed whether TGG1 transcript expression is altered in the triple mutant. Quantitative PCR showed that TGG1 transcript levels are at least 5-fold higher in the mns1 mns2 mns3 mutant (Fig. 6B) suggesting that under conditions of aberrant glycosylation increased amounts of TGG1 are beneficial for the plant. To investigate the effect of TGG1 deficiency in the mns1 mns2 mns3 line, we generated tgg1 mns1 mns2 mns3 quadruple knockouts and compared its growth and development to mns1 mns2 mns3. Under our growth conditions seedlings as well as the areal parts of soil grown quadruple knockouts were indistinguishable from mns1 mns2 mns3 (Fig. 6C and D) being consistent with the current model that TGG1 is involved in glucosinolate breakdown to deter herbivores and pathogens but plays no role for the normal development of A. thaliana (Barth and Jander, 2006; Husebye et al., 2002; Ueda et al., 2006).

2.5. TGG1 and TGG2 N-glycans are not altered in the mvp1 mutant

Recently, a protein trafficking mutant was identified, which displayed mislocalization of TGG1 and TGG2 to aggregates within the cell (Agee et al., 2010). Apart from mistargeting of vacuolar proteins the modified vacuole phenotype1 (mvp1-1 and mvp1-2) mutants displayed also aberrant localization of Golgi-resident proteins as observed for a fluorescent protein tagged N-acetylglucosaminyltransferase I (GnTI or NAG1), which was used as a Golgi marker (Agee et al., 2010). Since GnTI is the key enzyme for the initiation of complex N-glycan formation in plants (Strasser et al., 1999; von Schaewen et al., 1993) its aggregate formation in the mvp1 mutants might cause aberrant glycosylation of proteins and could also affect the two myrosinases. To address whether mvp1 plants display overall changes in N-glycan processing we first analyzed the total N-glycosylation profile of endogenous proteins. Despite the reported global defects in subcellular localization of proteins, the N-glycan profile from mvp1-2 was essentially the same when compared with wild-type derived N-glycans (Fig. S4). In addition, aggregate formation of TGG1 and TGG2 in mvp1-2 did not have an effect on their N-glycosylation profile. As shown in Fig. S5 the processed oligomannosidic N-glycans on TGG1 and TGG2 glycopeptides resemble the oligosaccharides found on in wild-type derived myrosinases showing that neither the aberrant localization of the TGG1 and TGG2 nor global defects in protein trafficking result in changes of N-glycosylation in MVP1 deficient plants.

3. Conclusions

The fundamental role of N-glycosylation for development and response to adverse environmental conditions is very well documented in the plant model organism A. thaliana. Remarkably, very little is known about the N-glycosylation of individual A. thaliana proteins and the specific function of their N-glycans. Here, we analyzed the glycans of two myrosinases expressed in leaves of wild-type plants and in four different mutant backgrounds. To date, this is the first comprehensive N-glycan analysis of endogenous A. thaliana glycoproteins with multiple N-glycosylation sites. Our structural analysis reveal that both myrosinases contain exclusively oligomannosidic N-glycans with the (M6M3)M Man5GlcNAc2 isomer as predominant structure. The data from the mns mutants confirm that this oligomannosidic N-glycan is processed by the α-mannosidases MNS1 to MNS3. Surprisingly, there is no further processing of TGG1 and TGG2 N-glycans by GnTI, which utilizes (M6M3)M as substrate to initiate hybrid and complex N-glycan formation in plants (Strasser et al., 1999; von Schaewen et al., 1993). MNS1 and MNS2 are like GnTI cis/medial Golgi-located enzymes (Liebminger et al., 2009; Kajiura et al., 2010a; Nebenführ et al., 1999; Saint-Jore-Dupas et al., 2006; Schoberer et al., 2009; Staehelin and Kang, 2008) which raises the question how this myrosinase-specific glycosylation pattern is generated. We can provide three possible explanations: (i) the N-glycans from TGG1 and TGG2 are not accessible for GnTI and are therefore not further processed. There are no clear data from secreted plant glycoproteins available that support this explanation, but in mammals it has been shown that the occurrence of oligomannosidic N-glycans on the envelope glycoprotein from HIV-1 is at least in part caused by steric constraints that prevent further processing in the Golgi (Doores et al., 2010). (ii) TGG1 and TGG2 are only expressed in specialized cells that lack GnTI activity. While this possibility cannot be ruled out it is commonly believed that most of the Golgi-located N-glycan processing enzymes are expressed ubiquitously in plants and with the exception of the Lewis a structures (Strasser et al., 2007) no cell- or organ-specific glycosylation profile has been described for A. thaliana so far. (iii) In TGG1 and TGG2 expressing cells MNS proteins and GnTI are located in different sub-Golgi compartments. If the final destination for both myrosinases is the vacuole (Ueda et al., 2006), this latter hypothesis would suggest a specific sorting from the cis-Golgi (or MNS compartment) to the vacuole without trafficking through other Golgi cisternae. Current models for subcellular localization of glycosylation enzymes suggest the existence of some kind of assembly line across the Golgi stack (Saint-Jore-Dupas et al., 2006; Schoberer and Strasser, 2011). However, additional studies using advanced imaging and protein–protein interaction technologies (Sparkes et al., 2011) are required to reveal the organization of Golgi-located glycosylation enzymes in plants.

The biological significance of TGG1 and TGG2 N-glycosylation remains elusive and the mechanisms as well as the consequences of the increased myrosinase expression in the mns1 mns2 mns3 mutant are unclear. Further studies are necessary to unravel a biological role for the processed oligomannosidic N-glycan structures on TGG1 and TGG2 and provide a link to myrosinase function during pathogen attack. In summary, our in depth structural analysis offers interesting new insights into N-glycan processing of individual plant proteins that are missed by mass spectrometry of N-glycans from large pools of glycoproteins.

4. Experimental

4.1. Plant material and growth conditions

A. thaliana wild-type (ecotype Columbia-0) and mutant plants were grown as described previously (Liebminger et al., 2009). Seeds of tgg1-1 (Barth and Jander, 2006) and mvp1-2 (Agee et al., 2010) were obtained from the Nottingham Arabidopsis Stock Centre (NASC). Line tgg1-1 was crossed with the mns1 mns2 mns3 triple knockout line (Liebminger et al., 2009) to generate the tgg1 mns1 mns2 mns3 quadruple mutant. Homozygous lines were confirmed by genotyping using the following primer combinations: At1g51590-1F/2R for mns1, At3g21160-1F/2R for mns2, At1g30000-3F/2R for mns3; At5g26000-1F/2R for tgg1 and MVP-1F/2R for mvp1-2. All primer sequences are presented in Table S1.

4.2. SDS–PAGE and LC–ESI-MS analysis of glycopeptides

Rosette leaf material from A. thaliana wild-type and mns single, double and triple knockouts was ground in liquid nitrogen and resuspended in 1× phosphate buffered saline (PBS). Samples were incubated for 10 min on ice and then centrifuged for 10 min at 9.600g. The supernatant was centrifuged for 5 min at 9.600g and mixed with SDS–PAGE loading buffer. The samples were incubated for 5 min at 95 °C, separated by SDS–PAGE (10%) under reducing conditions and polypeptides were detected by Coomassie Brilliant Blue staining. The corresponding bands were excised from the gel, destained, carbamidomethylated, in-gel digested with trypsin and GluC and analyzed by LC–ESI-MS as described previously (Stadlmann et al., 2008; Kolarich et al., 2012).

4.3. PGC-LC–ESI-MS analysis

For isomeric analysis, the above described glycopeptides were digested with peptide: N-glycosidase A (Proglycan, Vienna, Austria). The liberated glycans were reduced with sodium borohydride and subsequently purified using graphitic carbon cartridges (Pabst et al., 2012).

Reduced oligosaccharides were analyzed by PGC-LC–ESI-MS on a Hypercarb column (0.32 × 100 mm, Thermo Fisher Scientific) coupled to an Ultimate 3000 capillary HPLC (Thermo Fisher Scientific) and a Q-TOF Ultima MS (Waters) (Pabst et al., 2007) Briefly, the aqueous solvent was 0.3% formic acid buffered to pH 3.0 and a linear gradient from 8.8% to 17.2% acetonitrile was developed during 55 min. Detection was accomplished by positive mode ESI-MS. Assignment of isomers was based on reference oligosaccharides prepared from kidney beans and bovine ribonuclease B as described in detail previously (Pabst et al., 2012). For retention time normalization, the diantennary, complex-type asialo-N-glycan A4A4 was added to each sample.

4.4. Deglycosylation with PNGase F and Endo H

Protein extraction from wild-type and fuct11 fuct12 line (Strasser et al., 2004) was performed as described above. Prior to PNGase F/Endo H treatment, the protein extracts were mixed with 10x denaturation buffer (New England Biolabs, Frankfurt, Germany), boiled for 10 min at 98 °C and then cooled for 2 min on ice. For PNGase F digestion, the extract (10 μL) was mixed with 10× G7 reaction buffer containing 10% NP-40 (New England Biolabs), water and 0.5 U PNGase F (New England Biolabs) in a total volume of 15 μL. For Endo H digestion, the extract was mixed with 10× G5 reaction buffer (New England Biolabs), water and 0.5 U Endo H (New England Biolabs) in a total volume of 15 μL. Controls were treated as described above but the respective enzymes were replaced by water. After incubation of the reaction mixture for 3 h at 37 °C, SDS–PAGE loading buffer was added and the samples were heated to 95 °C for 5 min. Deglycosylated proteins and controls were subjected to SDS–PAGE (10%) followed by immunoblotting with anti-TGG1 and anti-TGG2 antibodies (kind gift of Ikuko Hara-Nishimura) (Ueda et al., 2006). A rabbit polyclonal antiserum against UDP-glucose pyrophosphorylase (anti-UGPase, Agrisera, Vännäs, Sweden) was used as a control for equal loading.

4.5. FM4-staining and imaging

A. thaliana seedlings (6 to 8-day-old) were incubated in 50 μM FM4-64 dye (Invitrogen, F34653) for an appropriate time. Seedlings were then mounted on a slide and imaging of stomata was performed using a Leica TCS SP2 confocal laser scanning microscope as described previously (Schoberer et al., 2009).

4.6. RNA Isolation, reverse transcription and qPCR

Total RNA from rosette leaves was isolated using a SV Total RNA Isolation kit (Promega). First strand cDNA was synthesized from 1.5 μg of total RNA at 42 °C using oligo dT primers and AMV reverse transcriptase (Promega) in a total volume of 20 μl. PCR reactions were performed in a Rotor-Gene RG-3000A (Corbett, Qiagen, Hilden, Germany). Reactions contained 4 μL 5× HOT FIREPol® EvaGreen® qPCR Mix Plus (no ROX) (Solis BioDyne, Tartu, Estonia), 0.1 μL cDNA and 300 nM of each gene-specific primer in a total volume of 20 μL. The following profile was used for all PCR reactions: 95 °C for 15 min, 40 cycles of 95 °C for 15 s, 60 °C for 20 s, 72 °C for 20 s. To detect TGG1 transcripts cDNA was amplified with primers At5g26000-11F/12R and normalized to the expression of the protein phosphatase 2A (PP2A) gene (primers At1g13320-1F/2R). The specificity of the PCR amplification was checked with a melting curve analysis from 65 to 99 °C and data were analyzed using Rotor-Gene software (version 6). PCR reactions were done in triplets and at least three independent biological experiments were performed.

Acknowledgments

We thank Ikuko Hara-Nishimura (Department of Botany, Graduate School of Science, Kyoto University, Kyoto, Japan) for the kind gift of anti-TGG1 and anti-TGG2 antibodies. We also thank Cornelia Konlechner (Department of Applied Genetics and Cell Biology) for help with qPCR experiments and Karin Polacsek (Department of Chemistry) for N-glycan analysis. This work was funded by the Austria Science Fund (FWF): Project P20817-B12 and by the Ph.D. programme “BioToP – Biomolecular Technology of Proteins” from the Austrian Science Funds (FWF): Project W1224-B09.

Footnotes

Appendix A

Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.phytochem.2012.08.023.

Appendix A. Supplementary data

Supplementary data 1

This document contains Supplementary Figs. S1–S5 and Table S1.

mmc1.pdf (515.8KB, pdf)

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Supplementary Materials

Supplementary data 1

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