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. 2010 Apr 13;153(2):693–702. doi: 10.1104/pp.109.152363

The Changing Fate of a Secretory Glycoprotein in Developing Maize Endosperm1,[C],[W]

Elsa Arcalis 1, Johannes Stadlmann 1, Sylvain Marcel 1,2, Georgia Drakakaki 1,3, Verena Winter 1, Julian Rodriguez 1, Rainer Fischer 1, Friedrich Altmann 1, Eva Stoger 1,*
PMCID: PMC2879800  PMID: 20388665

Abstract

Zeins are the major storage proteins in maize (Zea mays) endosperm, and their accumulation in zein bodies derived from the endoplasmic reticulum is well characterized. In contrast, relatively little is known about post-Golgi compartments or the trafficking of vacuolar proteins in maize endosperm, specifically the presence of globulins in structures resembling protein storage vacuoles that appear in early to mid-stage seed development. We investigated this pathway by expressing and analyzing a recombinant reporter glycoprotein during endosperm maturation, using a combination of microscopy and sensitive glycopeptide analysis. Specific N-glycan acceptor sites on the protein were followed through the stages of grain development, revealing a shift from predominantly paucimannosidic vacuolar glycoforms to predominantly trimmed glycan structures lacking fucose. This was accompanied by a change in the main subcellular localization of the protein from large protein storage vacuole-like post-Golgi organelles to the endoplasmic reticulum and zein bodies. The endogenous storage proteins corn α-globulin and corn legumin-1 showed a similar spatiotemporal profile both in transgenic plants expressing the reporter glycoprotein and in wild-type plants. This indicates that the shift of the intracellular trafficking route, as observed with our reporter glycoprotein, may be a common strategy in maize seed development.

Storage proteins in cereal seeds accumulate in different compartments of the endosperm cell, and their abundance and distribution varies according to the species. While in most cereals prolamins are the more abundant class of storage proteins, small-grain species (e.g. wheat [Triticum aestivum], oat [Avena sativa], and barley [Hordeum vulgare]) may contain variable proportions of both prolamins and globulins, and these are delivered to the protein storage vacuole (PSV) via Golgi-dependent and Golgi-independent pathways (Wettstein, 1980; Levanony et al., 1992; Herman and Schmidt, 2004; Takahashi et al., 2005; Cameron-Mills and von Tosi et al., 2009). In rice (Oryza sativa), where globulins and prolamins accumulate in distinct storage compartments, most globulins (mainly glutelins) accumulate in PSVs whereas prolamins aggregate into dense protein bodies within the rough endoplasmic reticulum (ER) and remain in ER-derived organelles (Okita and Rogers, 1996). Maize (Zea mays) stores mainly prolamins (zeins) comprised in three zein subfamilies (α, γ, and δ) that form ER-derived zein bodies. Mature zein bodies consist of a central core of α and δ zeins, while γ zeins are mainly found in the periphery (Lending and Larkins, 1989). Small amounts of globulins also accumulate in maize endosperm, i.e. corn α-globulin (CAG) and corn legumin-1 (CL-1; Woo et al., 2001). Unlike legumin homologs in other plant species including cereals, CL-1 lacks the canonical asparaginyl endopeptidase cleavage sequence (Woo et al., 2001), so it is not cleaved into α and β chains (Yamagata et al., 2003). CAG has been observed in small, PSV-like compartments within the maize endosperm cell (Woo et al., 2001) and a similar fate has been predicted for CL-1 (Yamagata et al., 2003). The identification and localization of globulins in maize indicates the presence of storage vacuoles in maize endosperm, but it does not address the question whether the size and number of these organelles is significant in maize, whether they change morphologically during seed maturation, and how proteins reach this destination.

Proteins may reach the PSV by different routes, and in some species storage protein trafficking appears to undergo changes during seed development. For example, in the context of 2S and 11S storage protein trafficking in pumpkin (Cucurbita pepo) and castor bean (Ricinus communis) it has been proposed that seed developmental stages may be important in determining the transport routes to the PSV (Vitale and Hinz, 2005). A seed-development-mediated change in the trafficking route of wheat prolamins has been suggested earlier as well (Shy et al., 2001; Tosi et al., 2009). One approach to study such change in trafficking routes along seed maturation is to scrutinize the glycosylation pattern of proteins destined to the PSV, taking advantage of the fact that the intracellular trafficking route of a glycoprotein determines its final N-glycan structures (Lerouge et al., 1998).

The first stage of N-glycosylation (which takes place in the ER) involves the cotranslational addition of a precursor oligosaccharide (Glc3Man9GlcNAc2) that is modified by various glycosidases and glycosyltransferases to form the final glycan structure as the protein migrates through the endomembrane system (Lis and Sharon, 1993; Lerouge et al., 1998). ER-resident glycoproteins contain high-Man-type N-glycans whereas proteins passing though the Golgi apparatus contain complex-type N-glycans that include α(1-3)-Fuc and/or β(1-2)-Xyl residues (Lerouge et al., 1998). While secreted glycoproteins contain terminal GlcNAc residues in addition to the core Fuc and Xyl, these terminal residues are trimmed off by enzymes either en route to the vacuole or within the vacuole (Lerouge et al., 1998). Thus the structure of N-glycans is a useful indicator for the intracellular pathway of a protein (Vitale and Hinz, 2005).

Unfortunately, most seed storage proteins, particularly those in cereals, are not glycosylated. However, information on N-glycan structures can be obtained from recombinant glycoproteins. For example, a KDEL-tagged antibody, which was located primarily in ER-derived zein bodies, was predominantly made up of molecules with single GlcNAc residues lacking Fuc (Rademacher et al., 2008). In contrast, recombinant human lactoferrin isolated from maize seeds was reported to contain pauci-Man-type N-glycans with β(1,2)-Xyl and α(1,3)-linked core Fuc (Samyn-Petit et al., 2001). Interestingly, this glycan pattern suggests a vacuolar location of this recombinant protein, and provides a second strong evidence for the presence of PSVs in maize, although the actual subcellular localization of lactoferrin in maize endosperm cells has not been confirmed.

In previous studies we have shown that recombinant glycoproteins can help to clarify questions about the intracellular trafficking of proteins in cereal endosperm, and we found that a recombinant fungal phytase, although secreted from leaf cells, is mainly localized in the PSVs of wheat and rice endosperm (Arcalis et al., 2004; Drakakaki et al., 2006). In this study we used recombinant phytase to facilitate the visualization and characterization of the PSVs in maize, and we followed the intracellular fate of recombinant phytase in developing endosperm using a combination of microscopy and N-glycan analysis, revealing that the trafficking of the protein does indeed change as the seed matures. This behavior is mirrored by the two endogenous (aglycosylated) globulins, CAG and CL-1, indicating that the diversion of storage proteins may be a common strategy in seed development.

RESULTS

Comparative Accumulation of Recombinant Phytase and Endogenous Storage Proteins

Transgenic maize plants expressing a fungal phytase under the control of the endosperm-specific rice glutelin-1 promoter (Drakakaki et al., 2005) were used as a model to study the trafficking of a recombinant glycoprotein carrying an N-terminal sequence for entry into the endomembrane system. As expected from the previously established activity of the promoter in this background, only small amounts of recombinant phytase were produced at 10 d after pollination (DAP), but larger amounts were detected between 15 and 20 DAP. Recombinant phytase extracted from maize seeds migrated as a blurred band with an apparent molecular mass of 60 to 65 kD, as observed earlier with glycosylated phytase from rice seeds (Drakakaki et al., 2006). Phytase remained at high levels in further developmental stages, close to maturity (30 DAP), appearing only slightly diluted by the progressive accumulation of starch toward the end of seed maturation (Fig. 1A).

Figure 1.

Figure 1.

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Accumulation of phytase and seed storage proteins during seed maturation. Total soluble protein was extracted from transgenic seeds at various developmental stages (10–30 DAP) at a fixed w/v ratio, and 10 μL were loaded per lane. Immunoblots were incubated with antisera against phytase (A), γ zein (B), CAG (C), and CL-1 (D).

Very similarly, in both transgenic and wild-type plants, the relative amount of three endogenous storage proteins (γ-zein, CAG, and CL-1) accumulated progressively during grain development and peaked at approximately 25 DAP before slightly declining at later stages (Fig. 1, B–D) due to dilution by the progressive accumulation of starch. Thus, the accumulation rate of the recombinant phytase closely resembles that of endogenous storage proteins, demonstrating a high stability of the protein.

The N-Glycan Structure of Phytase Changes during Endosperm Maturation

To monitor changes in the glycosylation pattern of phytase during development we analyzed the N-glycan profile of the protein isolated from maize endosperm at different stages of maturity. Since it was not possible to obtain sufficient amounts of the protein from seeds younger than 20 DAP, we compared the N-glycan structures of phytase isolated from maize endosperm at midmaturation stage (20 DAP), seeds close to maturity (30 DAP), and mature, dry seeds. Protein samples were separated by SDS-PAGE, and the phytase band was excised and digested with trypsin. Glycopeptide analysis allowed us to follow up specific glycosylation sites through different developmental stages. The results for the glycopeptides TYNYS LGADD LTPFG EQELV NSGIK and YSALIEEIQQNATTFDGK are shown in Figure 2 and Supplemental Figures S1 and S2. The absence of unglycosylated peptides at all stages confirms the efficiency of N-glycosylation. Figure 2 shows that the glycan patterns of different peptides are not identical, most likely due to different accessibility of the individual glycosylation sites, but the major changes in abundance of vacuolar glycoforms and trimmed GlcNAc structures during development are very similar for both glycopeptides. At 20 DAP, we identified large amounts of glycopeptide species containing the N-glycans Man3XylFucGlcNAc2 (Fig. 2, A and B) and Man3XylGlcNAc2 (Fig. 2B). These structures, also known as MMXF and MMX (Altmann, 2007), are typically found on vacuolar glycoproteins (Lerouge et al., 1998). Minor peaks corresponded to the peptide containing trimmed glycans, some of which contained Fuc on the core GlcNAc, representing degradation products of Golgi-modified complex glycans. At 30 DAP the relative amount of the formerly predominant glycan MMXF had decreased dramatically, and the predominant glycopeptide species had a molecular mass of 2,934.4 amu or 2,231 amu, respectively, indicating the presence of trimmed structures consisting of core GlcNAcs alone. Single GlcNAcs have been found previously on a KDEL-tagged recombinant protein from maize (Rademacher et al., 2008). After 30 DAP the N-glycan profile did not change significantly as shown with phytase isolated from fully mature seeds (40 DAP; Fig. 2).

Figure 2.

Figure 2.

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N-glycan structures attached to recombinant phytase glycopeptides TYNYS LGADD LTPFG EQELV NSGIK (A) and YSALIEEIQQNATTFDGK (B), derived from developing maize endosperm at 20, 30, and 40 DAP. The vacuolar-type glycans MMXF and MMX were major N-glycan structures in phytase derived from young seeds (20 DAP), whereas small structures consisting of only one or two core GlcNAc residues dominated in phytase isolated from more mature seeds (30 and 40 DAP). See http://www.proglycan.com for an explanation of N-glycan structure abbreviations.

Ultrastructural Changes in Maize Seeds during Development

Changes in the N-glycan profile of phytase during development suggested that the subcellular localization of the protein might also change as seed matured. We therefore investigated the subcellular organization of maize endosperm cells during development to monitor any changes in the different protein storage compartments. Based on the ultrastructural characteristics of the cells in the different stages studied, we defined three representative developmental stages, in which major changes in subcellular organization were observed: stage 1 (young seeds, around 10 DAP), stage 2 (midmaturation stage, seeds around 20 DAP), and stage 3 (seeds close to maturity, around 30 DAP).

Stage 1

In young seeds, there is no clear difference between the aleurone and the subaleurone layers. Thus, there is an outermost layer of cubical aleurone cells, with a high content in aleurone grains. Layers 2 to 4 could be considered as transition cells, as there is a progressive loss of aleurone grains, accompanied by a gain in starch grains of increasing size. Storage protein synthesis is in progress and zein bodies can be found from layer 5 onwards. Several PSV-like organelles, approximately spherical and similar in size to starch grains (approximately 2 μm), can be identified in the same layers (Fig. 3A).

Figure 3.

Figure 3.

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Development of maize seeds. Sections after embedding in Spurr's resin. A to C, Light microscopy, toluidine blue: stage 1 (A), stage 2 (B), and stage 3 (C). Aleurone (Al) and endosperm (En). Cells accumulate starch (s), some storage vacuole-like compartments are already seen (arrowheads), smaller refringent vacuolar compartments are also observed (B, arrow), zein bodies appear as small blue stained accretions, few nuclei (n) are still present. D and E, Electron microscopy, general nonspecific contrast: stage 2. Zein bodies (zb) and PSV. Note the globulin-like inclusion in the vacuole-like compartments (*). rER, Rough ER. Bars 20 μm (A–C), 0.5 μm (D and E). [See online article for color version of this figure.]

Stage 2

The aleurone layer presents its characteristic appearance of polyhedral cells with thick cell walls, clearly distinct from the rest of the endosperm. The starch content of the endosperm has increased significantly and also the number and size of the protein bodies (Fig. 3B). Although already observed in younger cells (Fig. 3A), vacuolar storage compartments become more prominent at this stage and they are more abundant and have reached a diameter of approximately 4 to 5 μm (Fig. 3, B, D, and E). Some small vacuole-like compartments, similar in size to the zein bodies can also be observed within the endosperm cells (Fig. 3B). In the electron microscope, vacuolar storage compartments in OsO4-fixed samples appear to contain a globulin-like inclusion, which is highly electron dense (Fig. 3, D and E). The electron-dense inclusion in the PSV-like structures consists of CAG (Woo et al., 2001) and probably CL-1 (Yamagata et al., 2003).

Stage 3

Endosperm cells are packed with starch and proteins, and the number of visible or identifiable PSVs has apparently decreased (Fig. 3C). The average size of the zein bodies has increased above 1 μm and that of the starch grains up to approximately 8 μm. In later developmental stages zein bodies and starch grains enlarge even further, but no significant morphological changes can be observed. The mature endosperm cells are tightly packed with enlarged zein bodies that fill the spaces between the starch grains (data not shown).

The Subcellular Localization of Phytase Changes during Seed Development

We determined the localization of recombinant phytase in sections of endosperm cells at the different developmental stages described above. The ultrastructural changes observed during development were reflected in the deposition of the recombinant protein as the distribution of phytase changed along maturation (Fig. 4, A–C).

Figure 4.

Figure 4.

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Localization of recombinant phytase. Sections after embedding in LRWhite resin. A to C, Fluorescence microscopy. D to F, Electron microscopy. Comparison of the distribution pattern of recombinant phytase between endosperm cells in stage 1 (A), stage 2 (B), and stage 3 (C). Note the storage vacuole-like compartments in each of the developmental stages (arrows) and the nonlabeled spherical zein bodies evenly spread within the cytoplasm (B and C). No significant labeling in the apoplast (arrowheads) or the starch granules (s). D, Stage 1. Note the heavy labeling on a small PSV. E, Stage 3. Gold particles decorating the periphery of a zein body (zb). F, Stage 3. Double labeling, α-zein (15 nm), and phytase (10 nm). Presence of recombinant phytase on the ER and also on the periphery of the protein body. Note that α-zein and phytase do not colocalize. rER, Rough ER. Bars 20 μm (A–C), 0.5 μm (D–H). [See online article for color version of this figure.]

Stage 1

Young endosperm cells showed strong labeling in PSV-like structures. Consistent with the ultrastructural observations, labeled putative PSVs in these cells were small (approximately 2 μm). In some PSV-like structures the signal was equally distributed, others showed labeled phytase deposits at the periphery only (Fig. 4, A and D). There was no significant labeling within the spherical zein bodies, but some signal was visible in the cytoplasm, probably reflecting the ongoing trafficking of recombinant phytase through the endomembrane system. No significant labeling was detected in the starch granules or in the apoplast (Fig. 4A).

Stage 2

Labeling was still concentrated in the putative PSVs in middevelopment endosperm cells (Fig. 4B). This localization is in agreement with the main N-glycan structure attached to phytase at this stage (MMXF), which is typically found on vacuolar glycoproteins (Lerouge et al., 1998). The labeled PSV-like structures had increased considerably both in size (approximately 5 μm) and in number, as described in the previous section for this developmental stage (Fig. 4B).

As in stage 1, recombinant phytase was detected almost exclusively in PSV-like organelles, to the exclusion of the clearly distinct zein bodies (Fig. 5, A–C).

Figure 5.

Figure 5.

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Characterization of the storage compartments. Sections after embedding in LRWhite resin. A to D, Stage 2. F and G, Fluorescence microscopy. E, Light microscopy. Lugol's iodine staining. A, Localization of phytase. s, Starch. B, Localization of α-zein. C, Merged. Note that phytase accumulates in storage vacuole-like compartments (arrows), but there is no colocalization with α-zein. D, Localization of phytase. Strong labeling in PSV-like organelles (arrows). E, Lugol's iodine staining of the same cell, vacuole-like compartments do not stain (arrows). s, Purple-stained starch grains. F, Detection of post-Golgi glycan modifications with bee venom anti-serum. Significant labeling in the storage vacuoles (arrows). s, Starch. G, Control: no signal after competition with bromelain. Bars 20 μm.

Stage 3

A striking change in the localization of recombinant phytase was observed in older endosperm cells, consistent with the shift from MMXF to GlcNAc glycans that was also observed in seeds close to maturation. Now only a few vacuolar compartments could be clearly identified, and these residual PSVs were still labeled (Fig. 4C), but a strong signal was also observed around the periphery of the zein bodies (Fig. 4, E and F). Moreover, significant labeling in the ER was also observed (Fig. 4F).

To investigate if the reduced Golgi-mediated transport that was observed for recombinant phytase during late seed development is due to a reduced availability of Golgi organelles we used a marker antibody to visualize and fluorescently label Golgi stacks in vibratome sections of developing endosperm (Horsley et al., 1993). Using confocal microscopy we determined the number of Golgi compartments per cell (Supplemental Fig. S3) and observed a 4-fold increase between stage 1 and stage 3.

Proteins in the PSV Contain Golgi-Modified N-Glycans

By developmental stage 2, the PSVs are similar in size to starch grains, but can be distinguished because they do not stain with Lugol's iodine (Fig. 5, D and E). The glycan composition of glycoproteins within the PSVs was confirmed by the strong labeling observed using an antiserum specific for fucosylated N-glycans (Fig. 5F). The labeling could be outcompeted by the addition of bromelain, an unrelated plant glycoprotein with complex paucimanosidic N-glycans, thus confirming the specificity of the signal (Fig. 5, F and G). This confirms that the putative PSVs contain glycoproteins that have undergone Golgi-specific modifications. PSVs reacting with the antiserum specific for fucosylated N-glycans are also present in wild-type seeds (data not shown), indicating that endogenous glycoproteins accumulate in a similar fashion.

Endogenous Maize Globulins Show Similar Changes in Subcellular Localization during Seed Development

To determine whether the developmental profile of phytase was mirrored by endogenous storage globulins, we carried out localization experiments as described above, this time using CL-1 and CAG. At stage 2 the distribution of both CAG and CL-1 is similar to that of phytase in this developmental stage, and both globulins could be found within the PSVs (Fig. 6, A and C). At stage 3 the shift in the localization of phytase is also observed for CL-1 and CAG, which are now found at the periphery of the protein bodies, forming a ring around the zein inclusion (Fig. 6, B and C).

Figure 6.

Figure 6.

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Deposition of maize globulins. Sections after embedding in LRWhite resin. Electron microscopy. A and B, Localization of CL-1. C and D, Localization of CAG. A and C, Stage 2. B and D, Stage 3. A and C, Significant labeling for CL-1 and CAG in the PSV. B to D, Note labeling for both CL-1 and CAG, now distributed around the periphery of the zein bodies (zb; arrows). Bars 0.25 μm.

DISCUSSION

Previous studies have suggested that the intracellular trafficking and distribution of storage proteins may change during seed development (Levanony et al., 1992; Shy et al., 2001; Vitale and Hinz, 2005). To investigate this phenomenon in maize endosperm, we tracked the recombinant model glycoprotein phytase and two endogenous storage globulins (CAG and CL-1) through the development of the maize seed using a combination of microscopy and N-glycan analysis. In all three cases, we observed a major developmental switch from post-Golgi PSV-like organelles to ER-derived zein bodies, confirming that seeds alter protein trafficking and accumulation as part of the developmental program.

PSV-like structures appear to be the major storage sites in young endosperm cells for all three proteins we investigated. Storage vacuoles have been the focus of numerous recent studies looking at structure, classification, and protein trafficking but most of these studies have concerned dicotyledonous plants (Wenzel et al., 2005; Hinz et al., 2007; Craddock et al., 2008). In cereals, studies of storage protein accumulation have focused mainly on the prolamins, and this is particularly the case in maize where the vast majority of storage protein is made up of prolamins (zeins) that accumulate in ER-derived zein bodies. In contrast, only small amounts of globulins are stored in maize endosperm, and only two previous studies have addressed their spatiotemporal expression and localization (Woo et al., 2001; Yamagata et al., 2003).

We have confirmed that both CAG and CL-1 accumulate in PSV-like compartments in early endosperm development. Woo et al. (2001) reported that the putative PSVs increased in size in maturing endosperm cells, eventually swelling to three times the size of zein bodies. Labeling with an antibody against CAG was less dense in the larger compartments, leading the authors to assume that CAG was being diluted and was therefore more difficult to detect. Since phytase is an easily detectable reporter protein, the labeling of phytase greatly facilitated the visualization of even large PSV-like organelles. Thus, we were able to show that PSVs at developmental stage 2 varied greatly in size, although they were on average much larger than those at earlier stages, and that they contained glycan structures with trans-Golgi-specific modifications, clearly identifying them as post-Golgi compartments. Since both CAG and CL-1 lack N-glycan chains, the fact that vacuolar N-glycans were also detected in the PSVs of stage-matched wild-type seeds indicates that these organelles must contain endogenous glycoproteins that remain to be identified. Indeed, the analysis of total soluble proteins from wild-type maize endosperm revealed the presence of glycoproteins with complex paucimannosidic N-glycans (data not shown), and those are probably residents of the PSV-like organelles.

In immature cereal endosperm cells PSVs appear to be a preferred accumulation site for recombinant proteins bearing an N-terminal signal peptide for entry into the endomembrane system, even though such proteins are normally secreted from vegetative tissues. This phenomenon has been reported for phytase expressed in rice and wheat (Arcalis et al., 2004; Drakakaki et al., 2006), and for a secretory antibody and lysozyme expressed in rice (Yang et al., 2003; Nicholson et al., 2005).

We found that in maize the number of identifiable PSVs decrease relative to the other organelles later in endosperm development, concomitant with a progressive change in the subcellular localization of recombinant phytase and both endogenous globulins. At this stage, a very clear label became visible around the periphery of the zein bodies in transgenic plants expressing phytase. This raises the question as to whether or not an interaction takes place between phytase and γ zein, which is also located in the outer part of the mature zein bodies (Lending and Larkins, 1989). However, phytase is localized exclusively around the periphery of the protein bodies, whereas deposits of γ zein are sometimes also found in the inner areas, suggesting that the proteins do not colocalize (Fig. 4E).

It was conceivable that the developmental change in the subcellular localization of recombinant phytase and the two native storage proteins could perhaps be related to the induction of an unfolded protein response, triggered by the recombinant protein. However, in our transgenic plants, the levels of the chaperone BiP were not significantly increased as compared to wild-type levels (data not shown). In addition, although an influence of BiP cannot be ruled out, the fact that CAG and CL-1 accumulated in the same compartments with the same developmental profile in wild-type plants and transgenics suggests that its role is not specific in transgenic plants.

Both of the endogenous globulins lack N-glycan acceptor sites so although their final destination can be determined it is more difficult to follow their pathway through the cell. In contrast, the N-glycan acceptor sites on phytase allow the route taken by this protein to be determined by examining the N-glycan structures. The developmental switch from PSV to zein body as preferred accumulation site for phytase is mirrored by a corresponding change in N-glycan structures. Initially, the glycan residues were indicative of vacuolar localization, consistent with immunodetection in the PSV-like organelles. Later in development and simultaneous with the changing subcellular localization, the relative abundance of glycoforms with complex modifications including β(1,2)-Xyl and α(1,3)-linked core Fuc decreased while small glycans consisting only of one or two GlcNAc residues without Fuc became dominant. There is some evidence for the time-dependent trafficking of storage proteins in wheat, where the amount of transcripts for Golgi-associated proteins declines with seed maturation. Therefore, the role of Golgi vesicles in the trafficking of prolamins in wheat appears mainly restricted to the early stages of seed development (Parker, 1982; Galili et al., 1993; Shy et al., 2001). In maize we observed that the number of Golgi organelles per cell increases 4-fold between stage 1 and 3. Although the volume of the cells and the synthesis of storage proteins increase markedly at the same time, this result does not indicate a striking decline of Golgi activity during maize endosperm development. Nevertheless, we observed a time-dependent protein trafficking in maize resulting in two pools of phytase, an early pool that reflects trafficking through the ER and Golgi vesicles to the PSV, and a later pool accumulating in the ER and in ER-derived compartments. This is consistent with the changing N-glycan profiles of recombinant phytase and with the occurrence of trimmed glycoforms. We have previously observed the same N-glycan structures attached to a KDEL-tagged recombinant antibody, which accumulated in ER-derived zein bodies (Rademacher et al., 2008). Interestingly, recombinant human lactoferrin isolated from mature maize seeds was reported to contain almost exclusively (98%) pauci-Man-type N-glycans with β(1,2)-Xyl and α(1,3)-linked core Fuc (Samyn-Petit et al., 2001). However, it is important to note that in this study, as in most previous reports, glycans were enzymatically released from the protein prior to analysis, and this method would not have identified small glycan structures, such as single or double GlcNAcs. Thus, it is possible that the described vacuolar glycan pattern characterizes only one part of the lactoferrin, whereas a second part with single GlcNac structures would not have been noticed. In this study we also identified GlcNAcs with core Fuc residues on a small proportion of recombinant phytase molecules isolated from endosperm cells, apparently representing the result of glycan degradation in post-Golgi compartments. The trimmed glycan structures clearly indicate the presence of various glycanase activities in maize endosperm. Endoglycanase (ENGase and PNGase) activities in cereal seeds have been reported previously, but were thought to be mainly derived from the embryo (Chang et al., 2000; Vuylsteker et al., 2000).

In conclusion, the use of a model glycoprotein (in addition to two endogenous globulins) allowed us to visualize and characterize the vacuolar compartments in maize endosperm, and follow the developmental changes in protein trafficking and distribution. In contrast to earlier reports, we have shown that the vacuolar compartments are significant storage compartments in developing maize endosperm, particularly at early developmental stages. A combination of microscopy and N-glycan analysis provided evidence that protein trafficking in maize is time dependent, as previously suggested in wheat. Further experiments are under way to determine the fate of the PSVs in later developmental stages and the implications of our findings on the production of recombinant pharmaceutical proteins in cereal grains.

MATERIALS AND METHODS

Transgenic Plants

The transgenic maize (Zea mays) plants have been described previously (Drakakaki et al., 2005) and contained the construct pLPL-phyA (including the Aspergillus niger phyA gene preceded by the N-terminal signal peptide from the murine immunoglobulin κ chain) under the control of the rice (Oryza sativa) glutelin-1 promoter. Fourth generation plants (T4) homozygous for the transgene were maintained in a controlled growth room at 28°C/20°C day/night temperature with a 16-h photoperiod and 60% to 90% relative humidity. Nontransgenic control plants were regenerated from the same batch of callus material as used for transformation (Drakakaki et al., 2005), and propagated and grown under the same conditions as the transgenic lines. Developing ears were harvested at 10, 15, 20, 25, 30, and 40 DAP.

Immunoblot Analysis

Total soluble protein was extracted from seeds as described (Arcalis et al., 2004) using phosphate-buffered saline (PBS) containing 10 mm ascorbic acid, 500 mm NaCl, and 5% β-mercaptoethanol. After centrifugation at 17,500g, 4°C, the supernatants were boiled for 10 min and separated by 12% (w/v) SDS-PAGE under reducing conditions. Immunoblot analysis was carried out according to standard protocols. For each time point, five kernels from the same ear were pooled for each sample.

Protein Extraction and Liquid Chromatography-Mass Spectrometry Analysis

Proteins were extracted from ground seeds with PBS containing 10 mm ascorbic acid, 500 mm NaCl, and 5% β-mercaptoethanol to minimize enzymatic activities post extraction. Phytase was isolated from the extracts by sequential ammonium sulfate fractionation as described by Arcalis et al. (2004) and concentrated by ultrafiltration before separation by 12% (w/v) SDS-PAGE under reducing conditions. The bands corresponding to phytase were excised, destained, carbamidomethylated, digested with trypsin, and extracted from gel pieces as described (Kolarich and Altmann, 2000; Kolarich et al., 2006). Peptide fractionation and analysis using a Q-TOF Ultima Global (Waters Micromass) mass spectrometer was performed as described previously (Kolarich and Altmann, 2000; Van Droogenbroeck et al., 2007).

The mass spectrometry data from the tryptic peptides were compared to datasets generated by in silico tryptic digestion of A. niger phytase using the PeptideMass program (http://www.expasy.org/tools/ peptide-mass.html). Tryptic glycopeptide datasets were generated by the addition of glycan mass increments to the masses of the potential glycopeptides.

Light and Electron Microscopy

A minimum of 10 phyA-containing grains from three different ears were examined for each developmental stage. Seeds were bisected longitudinally and the embryo was removed. One half of the endosperm was processed for recombinant protein analysis by immunoblot. Thin slices were cut from the remaining half with a razor blade under phosphate buffer (0.1 m, pH 7.4). Tissue pieces were fixed in 2% (w/v) paraformaldehyde and 2.5% (v/v) glutaraldehyde in phosphate buffer (0.1 m, pH 7.4) for ultrastructural analysis or in 4% (w/v) paraformaldehyde and 0.5% (v/v) glutaraldehyde in phosphate buffer (0.1 m, pH 7.4) for immunolocalization studies and then processed as described previously (Arcalis et al., 2004). For light microscopy, 1-μm sections were stained in toluidine blue. Starch was stained by incubating 1-μm sections with Lugol's iodine solution for 5 min. For electron microscopy, sections showing silver interference colors were stained in 2% (w/v) aqueous uranyl acetate. The sections were observed using a Philips EM-400 transmission electron microscope.

Sections mounted either on glass slides for fluorescence microscopy or on gold grids for electron microscopy were preincubated in 5% (w/v) bovine serum albumin (BSA; fraction V) in phosphate buffer (0.1 m, pH 7.4) and then incubated with the appropriate dilution of polyclonal rabbit anti-phytase, anti-CL-1, and anti-CAG or monoclonal rat anti-α-zein. Sections were then treated with the secondary antibody diluted in phosphate buffer (0.1 m, pH 7.4). This was a goat anti-rabbit IgG Alexa Fluor 594 or goat anti-rat IgG Alexa Fluor 488 for fluorescence microscopy and a goat anti-rabbit IgG 10-nm gold and goat anti-rat IgG labeled with 15-nm gold particles for electron microscopy.

Immunofluorescence Analysis and Confocal Microscopy

Seeds of wild-type maize (10 and 25 DAP) were fixed in 2% (w/v) paraformaldehyde for 12 to 72 h. After washing with 0.1 m phosphate buffer (pH 7.4), 60 to 100 μm vibratome sections were prepared and placed on glass slides coated with 0.1% (w/v) poly-Lys (Sigma), respectively. Sections were dehydrated by an ethanol series and equilibrated in 0.1 m phosphate buffer. The cell wall was digested with 2% (w/v) cellulase (Onozuka R-10 from Trichoderma viride) in phosphate buffer (0.1 m, pH 7.4) for 1 h at room temperature. Following a treatment with 0.5% Triton X-100 in 0.1 m phosphate buffer for 1 h at room temperature, nonspecific binding sites were blocked with 3% (w/v) BSA (fraction V) in PBS for 10 min. Sections were incubated with monoclonal antibody JIM84 diluted 1:50 in phosphate buffer (0.1 m, pH 7.4) overnight at 4°C (Horsley et al., 1993). Antibody binding was visualized by Cy3 conjugated goat anti-rabbit IgM, μ-chain-specific antibody (Jackson Immunoresearch). The sections were mounted in 50% glycerol in phosphate buffer (0.1 m, pH 7.4) and observed in the confocal laser scanning microscope (TCS-SP2, Leica) via the Leica confocal software (version 2.61) and long-distance 40×, 63× water-immersion objectives. Excitation wavelengths were 488 nm (argon laser) for Cy3. Emission was detected between 551 and 616 nm. Z sections were collected and overlay pictures were generated using ImageJ. The Golgi compartments were counted per cell, excluding overlap areas between two adjoining cells. A minimum of five cells were counted for each sample.

Bromelain Competition

Polyclonal rabbit anti-bee venom serum, which recognizes core-linked α(1-3)-Fuc (Prenner et al., 1992), was diluted (1:100) in phosphate buffer (0.1 m, pH 7.4) with or without 10% (w/v) bromelain (preheated to destroy enzymatic activity) and incubated at room temperature for 30 min. Sections were preincubated in 5% (w/v) BSA (fraction V) and then incubated with the anti-bee venom antiserum, which binds to core Fuc glycan residues. Samples were then treated with the secondary antibody (goat anti-rabbit Alexa Fluor 594) diluted in phosphate buffer (0.1 m, pH 7.4).

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Figure S1. Deconvoluted liquid chromatography-mass spectrometry spectrum of the glycopeptide TYNYS LGADD LTPFG EQELV NSGIK.

  • Supplemental Figure S2. Deconvoluted liquid chromatography-mass spectrometry spectra of the glycopeptide YSALIEEIQQNATTFDGK.

  • Supplemental Figure S3. Golgi bodies visualized in endosperm cells at different stages.

Supplementary Material

[Supplemental Data]
pp.109.152363_index.html (707B, html)

Acknowledgments

The authors would like to thank Dr. Richard M. Twyman for critical assessment and help with manuscript preparation and Dr. Rudolf Jung for helpful discussions and for providing antibodies against corn legumin, CAG, and α-zein. We would also like to thank Dr. A. Ohmann for providing antibodies against phytase, and the Pathology Department University Hospital of the Rheinisch-Westfälische Technische Hochschule Aachen for allowing us to use their microscopy facilities.

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Associated Data

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Supplementary Materials

[Supplemental Data]
pp.109.152363_index.html (707B, html)
pp.109.152363_1.pdf (713.1KB, pdf)

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