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(American Journal of Botany. 2008;95:1049-1062.) doi: 10.3732/ajb.2007404 © 2008 Botanical Society of America, Inc. |
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Developmental Biology and Developmental Genetics |
2 Technische Universität München, Lehrstuhl für Botanik, Biologikum-Weihenstephan, Am Hochanger 4, D-85350 Freising, Germany 3 Ohio University, Department of Chemistry and Biochemistry, Athens, Ohio 45701 USA 4 Max-Plank-Institut für Biochemie, Am Klopferspitz, D-82152 Martinsried, Germany
Received for publication 10 December 2007. Accepted for publication 10 June 2008.
ABSTRACT
KDEL-tailed cysteine endopeptidases are a group of papain-type peptidases found in senescing tissue undergoing programmed cell death (PCD). Their genes have so far been cloned and analyzed in 12 angiosperms. They are synthesized as proenzymes with a C-terminal KDEL endoplasmatic reticulum retention signal, which is removed with the prosequence to activate enzyme activity. We previously identified three genes for KDEL-tailed cysteine endopeptidases (AtCEP1, AtCEP2, AtCEP3) in Arabidopsis thaliana. Transgenic plants of A. thaliana expressing β-glucuronidase (GUS) under the control of the promoters for the three genes were produced and analyzed histochemically. GUS activity was promoter- and tissue-specific GUS activity during seedling, flower, and root development, especially in tissues that collapse during final stages of PCD, and in the course of lateral root formation. KDEL-tailed cysteine endopeptidases are unique in being able to digest the extensins that form the basic scaffold for cell wall formation. The broad substrate specificity is due to the structure of the active site cleft of the KDEL-tailed cysteine endopeptidase that accepts a wide variety of amino acids, including proline and glycosylated hydroxyproline of the hydroxyproline rich glycoproteins of the cell wall.
Key Words: Arabidopsis thaliana • Brassicaceae • cell wall degradation • development in generative and vegetative tissues • Euphorbiaceae • β-glucuronidase (GUS) • KDEL-tailed cysteine endopeptidases • programmed cell death • ricinosome • Ricinus communis
Programmed cell death (PCD) is a highly regulated, genetically determined process in all multicellular organisms. In plants, PCD occurs during development, under stress conditions, during senescence, and in response to pathogen infection. Studies of PCD in the form of apoptosis in animals have identified a central role for a conserved family of cysteine proteases (caspases), which cleave at an aspartate residue at the P1 position of their substrate recognition site (Salvesen and Boatright, 2004
). Apoptosis in animals is characterized by nuclear condensation, cellular fragmentation into apoptotic bodies, formation of nucleosomal fragments, nicking of DNA as demonstrated by the TUNEL assay (terminal deoxynucleotidyltransferase-mediated dUTP nick end labeling). In the end stage of apoptosis, the dying cells are engulfed by neighboring cells (Cuervo, 2004). Apoptosis in animal cells differs from PCD in plants by the involvement of phagocytosis, a process not present in plants.
PCD in plants is effected by a unique group of papain-type cysteine endopeptidases with a C-terminal KDEL endoplasmic reticulum (ER) retention signal (Fig. 1; Gietl et al., 2000; Gietl and Schmid, 2001; Beers et al., 2004). The plant species from which genes for KDEL-tailed cysteine endopeptidases have been cloned and sequenced are listed in Fig. 1. The high homology of the deduced amino acid sequences is obvious. The N-terminal signal peptide that transfers the precursor protein into the lumen of the ER is, as generally observed, more variable. The phylogenetic tree discloses distinct groups among KDEL-tailed cysteine endopeptidases between dicots and monocots (Gietl et al., 2000; Gietl and Schmid, 2001; Beers et al., 2004). These enzymes have no structural relationship to the caspases and homologous genes have not been found in mammals or yeast. The biological features and the organ-specific expressions of these peptidases in different plant species have been reviewed (Gietl and Schmid, 2001). The presence of a rigid cell wall and absence of phagocytosis in plants poses a distinct problem regarding how the cell wall is degraded and how the degradation products of the cellular contents are transferred into neighboring cells.
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, 1999). The blebbing of ricinosomes from the ER takes place after the storage proteins have been mobilized and the resulting amino acids and peptides transferred to the germinating seedling. The final stage of PCD in the degenerating endopserm is characterized by destruction of the nucleus, disruption of the large vacuole releasing nucleases and proteases, acidification of the cytoplasm and disruption of the ricinosomes, which release CysEP. The N-terminal propeptide and the C-terminal KDEL are cleaved off, and the activated CysEP degrades the cytosolic macromolecules (Schmid et al., 1998, 1999, 2001). KDEL-tailed cysteine endopeptidases have been identified in ricinosomes by immunoelectron microscopy in the nucellus undergoing PCD in maturing seeds of Ricinus (Greenwood et al., 2005) and in senescing flower petals of Hemerocallis (Schmid et al., 1999). Destruction of the chromosomes is evidenced by positive TUNEL assays and collapse of cellular components. Digestion products are presumed to be taken up by the adjacent developing endosperm cells, which are transfer cells exhibiting a special morphology with extensive cell wall folds. The endosperm cell walls of germinating seeds as well as the nucellus cell walls of maturing seeds remain as crushed layers (Schmid et al., 1999; Greenwood et al., 2005). This state can be tolerated because there is no need for recycling carbon in a photosynthetic plant.
In Arabidopsis thaliana, we have identified three KDEL-tailed cysteine endopeptidases with homology to the castor bean CysEP, designated AtCEP1 (At5g50260), AtCEP2 (At3g48340) and AtCEP3 (At3g48350) (Fig. 1). Because KDEL-tailed cysteine endopeptidases from Arabidopsis cannot be isolated in large amounts, we purified CysEP from the ricinosomes of Ricinus, crystallized it, and determined the structure by molecular replacement at a 2.0-Å resolution (Than et al., 2004). CysEP prefers substrates with a basic amino acid in the P1 position (position –1 relative to the cleavage site) and neutral amino acids with large aliphatic and nonpolar or aromatic side chains in the P2 position (position –2 relative to the cleavage site). The open pocket of the Ricinus CysEP correlates with the extended variety of substrate amino acid residues accommodated by this enzyme, including proline at the P1 and P1' positions (positions –1 and +1 relative to the cleavage site). This more open pocket may allow the enzyme to cleave a greater variety of proteins during PCD (Than et al., 2004).
It is obvious that KDEL-tailed cysteine endopeptidases (Fig. 1) are found in tissues undergoing PCD, especially in cells that finally collapse, such as the hypogeous cotyledons of Vicia sativa (Becker et al., 1997), the maturing pods of Phaseolus vulgaris (Tanaka et al., 1991), the unpollinated ovaries of Pisum sativum (Cercos et al., 1999), the outer integument developing into the seed coat of Phalaenopsis (Nadeau et al., 1996), the senescing flower petals of Hemerocallis (Valpuesta at al., 1995) and Sandersonia aurantiaca (O’Donoghue et al., 2002), the megagametophyte cells after germination of Picea glauca seeds (He and Kermode, 2003), and the epigeous cotyledons of Vigna mungo (Toyooka et al., 2000). Furthermore, castor bean CysEP as the prototype for this subgroup of papain-type cysteine endopeptidases accepts proline near the cleavage site, which is highly unusual among endopeptidases (Cunningham and O’Connor, 1997; Simpson, 2001). At the final stages of PCD, the cell walls generally collapse into crushed layers, which requires the structural organization of the walls to have undergone changes. This observation led us to the hypothesis that KDEL-tailed cysteine endopeptidases participate in the final cell collapse during PCD by attacking the structural hydroxyproline-rich glycoproteins of the cell wall.
While the biosynthesis of cell walls in plants is increasingly understood in biochemical and structural terms, wall degradation during elimination of cells and restructuring during intercalation of additional cells is little understood. The plant cell wall comprises three interpenetrating networks, two of structural polysaccharides and one of structural glycoproteins (Lamport, 1966). Each network is internally crosslinked: cellulose microfibrils by xyloglucans, pectins by calcium salt bridges and apiosyl borate esters, and structural hydroxyproline (Hyp)-rich glycoproteins (HRGPs), notably the extensins, via tyrosine (Epstein and Lamport, 1984). Extensins encoded by a multigene family are best studied in the case of the HRGPs in Arabidopsis and Nicotiana tabacum. Self-assembly of the plant cell wall requires an extensin scaffold (Cannon et al., 2008). This scaffold is formed by crosslinking peptide modules extensively O-glycosylated at the Hyp residues. The patterns of Hyp-O-glycosylation are coded by the primary amino acid sequence, as described by the Hyp contiguity hypothesis, which predicts that contiguous Hyp residues be the attachment sites of small arabino-oligosaccharides (1–5 Ara residues/Hyp), while clustered, noncontiguous Hyp residues are the preferred sites for the addition of arabinogalactan polysaccharides. In addition, monogalactose residues are bound to Ser in Ser-(Hyp)4 repeats (Shpak et al., 1999; Kieliszewski and Shpak, 2001). Three major types of monomeric extensin precursors are widespread in dicots: the P1-type (precursor 1) extensins characterized by the repetitive Ser-(Hyp)4-Thr-Hyp-Val-Tyr-Lys motif; the P2-type (precursor 2) extensins containing repeats of the Ser-(Hyp)4-Val-Tyr-Lys-Tyr-Lys motif; and the P3 (precursor 3)-type extensins containing variations of the Ser-(Hyp)4-Ser-Hyp-Ser-(Hyp)4-Tyr-Tyr-Tyr-Lys repeat. In addition to intermolecular crosslinking, both P2 and P3 type extensins undergo intramolecular crosslinking of Tyr residues within the cross linking module Tyr-Xxx-Tyr (underlined in the previous sequences) (Held et al., 2004). Thus, P1-type extensins differ from P2- and P3-type extensins in that they offer possible cleavage sites for a KDEL-tailed cysteine endopeptidase such as the Ricinus CysEP within the Thr-Hyp-Val-Tyr-Lys motif or the Lys-Lys-Pro-Tyr-Tyr-Pro-Hyp-His-Thr-Hyp-Val-Tyr-Lys-sequence C-terminal to the Ser-(Hyp)4-motif (Fig. 2). P2- and P3-type extensins, on the other hand, offer no cleavable amino acid stretches for a "classical" protease and might hinder an attack by proteases as a result of the intramolecular cross-linking of Tyr residues (Fig. 3).
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The availability of the KDEL-tailed cysteine endopeptidase from Ricinus communis and extensin substrates from Nicotiana tabacum enabled us to demonstrate that this enzyme cleaved the P1-type extensin not only at the "classical" protease cleavage sites but also at the C-terminus of the Ser-(Hyp)4-Lys module and at the N-terminus of the Lys-Ser-(Hyp)4 module. P3-type extensin, on the other hand, was completely resistant to the enzyme.
MATERIALS AND METHODS
Tissue-specific expression of AtCEP1, AtCEP2, and AtCEP3
Total RNAs from roots, leaves, stems, cauline leaves, buds, and green siliques of 3–6-wk-old Arabidopsis wild-type plants were isolated using the RNeasy-Kit (Quiagen, Hilden, Germany) and assayed by RT-PCR (Promega, Mannheim, Germany) with the following specific pairs of primers for the presence of AtCep1, AtCep2, or AtCep3 mRNAs. Actin 1 (ACT1; accession M20016) was used as a positive control. The primers were as follows: AtCEP1 sense (beginning of 1. exon), 5' TAT ACG AAC GGT GGA GGA GTC ACC 3'; AtCEP1 antisense (beginning of 3. exon), 5' TCC GCA TCT CCC GGT AAA CAC TCC 3'; AtCEP2 sense (end of 1. exon), 5' GTG CTG TCA CTG AAA TCA AGA ATC AAG G 3'; AtCEP2 antisense (beginning of 3. exon), 5' GTT CAG CTC TGT TCC ACA AGA TCC CG 3'; AtCEP3 sense (end of 1. exon), 5' TTC TGT TGA TTG GCG AGA GAA AGG AGC 3'; AthCEP3 antisense (beginning of 3. exon), 5' TCC ATA CCC AAC AAT CAC CAC CCC G 3'.
Primers for Actin1 (chromosome 2) were as follows: ACT1-sense (end of 2. exon), 5' CCT GCT ATG TAT GTG GCT ATT CAG GC 3'; ACT1-antisense (within 4. exon), 5' TCG TCA TACT CT GCC TTTGCG ATC 3'.
RT-PCR products were separated by standard 1% agarose gel electrophoresis.
For sequence and position of CEP primers, see Appendix S1 (see Supplemental Data with the online version of this article).
CEP1, CEP2, and CEP3 promoter::glucuronidase (GUS) reporter plants
AtCEP1 is found on chromosome V (TAC clone K6A12; AB024031). Between the cDNA sequence of AtCEP1 (bases 50038–51750) and the next upstream cDNA sequence are 1300 bp. AtCEP2 and AtCEP3 are found in tandem on chromosome III (BAC clone T29H11; AL049659). Between the coding sequence of AtCEP2 and the next upstream coding sequence are 
3700 bp, and between the coding sequence of AtCEP2 and AtCEP3 are 6500 bp, both of which should be taken into consideration as the respective putative promoters (Appendix S2; see online Supplemental Data). However, both sequences would be unusually long for an Arabidopsis promoter (Lee et al., 2006). We used promoters with of aproximately 1360 bp for CEP1, 1910 bp for CEP2, and 1865 bp for CEP3 and amplified them by PCR from genomic Arabidopsis DNA as SacII-XbaI-fragments, which were fused to the uidA gene for the expression of β-glucuronidase (GUS) and cloned into the binary Agrobacterium vector pBI121 (Clontech, Palo Alto, CA). The primers were as follows: Pcep1-sense, 5' AAG CCG CGG AAC GAA TAC ATT TAT TTT TC 3'; Pcep1-antisense: 5' TAC TCT AGA TTG TGA TTG AGT TTG TTG ATG 3'; Pcep2-sense, 5' AAA CCG CGG CGT TGC ATT TTG GGG ATC 3'; Pcep2-antisense, 5' GGT TCT AGA TTT GAG GTT GAA AGT GCA ATG 3'; Pcep3-sense, 5' CTA GAC CGC GGT CAC AAA ACA GAG GAG ACT G 3'; Pcep3-antisense, 5' TTG TCT AGA TGG TTG GTT CTG TTT ATG 3'.
For sequence and position of CEP primers, see Appendix S1 (see online Supplemental Data).
The transition between the respective promoters and the coding region for GUS were sequenced for the control. Transformation of A. tumefaciens strain C58 with Ti plasmid pGV3580 was carried out by electroporation. T-DNAs carrying the promoter-GUS constructs, AtCEP1::GUS, AtCEP2::GUS and AtCEP3::GUS were transformed into Arabidopsis flowers ecotype Columbia (Col-0) by vacuum infiltration (Meyer et al., 1994). T0 plants were generated by selection using kanamycin (0.2% of the seeds in the T0 generation). Genomic leaf DNA was isolated (Lukowitz et al., 2000), and PCR with appropriate primers verified transgenic plants containing the promoter::GUS constructs. For each construct (AtCEP1::GUS, AtCEP2::GUS and AtCEP3::GUS), five different transgenic lines were isolated and grown for further generations to obtain homozygous plants.
Plant growth
The seeds of the promoter::GUS reporter lines were sterilized with 80% ethanol–0.1% TritonX100 and grown on Murashige–Skoog agar (Murashige and Skoog, 1962) with microsalts and vitamins (Gamborg et al., 1968) at 22°C under continuous light (100 µE•m–2•s–1). Vegetative tissues of 4–10-d-old seedlings were analyzed. Seedlings were transferred to soil for further growth in phytochambers (Conviron, Canada) under long day conditions (16 h light at 22°C and 65% RH; 8 h dark at 18°C and 75% RH). Alternatively, plants were grown in a greenhouse. Generative tissues (inflorescences) were analyzed from the emergence of flowers in 4-wk-old plants until seed maturation in 10–12-wk-old plants.
Histochemical GUS analysis
Arabidopsis transformants expressing the β-glucuronidase (GUS) reporter gene under the control of the three different Arabidopsis KDEL-tailed cysteine endopeptidase promoters were histochemically assayed with either 5-bromo-4-chloro-3-indolyl-β-D-glucuronide (X-Gluc; AppliChem, Darmstadt, Germany) using light microscopy or with ELF97-β-D-glucuronide (ELF97-Gluc; Molecular Probes, Leiden, The Netherlands) using fluorescence microscopy or confocal laser-scanning microscopy (LSM; Zeiss, Jena, Germany). Seedlings and influorescences were analyzed.
Staining with X-Gluc was performed as described (Jefferson et al., 1987; Rodrigues-Pousada et al., 1993; An et al., 1996). For better visualization of the indigo stain, tissue blocks were cleared in graded ethanol solutions (30%, 50%, then 70%) to remove chlorophyll. Nuclear DNA was counterstained with 0.1 µg/mL of 4,6-diamidino-2-phenylindole (DAPI; Sigma, München, Germany) for 5 min at room temperature. Thin sections of positively stained plant material were prepared after embedding in Paraplast Plus (melting point 56°C; Sigma) (Schmid et al., 1999) with the following modifications: X-Gluc-stained tissues were transferred to FAA (3.7% formaldehyde, 50% ethanol, 5% acetic acid) instead of 50% ethanol for 30 min (seedlings) or 2 h (influorescences). Tissue blocks were dehydrated by immersion into a series of aqueous alcohol solutions (70%, 2 x 90%, 96%, 2 x 100% ethanol; 2 x 100% isopropanol) for 1 h each. Subsequently, the tissue blocks were infiltrated with Rotihistol (Roth, Karlsruhe, Germany) as the carrier, 3 times for 1 h each at room temperature. Infiltration with Paraplast was at 60°C. Tissue blocks were infiltrated with Rotihistol-Paraplast-mixture (1:1) for 1 h, with Rotihistol-Paraplast-mixture (1:10) overnight with 100% Paraplast 2 times for 1 h each. Polymerization was done in aluminum moulds. After hardening, the specimens were cut into 8–10 µm sections using a rotary microtome (Leitz, Jena, Germany).
Plant material was washed, vacuum-infiltrated on ice for 10 min in fixative (0.15 M Na phosphate buffer pH 7.0, 0.2% (v/v) Triton-X 100, 1% formaldehyde) followed by further gentle agitation for 1 h on ice, equilibration on ice for 15 min in equilibration buffer (0.15 M Na phosphate buffer pH 7.0, 0.2% (v/v) Triton-X 100) and incubation at 37°C for 1 h in reaction buffer (0.15 M Na phosphate buffer pH 7.0, 0.2% (v/v) Triton-X- 100, 25 µm ELF97; that is 1:200 dilution of 5 mM ELF97-Gluc in dimethyl sulfoxide (DMSO) into equilibration buffer). Microscopy was carried out with a Zeiss fluorescence microscope (Axioskop) and the ELF97 filter (excitation 365 nm/ beam splitter, 395 nm/ emission filter 520 nm LP); emission filter 535/50 HQ was used for chlorophyll autofluorescence.
Isolation and characterization of extensins from tobacco BY2 cell suspension cultures
Nicotianum tobacum L. cv. Bright Yellow 2 (BY2) cells were subcultured, and cell surface HRGPs, including the P1-type extensin (accession CAA50603; Fig. 2), were purified and biochemically characterized as described earlier (Lamport and Miller 1971; Smith et al., 1984; Held et al., 2004). The gene encoding the arabinogalactan protein P3-type extensin analogue, designated AP-YK20 (Fig. 3), was inserted between the XmaI/BbsI site in pUC-(YK)20, described earlier (Held et al., 2004) as the XmaI/BbsI fragment below:
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The resulting synthetic gene, (AlaPro)4(YK)20, was ligated into pUC-SStob(AP)51EGFP replacing the (AP)51 gene (Tan et al., 2003). The SStob(AlaPro)4(YK)20EGFP fragment was then inserted into pBI121 to give pBI-SStob(AlaPro)4(YK)20EGFP. EGFP was removed from AP-YK20-EGFP by tryptic digestion, and AP-YK20 was purified by HPLC using methods described earlier (Held et al., 2004).
The resulting purified monomeric P1-type and P3-type extensins associate on their own in solution to create scaffolds (Cannon et al., 2008).
Digestion of cell wall proteins with Ricinus CysEP
Ricinosomes were isolated from 5 day-old germinating castor bean endosperm on a discontinuous sucrose gradient as described (Schmid et al., 2001). Purified ricinosomes were diluted 10-fold with buffer (10 mM MES, 10 mM dithiothreitol, pH 5.5) and incubated at 20°C for 30 min to disrupt the organelles and autoactivate CysEP. The membranes were separated from the matrix proteins by high speed centrifugation.
The activated Ricinus CysEP from isolated ricinosomes was added to purified P1-type extensin or to AP-YK20 at a substrate to enzyme ratio of 25:1 (w/w). The reaction was either terminated within 15 s or after 5 or 10 min at 25°C. P1-type and P3-type extensins incubated with buffer were used as a control. The cleavage products were analyzed by SDS-PAGE (12.5% polyacrylamide gel) and silver staining (Blum et al., 1987). For further analysis of cleavage products, digestion times of 15 s, 2 min, 10 min, 1 h, 6 h, and 24 h were used. Peptides were separated by reversed phase HPLC prior to N-terminal sequencing and matrix-assisted laser desortpion/ionization (MALDI) analysis as previously described (Than et al., 2004).
RESULTS
Tissue-specific expression of AtCEP1, AtCEP2, and AtCEP3
RT-PCR with specific pairs of primers detected the presence of mRNA for AtCEP1, AtCEP2, and AtCEP3 in roots, stems, flowers and buds, and green siliques of wild-type Arabidopsis. In addition, AtCEP2, but not AtCEP1 or AtCEP3, was detectable in rosette and cauline leaves (Fig. 4). A gene expression map of Arabidopsis thaliana development largely confirmed these results for CEP1 and CEP3; no data were available for CEP2 (Schmid et al., 2005; Appendix S3, see Supplemental Data with the online version of this article). The data presented were analyzed using the algorithm gcRMA, which differs from the algorithm implemented in the software package Affymetrix MAS5/GCOS. We reanalyzed the original data using the MAS5 software, which allowed us to determine at which gcRMA expression level a given gene is reliably detectable, i.e., present in all three arrays that were hybridized for each of the 79 tissues in the expression atlas. In the case of AtCEP1 (At5g50260; Affy probe Id. 248545_at), we found the gene was stably expressed in 21 of the 79 tissues according to MAS5/GCOS analysis. Superimposition of MAS/GCOS onto gcRMA analysis indicates that AtCEP1 (At5g50260) can be considered stably expressed for gcRMA values >30. Similarly, AtCEP3 (At3g48350; Affy probe Id. 252365_at) was reliably detectable in 63 tissues, but absent from 16 tissues. AtCEP3 (At3g48350) could be considered stably expressed for gcRMA values >13. These results indicated that the three CEP genes were expressed quite broadly during Arabidopsis development, but nevertheless showed some evidence of tissue and stage specific expression. All samples that were analyzed by RT-PCR or that were part of the AtGenExpress gene expression map were composed of several different tissues and cell types. To obtain a more detailed picture of the expression of the CEP genes during seedling and flower development, we used histochemical GUS assays to analyze transgenic Arabidopsis plants expressing β-glucuronidase (GUS) under the control of the promoters for AtCEP1, AtCEP2 and AtCEP3.
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Roots
CEP1 promoter activity was found within columella or beginning root cap (Fig. 5) and in lateral root cap cells of 4–10-d-old seedlings (Fig. 6) and in the course of lateral root formation. CEP1::GUS activity was found in the endodermis, the cortex, and the epidermis. The first lateral roots are normally formed at the hypocotyl-root transition (Fig. 7). A series of developmental stages during lateral root formation (Malamy and Benfey, 1997) is shown in Figs. 8–11. CEP1::GUS activity was found in endodermis cells at stage II during transversal divisions (not shown) and at stage III during periclinal divisions (Fig. 8) of the pericycle cells. GUS-activity was found in endodermal cells pushed aside as the lateral root primordium penetrated the endodermis (stage IV) (Fig. 9) and in penetrated cortex cells (Fig. 10) (stage VII). Finally, CEP1::GUS activity was found at the basis of the lateral root as the primordium penetrates the epidermis (stage VIII) (Fig. 11). Thus CEP1 promoter activity was localized at collapsing cells such as the calyptra and in tissues where the cells need to make room, for example, for a growing lateral root.
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CEP3 promoter activity in roots was found at the hypocotyl–root transition zone (Fig. 20) and in the vascular tissue and possibly in the endodermis (Figs. 20–22).
Stomata of cotyledons and trichomes of primary leaves
Promoter activity of CEP1, CEP2, or CEP3 was not found in leaves with the exception of the trichomes. Interestingly, CEP3 promoter activity was localized in the trichomes of true leaves (Figs. 23 and 24), at the basis of cotyledons near vascular tissue (Figs. 23 and 25), and especially near the stomata of cotyledons but not of true leaves (Figs. 23 and 26).
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, Smyth et al. (1990) and Roeder and Yanowski (2006) (Appendix S4, see Supplemental Data with online article). Fruit development is defined to begin after fertilization of the egg cell and endosperm during stage 14. CEP1 promoter activity was first found at stage 15, when the gynoecium elongates and extends beyond the top of the stamens, at stage 16, when the sepals, petals, and stamens wither and separate from the fruit, and at stage 17, when all organs separate from green siliques, and when the fruit elongates completely and protects the seeds throughout their development. At stage 15, CEP1::GUS expression was visible in anthers near the end of the filament (Fig. 27). At stage 16, this signal became stronger in anthers and was also found in the stigma (Figs. 28–30). Stage 17 is a long stage, which starts when the floral organs separate from the fruit and ends when the fruit starts to yellow. CEP1 promoter activity was found in a ribbon along the abscission zones of the flower organs. Here, expression was especially conspicuous in the area of the lateral nectaries and in the abscission zones of sepals and petals (Figs. 31–33). In young siliques during late stage 16 and early stage 17, maturing seeds had CEP1 promoter activity (Fig. 34), which ceased in the course of seed maturation during late stage 17 (Fig. 35) and disappeared in stage 18, when the siliques turned yellow, corresponding to the time when the inner integument developed into the seed coat and the outer integument became slimy (Fig. 36). CEP1 promoter activity was strong in unpollinated, degrading ovules (Figs. 37 and 38). A cross-section through an unpollinated and a pollinated ovule (Fig. 38) showed that the pollinated maturing seed stained with DAPI, indicating an intact nucleus, but no GUS expression, whereas the unpollinated ovule had no DAPI staining with CEP1::GUS expression, indicating collapsing tissue (Figs. 39 and 40). The degrading ovules became smaller with progressive silique maturation and showed CEP1::GUS activity till the end. CEP1 promoter activity was found in collapsing endosperm cells upon germination (Figs. 41 and 42) and at two sites of lateral root formation (Fig. 41; for comparison see Figs. 8–11 and Fig. 19).
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CEP3 promoter activity was strongest during silique development in stages 14 and 15 and became weaker in later stages. GUS expression was limited to the carpels and the pedicel (Figs. 43–45). CEP3 promoter activity was never found in the abscission zone of flower organs or in the dying nectaries or in the stigma, as was characteristic of CEP1.
Ricinus CysEP digests the cell wall protein P1-type extensin, but not the P3-type extensin AP-YK20
GUS expression under the control of the AtCEP1, AtCEP2, and AtCEP3 promoters showed that the KDEL-tailed cysteine endopeptidases were expressed in tissues where cells were making way, for example, for a growing lateral root, and especially where the cells were about to collapse in the course of developmentally regulated PCD. Consequently, we investigated if Ricinus CysEP was able to digest proline/hydroxyproline-rich proteins such as the extensins. Extensins form positively charged scaffolds in the cell plate and subsequently provide the template for deposition of the other cell wall components (Cannon et al., 2008). This scaffold may uphold the cells three-dimensional structure even after breakdown of the cells turgor. Conversely, its digestion may allow modulations to accommodate intercalation of new cells and cell wall adjustments.
The Hyp residues of tobacco P1-type extensin (Fig. 2) and P3-type extension AP-YK20 (Fig. 3) were highly arabinosylated (93 mol % Ara, 7 mol % Gal for P1 and 80 mol % Ara, 12 mol % Gal for AP-YK20). Tobacco P1 extensin contained no arabinogalactosylated Hyp, all Hyp residues being exclusively arabinosylated (Fig. 2). Consistent with the presence of three clustered noncontiguous Hyp residues in AP-YK20, we calculated from the protein sequence and Hyp glycoside profiles (Fig. 3) that each glycoprotein contained 3 arabinogalactan polysaccharides, the bulk of the Hyp residues being arabinosylated, consistent with predictions of the Hyp contiguity hypothesis (Shpak et al., 1999; Held et al., 2004; Tan et al., 2003).
Undigested P1-extensin and AP-YK20 are very high molecular weight basic molecules and only entered the stacking gel (5% polyacrylamide) (Fig. 46, lanes 1 and 5). Upon incubation with Ricinus CysEP for up to 10 min, AP-YK20 remained in the stacking gel (Fig. 46, lanes 6–8), whereas P1-extensin was reduced to a mixture of cleavage products within 15 s (Fig. 46, lanes 2–4). AP-YK20, composed mainly of the the repeated sequence (SOOOOSOSOOOOYYYK)20 (O = Hyp), remained undigested after up to 24 h, and sequencing revealed only the original N-terminus PAOAOAO (Fig. 3). P1 extensin was digested within 2 min, the main cleavage products being KPYYPOH... and KS*OOOO... and possibly S*OOOO (S* = galactosylated Ser). Arabinofuranosides are extremely acid-labile and were removed from Hyp by the concentrated trifluoroacetic acid used in Edman degradation. Ser-Gal is more stable and was inferred from blank cycles that coincided with Ser residues predicted from the gene sequence (cf. Fig. 2) and from earlier sequencing of the glycoprotein (Terneus, 2006). Due to the blank cycle, it was difficult to determine if the peptide S*OOOO... was a cleavage product in addition to the peptides KS*OOOO. TOVYKSO... and HPVYKSO... were also found as by-products within the 2 min of digestion. An additional cleavage product YPOHTOVY..., although not present within the first 2 min digestion time, accumulated after 24 h.
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KPYYPOH (6x) and OOOHKKPYYPOH (1x). Looking at the cleavage sites from positions P4 to P4' (positions –4 to +4 relative to the cleavage site), it seems likely that Ricinus CysEP accepts glycosylated Hyp at the P2 and P3 positions (positions –2 and –3 relative to the cleavage site). Although approximately nine nonglycosylated Hyp residues occur in the P1-type extensin, it is unlikely they occur within the blocks of contiguous Hyp residues (Terneus, 2006; Held et al., 2004). Sequences leading to the digestion product KS*OOOO... (and S*OOOO...) are found 16 times: TOVYK()S*OOO (14x), KKLYK()S*OOO (1x) and HPVYK()S*OOO (1x). There is only one sequence motif that would lead to the cleavage product S*OOOO...: YQYS()S*OOOO. It can, therefore, be deduced that Ricinus CysEP also accepts glycosylated amino acids at the P2' and P3' positions (positions +2 and +3 relative to the cleavage site). The by-product TOVYKSO... can result from cleavage of the sequence YPOHTOVY (7x) and possibly also from OOOOTOVY (7x), if the Ricinus CysEP accepts arabinosylated Hyp at the P1 position. However, because the Hyp-Lys bonds in the P1-extensin S*OOOK repeats were resistant to cleavage, cleavage of the module OOOOTOVY might not be the case. The cleavage products HPVYKSO... and YPOHTOVY... result from sequences with glycosylated amino acids in the P3 position or with no glycosylated amino acids near the cleavage site: OOVHHPVY (1x) and KKPYYPOH (7x).
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Germinating castor bean seeds were used previously to study the role of glyoxysomes in storage oil mobilization (Beevers, 1979). Vigil (1970) reported that the isolation of glyoxysomes on a sucrose gradient resulted in coenrichment of another organelle called "dilated cisternae" because it seemed to develop from the endoplasmic reticulum. Mollenhauer and Totten (1970) named this organelle "ricinosome" because it was found at that time only in the endosperm of germinating Ricinus communis seeds. The lack of a known marker enzyme prevented attempts at the time to biochemically characterize ricinosomes. Ricinosomes were rediscovered with the identification of a KDEL-tailed cysteine endopeptidase (CysEP) as their marker enzyme (Schmid et al., 1998) and were found to play an important role in the final stage of developmentally regulated PCD (Gietl and Schmid, 2001). Ricinosomes are not only found in the senescing Ricinus endosperm after the completion of storage mobilization. Ricinosomes were likewise identified by immunolocalization of the CysEP in other senescing tissues undergoing developmentally regulated PCD, such as the collapsing nucellus cells of maturing Ricinus seeds (Greenwood et al., 2005) and in the senescing flower daylily petals (Schmid et al., 1999).
Ricinus communis is an important plant for biochemical and molecular studies of the ricinosome and its enzymes because of the methods developed for the purification of this organelle. However, no genomic tools are available for this plant. We have therefore surveyed the occurrence of the ricinosome- and PCD-marker KDEL-tailed cysteine endopeptidase on other plants (Fig. 1) and presented their phylogenetic tree (Gietl et al., 2000; Gietl and Schmid, 2001; Beers et al., 2004). For further analysis, we selected the three KDEL-tailed cysteine endopeptidases in the genome of Arabidopsis thaliana and made use here of the genomic tools available for this organism.
The three KDEL-tailed cysteine endopeptidases CEP1, CEP2, and CEP3 of Arabidopsis are expressed in tissues undergoing PCD
Three KDEL-tailed cysteine endopeptidases are found in Arabidopsis. They had an expression pattern comparable to the tissue-specific expression during PCD, whereby especially CEP1 and also CEP3 play a prominent role in developmentally regulated PCD of generative tissues during distinct stages of flower development. Similar to the expression of the RDEL-tailed cysteine endopeptidase O141 of Phalaenopsis, CEP1::GUS activity was found in maturing seeds (Nadeau et al., 1996), where the integuments develop into the seed coat and the outer cell layer of the seed coat contains layers of mucilage. In analogy to the expression of the KDEL-tailed thiolprotease TPE4A in the unpollinated ovules of Pisum sativum (Cercos et al., 1999), CEP1::GUS activity was high in unpollinated, degrading ovules.
As found with the KDEL-tailed peptidases expressed in the senescing storage tissues of Ricinus, Vicia, and Picea after completion of oil and protein mobilization (Schmid et al., 1999; Becker et al., 1997; He and Kermode, 2003), CEP1 promoter activity was also found in the remnants of the Arabidopsis endosperm after germination. In maturing Arabidopsis seeds, the cells of the transient nuclear endosperm are mainly exploited as the major storage organ, the cotyledons, expand and begin to accumulate reserves. Continued cell expansion throughout the embryo causes it to fill most of the embryo sac, crushing the endosperm formed earlier during embryogenesis and presumably absorbing it. By maturity, the cotyledons are tightly appressed to the radicle, leaving little open space in the embryo sac, with only a persistent layer of endosperm remaining (Bowman, 1993). Arabidopsis stores significant amounts of lipids in the endosperm, and carbohydrates derived from this lipid are required for postgerminative seedling growth in the dark (Penfield et al., 2004). Thus the endosperm undergoes PCD during germination after storage material mobilization, similar to castor bean endosperm.
CEP1 exhibits a striking activity in the abscission zones of old postsecretory flowers in stage 17, when the nectaries shrink and collapse, and when the sepals, petals, and stamens senesce and fall from the silique within a few days after fertilization.
As with the expression of the KDEL-tailed cysteine endopeptidase (EP-C1) in maturing pods of P. vulgaris (Tanaka et al., 1991), CEP3::GUS activity was high during silique development in the carpels and in the pedicel of stage 15. Siliques remain green for more than a week after fertilization, but turn yellow (stage 18), and then brown at maturity. At that time, the valves of the dry siliques separate, releasing the seeds (stage 19 and 20). CEP3::GUS expression ceased in the course of seed maturation and disappeared in stage 18.
The cotyledons of V. sativa are a storage tissue to support embryo growth during germination similar to the cotyledons of Arabidopsis. Afterward, however, V. sativa cotyledons are committed to PCD, whereas Arabidopsis cotyledons rather become green in the light and perform photosynthesis. Senescence and PCD are expected to take place later in development. Interestingly, CEP3::GUS expression was found in guard cells of Arabidopsis cotyledons, but not in those of primary leaves. Possibly, cotyledon senescence starts within the stomata. CEP3 promoter activity in guard cells of cotyledons and in trichomes of rosette leafs might alternatively indicate cell type specific wall alterations.
The Arabidopsis root has a relatively simple anatomy composed of single layers of epidermal, cortical, and endodermal cells surrounding the vascular tissues. Four types of stem cells (or initial cells) at the root tip generate these tissues (Scheres et al., 2002). The epidermal/lateral root cap initials give rise to the epidermis and the outer portion of the root cap known as the lateral root cap. The central portion of the root cap, the columella, has its own set of initials. The basal tissue cells, the cortex and endodermis, are generated by division of the cortex/endodermis initials. Finally, the vascular tissue and pericycle have their own initials. CEP2::GUS expression is especially found in the root cap domain, where the outermost, oldest cells of the calyptra finally collapse and slough off. CEP2 seems to be involved in the developmental PCD of these short-lived cells that are continually replaced by the root meristem. CEP1 promoter activity was found within the columella or beginning root cap and in lateral root cap cells. CEP1::GUS expression, however, seems to be a slightly different phenomenon during lateral root development. Lateral roots arise from complex cell division patterns within the pericycle, and eventually the lateral root pushes through the endodermis cortex and epidermal cell layers to erupt through the side of the root. The CEP1 peptidase seems to be involved in loosening the contact between the cells rather then in the final collapse of cells, to make room for the growing lateral root.
Even though AtCEP3 is expressed in senescing leaves according to the AtGenExpress gene expression map, CEP::GUS activity was not detected in this tissue in any of the reporter lines analyzed, indicating that the promoter constructs might lack certain regulatory elements that are required for CEP expression during senescence. Also, we cannot exclude the possibility that more thorough observations will disclose the expression of KDEL-tailed cysteine peptidases during a narrow time window in special tissues such as the leaf abscission zone. In principle, however, the degraded cytosolic constituents during the final stage of cell death might be transferred through the vascular system for resource allocation, thus making the collapse of leaf cells and their absorption by the surrounding cells unnecessary.
While we have demonstrated that the three KDEL-tailed cysteine endopeptidases of Arabidopsis are major enzymes in degradation of proteins during PCD, they are not the only peptidases participating in this process; and formation of ricinosomes is not the only pathway to transport cysteine proteases to the site of action.
So far ricinosomes have not been identified in Arabidopsis, but a detailed analysis of the endothelium tissue during seed maturation has identified an alternative pathway for the transfer of enzymes into the lytic vacuoles (LV) and into the protein storage vacuoles (PSV) prior to PCD (Ondzighi et al., 2008). A protein disulfide isomerase (PDI5) with a C-terminal KDEL retention signal is transferred together with a cysteine protease without a KDEL motif via the Golgi apparatus into the two types of vacuoles. The protein of the PVS is degraded, and the PVSs become the lytic vacuoles for PCD of the endothelium cells. It is suggested that the KDEL-tailed PDI5 functions as a chaperone for transporting the proteases from the ER through the Golgi apparatus to the sites of action and prevents their premature function. The inhibition of the activity of the cysteine proteases by PDI5 is shown in vitro, and a knock-out mutant of Arabidopsis for the structural gene of PDI5 leads to premature onset of PCD of the endothelium. Because the PDI5 carries a KDEL retention signal, it remains to be seen how the ER retention signal is silenced during the transport of the PDI5 through the Golgi apparatus (Ondzighi et al., 2008).
KDEL-tailed cysteine endopeptidases digest extensin, thus supporting the final cell collapse during PCD
During plant PCD, the breakdown products of the protoplast are thought to be absorbed by the surrounding cells, but the cell wall may be partially or completely degraded or left intact as a corpse-like remnant (Jones, 2001). For example, lateral walls are partially degraded during xylem vessel element differentation (Fukuda, 2000). The disintegration and final collapse of the endosperm in germinating seedlings or the nucellus in maturing seeds results in crushed and folded cell wall residues in the apoplastic space (Schmid et al., 1999; Greenwood et al., 2005).
It is noteworthy, that the KDEL-tailed proteases are specifically involved in the PCD of tissues that finally collapse, serving a dual function in digesting cellular constituents and weakening the cell wall. The KDEL-tailed proteases possess the rare ability to accept proline at the P1 and P1' positions of the cleavage site (Than et al., 2004), even accepting highly glycosylated hydroxyprolines at the P2 and P2' positions. The crystal structure and the peptide cleavage specificity of the Ricinus KDEL-tailed cysteine endopeptidase led to the investigation of its substrate specificity toward hydroxyproline-rich glycoproteins (HRGPs) such as the P1-type extensin from tobacco. As is presented in Fig. 47, the Ricinus KDEL-tailed cysteine endopeptidase cleaved the P1-type extensin into 17 peptides containg the Ser-(Hyp)4 modul. The importance of extensins in the self assembly of the plant cell wall was recently underscored by the analysis of the knockout Arabidopsis mutant rsh (Cannon et al., 2008). The gene encodes the P3 extensin and in its absence disorganized cell walls are formed. Sequencing after deglycosylation revealed that extensin had 11 identical 28 residues peptide repeats each with a Tyr-Val-Tyr repeat for isodityrosine crosslinks. The purified monomers were crosslinked in vitro by extensin peroxidase. A staggered alignment of the repetitive peptides by 12 amino acids allows the formation of dityrosine bonds formed between neighboring Tyr-Val-Tyr and lone Tyr residues (pulcherosine crosslink motifs). This staggered alignment allows growth of the polymer into a two-dimensional scaffold, which was demonstrated by atomic force microscopy (Cannon et al., 2008). The authors conclude that the positively charged extensin scaffolds form in the cell plate of newly dividing cells. These react with negatively charged pectins to create an extensin pectate coacervate that may form the template for further orderly deposition of the new cross wall at cytokinesis. The CEP1 Arabidopsis KDEL-tailed cysteine endopeptidase seems to be expressed during cell division during the formation of lateral roots (Figs. 8–11). Partial degradation of the extensins of the mother cell wall might be performed by CEP1 when a new wall has to be attached during cytokinesis.
Conclusions
Hydrolytic enzymes, including the KDEL-tailed cysteine endoproteases execute programmed cell death in plants, catalyzing the hydrolysis of cellular constituents for resource reallocation. Through mapping the expression of the reporter protein GUS attached to the promoters of three KDEL-tailed CEP proteases in Arabidopsis transformants, the activities of the individual protease involved in programmed cell death seem to be highly cell-, tissue- and organ-specific. Additionally, KDEL-tailed proteases cleave structural cell wall proteins such as extensins, thus contributing to the final cell collapse. When tested on a major glycosylated hydroxyproline containing cell wall protein, P1 extensin, the homologous KDEL-tailed cysteine endopeptidase from Ricinus communis degrades this protein, thus introducing a new tool for studies of cell wall synthesis and degradation.
FOOTNOTES
1 The authors thank H. Cochran (Washington State University, USA) for artwork, J. S. Greenwood (University of Guelph, Canada) for valuable discussions, and D. Simpson (Denmark) for critically reading the manuscript. This work was supported by the Deutsche Forschungsgemeinschaft (Gi154/11-5). 
5 Present address: Max-Plank-Institut für Entwicklungsbiologie, Spemannstr. 37-39, D-72076 Tübingen, Germany
6 Author for correspondence (e-mail: christine.gietl{at}wzw.tum.de)
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