Plant Physiology

Minorsky, Peter V.

doi: 10.1104/pp.104.900181pmid: N/A

Transcriptome of the Female Gametophyte Because the female gametophytes of flowering plants are small and embedded within the maternal tissues of the ovule it is difficult to isolate them free of contaminating maternal tissue. Thus, relatively little is known about the genes that are expressed during female gametophyte development. It is not even known, for example, whether the female gametophyte transcriptome contains a major set of genes that are not expressed in the sporophyte or whether it is primarily a subset of the sporophytic transcriptome. Yu et al. (pp. 1853–1869) have sidestepped the problem of the female gametophyte being embedded within the maternal ovule tissue by utilizing the Arabidopsis (Arabidopsis thaliana) mutant sporocyteless that produces ovules lacking female gametophytes. Using an Arabidopsis whole-genome oligonucleotide array, they were able to identify a minimum of 225 genes as female gametophyte genes. Nearly 45% of the identified genes have not previously been detected by sporophytic expression profiling, suggesting that the female gametophyte transcriptome may contain a significant fraction of transcripts restricted to the gametophyte. Plant Defense against Aphids The green peach aphid (Myzus persicae; Fig. 1 Figure 1. Open in new tabDownload slide Arabidopsis plants shed leaves prematurely to defend themselves against green peach aphids (photo courtesy of Emma Hunt). Figure 1. Open in new tabDownload slide Arabidopsis plants shed leaves prematurely to defend themselves against green peach aphids (photo courtesy of Emma Hunt). ) has a wide host range covering greater than 50 families of plants and is the vector for more than 100 plant viruses. Unlike the better studied chewing insects, aphids do not cause extensive wounding to the plant host, suggesting that plant responses to phloem-feeding insects may differ significantly from those elicited by chewing insects. Aphid feeding causes changes in resource allocation in the host, resulting in increased nutrient flow to the insect-infested tissue. Pegadaraju et al. (pp. 1927–1934) have hypothesized that premature leaf senescence would be useful in limiting aphid growth. The authors report that green peach aphid feeding upon Arabidopsis leaves induced premature chlorosis and cell death and increased the expression of SENESCENCE ASSOCIATED GENES, all hallmarks of leaf senescence. Premature senescence was accompanied by enhanced resistance against green peach aphid in two mutants of Arabidopsis that have elevated expression of SENESCENCE ASSOCIATED GENES. In contrast, resistance against green peach aphid was compromised in the pad4 mutant plant. PAD4 is associated with the synthesis of camalexin, an antimicrobial phytoalexin, and with salicylic acid (SA) signaling. Studies with other mutants that are defective in camalexin synthesis or SA signaling, however, indicated that camalexin and SA have no bearing on aphid resistance. Hence, the involvement of PAD4 in Arabidopsis defense against green peach aphid is most likely independent of its role in SA signaling and camalexin biosynthesis. Metabolite Profiling of Chlamydomonas Metabolic profiles reflect the dynamic response of biochemical pathway networks to environmental, genetic, or developmental signals. As such, they provide valuable information concerning how a living system adjusts to its changing environment. Bölling and Fiehn (pp. 1995–2005) have developed a metabolite-profiling technique for Chlamydomonas reinhardtii cells. The experimental protocol was optimized to quickly inactivate enzymatic activity, achieve maximum extraction capacity, and process large sample quantities. As a result of the rapid sampling, extraction, and analysis by gas chromatography coupled to time-of-flight mass spectrometry, more than 800 analytes from a single sample can be measured. To demonstrate the power of their technique, the authors analyzed how metabolite profiles change under conditions of nitrogen, sulfur, phosphorus, and iron depletion, respectively. Sulfur-depleted cells exhibited the largest increase of any single compound: 4-Hyp accumulated more than 50-fold compared to control conditions. Hyp is a prominent constituent in the Hyp-rich glycoprotein framework forming the Chlamydomonas cell wall. Cell wall proteins are extensively sulfated and rearranged during sulfur starvation while several prolyl 4-hydroxylases are down-regulated. Thus, the rise in 4-Hyp could be the result of enhanced degradation of cell wall proteins. Two other findings of note were that phosphate removal results in a 25-fold rise in intracellular Cys, whereas sulfate removal causes 2-ketovaline levels to plummet to 2% of control levels. Alternative Oxidase and Oxidative Stress The cyanide-resistant alternative oxidase (AOX) of plant mitochondria accepts electrons from the ubiquinone pool and uses them to reduce oxygen to water, with no conservation of energy by the formation of proton gradients across the inner mitochondrial membrane. This raises the question of why a seemingly energy-wasteful pathway operates in plant mitochondria. One hypothesis is that AOX may be involved in acclimation to oxidative stress: AOX may function to prevent the formation of reactive oxygen species (ROS) by diverting reductants in excess of cytochrome pathway capacity down the AOX pathway. To date, the AOX connection with mitochondrial ROS has been investigated only in isolated mitochondria and suspension culture cells. To study ROS and AOX in whole plants, Umbach et al. (pp. 1806–1820) generated three classes of transformed lines of Arabidopsis: AtAOX1a overexpressors, AtAOX1a anti-sense plants, and overexpressors of a mutated, constitutively active AtAOX1a. In the presence of KCN, leaf tissue of AOX overexpressors showed no increase in oxidative damage, whereas anti-sense lines had levels of damage greater than those observed for untransformed leaves. Similarly, ROS production increased markedly in response to KCN treatment in anti-sense and untransformed, but not overexpressor, roots. Thus, AOX functions in leaves and roots, as in suspension cells, to ameliorate ROS production when the cytochrome pathway is chemically inhibited. In contrast to previous reports using suspension culture cells, no changes in leaf transcript levels of selected electron transport components or oxidative-stress-related enzymes were detected. Furthermore, a microarray study using an anti-sense line showed AOX influences processes outside mitochondria, particularly in chloroplasts and several carbon metabolism pathways. These results illustrate the value of expanding AOX transformant studies to whole tissues. The different transcriptional responses of leaves and cultured cells indicate that the effects of altered AOX levels need to be examined in intact tissues to achieve a better understanding of AOX function and its interaction with other metabolic systems in whole plants. In a companion paper, Fiorani et al. (pp. 1795–1805) tested the hypothesis that AOX plays a role in ameliorating plant stress under cold conditions by preventing excess accumulation of ROS. In plants grown at 12°C, AOX anti-sense plants showed 27% reduced leaf area and 25% smaller rosettes. In AOX overexpressors, the leaf areas and rosette size were 30% and 33% larger, respectively. These phenotypic differences were not the result of major alterations in tissue redox state because the changes in levels of lipid peroxidation products, reflecting oxidative damage, and the expression of genes encoding antioxidant and electron transfer chain redox enzymes did not correspond with the shoot phenotypes. These results demonstrate that: (1) AOX activity plays a role in shoot acclimation to low temperature in Arabidopsis and (2) AOX not only functions to prevent excess ROS formation in whole tissues under stressful environmental conditions but also affects metabolism through more pervasive effects, including some that are extramitochondrial. Rapamycin Sensitivity in C. reinhardtii The antibiotic rapamycin is a potent antifungal agent that also exhibits immunosuppressive activity due to its capacity to block the growth and proliferation of T cells. Although the vegetative growth of higher plants is unaffected by rapamycin, many players in the rapamycin signal transduction pathway are present in higher plants. Rapamycin's mechanism of action, based on research with Saccharomyces cerevisiae, involves the binding of rapamycin to the FK506-binding protein (FKBP12). This complex then inhibits a Ser/Thr kinase called TOR (target of rapamycin). Studies have identified FKBP12 in bacteria, fungi, animals, plants, and more recently in the green alga C. reinhardtii. However, the physiological function of this protein is still poorly understood in plants. In Arabidopsis, FKBP12 interacts with AtFIP37, a phosphatidylinositol kinase essential for development, whereas TOR kinase plays an important role in controlling plant cell growth and disruption of its gene leads to the premature arrest of endosperm and embryo development. Whereas Arabidopsis is insensitive to rapamycin, this drug does inhibit the growth of Chlamydomonas. Crespo et al. (pp. 1736–1749) report that Chlamydomonas lacking FKBP12 are fully resistant to the drug, indicating that this protein mediates rapamycin's inhibitory effect on cell growth. The authors also demonstrate that Chlamydomonas FKBP12, unlike its higher plant homolog, exhibits high affinity to rapamycin in vivo, and that TOR binds FKBP12 in the presence of rapamycin. It is also reported that rapamycin treatment results in a pronounced increase of vacuole size that resembles autophagy. These findings suggest that Chlamydomonas cell growth is positively controlled by a conserved TOR kinase. Author notes www.plantphysiol.org/cgi/doi/10.1104/pp.104.900181. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Ort, Donald R.

doi: 10.1104/pp.104.900180pmid: N/A

The average length of research articles published in Plant Physiology has been growing and reached an average of 11 pages over last 2 years. As shown in the histogram below (Fig. 1 Figure 1. Open in new tabDownload slide Plant Physiology article length. Figure 1. Open in new tabDownload slide Plant Physiology article length. ), in 2003 and 2004, about 35% of the research articles published were over 11 pages. The research sections “Biochemical Processes and Macromolecular Structures,” “Genetics, Genomics, and Molecular Evolution,” and “Environmental Stress and Adaptation to Stress” typically have had the longest papers. While article length in and of itself is not a concern, many of the longest articles that Plant Physiology has published are ones in which authors are not taking advantage of the supplemental data option. Papers appearing in the “Genetic, Genomics, and Molecular Evolution” section have used the online supplemental data option most frequently. Our review of research articles published in 2003 and 2004 also shows that many of the longer papers were not tightly written and, in particular, contained significant repetition between the “Results” and “Discussion” sections. In addition to perhaps lowering the impact of these papers, significant costs are associated with printing and mailing the “extra/unnecessary” pages. Because the editorial board and I believe that concisely written papers improve the impact of what we publish, there will be a limit of 10 journal pages for submissions after January 1, 2006. While longer papers will be permitted at a higher page charge, we believe that the 10-page target will be a positive incentive for tight composition, reduced repetition, and appropriate use of supplemental data files. Beginning on January 1, 2006, page charges will be $75 per page for the first 10 pages ($55 per page when the corresponding author is a member of ASPB). The charge for the 11th page and above will be doubled. As with other publication costs, Plant Physiology authors may request a waiver of this extra page surcharge if special circumstances justify. Plant Physiology has developed a user-friendly page calculator that will determine the number of printed pages your article will run if it is accepted for publication. Upon submission the author will be asked for the following information: Number of characters in the manuscript (including spaces) Number of one-column figures Number of two-column figures Number of lines in one-column tables Number of lines in two-column tables The resulting page number projection will be returned to the author as well as automatically inserted into a field with the other article data. Our testing shows that, even with these simple inputs, the calculator is quite accurate in estimating the length of printed articles. Should there be any inaccurate estimates, authors will of course not be held responsible for them. The calculator will also be accessible from the Instructions for Authors Web page (http://www.plantphysiol.org/misc/ifora.shtml) as an author tool that can be used during the preparation of manuscripts. I would like to close by reminding all of the opportunity to publish your article in Plant Physiology with full and immediate Open Access (http://www.aspb.org/openaccess/). This option allows immediate free access for everyone upon online publication of accepted manuscripts. Open Access articles will be so indicated on eToCs (our electronic content alerts [sign up at http://www.plantphysiol.org/cgi/alerts/etoc and http://www.plantcell.org/cgi/alerts/etoc]), on the print and online table of contents, and on the opening page of the article itself, both online and in print. In this, the inaugural issue for the Open Access option, several groups of your colleagues have selected the Open Access option and in doing so invested in the increased visibility and impact of their published work. Author notes www.plantphysiol.org/cgi/doi/10.1104/pp.104.900180. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Grennan, Aleel K.

doi: 10.1104/pp.104.900179pmid: 16339803

THIS MONTH'S SELECTION This month's selection for High Impact is “The putative Arabidopsis Arp2/3 complex controls leaf cell morphogenesis” by Li et al. (2003), which appeared in our August 2003 issue and as of November 2005 had been cited 44 times (Thompson ISI Web of Science, http://www.isinet.com). The research presented in the article has furthered the understanding of the dynamic regulation of the actin cytoskeleton and its involvement in cell shape determination. BACKGROUND The actin cytoskeleton is involved many cellular processes, including signal transduction, organelle positioning, and determination of cell shape. Actin filaments have also been shown to accumulate in the leading edge of tip-growing cells, such as root hairs and pollen tubes, as well as diffuse growing cells like epidermal pavement cells. Actin has no enzymatic activity; instead, it provides a dynamic structure for interaction between other proteins. The actin cytoskeleton needs to be rapidly assembled and disassembled in order for it to provide this scaffolding where and when it is needed. One way this process is controlled and the dynamic nature of actin is maintained is by actin-nucleating proteins such as the actin-related proteins (ARPs) like Arp2/3. Arp2/3 is a complex of seven subunits and is found in all eukaryotic kingdoms. The Arp2/3 complex's ability to initiate actin polymerization is achieved by binding an actin filament and nucleating a “daughter” filament from the side. Arp2/3 was originally discovered in amoeba cells (Machesky et al., 1994) and has been intensively studied in a variety of diverse systems, including yeast (Saccharomyces cerevisiae), Caenorhabditis elegans, Drosophila, and mammals (for review, see Deeks and Hussey, 2003; Mathur, 2005). The first plant homolog/subunit (ARP2) was cloned in 1999 by Klahre and Chua (1999), and the remaining six putative genes were genetically identified and described in the article by Li et al. (2003). Concurrently, the labs of Dan Szymanski (Le et al., 2003) and Martin Hulskamp (Mathur et al., 2003a, 2003b) independently identified subunits of the Arp2/3 complex and confirmed the importance of this complex in cell morphogenesis. WHAT WAS SHOWN Li et al. (2003) identified subunits of Arabidopsis (Arabidopsis thaliana) Arp2/3 from the Arabidopsis genome database based on similarity to yeast Arp2/3 subunits. T-DNA mutants were identified for three of the subunit genes, arp2, arpc5, and arp3, allowing functional characterization. Although no obvious changes to the whole-plant morphology or to tip-growing cells were observed in these mutants, inactivation of any of the three genes produced identical defects in the development of epidermal pavement cells and leaf trichomes. Pavement cells, nonspecialized epidermal cells, were observed to have less wavy margins in the mutants than wild type, and closer investigation of these cells demonstrated reduced height of the lobes with no change in the neck width. The aberrant trichomes of the Arp2/3 subunit mutants were similar to what is observed in the “distorted” class of mutants in which the trichomes have reduced branch length and are stunted. The application of drugs disrupting actin cytoskeleton phenocopy these mutants, leading to the hypothesis that the mutant phenotypes could be defects in the formation of a fine actin network. Investigation of actin cytoskeleton structure in leaf epidermal cells with transiently expressed green fluorescent protein-tagged actin-binding domain of mouse talin revealed no obvious defects in the actin structure until the pavement cells started expanding. At that stage, a change in the localization of actin patches in the lobes occurred. Patches of diffuse F-actin, found at the expanding lobes in wild-type cells, were shown to be more evenly distributed throughout the cell edges rather than in the lobes of the mutant cells. This work demonstrates the role of the Arp2/3 complex in cell morphogenesis in the formation of actin patches involved in diffuse polar growth in plants. The authors also suggest a novel function for Arp2/3, the possibility of cell-specific actin polymerization, and the regulation of actin spatial distribution. THE IMPACT The identification of the Arp2/3 complex in plants lead to the next question: How is it regulated? By itself the complex is inactive; nucleation-promoting factors are required for its activation. In animal systems, the regulation of the Arp2/3 complex has been studied and was determined to involve WASP (Wiskott-Aldrich syndrome protein)/Scar (suppressor of cAMP receptor defects)/WAVE (WASP family verprolin homologous protein) family members. Utilizing a variety of approaches, Arp2/3 regulation was explored in numerous subsequent papers, including those by Frank et al. (2004) and Basu et al. (2005), leading to the identification of plant homologs of the WASP/Scar/WAVE regulatory complex. In Arabidopsis and maize (Zea mays), a family of Scar/WAVE-related proteins, AtSCAR1 to 4 and ZmSCAR1, were identified by Frank et al. (2004) and were shown to interact with BRICK1, a WAVE-binding protein believed to be involved in Scar/WAVE complex regulation. AtSCAR was found to activate the bovine Arp2/3 complex in vitro, supporting the idea that they have a similar function of activating Arp2/3 in plants. Another piece of the regulation puzzle was uncovered by Basu et al. (2005) when the gene responsible for the Arabidopsis mutant DISTORTED3 (DIS3) was identified as SCAR2, a member of the Scar/WAVE complex. DIS3/SCAR2 was shown to function within a WAVE-Apr2/3 complex in vivo and in a polymerization assay with bovine Arp2/3 to activate the complex, inducing actin filament nucleation and branching activity. The identification of the Arabidopsis Arp2/3 complex also aided in the further understanding of the establishment of zygote polarity in a paper by Hable and Kropf (2005). Default polarity in fertilized zygotes of fucoid brown algae is established at the site of sperm entry, where an F-actin patch forms. However, depending upon spatial cues from the local environment, this axis can be re-established by actin patch disassembly at the initial site and reassembly at the new axis, altering the polarity. To understand the polymerization of the new actin arrays in the zygotes, the relationship between actin and Arp2/3 complex during actin-dependent processes was observed using antibodies to the C-terminal peptide of brown algae ARP2 ortholog. Both actin and Arp2/3 were observed colocalized in a patch at the rhizoid pole as well as around the nucleus, indicating that they are associated together, suggesting that Arp2/3 is involved in nucleating dynamic actin arrays that function in polarity establishment, as was shown by Li et al. (2003). SUMMARY The 2003 article by Li et al. identified subunits of the plant Arp2/3 complex and functionally characterized three of these subunits. The foundation laid by this article aided in the identification of Arp2/3 regulatory proteins, which suggests that plant Arp2/3 may be regulated in a similar manner to animal systems. In addition, this article furthered the understanding of polarity establishment in fucoid zygotes. LITERATURE CITED Basu D, Le J, El-Essal SE, Huang S, Zhang C, Mallery EL, Koliantz G, Staiger CJ, Szymanski DB ( 2005 ) DISTORTED3/SCAR2 is a putative Arabidopsis WAVE complex subunit that activated the Arp2/3 complex and is required for epidermal morphogenesis. Plant Cell 17 : 502 –524 Deeks MJ, Hussey PJ ( 2003 ) Arp2/3 and ‘the shape of things to come’. Curr Opin Plant Biol 6 : 561 –567 Frank M, Egile C, Dyachok J, Djakovic S, Nolasco M, Li R, Smith LC ( 2004 ) Activation of Arp2/3 complex-dependent actin polymerization by plant proteins distantly related to Scar/WAVE. Proc Natl Acad Sci USA 101 : 16379 –16384 Hable WE, Kropf DL ( 2005 ) The Arp2/3 complex nucleates actin arrays during zygote polarity establishment and growth. Cell Motil Cytoskeleton 61 : 9 –20 Klahre U, Chua NH ( 1999 ) The Arabidopsis actin-related protein 2 (AtARP2) promoter directs expression in xylem precursor cells and pollen. Plant Mol Biol 41 : 65 –73 Le J, El-Assal SE, Basu D, Saad ME, Szymanski DB ( 2003 ) Requirements for Arabidopsis ATARP2 and ATARP3 during epidermal development. Curr Biol 13 : 1341 –1347 Li S, Blanchoin L, Yang Z, Lord EM ( 2003 ) The putative Arabidopsis Arp2/3 complex controls lead cell morphogenesis. Plant Physiol 132 : 2034 –2044 Machesky LM, Atkinson SJ, Ampe C, Vandekerckhove J, Pollard TD ( 1994 ) Purification of a cortical complex containing two unconventional actins from Acanthamoeba by affinity chromatography on profilin-agarose. J Cell Biol 127 : 107 –115 Mathur J ( 2005 ) The ARP2/3 complex: giving plant cells a leading edge. Bioessays 27 : 377 –387 Mathur J, Mathur N, Kernebeck B, Hulskamp M ( 2003 a) Mutations in actin-related proteins 2 and 3 affect cell shape development in Arabidopsis. Plant Cell 15 : 1632 –1645 Mathur J, Mathur N, Kirik V, Kernebeck B, Srinivas BP, Hulskamp M ( 2003 b) Arabidopsis CROOKED encodes for the smallest subunit of the ARP2/3 complex and controls cell shape by region specific fine F-actin formation. Development 130 : 3137 –3146 Author notes www.plantphysiol.org/cgi/doi/10.1104/pp.104.900179. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Meagher, Richard B.; Deal, Roger B.; Kandasamy, Muthugapatti K.; McKinney, Elizabeth C.

doi: 10.1104/pp.105.072447pmid: 16339804

Complex regulatory networks control cell fate and the development of organs and tissues in multicellular organisms. But what mechanisms initiate the necessary global changes in patterns of gene expression? What regulates the regulators of organismal development? The nuclear actin-related proteins (ARPs) participate in macromolecular chromatin-remodeling machines that regulate the transcription of developmentally important genes. In Arabidopsis (Arabidopsis thaliana), ARP4, ARP6, and ARP7, which are predominantly localized in the nucleus, participate in the regulation of several pathways affecting cell proliferation and organ development. The diverse plant phenotypes resulting from deficiencies in these nuclear ARPs include reduced cell size or numbers, photoperiod-dependent and -independent early flowering, delayed floral senescence, altered leaf, stem, and flower organ morphology, embryo lethality, and an assortment of male and female reproductive defects. A working hypothesis emerging from these and other plant and animal data is that diverse isoforms of nuclear ARP-containing chromatin-modifying complexes exert epigenetic control over global regulators of multicellular development. In support of this hypothesis, we herein examine nuclear ARP phylogeny, the chromatin-remodeling activities of ARP-containing complexes that lead to epigenetic control of gene expression, the expanding developmental roles assigned to several putative plant ARP-containing complexes, as well as the evidence that a large number of ARP complex isoforms may have evolved in concert with the significant demands of multicellular development. The ARPs share limited sequence identity (15%–60%) with conventional actins, but they appear to maintain the actin fold, a nucleotide-binding pocket and hinge region that enables a conformational change in actin. Eight to 10 ancient classes of ARPs are found in most eukaryotes that have been examined, and all appear to participate in protein complexes (McKinney et al., 2002; Blessing et al., 2004; Kandasamy et al., 2004). Several of the more divergent ARPs are homologs of yeast (Saccharomyces cerevisiae) Arp4, 5, 6, and 8, which are found primarily in the nucleus. This location distinguishes the nuclear ARPs from actin or Arp2 and Arp3, which have been found in the nucleus, but are primarily concentrated in the cytoplasm. None of the nuclear ARPs are believed to form polymers such as actin microfilaments or the short filaments formed from ARP1 in the centractin complex, which contains both ARP1 and ARP10. Nuclear ARPs act as essential subunits of macromolecular machines that remodel chromatin structure, and their only demonstrated functions are within such complexes. Specifically, ARP-containing complexes are involved in nucleosome phasing and movement, histone acetylation, and exchange of histone subunit isovariants within nucleosomes. The chromatin-modifying activities of these complexes can serve the basal regulatory function of reinforcing or alleviating the nucleosomal suppression of transcription that affects most genes (Yuan et al., 2005). However, they may also exert more precise epigenetic control over development via the particular activities of ARP-containing complexes on the transcription of a small subset of regulatory genes. The activities of nuclear ARP complexes on genes that direct global changes in cell proliferation and ontogeny of organs and tissues are the subjects of our working hypothesis and the major focus of this Update article. NUCLEAR ARP PHYLOGENY The majority of genes encoding the nuclear ARPs evolved from a common ancestral actin gene prior to the divergence of the four eukaryotic kingdoms, yet the nuclear ARP proteins share 15% to 40% amino acid sequence identity with conventional actin (Blessing et al., 2004). Because the initial characterization of the entire family of ARPs was performed in yeast, most nuclear ARPs in other organisms are named relative to the yeast nuclear ARPs (Arp4, 5, 6, 7, 8, and 9; Poch and Winsor, 1997). The increasing numbers in this ARP nomenclature represent increasing divergence from conventional actin, with ARP4 being most closely related (approximately 40% amino acid identity to actin) and ARP9 the most divergent (approximately 15% identity; McKinney et al., 2002; Kandasamy et al., 2004). While the nuclear ARPs share the backbone of actin sequence and are predicted to retain the actin fold, they contain several, sometimes large, insertions and small deletions that distinguish each ARP clade. The phylogenetic relationships of the yeast nuclear ARPs to those in humans, Arabidopsis, and rice (Oryza sativa) are illustrated in Figure 1 Figure 1. Open in new tabDownload slide Phylogenetic relationships among the nuclear ARPs. The phylogenetic relationships among the nuclear ARPs from yeast (purple), Arabidopsis (dark green), rice (light green), and human (red) are shown. The four clades of nuclear ARPs that are clearly conserved in all eukaryotes and named after the closest yeast ARP homolog are indicated (uppercase blue font). ClustalW was used to align the sequences. The phylogram presented used Parsimony to create the tree's topography based on sequence similarity and a heuristic search with 100 replicates to generate bootstrap values within PAUP (Swofford, 1998). Bootstrap values below 50% are not indicated. A tree with very similar branching was obtained using the neighbor-joining tree-building method. Figure 1. Open in new tabDownload slide Phylogenetic relationships among the nuclear ARPs. The phylogenetic relationships among the nuclear ARPs from yeast (purple), Arabidopsis (dark green), rice (light green), and human (red) are shown. The four clades of nuclear ARPs that are clearly conserved in all eukaryotes and named after the closest yeast ARP homolog are indicated (uppercase blue font). ClustalW was used to align the sequences. The phylogram presented used Parsimony to create the tree's topography based on sequence similarity and a heuristic search with 100 replicates to generate bootstrap values within PAUP (Swofford, 1998). Bootstrap values below 50% are not indicated. A tree with very similar branching was obtained using the neighbor-joining tree-building method. . Humans contain clear homologs of yeast ARP4 (HsaBAF53), Arp5 (HsARP5), Arp6 (HsARP6x), and Arp8 (HsARP8). Arabidopsis and rice ARP4, ARP5, ARP6, and ARP9 are clear plant homologs of ARP4 (Baf53), Arp5, Arp6, and Arp8, respectively, in animals and yeast. Thus, these four clades of ARPs (labeled in Fig. 1) certainly predate the divergence of the three kingdoms represented from a common ancestor. We have shown that Arabidopsis ARP4, ARP6, and ARP7 are localized to the interphase nucleus (Kandasamy et al., 2003; Deal et al., 2005), and by their phylogenetic position in the sequence tree the plant homologs of yeast ARP5, ARP8, and ARP9 are predicted to encode nuclear ARPs (Kandasamy et al., 2004). The six phylogenetically paired sets of plant ARP proteins from Arabidopsis and rice are 52% to 87% identical at the amino acid sequence level. Because these two angiosperms have not shared a common ancestor for 200 million years, it is likely that homologs to these six ARPs are reasonably well conserved and may be universally present in all higher plants (McKinney et al., 2002; Kandasamy et al., 2004). The phylogenetic relationships of Arabidopsis ARP7 and ARP8 to ARPs in other eukaryotic kingdoms are not clear, and hence, have an orphaned status (Blessing et al., 2004). THE ACTIVITIES OF NUCLEAR ARPS AND ARP-CONTAINING CHROMATIN-MODIFYING COMPLEXES All of the nuclear ARPs that have been examined to date are constituents of either ATP-dependent nucleosome-remodeling (NR) complexes, histone acetyltransferase (HAT) complexes, or histone variant exchange (HVE) complexes, all of which are involved in the modification of chromatin structure (Olave et al., 2002; Mizuguchi et al., 2004). The ATP-dependent NR and HVE complexes all contain a DNA-dependent ATPase and use the energy of ATP hydrolysis to drive the repositioning of nucleosomes on DNA or the exchange of one histone type for another within nucleosomes. The NR and HVE complexes share other similar subunits, and there are mechanistic arguments that all NR complexes can carry out HVE activities (Flaus and Owen-Hughes, 2004). In contrast, the HAT complexes transfer an acetyl group onto histone N-terminal tails within nucleosomes, thereby altering chromatin structure indirectly, but they do not contain a DNA-dependent ATPase. The modifications that HAT and related complexes perform on the termini of histone polypeptides generate a histone code that can be decoded to define active and inactive regions of chromatin (Margueron et al., 2005). Surprisingly, the majority of the well-characterized chromatin-modifying complexes include monomeric actin as a subunit in addition to one or more ARPs (Olave et al., 2002). A number of important themes have emerged regarding the biochemical functions of the ARPs and actins in NR, HVE, and HAT complexes, but no single function stands out as universally conserved in all complexes (Blessing et al., 2004). Recalling that ARPs belong to the actin superfamily of proteins that includes cytoskeletal actins, heat shock proteins, ATPases, and sugar kinases, most models for ARP and actin function center on the actin fold and hinge region that impart the ability to shift between distinct conformational states upon nucleotide binding, ATP hydrolysis, and ATP/ADP exchange (Sunada et al., 2005). Through this change in conformation, the ARP and actin subunits may act as molecular switches controlling the assembly, stability, and/or the activity of these machines. However, to date, only ARP2 and ARP4 are known to bind and hydrolyze nucleotides (Sunada et al., 2005). Thus, ARPs may possess some of their own ATPase activity, but they are more likely to act indirectly by activating the DNA-dependent ATPase subunit found in NR and HVE complexes, given that mutations in the predicted ATPase domains of yeast ARP7 and ARP9 did not alter the ATPase activity of the RSC (remodel the structure of chromatin) or SWI/SNF (mating-type switch/Suc nonfermenting)-related complexes (Cairns et al., 1998; Szerlong et al., 2003). In addition, yeast ARP4 and ARP8 can bind histones directly, and thus, may recruit chromatin complexes directly to nucleosomes (Shen et al., 2003). Finally, it has been proposed that ARPs and actins may connect one chromatin complex to another to create higher order chromatin complexes, in this way acting similarly to conventional actin, but in the polymerization of higher order structures instead of filaments (Blessing et al., 2004). The activities of several nuclear ARP-containing chromatin-remodeling complexes are known. In yeast, for example, ARP4 is an essential gene and ARP4 protein is a component of the NUA4 (nucleosomal acetyltransferase of H4, HAT), the INO80 (inositol requiring) NR, and the SWR1 HVE complexes (Galarneau et al., 2000; Krogan et al., 2004; Mizuguchi et al., 2004). The 1.3-megadalton NUA4 complex contains 11 subunits, including ARP4 and conventional actin, and it is known to primarily acetylate histone H4. In temperature-sensitive arp4 mutants the NUA4 complex is absent (Galarneau et al., 2000), indicating that ARP4 is required to assemble or stabilize the complex. These same mutants display defects in the transcription of certain target genes at the restrictive temperature, coincident with changes in the chromatin structure around those genes (Jiang and Stillman, 1996; Harata et al., 2002). In addition, ARP4 has been shown to bind all four core histones in vitro (Harata et al., 1999). Thus, ARP4 may serve to recruit components and/or stabilize complexes and could also provide a targeting function by interacting with core histones. In addition to yeast, ARP4 orthologs are also found in at least three different mammalian ATP-dependent NR complexes, in at least one remodeling complex in Drosophila, and in one HAT complex in mammals (Olave et al., 2002). In addition to ARP4, the yeast INO80 NR complex also contains ARP5, ARP8, monomeric actin, and seven other subunits as shown in Table I Table I. Predicted Arabidopsis subunit isovariant diversity for three chromatin-remodeling complexes Yeast Complex-Protein Subunitsa . Genes in Arabidopsis Encoding Isovariantsbc . Accession Nos. of Arabidopsis Genes . SWI/SNF     Arp7 (YPR034W) 1 (ARP7d) At3g60830b     Arp9 (YMR033W) N.D.e     Snf2 (YOR290C) 4 At5g19310, At3g06010, At2g28290, At3g06400, At5g18620b     Snf5 (YBR289W) 1 At3g17590b     Snf12 (YNR023W) 2 At5g14170, At3g01890b     Swi1 (YPL016W) 2 At1g79000, At1g16710b     Swi3 (YJL176C) 4 At2g33610, At1g21700, At4g34430, At2g47620b     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Rtt102 (YGR275W), Snf11 (YDR073W), Snf6 (YHL025W), N.D.e SWR1/SWR-C     Arp4 (YJL081C) 1 (ARP4) At1g18450     Arp6 (YLR085C) 1 (ARP6) At3g33520     Actin ACT1 (YFL039C) 8f     Swr1 (YDR334W) Snf2 homolog 4 At3g12810, At3g57300, At3g06400, At5g19310b     Swc3 (YAL011W) 5 At4g23800, At1g15340, At3g16000, At4g11080, At1g65470b     Swc5 (YBR231C) 3 At3g28730, At1g08600, At1g58025b     Vps72 (YDR485C) 5 At4g23800, At5g22650, At2g06210, At4g11080, At5g18620b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Swc4 (YGR002C) 1 At2g47210c     Vps71 (YML041C) 1 At5g37055c     Yaf9 (YNL107W) 2 At5g45600, At2g18000c     Swc7 (YLR385C) N.D.e INO80     Arp4 (YJL081C) 1 (ARP4) At1g18450b     Arp5 (YNL059C) 1 (ARP5) At3g12380b     Arp8 (YOR141C) 1 (ARP9) At5g43500b     Actin ACT1 (YFL039C) 8f     Ino80 (YGL150C) snf2 homolog 3 At3g57300, At3g12810, At5g66750b     Nhp10 (YDL002C) 7 At4g23800, At4g11080, At1g20693, At3g51880, At1g20696, At3g28730, At5g23420b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Ies1 (YFL013C), Ies3 (YLR052W) N.D.e Yeast Complex-Protein Subunitsa . Genes in Arabidopsis Encoding Isovariantsbc . Accession Nos. of Arabidopsis Genes . SWI/SNF     Arp7 (YPR034W) 1 (ARP7d) At3g60830b     Arp9 (YMR033W) N.D.e     Snf2 (YOR290C) 4 At5g19310, At3g06010, At2g28290, At3g06400, At5g18620b     Snf5 (YBR289W) 1 At3g17590b     Snf12 (YNR023W) 2 At5g14170, At3g01890b     Swi1 (YPL016W) 2 At1g79000, At1g16710b     Swi3 (YJL176C) 4 At2g33610, At1g21700, At4g34430, At2g47620b     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Rtt102 (YGR275W), Snf11 (YDR073W), Snf6 (YHL025W), N.D.e SWR1/SWR-C     Arp4 (YJL081C) 1 (ARP4) At1g18450     Arp6 (YLR085C) 1 (ARP6) At3g33520     Actin ACT1 (YFL039C) 8f     Swr1 (YDR334W) Snf2 homolog 4 At3g12810, At3g57300, At3g06400, At5g19310b     Swc3 (YAL011W) 5 At4g23800, At1g15340, At3g16000, At4g11080, At1g65470b     Swc5 (YBR231C) 3 At3g28730, At1g08600, At1g58025b     Vps72 (YDR485C) 5 At4g23800, At5g22650, At2g06210, At4g11080, At5g18620b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Swc4 (YGR002C) 1 At2g47210c     Vps71 (YML041C) 1 At5g37055c     Yaf9 (YNL107W) 2 At5g45600, At2g18000c     Swc7 (YLR385C) N.D.e INO80     Arp4 (YJL081C) 1 (ARP4) At1g18450b     Arp5 (YNL059C) 1 (ARP5) At3g12380b     Arp8 (YOR141C) 1 (ARP9) At5g43500b     Actin ACT1 (YFL039C) 8f     Ino80 (YGL150C) snf2 homolog 3 At3g57300, At3g12810, At5g66750b     Nhp10 (YDL002C) 7 At4g23800, At4g11080, At1g20693, At3g51880, At1g20696, At3g28730, At5g23420b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Ies1 (YFL013C), Ies3 (YLR052W) N.D.e a Systematic yeast gene name (dp.yeastgenome.org). b The Plant Chromatin Database (www.chromdb.org) and cThe Arabidopsis Information Resource were searched for various protein homologs of the yeast proteins. The phylogeny of related gene sequences was examined to make an estimate of the number of hits in the databases with scores equal to or less than (more significant than) E = 0.001 (the expectation of finding two sequences with a given amount of similarity by chance). An E value of 0.001 is considered statistically significant in large databases (Gerstein, 1998; Pearson, 2000). d Arabidopsis has no unambiguous homolog of yeast ARP7, but the orphaned plant ARP7 could be its immediate ortholog (Kandasamy et al., 2005b). In animals, ARP4 homologs have been found in most SWI/SNF complexes and the same could be true in plants. e None detected based on a homology score of less than 0.001. f The Arabidopsis genome encodes eight isovariants of conventional actin that were not included in calculations of isoform diversity. Open in new tab Table I. Predicted Arabidopsis subunit isovariant diversity for three chromatin-remodeling complexes Yeast Complex-Protein Subunitsa . Genes in Arabidopsis Encoding Isovariantsbc . Accession Nos. of Arabidopsis Genes . SWI/SNF     Arp7 (YPR034W) 1 (ARP7d) At3g60830b     Arp9 (YMR033W) N.D.e     Snf2 (YOR290C) 4 At5g19310, At3g06010, At2g28290, At3g06400, At5g18620b     Snf5 (YBR289W) 1 At3g17590b     Snf12 (YNR023W) 2 At5g14170, At3g01890b     Swi1 (YPL016W) 2 At1g79000, At1g16710b     Swi3 (YJL176C) 4 At2g33610, At1g21700, At4g34430, At2g47620b     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Rtt102 (YGR275W), Snf11 (YDR073W), Snf6 (YHL025W), N.D.e SWR1/SWR-C     Arp4 (YJL081C) 1 (ARP4) At1g18450     Arp6 (YLR085C) 1 (ARP6) At3g33520     Actin ACT1 (YFL039C) 8f     Swr1 (YDR334W) Snf2 homolog 4 At3g12810, At3g57300, At3g06400, At5g19310b     Swc3 (YAL011W) 5 At4g23800, At1g15340, At3g16000, At4g11080, At1g65470b     Swc5 (YBR231C) 3 At3g28730, At1g08600, At1g58025b     Vps72 (YDR485C) 5 At4g23800, At5g22650, At2g06210, At4g11080, At5g18620b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Swc4 (YGR002C) 1 At2g47210c     Vps71 (YML041C) 1 At5g37055c     Yaf9 (YNL107W) 2 At5g45600, At2g18000c     Swc7 (YLR385C) N.D.e INO80     Arp4 (YJL081C) 1 (ARP4) At1g18450b     Arp5 (YNL059C) 1 (ARP5) At3g12380b     Arp8 (YOR141C) 1 (ARP9) At5g43500b     Actin ACT1 (YFL039C) 8f     Ino80 (YGL150C) snf2 homolog 3 At3g57300, At3g12810, At5g66750b     Nhp10 (YDL002C) 7 At4g23800, At4g11080, At1g20693, At3g51880, At1g20696, At3g28730, At5g23420b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Ies1 (YFL013C), Ies3 (YLR052W) N.D.e Yeast Complex-Protein Subunitsa . Genes in Arabidopsis Encoding Isovariantsbc . Accession Nos. of Arabidopsis Genes . SWI/SNF     Arp7 (YPR034W) 1 (ARP7d) At3g60830b     Arp9 (YMR033W) N.D.e     Snf2 (YOR290C) 4 At5g19310, At3g06010, At2g28290, At3g06400, At5g18620b     Snf5 (YBR289W) 1 At3g17590b     Snf12 (YNR023W) 2 At5g14170, At3g01890b     Swi1 (YPL016W) 2 At1g79000, At1g16710b     Swi3 (YJL176C) 4 At2g33610, At1g21700, At4g34430, At2g47620b     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Rtt102 (YGR275W), Snf11 (YDR073W), Snf6 (YHL025W), N.D.e SWR1/SWR-C     Arp4 (YJL081C) 1 (ARP4) At1g18450     Arp6 (YLR085C) 1 (ARP6) At3g33520     Actin ACT1 (YFL039C) 8f     Swr1 (YDR334W) Snf2 homolog 4 At3g12810, At3g57300, At3g06400, At5g19310b     Swc3 (YAL011W) 5 At4g23800, At1g15340, At3g16000, At4g11080, At1g65470b     Swc5 (YBR231C) 3 At3g28730, At1g08600, At1g58025b     Vps72 (YDR485C) 5 At4g23800, At5g22650, At2g06210, At4g11080, At5g18620b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Swc4 (YGR002C) 1 At2g47210c     Vps71 (YML041C) 1 At5g37055c     Yaf9 (YNL107W) 2 At5g45600, At2g18000c     Swc7 (YLR385C) N.D.e INO80     Arp4 (YJL081C) 1 (ARP4) At1g18450b     Arp5 (YNL059C) 1 (ARP5) At3g12380b     Arp8 (YOR141C) 1 (ARP9) At5g43500b     Actin ACT1 (YFL039C) 8f     Ino80 (YGL150C) snf2 homolog 3 At3g57300, At3g12810, At5g66750b     Nhp10 (YDL002C) 7 At4g23800, At4g11080, At1g20693, At3g51880, At1g20696, At3g28730, At5g23420b     Rvb1 (YDR190C) 1 At5g22330c     Rvb2 (YPL235W) 2 At5g67630, At3g49830c     Taf14 (YPL129W) 2 At2g18000, At5g45600c     Ies1 (YFL013C), Ies3 (YLR052W) N.D.e a Systematic yeast gene name (dp.yeastgenome.org). b The Plant Chromatin Database (www.chromdb.org) and cThe Arabidopsis Information Resource were searched for various protein homologs of the yeast proteins. The phylogeny of related gene sequences was examined to make an estimate of the number of hits in the databases with scores equal to or less than (more significant than) E = 0.001 (the expectation of finding two sequences with a given amount of similarity by chance). An E value of 0.001 is considered statistically significant in large databases (Gerstein, 1998; Pearson, 2000). d Arabidopsis has no unambiguous homolog of yeast ARP7, but the orphaned plant ARP7 could be its immediate ortholog (Kandasamy et al., 2005b). In animals, ARP4 homologs have been found in most SWI/SNF complexes and the same could be true in plants. e None detected based on a homology score of less than 0.001. f The Arabidopsis genome encodes eight isovariants of conventional actin that were not included in calculations of isoform diversity. Open in new tab . The loss-of-function mutations for the Ino80 subunit, a DNA-dependent ATPase, are defective in transcribing the genes involved in inositol biosynthesis and are hypersensitive to DNA-damaging agents (Ebbert et al., 1999; Shen et al., 2003), suggesting a role for the INO80 complex not only in transcriptional control, but also in DNA damage repair. Deletion mutants lacking the ARP5 and ARP8 genes also display the ino80− inositol requiring phenotype, indicating that these ARP proteins are critical to the function of the complex. INO80 complexes purified from arp5Δ or arp8Δ yeast strains still retain most other subunits, but are deficient in Ino80 ATPase activity, DNA binding, and NR functions. In addition, the complex purified from arp8Δ cells also lacks ARP4 and actin, indicating that ARP8 is needed to recruit these components to the complex (Shen et al., 2003). The finding that ARP8 also binds histones H3 and H4 in vitro (Shen et al., 2003) suggests that, in addition to stimulating the ATPase activity of the Ino80 subunit, the ARPs may also serve as points of contact between the complex and chromatin. For some time, our knowledge of ARP6 function was limited to the observation that this protein was localized to the nucleus in both yeast and Drosophila. However, two groups have recently isolated the yeast SWR1 complex and found that it contains not only ARP6, but also ARP4, actin, and 10 other subunits as listed in Table I (Krogan et al., 2004; Mizuguchi et al., 2004). This complex carries out a new class of chromatin-remodeling activity: the exchange of one histone variant for another within the nucleosome. The SWR1 HVE complex replaces histone H2A with the variant H2A.Z at specific chromosomal locations (Krogan et al., 2004; Mizuguchi et al., 2004). The H2A.Z histone variant is conserved among eukaryotes, and thus the HVE activity of ARP6-containing complexes is probably ancient and present in most eukaryotes. Histone H2A.Z-containing nucleosomes act partly to antagonize the spread of silent heterochromatin into euchromatic regions, but they also have important heterochromatic functions (Meneghini et al., 2003; Dryhurst et al., 2004). Yeast SWI/SNF and RSC are two related ATP-dependent NR complexes containing 11 and 15 subunits, respectively, including ARP7 and ARP9. Table I shows the composition of the yeast SWI/SNF complex. Unlike the complexes described above, neither SWI/SNF nor RSC contain monomeric actin. The SWI/SNF complex was identified independently in genetic screens for genes involved in mating-type switching and Suc fermentation, and the RSC complex was later isolated based on homology to SWI/SNF complex components. Yeast strains lacking ARP7 or ARP9 show typical swi/snf− phenotypes, indicating that these proteins play an essential role in the function of SWI/SNF. In addition to the swi/snf− phenotype, mutations in ARP7 or ARP9 also lead to other transcriptional defects not related to SWI/SNF function, indicating that RSC plays a role in transcriptional regulation as well (Cairns et al., 1998). Surprisingly, RSC complexes isolated from arp7Δ/arp9Δ cells are fully intact and retain the ability to remodel nucleosomes in vitro. However, a screen for suppressors of arp7 and arp9 mutations identified the transcription factor Nhp6, which interacts physically with RSC and enhances the activity of the complex in vitro (Szerlong et al., 2003). This finding suggests that ARP7 and ARP9 serve to connect the RSC complex to interacting proteins or other complexes, allowing functionality in vivo. The animal SWI/SNF NR complexes that have been characterized contain homologs of yeast ARP4 and actin instead of homologs of ARP7 or ARP9 found in the yeast complex (Olave et al., 2002). This suggests at least a functional relationship if not an undetected phylogenetic relationship exists between these more divergent ARPs and the reasonably well conserved ARP4. In summary, nuclear ARP functions are believed to be central to the activity of the majority of the known chromatin-remodeling complexes and to NR, HAT, and HVE complexes in particular. The six yeast nuclear ARPs (ARP4–9) are all found in these three classes of complexes. Thus, the subunit compositions of the yeast NR, HAT, and HVE complexes, such as those examples presented in Table I, are likely to be predictive of the basic subunit compositions of ARP-containing complexes in other kingdoms. THE ROLES OF NUCLEAR ARPS IN PLANT DEVELOPMENT During the last few years, chromatin remodeling has been directly linked to numerous pathways of plant development (Wagner, 2003; Gendrel and Colot, 2005). Although it is known that ARPs are essential components of all SWI/SNF and most other chromatin-modifying complexes in yeast, there were until recently no data directly linking ARP functions to plant development. Therefore, it was exciting to find that knocking down or knocking out the Arabidopsis nuclear ARPs ARP4, ARP6, and ARP7 was associated with defects in several developmental pathways. A sampling of the developmental abnormalities associated with deficiencies in these three ARPs is illustrated in Figure 2 Figure 2. Open in new tabDownload slide Deficiencies in Arabidopsis ARPs ARP4, ARP6, and ARP7 alter cell proliferation and plant development. A, Wild-type Arabidopsis plant. ARP4, 6, and 7 are localized to the interphase nucleus. In mitotic cells lacking intact nuclear membrane (*) they are dispersed throughout the cytoplasm. For example, the insert depicts the subcellular distribution of ARP4 in wild-type root apical cells. B, Knockout arp7-1 mutation causes homozygous embryo lethality. Mutant embryos (right) are arrested at the heart (top) or torpedo (bottom) stage of development. A wild-type embryo at the cotyledon stage from the same silique (fruit) is shown at the left. C, Knocking out or knocking down ARP4, 6, and 7 causes poor seed germination or early seedling development, depending upon the level of reduction in the expression of these proteins. Dwarf ARP4RNAi seedlings (10-d-old) with stunted cotyledons are shown with three wild-type seedlings (arrow heads) in the top section. Two-week-old normal wild-type (WT) and arrested ARP7RNAi seedlings from a strong line showing more than 85% reduction in ARP7 expression are shown in the bottom section. D, Deficiencies in the expression of ARP4, 6, and 7 affect leaf size, number, and/or morphology. The arp6-1 knockout mutants produce narrow and highly serrated leaves under short-day photoperiod (bottom section). All the leaves from an adult ARP4RNAi plant are compared with those from wild type (top section). E, Scanning electron microscopy of the smaller leaves of ARP7RNAi plants reveals similar number, but smaller sized leaf epidermal cells. For comparison, knocking out ARP6 in the arp6-1 mutant results in dwarfed leaves with a smaller number of normally sized cells (not shown). F, A knockdown of ARP7 in RNAi plants severely affects root growth. The retarded roots of RNAi plants have highly reduced cell elongation zone compared to wild type (WT). G, Deficiencies in ARP4 and ARP6 expression affect flowering time. ARP4RNAi plants flower early only under long-day (LD) conditions (bottom section), whereas the arp6-1 mutants flower early both under long- and short-day (SD) conditions (top section). Thus, ARP4 and ARP6 are involved in photoperiod-dependent and photoperiod-independent flowering pathways, respectively. H, A knockdown in ARP4 and ARP7 in the RNAi plants causes delayed senescence and abscission of floral organs. Wild-type inflorescences each contain five or six flowers with intact sepals and petals, whereas the RNAi plants have 15 to 20 flowers in each inflorescence with intact sepals and petals. I, A strong reduction in ARP4 and ARP7 expression affects stamen development and male fertility. The aberrant heart-shaped anthers in the arp4-1 mutant contain less pollen than wild type. J, Deficiencies in ARP4 and ARP6 expression affect female fertility. Defects in pistil development and pollination results in stunted fruits with reduced seed set compared to wild type. K, Knocking out or knocking down ARP4, 6, and 7 expression affect flower morphology and/or organ number. Figure 2. Open in new tabDownload slide Deficiencies in Arabidopsis ARPs ARP4, ARP6, and ARP7 alter cell proliferation and plant development. A, Wild-type Arabidopsis plant. ARP4, 6, and 7 are localized to the interphase nucleus. In mitotic cells lacking intact nuclear membrane (*) they are dispersed throughout the cytoplasm. For example, the insert depicts the subcellular distribution of ARP4 in wild-type root apical cells. B, Knockout arp7-1 mutation causes homozygous embryo lethality. Mutant embryos (right) are arrested at the heart (top) or torpedo (bottom) stage of development. A wild-type embryo at the cotyledon stage from the same silique (fruit) is shown at the left. C, Knocking out or knocking down ARP4, 6, and 7 causes poor seed germination or early seedling development, depending upon the level of reduction in the expression of these proteins. Dwarf ARP4RNAi seedlings (10-d-old) with stunted cotyledons are shown with three wild-type seedlings (arrow heads) in the top section. Two-week-old normal wild-type (WT) and arrested ARP7RNAi seedlings from a strong line showing more than 85% reduction in ARP7 expression are shown in the bottom section. D, Deficiencies in the expression of ARP4, 6, and 7 affect leaf size, number, and/or morphology. The arp6-1 knockout mutants produce narrow and highly serrated leaves under short-day photoperiod (bottom section). All the leaves from an adult ARP4RNAi plant are compared with those from wild type (top section). E, Scanning electron microscopy of the smaller leaves of ARP7RNAi plants reveals similar number, but smaller sized leaf epidermal cells. For comparison, knocking out ARP6 in the arp6-1 mutant results in dwarfed leaves with a smaller number of normally sized cells (not shown). F, A knockdown of ARP7 in RNAi plants severely affects root growth. The retarded roots of RNAi plants have highly reduced cell elongation zone compared to wild type (WT). G, Deficiencies in ARP4 and ARP6 expression affect flowering time. ARP4RNAi plants flower early only under long-day (LD) conditions (bottom section), whereas the arp6-1 mutants flower early both under long- and short-day (SD) conditions (top section). Thus, ARP4 and ARP6 are involved in photoperiod-dependent and photoperiod-independent flowering pathways, respectively. H, A knockdown in ARP4 and ARP7 in the RNAi plants causes delayed senescence and abscission of floral organs. Wild-type inflorescences each contain five or six flowers with intact sepals and petals, whereas the RNAi plants have 15 to 20 flowers in each inflorescence with intact sepals and petals. I, A strong reduction in ARP4 and ARP7 expression affects stamen development and male fertility. The aberrant heart-shaped anthers in the arp4-1 mutant contain less pollen than wild type. J, Deficiencies in ARP4 and ARP6 expression affect female fertility. Defects in pistil development and pollination results in stunted fruits with reduced seed set compared to wild type. K, Knocking out or knocking down ARP4, 6, and 7 expression affect flower morphology and/or organ number. . Reduction in ARP4 expression produces phenotypes affecting numerous aspects of plant growth and development (Kandasamy et al., 2005a). For example, the leaky arp4-1-deficient mutant allele is partially male sterile. Anthers retain a small, immature heart shape and make a small number of mature pollen grains (Fig. 2I), reducing the efficiency of selfpollination. Targeting the distinct 3′ untranslated region of Arabidopsis ARP4 transcripts with RNA interference generates an epiallelic series with strong, moderate, or weak ARP4 deficiency phenotypes (Fig. 2). Strong lines with barely detectable ARP4 protein levels have smaller than normal cotyledons, leaves, inflorescence stems, flowers, and fruits (Fig. 2, C, D, and J). Under long-day but not short-day growth conditions, the strong RNAi lines flower a week early with only six rosette leaves compared to 12 leaves for wild type (Fig. 2G), suggesting a defect in the photoperiod-dependent flowering pathway. The plants with strong epialleles also show an extreme delay in floral organ senescence and abscission, with 20 or more flowers retaining their petals and sepals (Fig. 2H), whereas a wild-type inflorescence seldom retains these floral organs on more than six flowers. ARP7 is an essential protein whose knockdown results in aberrant cell expansion and retarded plant development (Kandasamy et al., 2005b). Knocking out the expression of Arabidopsis ARP7 protein in the homozygous arp7-1 T-DNA mutant produces morphologically abnormal embryos that are arrested early in development (torpedo stage, Fig. 2B). Moreover, knocking down the expression of ARP7 protein levels with RNA interference produces an epiallelic series of plant lines with dosage-dependant, heritable defects in multiple developmental pathways. ARP7-defective plants are severely to moderately dwarfed, with highly retarded roots having reduced elongation zones at their tips (Fig. 2F). Moreover, the dwarf plants contain fewer rosette leaves that were small and severely curled as compared to wild type. These smaller leaves have a similar number of cells to wild type, but the cells are about one-half the size of wild-type cells (Fig. 2E; Kandasamy et al., 2005b). ARP7-deficient plants show a significant decrease in fertility due to defects in pistil and anther development, and poor pollen production. Silenced lines also show delayed abscission of the petal and sepals (Fig. 2H). Null alleles lacking ARP6 protein (arp6-1, arp6-2) display defects during several stages of Arabidopsis development (Choi et al., 2005; Deal et al., 2005). Rosette and cauline leaves, inflorescence stems, flowers, and fruits are dwarfed due to poor cell proliferation, with fewer but normal-sized cells constituting each organ. Under long-day growth conditions the leaves are smaller and narrower than wild type, while under short-day growth the leaves have normal length but are narrower than wild type (Fig. 2G, top), and the mutant leaves are deeply serrated along the margins (Fig. 2D). The arp6− mutants also show defects in inflorescence and flower organ morphologies. Some organs appear underdeveloped relative to mature wild-type organs (Fig. 2K), and under some growth conditions the flowers contain extra petals. There is also a loss of apical dominance, a trait often associated with the loss of normal hormone-stimulated stem cell development. Mutant plants flower earlier than wild type, and early flowering occurs in both long- and short-day photoperiods, implying defects in the photoperiod-independent flowering pathway (Fig. 2G). The mutants are relatively infertile due to a defect in female fertility but not pollen fertility. In plants, the mechanisms by which ARP-containing chromatin-remodeling complexes are targeted to specific genes are poorly understood, but there are several examples of the subunits of these complexes acting as high-level regulators of cell proliferation and development. The control of flowering time in Arabidopsis is the most thoroughly studied plant developmental pathway regulated by changes in chromatin structure, as summarized in Figure 3 Figure 3. Open in new tabDownload slide Regulatory networks controlling flowering time in Arabidopsis. Flowering time is controlled by the activities of the global regulator, FLC. A large number of environmental factors and several plant signaling molecules stimulate or repress the transcription of the FLC gene (underlined) via chromatin remodeling. Figure 3. Open in new tabDownload slide Regulatory networks controlling flowering time in Arabidopsis. Flowering time is controlled by the activities of the global regulator, FLC. A large number of environmental factors and several plant signaling molecules stimulate or repress the transcription of the FLC gene (underlined) via chromatin remodeling. (He and Amasino, 2005). ARP4- and ARP6-deficient Arabidopsis lines revealed distinct photoperiod-dependent and photoperiod-independent early-flowering phenotypes, respectively. Defective ARP4 and ARP6 alleles flowered in approximately half the time required for wild-type plants, when grown under long-day growth conditions. The complex regulation of a master repressor of flowering time, FLOWERING LOCUS C (FLC; Fig. 3) and other related transcription factors in this pathway, are controlled by modulations of chromatin structure (He et al., 2004; Oh et al., 2004). Nearly two-dozen genes in several pathways are known to exert competing signals to stimulate or repress chromatin-mediated FLC expression and therefore repress or activate flowering, respectively (He et al., 2003). FLC and a few other related transcription factors appear to be the primary global regulators of flowering time that integrate signals from the various information pathways. Thus, the flowering-time signaling pathway in Arabidopsis is relevant to dissecting subsets of ARP4 and ARP6 functions. The photoperiod-independent early flowering of ARP6 knockout mutants is associated with reduced expression of FLC as well as MADS AFFECTING FLOWERING 4 and MADS AFFECTING FLOWERING 5 (Deal et al., 2005). Consistent with the reduced expression of FLC, the downstream targets of FLC, FT, and SOC are increased in expression in arp6 mutants. In addition, arp6 mutations suppress the FLC-mediated late flowering of a dominant FRIGIDA allele, indicating that ARP6 is required for the activation of FLC expression to levels that inhibit flowering (Choi et al., 2005; Deal et al., 2005). Finally, strong overexpression of ARP6 resulted in no abnormal plant phenotypes, consistent with ARP6 activity being limited by other components of a chromatin-modifying complex. Thus, FLC expression appears to be directly controlled by chromatin remodeling that is dependent upon the normal expression of ARP6. PHOTOPERIOD-INDEPENDENT EARLY FLOWERING1 and ARP6 encode the closest Arabidopsis homologs of two subunits in the yeast SWR1 complex. The photoperiod-independent early flowering1 and arp6 mutants have many of the same diverse developmental phenotypes, suggesting they participate in the same chromatin-remodeling complex(es) (Noh and Amasino, 2003). In addition, Arabidopsis encodes putative homologs of nearly all of the subunits of the yeast SWR1 complex, consistent with the existence of such complexes in plants (Table I). In summary, there are abundant examples of chromatin remodeling being required for normal plant development. Deficiencies in Arabidopsis ARP4, ARP6, and ARP7 each result in multiple distinct developmental abnormalities including defects in cell expansion and proliferation, dwarfing, and/or alterations in the shape of every vegetative and reproductive organ, and alterations in the timing of developmental pathways such as flowering and floral senescence. The defects observed are consistent with their roles in chromatin-remodeling complexes effecting epigenetic changes in the expression of global regulators like FLC. The possible roles of putative plant nuclear ARPs, ARP5, ARP8, and ARP9 in plant development are as yet unknown. A COMBINATORIAL ARGUMENT FOR THE EXISTENCE OF MULTIPLE ISOFORMS OF ARP-CONTAINING CHROMATIN-MODIFYING COMPLEXES There is substantial evidence that diverse isoforms of ARP-containing chromatin-modifying complexes exist with the potential to exert epigenetic control over development. To define what is meant by isoforms of chromatin complexes, consider the ARP-containing SWI/SNF, SWR1, and INO80 complexes. All three complexes contain 12 to 13 different protein subunits, summarized in Table I. Protein isovariants are classically defined as closely related polypeptides with altered sequences encoded by different gene family members. In plants and animals, divergent gene families encode multiple isovariants of several subunits in each complex. Substituting a single subunit with a different isovariant would generate a new complex isoform with the potential to recognize a new target gene or carry out a slightly different chromatin-modifying reaction (i.e. different phasing of nucleosomes or different histone modifications). Sequentially substituting three different isovariants of, for example, the Snf2 DNA-dependent ATPase subunit of a SWI/SNF complex would generate three isoforms of the complex. By extension, substituting three different isovariants of two different subunits has the potential to generate nine isoforms, and so on. Besides being derived from gene families, protein isovariants could be generated from single genes by alternate splicing or polyadenylation site selection, alternate initiation and termination of translation, and posttranslational protein modifications. Direct and indirect support for the existence of multiple isoforms of chromatin-remodeling complexes can be found in the plant and animal literature and from further examination of their complex genomes. Several human and mouse chromatin-remodeling complexes were first isolated as mixtures of isoforms. For example, the purified mammalian BAF or SWI/SNF complex was shown to have a basic composition of about nine to 12 proteins, but it existed in several isoforms. Isoforms purified from various organs varied in their Baf60 subunit and Swi2/Snf2 subunit compositions (Wang et al., 1996; Debril et al., 2004). BAF60a is constitutively expressed in all organs, whereas BAF60b and BAF60c are expressed in neural and muscle tissues and the pancreas. These Baf60 protein isovariants appear to contribute to the target gene specificity of the complex. Phosphorylation of BAF60 subunits creates an additional isovariant that appears to target a subset of BAF complex isoforms to particular target genes required for muscle development (Simone et al., 2004). Evidence for large numbers of isoforms of ARP-containing complexes in plants comes indirectly from the existence of small gene families encoding subunit isovariants, which we will discuss briefly in the context of the Arabidopsis genome. The three yeast complexes SWI/SNF, SWR1, and INO80 collectively contain all the yeast nuclear ARPs: ARP4, ARP5, ARP6, ARP7, ARP8, and ARP9 (www.yeastgenome.org). As a case study, the plant chromatin (www.chromdb.org) and The Arabidopsis Information Resource (www.arabidopsis.org) databases were searched for homologs of the subunits of these three yeast complexes. Table I summarizes this preliminary examination of Arabidopsis subunit isovariants. The yeast SWI/SNF, SWR1, and INO80 complexes each contain a single, high-Mr snf2-related ATPase subunit, Snf2, Swr1, and Ino80, respectively. The yeast genome encodes a total of 10 distinct Snf2-related proteins (www.yeastgenome.org), and all 10 ATPases are known to participate in a different chromatin-remodeling complex. By contrast, Arabidopsis encodes 42 genes that can be classified as homologs of these ATPases, with one or more being closely allied with each of the 10 yeast sequences in a protein sequence tree (www.chromdb.org). The Arabidopsis clade encoding the immediate homologs of yeast Snf2 contains four genes (Table I). Because the Snf2-related ATPases are not known to participate in any processes other than chromatin modification and have been found primarily in ARP-containing chromatin-remodeling complexes, it is likely that these four closest relatives of the Snf2-ATPase will participate in different isoforms of the Arabidopsis SWI/SNF complex. A similar analysis shows that in addition to the four Snf2 isovariants, Arabidopsis encodes two, two, four, and two isovariants of yeast Snf12, Swi1, Swi3, and Taf14 subunits, respectively (Table I). If these 14 subunit isovariants from the five Arabidopsis gene families are freely interchangeable in SWI/SNF complexes, 128 isoforms of the Arabidopsis complex (i.e. 4 × 2 × 2 × 4 × 2 = 128) would be generated. Each would have the potential to target different genes or carry out variations in NR complex function. A recent analysis of mutants in the four Arabidopsis Swi3 (Table I) isovariants provides direct support for the functional significance of four SWI/SNF complex isoforms each with a different Swi3 isovariant (Zhou et al., 2003; Sarnowski et al., 2005). Mutants in each isovariant subunit displayed a distinct set of phenotypes, including embryo lethality and vegetative and reproductive defects. The sum of the phenotypes described for these four mutants include most of the phenotypes we described for plants deficient in ARP7, which is likely to be a universal member of this complex (Kandasamy et al., 2005b). For example, the swi3a and swi3b mutants display an embryo-lethal phenotype similar to the homozygous arp7-1 mutant (Fig. 2B). In addition, the swi3d mutations cause severe dwarfism and male and female infertility like the ARP7 knockdown alleles (Figs. 2K and 3H). Plants carrying atswi3c mutations shared the retarded root and plant growth, curly leaf, and reduced fertility phenotypes of the ARP7-silenced lines. Moreover, homologs of several subunit proteins within the yeast SWR1 and INO80 complexes are also encoded by small gene families in Arabidopsis, as shown in Table I. There are 21 subunit isovariants in six families with the potential to produce 1,200 SWR1 isoforms. Similarly, there are 14 isovariants in four families with the potential to produce 84 INO80 isoforms. Thus, Arabidopsis ARP4, ARP5, ARP6, and perhaps ARP7 and ARP8 are each predicted to participate in numerous distinct isoforms of complexes distinguished by their isovariant subunit compositions. Each individual ARP complex with a different isovariant composition might control only one or a few target genes participating in the global regulation of development. It seems reasonable to ask how diverse isoforms of ARP-containing chromatin-remodeling complexes became functionally linked to the macroevolution of multicellular development. One evolutionary view would be that as plants and animals diversified from single-celled protist ancestors, there was a combinatorial expansion in the number of nuclear ARP complex isoforms following an expansion and diversification of the gene families encoding their various subunit isovariants. An increase in chromatin-modifying complex isoforms allowed the natural selection of more specialized control over chromatin dynamics and target gene transcription, which generated more specialized epigenetic control over multicellular development. It is logical to assume that greater target gene specificity and more finely tuned epigenetic control were selective advantages to multicellular organisms. SUMMARY Eukaryotic genomes encode several ancient classes of nuclear ARPs that participate in macromolecular machines affecting chromatin dynamics. Nuclear ARPs are variously required for assembly, chromatin binding, and/or the enzymatic activities of different complexes. ARP-containing chromatin-modifying complexes carry out NR, chemical modifications of histones, or HVE reactions. These activities exert epigenetic control over cell proliferation and multicellular development along with controlling basal levels of transcription. 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GM 07103–29 to R.B.D.). * Corresponding author; e-mail [email protected]; fax 706–542–1387. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Richard B. Meagher ([email protected]). www.plantphysiol.org/cgi/doi/10.1104/pp.105.072447. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Matzke, Antonius J.M.; Huettel, Bruno; van der Winden, Johannes; Matzke, Marjori

doi: 10.1104/pp.105.071068pmid: 16339805

Abstract Sixteen distinct sites distributed on all five Arabidopsis (Arabidopsis thaliana) chromosomes have been tagged using different fluorescent proteins and one of two different bacterial operator-repressor systems: (1) a yellow fluorescent protein-Tet repressor fusion protein bound to tet operator sequences, or (2) a green or red fluorescent protein-Lac repressor fusion protein bound to lac operator sequences. Individual homozygous lines and progeny of intercrosses between lines have been used to study various aspects of interphase chromosome organization in root cells of living, untreated seedlings. Features reported here include distances between transgene alleles, distances between transgene inserts on different chromosomes, distances between transgene inserts on the same chromatin fiber, alignment of homologous chromosomes, and chromatin movement. The overall findings are consistent with a random and largely static arrangement of interphase chromosomes in nuclei of root cells. These transgenic lines provide tools for in-depth analyses of interphase chromosome organization, expression, and dynamics in living plants. Although the arrangement of interphase chromosomes is thought to be important for regulating nuclear function and gene expression (Misteli, 2004; Bolzer et al., 2005; Wegel and Shaw, 2005), little information is currently available about interphase chromosome organization in living plant cells (Lam et al., 2004; Tessadori et al., 2004). Most studies so far have used fluorescence in situ hybridization (FISH) to visualize interphase chromosomes in nonliving, fixed material (e.g. Fransz et al., 2002; Pecinka et al., 2004). As an alternate technique, bacterial operator-repressor systems combined with fluorescent proteins offer a unique opportunity to visualize fluorescence-tagged loci in nuclei of living, unfixed cells. The operator repeats are integrated into the genome as a transgene array, which then specifically binds the respective nuclear-localized repressor protein that is fused with a fluorescent protein such as green fluorescent protein (GFP). The tagged loci appear as bright fluorescent dots when viewed with appropriate filters under a fluorescence microscope. Using a fluorescence microscope equipped with a motorized z axis and image-processing software, it is possible to make optical sections through nuclei and reconstruct them in three dimensions to determine spatial relationships among fluorescence-tagged loci. This technique has been employed in yeast, Drosophila, and mammalian cells to analyze interphase chromosome organization and dynamics (Gasser, 2002; Spector, 2003), but so far has been used to study only a limited number of genomic insertion sites in plant cells (Kato and Lam, 2001, 2003; Esch et al., 2003; Matzke et al., 2003; Pecinka et al., 2005). We report here on 16 transgenic Arabidopsis (Arabidopsis thaliana) lines in which distinct chromosomal sites carrying arrays of either tet or lac operators are tagged with the corresponding repressor proteins fused with either yellow fluorescent protein (YFP), red fluorescent protein (dsRed), or GFP. We have used these lines to analyze various aspects of interphase chromosome arrangement and movement in root cells of living, untreated seedlings. RESULTS AND DISCUSSION Constructs, Generation, and Screening of Transgenic Lines Three transgene constructs comprising tet or lac operator repeats and genes encoding the respective nuclear-localized repressor protein-fluorescent protein (RP-FP) fusion proteins were used in this study. Two of these constructs have been reported previously: Tet repressor-enhanced YFP (EYFP), which binds to tet operators (Fig. 1 Figure 1. Open in new tabDownload slide Constructs. Each construct contains a neomycin phosphotransferase II (NPTII) gene under the control of the NOS promoter (Np) and terminator (nt) for selection of transformed plant cells on kanamycin, an NPTIIB gene for selection of bacteria on kanamycin, and an intact NOS gene. Construct 5 encodes a Tet-repressor (TetR)-EYFP fusion protein and contains 112 copies of the tet operator; 16 encodes a Lac-repressor (LacI)-dsRed fusion protein and contains 256 copies of the lac operator; 25 encodes a LacI-EGFP and contains 256 copies of the lac operator. The fusion proteins are under the control of the 35S promoter and terminator (t) and contain three tandem copies of the SV40 nuclear localization signal (narrow vertical black bar). Arrows show the direction of transcription. LB, Left T-DNA border; RB, right T-DNA border; S, SalI; H, HindIII. Figure 1. Open in new tabDownload slide Constructs. Each construct contains a neomycin phosphotransferase II (NPTII) gene under the control of the NOS promoter (Np) and terminator (nt) for selection of transformed plant cells on kanamycin, an NPTIIB gene for selection of bacteria on kanamycin, and an intact NOS gene. Construct 5 encodes a Tet-repressor (TetR)-EYFP fusion protein and contains 112 copies of the tet operator; 16 encodes a Lac-repressor (LacI)-dsRed fusion protein and contains 256 copies of the lac operator; 25 encodes a LacI-EGFP and contains 256 copies of the lac operator. The fusion proteins are under the control of the 35S promoter and terminator (t) and contain three tandem copies of the SV40 nuclear localization signal (narrow vertical black bar). Arrows show the direction of transcription. LB, Left T-DNA border; RB, right T-DNA border; S, SalI; H, HindIII. , construct 5), and Lac repressor-enhanced GFP (EGFP), which binds to lac operators (Fig. 1, construct 25; Matzke et al., 2003). A third construct, Lac repressor-dsRed, which binds to lac operators, is reported here (Fig. 1, construct 16). In the three constructs, the genes encoding the RP-FP fusion proteins are under the control of the nominally constitutive 35S promoter of the Cauliflower mosaic virus. This allows visualization of fluorescent signals in nuclei of living plants that have not undergone any inducing treatments, which can potentially disturb interphase nuclear organization. The constructs were introduced into the Arabidopsis ecotype Columbia (Col-0) genome by the floral-dip method and transformants were selected on kanamycin-containing medium. Following prescreening of kanamycin-resistant T1 seedlings for nuclei with distinct signals appearing as bright, fluorescent dots (Matzke et al., 2003), 16 independent transgenic lines that contained T-DNAs segregating as single loci (nine YFP, six dsRed, and one GFP) were chosen for further analysis. The sites of T-DNA insertion in the 16 lines were determined initially by genetic mapping (Supplemental Fig. 1). The integration sites were confirmed and defined in more detail by using cosmid rescue cloning, lambda cloning, or thermal asymmetric interlaced (TAIL)-PCR to isolate the T-DNA inserts and then sequencing the flanking plant DNA. With the exception of the short arm of chromosome 2, each chromosome arm contains at least one T-DNA insertion site, and, with the exception of chromosome 4, each chromosome has at least one YFP and one dsRed insert (Fig. 2 Figure 2. Open in new tabDownload slide Genomic insertion sites of fluorescence-tagged T-DNAs. Nine YFP-tagged sites are shown in green and numbered beginning with the prefix 5. Six dsRed-tagged sites are shown in red and numbered beginning with the prefix 16. One GFP-tagged site is shown in green and numbered beginning with the prefix 25. Bacterial artificial chromosome numbers containing the insertions are shown in parentheses. White ovals indicate centromeres. Black knobs represent nucleolar organizing regions. All lines display essentially a wild-type phenotype; the exceptions are lines 5:75 and 16:125, which have a somewhat bushy phenotype. Figure 2. Open in new tabDownload slide Genomic insertion sites of fluorescence-tagged T-DNAs. Nine YFP-tagged sites are shown in green and numbered beginning with the prefix 5. Six dsRed-tagged sites are shown in red and numbered beginning with the prefix 16. One GFP-tagged site is shown in green and numbered beginning with the prefix 25. Bacterial artificial chromosome numbers containing the insertions are shown in parentheses. White ovals indicate centromeres. Black knobs represent nucleolar organizing regions. All lines display essentially a wild-type phenotype; the exceptions are lines 5:75 and 16:125, which have a somewhat bushy phenotype. ). The chromosome sequence coordinates of the T-DNA insertions and primers for detection of the inserts by PCR are listed for 14 lines in Supplemental Table I; for two inserts on chromosome 1, 5:56 and 16:52, only the identity of the bacterial artificial chromosome clone containing the T-DNA is currently available. Additional features of the genomic insertion sites are presented in Supplemental Table II. A Southern-blot analysis demonstrated that all lines contained the appropriate operator arrays, which were usually present in scrambled and/or multiple copies (Matzke et al., 2003; data not shown). Root cells were chosen to investigate the fluorescent signals in each line because they are nonphotosynthetic and hence have a low background fluorescence. An initial screen at low magnification of living seedlings mounted on indented microscope slides was used to locate a region of the root that produces optimal signals, which are visualized as bright fluorescent dots in a nucleoplasm of low diffuse background fluorescence (Matzke et al., 2003). Often a suitable region is followed by a region where the RP-FP fusion protein is overexpressed, filling the nucleoplasm and often obscuring the fluorescent dots (Supplemental Fig. 2). Despite the use of the nominally constitutive 35S promoter, the gene encoding the RP-FP fusion protein does not seem to be expressed in every cell because signals are not observed in all nuclei. Moreover, the frequency of RP-FP fusion gene expression varies among seedlings within a line and among individual lines as do the intensities of the signals (Supplemental Table III). It is not known whether the lower-than-expected frequencies result from cell-to-cell variation in the activity of the 35S promoter, gene silencing, or other factors that remain to be identified. Distances between Transgene Alleles The 16 transgenic lines were bred to homozygosity. The relatively uniform distribution of the transgene inserts throughout the Arabidopsis genome (Fig. 2) provided an opportunity to investigate whether alleles at any of the insertion sites showed preferential associations, such as pairing or fixed interallelic distances (Fig. 3A Figure 3. Open in new tabDownload slide Aspects of interphase chromosomes studied here. For illustrative purposes, interphase chromosomes are depicted as black lines. A, Distance between transgene alleles in homozygous lines (d) was determined for all 16 lines (YFP, dsRed, GFP; one color shown for simplicity). B, Splitting of signal might be due to separation of sister chromatids in polytene nuclei. C, Double hemizygous transgene insertions (dsRed and YFP/GFP) on nonhomologous chromosomes. D, Double homozygous transgene insertions (dsRed and YFP/GFP) on nonhomologous chromosomes. E, Double homozygous YFP insertions on homologous chromosomes (left). A backcross to untransformed plants produces double hemizygous YFP insertions on the same homolog (right). F, Double homozygous transgene insertions (dsRed and YFP/GFP) on homologous chromosomes. G, Transgene insertion sites depicted in F were used to study 3D arrangements. When connected by lines, the four signals can be arranged as a tetrahedron. This was defined here as a 3D space in which the smallest height (observed by rotating the connected dots 360° vertically and horizontally) exceeded 15% of the nuclear diameter. The four signals from the lines depicted in F can also be flat and lie in a plane. This was defined here as any arrangement in which the smallest height was less than 15% of the nuclear diameter. H, When transgene inserts are in the same plane (flat), chromosomes containing the signals can be in parallel/crossed or antiparallel orientations. These are extremes of a continuum that can also include perpendicular orientations (Supplemental Table IV). I, For flat parallel signals in the same plane, the distances between the YFP and dsRed alleles (d1 and d2, respectively) can provide information about alignment of homologs in the region between the YFP and dsRed inserts (d1 = d2). J, Lines containing transgene insertion sites depicted in F were backcrossed to untransformed plants to produce double hemizygous insertions on the same homolog (see also 3E, right). These lines can be used to measure cis-distances between insertions on the same chromatin fiber, which provide clues about chromosome folding and/or compaction. Figure 3. Open in new tabDownload slide Aspects of interphase chromosomes studied here. For illustrative purposes, interphase chromosomes are depicted as black lines. A, Distance between transgene alleles in homozygous lines (d) was determined for all 16 lines (YFP, dsRed, GFP; one color shown for simplicity). B, Splitting of signal might be due to separation of sister chromatids in polytene nuclei. C, Double hemizygous transgene insertions (dsRed and YFP/GFP) on nonhomologous chromosomes. D, Double homozygous transgene insertions (dsRed and YFP/GFP) on nonhomologous chromosomes. E, Double homozygous YFP insertions on homologous chromosomes (left). A backcross to untransformed plants produces double hemizygous YFP insertions on the same homolog (right). F, Double homozygous transgene insertions (dsRed and YFP/GFP) on homologous chromosomes. G, Transgene insertion sites depicted in F were used to study 3D arrangements. When connected by lines, the four signals can be arranged as a tetrahedron. This was defined here as a 3D space in which the smallest height (observed by rotating the connected dots 360° vertically and horizontally) exceeded 15% of the nuclear diameter. The four signals from the lines depicted in F can also be flat and lie in a plane. This was defined here as any arrangement in which the smallest height was less than 15% of the nuclear diameter. H, When transgene inserts are in the same plane (flat), chromosomes containing the signals can be in parallel/crossed or antiparallel orientations. These are extremes of a continuum that can also include perpendicular orientations (Supplemental Table IV). I, For flat parallel signals in the same plane, the distances between the YFP and dsRed alleles (d1 and d2, respectively) can provide information about alignment of homologs in the region between the YFP and dsRed inserts (d1 = d2). J, Lines containing transgene insertion sites depicted in F were backcrossed to untransformed plants to produce double hemizygous insertions on the same homolog (see also 3E, right). These lines can be used to measure cis-distances between insertions on the same chromatin fiber, which provide clues about chromosome folding and/or compaction. ). For this, three-dimensional (3D) reconstructions and allelic distance measurements were made for all lines on approximately 10 to 20 root nuclei in which two fluorescent dots could be observed (see Supplemental Fig. 3 for examples). The allelic distances were plotted against nuclear diameter (determined as described in “Materials and Methods”). Complicating the analysis to some extent was the fact that not all nuclei in root cells are round. Most round nuclei were between 3 to 7 μm in diameter, whereas larger nuclei were often irregularly or spindle shaped. In these cases, the estimated nuclear diameter was based on the longest nuclear end-to-end distance. The allelic distances in all lines vary in a similar manner, with a general trend toward larger interallelic distances as nuclear diameter increases, although low values in large nuclei can also be observed (Supplemental Fig. 4). This relationship applies even for insertion sites that are close to telomeres (16:101 on chromosome 1, 16:107 on chromosome 3, and 5:65 on chromosome 4) or in the vicinity of heterochromatic regions, in particular 5:25, which is between the centromere and the heterochromatic knob of chromosome 4 (Fig. 2). The shortest interallelic distance measured in this study was approximately 0.5 μm (Supplemental Fig. 4). This approaches the resolution limit of this technique in budding yeast (Saccharomyces cerevisiae), where two GFP spots closer than 300 nm cannot be resolved (Bystricky et al., 2004). Whether 0.5 μm can be considered to represent allelic pairing is unclear. Indeed, the analysis highlighted the uncertainty about how pairing should be assessed with this method, which allows visualization of transgene inserts from all angles in three dimensions. For example, would pairing be visualized from all viewing angles as a single large dot, approximately double the size of two smaller dots, or as two smaller dots side by side with little or no empty space between them? Conceivably, pairing could potentially lead to loss of signal if close proximity of alleles results in silencing of genes encoding RP-FP fusion proteins. Resolving these issues will require further analysis and an agreement on what constitutes pairing as assessed by this technique. Despite the uncertainties, the observations that most interallelic distances are greater than 1 μm and that they tend to increase as nuclear diameter increases suggest that if pairing of active (i.e. visible) loci occurs, it is not a constant feature of all nuclei within the region of the root examined. Furthermore, alleles throughout the genome do not appear to be at fixed distances relative to each other; instead, the interallelic distances can accommodate changes in nuclear size. Whether these conclusions extend to nuclei in other cell types and to other insertion sites in the genome remains to be investigated. A recent FISH study examining two tagged T-DNAs on the top arm of chromosome 3 in leaf nuclei suggested that pairing, which was thought to be mediated by common lac operator repeats, occurred more often at these sites than expected from a random arrangement (Pecinka et al., 2005). Signal Splitting In a number of homozygous lines (16:52, 5:87, 25:26, 5:117, 16:107, 5:25, 5:27, 5:65; Supplemental Table III), signals from individual alleles were often observed to split into two (or more) smaller signals. Splitting could affect only one allele or both alleles simultaneously (Fig. 4 Figure 4. Open in new tabDownload slide Splitting of signals. Arrows point to split signals affecting one allele only in homozygous lines 5:87, 5:27, and 16:79, and simultaneous splitting of both YFP alleles in the double homozygous line 5:25/16:101. White bar in 5:87 indicates 5 μm and is the same for all images shown. Figure 4. Open in new tabDownload slide Splitting of signals. Arrows point to split signals affecting one allele only in homozygous lines 5:87, 5:27, and 16:79, and simultaneous splitting of both YFP alleles in the double homozygous line 5:25/16:101. White bar in 5:87 indicates 5 μm and is the same for all images shown. ). Splitting reduced the strength of the signal and hence was often detected only after deconvolution, which removes out-of-focus haze and blur from stacks of optical sections and enhances the sharpness and clarity of the image. Understanding the basis of the splitting phenomenon will require more detailed investigation. One possibility is that splitting reflects stochastic separation of sister chromatids in polytene chromosomes of some root nuclei (Fig. 3B). Alternatively, cells could be in the G2 stage of the cell cycle, when sister chromatids split visibly in some chromosomal regions as judged by in situ labeling (Schubert et al., 2005). Distances between Transgene Inserts on Different Chromosomes (Ectopic Sites) To determine whether there is any preferential association between unlinked transgene inserts, intercrosses were made to produce lines containing dsRed and YFP signals on different chromosomes. For this, the homozygous dsRed line 16:101 (chromosome 1) was intercrossed with six homozygous YFP lines: 5:75 (chromosome 1), 5:87 (chromosome 2), 5:117 (chromosome 3), 5:25 (chromosome 4), 5:123 (chromosome 4), and 5:106 (chromosome 5; Fig. 2). F1 progeny of these crosses are hemizygous for the two transgene inserts (Fig. 3C, which depicts the two inserts on nonhomologous chromosomes; for the 5:75/16:101 combination, the two inserts are on homologs of chromosome 1). The double hemizygous progeny were analyzed with respect to the distance between the dsRed and YFP signals. As with the interallelic distances, the ectopic distances were variable, with a trend toward increased values as the nuclear diameter increased (Supplemental Fig. 5). There was no evidence for preferential associations of any ectopic T-DNA insertions. Note that, in these cases, the YFP and dsRed inserts contain heterologous operator arrays (tet or lac, respectively) and hence would not be prone to ectopic pairing of operator elements. Selfing of double hemizygous plants produced some F2 progeny that were doubly homozygous for both transgene inserts (Fig. 3D). Distance measurements between the two YFP alleles and the two dsRed alleles in the double homozygotes were similar to those obtained with the single homozygous lines and continued to indicate no preferential allelic associations (data not shown). Overall, the results on distances between ectopic insertions are similar to those observed here for alleles and are compatible with a random arrangement of interphase chromosomes. 3D Arrangements Some of the most interesting material came from lines that were doubly homozygous for either two YFP (or two dsRed) inserts or one dsRed and one YFP (or GFP) insert on the same chromosome (Fig. 3E, left, and F, respectively). The four signals define a space and hence give information about 3D spatial relationships among loci. To obtain these lines, intercrosses were made to generate lines with the following combination of dsRed and YFP (or GFP) signals: 5:75/16:101 (chromosome 1), 5:87/16:125 (chromosome 2), 25:26/16:125 (chromosome 2), and 5:106/16:112 (chromosome 5). In the case of chromosome 4, which did not have any dsRed inserts, two YFP signals were combined (5:25/5:123). Four signals could be observed in nuclei of these five double homozygous lines (Fig. 5 Figure 5. Open in new tabDownload slide Double homozygous lines containing two transgene inserts on the same chromosome. The combinations of transgene inserts are shown in white letters and their positions on a given chromosome are depicted below the images. The top images are top views of the maximal projection (all deconvoluted planes from the stack collapsed into a single plane); the bottom images are rotated 90° to give a side view. White bar in 5:75/16:101 indicates 5 μm and is the same for all images shown. Figure 5. Open in new tabDownload slide Double homozygous lines containing two transgene inserts on the same chromosome. The combinations of transgene inserts are shown in white letters and their positions on a given chromosome are depicted below the images. The top images are top views of the maximal projection (all deconvoluted planes from the stack collapsed into a single plane); the bottom images are rotated 90° to give a side view. White bar in 5:75/16:101 indicates 5 μm and is the same for all images shown. ), although the frequency was usually lower than the frequencies of observing two signals in each of the homozygous parental lines (Supplemental Table III; Supplemental Fig. 6). With the 25:26/16:125 combination on chromosome 2, in which both T-DNA inserts contain lac operator arrays but encode either a GFP- or dsRed-Lac repressor fusion protein, only the GFP-Lac repressor fusion protein was expressed such that four green signals (and no red signals) were usually observed (Fig. 5). For chromosome 3, the two possible YFP-dsRed combinations (5:117/16:79 and 5:117/16:107) resulted in signals that were too weak for proper imaging. Therefore, we used a double combination comprising two dsRed inserts, 16:79/16:107, in which four dots were visible (although only in a few nuclei per seedling root; Fig. 5; Supplemental Table III). The lower-than-expected frequencies in the double homozygous lines are currently unexplained, but they might be due to interactions between T-DNA inserts that lead to gene silencing. The analysis of 3D relationships is facilitated in double homozygous lines containing dsRed and YFP signals on the same chromosome because the two loci can be distinguished by color. Three such lines are available in the present collection: 5:75/16:101 (chromosome 1), 5:87/16:125 (chromosome 2), and 5:106/16:112 (chromosome 5; Fig. 5). Several features of the 3D arrangement could be discerned with these lines. First, the use of image-processing software to connect the four dots and rotate them 360° in all directions revealed that they either formed the corners of a tetrahedron or were more-or-less flat in a single plane (Fig. 3G). Second, in the case of a plane, the signals could be arranged in a way that was consistent with either a more-or-less parallel/crossed, perpendicular, or antiparallel orientation of homologs (Fig. 3H). Examples of parallel/crossed and antiparallel for 5:75/16:101 (chromosome 1) and 5:106/16:112 (chromosome 5) are shown in Figure 6 Figure 6. Open in new tabDownload slide Parallel/crossed and antiparallel orientations. Examples are shown for the YFP and dsRed inserts on chromosome 1 (5:75/16:101) and on chromosome 5 (5:106/16:112). In the four cases shown (six boxes each), the signals are shown in the nucleus (bounded in yellow), then from the top and side views. The dots are connected by white lines, then turned in three dimensions to see whether they fall into a line or a very shallow tetrahedron (<15% of nuclear diameter) and are thus considered flat arrangements (Fig. 3G; Supplemental Table IV). The drawings show that it is only possible to connect the dots in either a parallel/crossed or antiparallel orientation. White bar in 5:75/16:101 indicates 5 μm and is the same for all images shown. Figure 6. Open in new tabDownload slide Parallel/crossed and antiparallel orientations. Examples are shown for the YFP and dsRed inserts on chromosome 1 (5:75/16:101) and on chromosome 5 (5:106/16:112). In the four cases shown (six boxes each), the signals are shown in the nucleus (bounded in yellow), then from the top and side views. The dots are connected by white lines, then turned in three dimensions to see whether they fall into a line or a very shallow tetrahedron (<15% of nuclear diameter) and are thus considered flat arrangements (Fig. 3G; Supplemental Table IV). The drawings show that it is only possible to connect the dots in either a parallel/crossed or antiparallel orientation. White bar in 5:75/16:101 indicates 5 μm and is the same for all images shown. ; the complete set of data for all three lines is shown in Supplemental Table IV. For a parallel orientation, the distances between the YFP and dsRed alleles should be similar, essentially reflecting alignment of homologs in the region between the two T-DNA insertion sites (d1 ≅ d2; Fig. 3I). This was indeed the case for line 5:87/16:125 (Supplemental Fig. 7, middle), in which the YFP and dsRed inserts are separated by only 2.4 Mb on chromosome 2 (Fig. 2). The equivalence of d1 and d2 was independent of their absolute values, such that d1 and d2 could range from approximately 1.5 to 8 μm. These findings serve as a control to validate the distance measurements, since one would expect two adjacent loci separated by a relatively short physical distance to have similar interallelic distances in a given nucleus with a flat, parallel arrangement of homologs. There was less frequent equivalence of d1 and d2 as the YFP and dsRed inserts became separated by much larger distances in lines 5:75/16:101 (chromosome 1) and 5:106/16:112 (chromosome 5; Supplemental Fig. 7, top and bottom, respectively). Thus, when considering interallelic distances at two widely spaced loci, d1 and d2 could be quite different. These data suggest that alignment of homologs (defined here as a flat, parallel arrangement in which d1 = d2, regardless of the absolute value of d1 and d2; Fig. 3I) is most often detected over relatively short regions (e.g. 2.4 Mb in line 5:87/16:125). The models shown in Figure 3 depict the chromosomes as linear entities, which is an oversimplification. However, the models provide a basis for interpreting the 3D results, which demonstrate that the relative orientation of the dsRed and YFP signals on homologous chromosomes is not fixed. Indeed, evidence for all possible variations (Fig. 3, G–I) was obtained in root nuclei of a given line, supporting a random arrangement of interphase chromosomes in root cells of living, untreated seedlings. cis-Distances between Transgene Inserts on the Same Chromatin Fiber Although the distances between YFP alleles or dsRed alleles can be measured in plants that are doubly homozygous for the respective inserts (previous section), it was not possible in those cases to discern which YFP signal and which dsRed signal were together on the same chromatin fiber (e.g., parallel could not be distinguished from crossed; Fig. 3H). To address the question of distances between transgene inserts on the same chromatin fiber, we backcrossed all six lines that are doubly homozygous for two transgene inserts on the same chromosome (Fig. 5) to nontransgenic plants. The progeny of these crosses are double hemizygous on the same homolog for either two YFP inserts (Fig. 3E, right) or one YFP (or GFP) and one dsRed insert (Fig. 3J). With these plants, it is possible to measure the distance in cis between two transgene inserts that are known to be present on the same chromatin chain. This can potentially provide information on chromosome folding or compaction (Fig. 3J) since the approximate linear distance (in megabase pairs) between the two signals is known (Table I Table I. Possible fold compaction of DNA between fluorescence-tagged transgene inserts on the same chromosome based on maximal interlocus distance measurements in interphase nuclei of root cellsa Conversions: 1 bp double helical DNA (B form) = 0.34 nm (Lehninger, 1975); 1,000 bp (1 kb) = 340 nm; 3 kb equals approximately 1 μm; and 3 Mb equals approximately 1 mm. Chromosome . Inserts . Approximate Distance between Inserts . Approximate DNA Length . Maximal Observed Distance . Fold Compaction . Mb mm μm 1 5:75/16:101 25 8.3 8.7 960× 2 5:87/16:125 2.4 0.8 2.1 380× 2 25:26/16:125 6.7 2.2 5.1 430× 3 16:79/16:107 17.5 5.8 6.7 865× 4 5:25/5:123 12.3 4.1 7.9 520× 5 5:106/16:112 16.4 5.5 7.4 740× Average 650× Chromosome . Inserts . Approximate Distance between Inserts . Approximate DNA Length . Maximal Observed Distance . Fold Compaction . Mb mm μm 1 5:75/16:101 25 8.3 8.7 960× 2 5:87/16:125 2.4 0.8 2.1 380× 2 25:26/16:125 6.7 2.2 5.1 430× 3 16:79/16:107 17.5 5.8 6.7 865× 4 5:25/5:123 12.3 4.1 7.9 520× 5 5:106/16:112 16.4 5.5 7.4 740× Average 650× a Figure 7. Open in new tab Table I. Possible fold compaction of DNA between fluorescence-tagged transgene inserts on the same chromosome based on maximal interlocus distance measurements in interphase nuclei of root cellsa Conversions: 1 bp double helical DNA (B form) = 0.34 nm (Lehninger, 1975); 1,000 bp (1 kb) = 340 nm; 3 kb equals approximately 1 μm; and 3 Mb equals approximately 1 mm. Chromosome . Inserts . Approximate Distance between Inserts . Approximate DNA Length . Maximal Observed Distance . Fold Compaction . Mb mm μm 1 5:75/16:101 25 8.3 8.7 960× 2 5:87/16:125 2.4 0.8 2.1 380× 2 25:26/16:125 6.7 2.2 5.1 430× 3 16:79/16:107 17.5 5.8 6.7 865× 4 5:25/5:123 12.3 4.1 7.9 520× 5 5:106/16:112 16.4 5.5 7.4 740× Average 650× Chromosome . Inserts . Approximate Distance between Inserts . Approximate DNA Length . Maximal Observed Distance . Fold Compaction . Mb mm μm 1 5:75/16:101 25 8.3 8.7 960× 2 5:87/16:125 2.4 0.8 2.1 380× 2 25:26/16:125 6.7 2.2 5.1 430× 3 16:79/16:107 17.5 5.8 6.7 865× 4 5:25/5:123 12.3 4.1 7.9 520× 5 5:106/16:112 16.4 5.5 7.4 740× Average 650× a Figure 7. Open in new tab ). Of the six lines tested, four of them have the signals on opposite sides of the centromere (5:75/16:101 on chromosome 1, 16:79/16:107 on chromosome 3, 5:25/16:123 on chromosome 4, and 5:106/16:112 on chromosome 5; Fig. 5). In these lines, the cis-distances were quite variable (Fig. 7 Figure 7. Open in new tabDownload slide cis-Distances between hemizygous transgene inserts on the same chromatin fiber. The identity of the lines and the approximate distance (d) separating the transgene inserts on the same chromatin chain in megabase pairs are shown at the top of each graph and in Table I. Each point represents the cis-distance in a single nucleus determined after optical sectioning, deconvolution, and 3D reconstruction. Figure 7. Open in new tabDownload slide cis-Distances between hemizygous transgene inserts on the same chromatin fiber. The identity of the lines and the approximate distance (d) separating the transgene inserts on the same chromatin chain in megabase pairs are shown at the top of each graph and in Table I. Each point represents the cis-distance in a single nucleus determined after optical sectioning, deconvolution, and 3D reconstruction. , top and middle), indicating considerable nucleus-to-nucleus variability in chromosome folding and/or compaction. The cis-distances also appear to be proportional to some extent on nuclear diameter, as indicated by a tendency to increase as nuclear diameter increased, but lower values can also be seen in some larger nuclei, possibly as a consequence of folding (Fig. 3J). These results suggest that individual chromosome arms can be quite flexible when adopting their positions in interphase nuclei. A more constrained pattern was obtained with line 5:87/16:125 (chromosome 2), in which the YFP and dsRed signals are on the same side of the centromere and separated by only 2.4 Mb (Fig. 5). Here the cis-distance remained in most cases below 2 μm (Fig. 7, bottom right). The YFP and dsRed signals in this line are distinguishable in the fluorescence microscope but always close together (Fig. 5; Supplemental Fig. 8), indicating that 2.4 Mb is still above the minimum resolvable cis-distance between two differently colored inserts on the same chromatin fiber. A second chromosome 2 line that has the same dsRed insert (16:125), but a more distantly spaced GFP insert (25:26; distance 6.7 Mb) on the same chromosome arm (Fig. 5), shows increasing cis-distances up to a maximum measurement of 5.1 μm but also low values (Fig. 7, bottom left). Whether some low values might represent enhanced pairing in cis of lac operator repeats associated with both the GFP and dsRed inserts in this line is not known. cis-Pairing of common operator repeats is also potentially possible in line 16:79/16:107 on chromosome 3, in which both inserts have lac operators, and line 5:25/5:123 on chromosome 4, in which both inserts have tet operators. However, unlike the T-DNA inserts in line 25:26/16:125, the inserts in these lines are on opposite sides of the centromere. Whether this might influence the chances of cis-pairing is not known. It should be emphasized that interpretations of the cis-distance measurements assume that no chromosome rearrangements, such as inversions (Pecinka et al., 2005), have taken place during T-DNA integration. Under this assumption, the fold compaction associated with the maximum measured distances ranged from 380- to 960-fold (Table I). These values are only very rough estimates, since it is likely that the degree of compaction and/or folding vary considerably along a chromatin fiber. A larger collection of more closely spaced inserts would be valuable for studying chromatin compaction in plant interphase nuclei. Based on a similar approach of using fluorescence-tagged tet and lac operators on the same chromosome, the average compaction ratio of interphase chromatin in budding yeast was determined to be approximately 40-fold (Bystricky et al., 2004). Movement of Chromosomes Fluorescence-tagging of transgenes has permitted analysis of interphase chromosome dynamics in cells of yeast, Drosophila, and mammals (Gasser, 2002). A study on Arabidopsis using fluorescence-tagged T-DNAs on chromosome 3 in diploid nuclei of guard cells indicated that the maximal radius of the confinement area was 0.21 μm (Kato and Lam, 2003). In the absence of a nuclear envelope marker, which can be used to control for movement of the nucleus itself, we studied chromosome dynamics with respect to whether the distance between alleles varies over a period of 80 min. A strong signal that is stable for the duration of the experiment is required. Homozygous line 16:112, in which the dsRed insert is present toward the end of the long arm of chromosome 5 (Fig. 2), fulfilled this requirement. Consistent with the previous study (Kato and Lam, 2003), we observed a maximal change in the allelic distance of 0.2 μm over the observation period (Supplemental Fig. 9). Although we cannot rule out movements over very short time frames (e.g. seconds), the allelic arrangement appears overall to be quite stable. Additional studies are needed to determine whether similar results will be obtained with transgenes inserted at other chromosomal locations and in nuclei from other cell types. Interphase chromosomes might also change their positions during development or under various inducing or environmental stress conditions (Tumbar et al., 1999; Dietzel et al., 2004). The ability to view the fluorescence-tagged loci in living cells provides a means to study these and other problems of interphase chromosome organization. CONCLUSION We have developed 16 transgenic lines that have fluorescence-tagged T-DNAs at different locations in the Arabidopsis genome. The precise insertion sites have been determined in most cases so that the transgene inserts can be detected by PCR analysis. The lines have been used to study various features of interphase chromosome disposition and 3D spatial relationships. The results obtained so far indicate considerable variation in interphase chromosome arrangement in root cells of living, intact seedlings and generally support previous findings from FISH analyses of interphase chromosomes in Arabidopsis (Fransz et al., 2002; Pecinka et al., 2004). The types of analyses performed in this study can be extended to other cell types, developmental stages, or environmental conditions. There are several advantageous features of the transgenic lines reported here. The combination of two easily distinguishable fluorescent proteins (YFP and dsRed) that bind to heterologous operator repeats (tet and lac, respectively) eliminates the possibility that unlinked arrays of operator repeats might pair (Pecinka et al., 2005). Thus, these combinations allow 3D analyses of chromosomal sites under conditions in which interphase nuclear architecture is largely unperturbed, although local distortions induced by the operator repeat arrays cannot be ruled out. Minimal disruption of nuclear organization is also ensured by the fact that no treatments are required to induce expression of the RP-FP proteins in these lines. Living seedlings can simply be removed from growth medium, mounted on microscope slides, immediately viewed and optically sectioned under the fluorescence microscope, and then replaced on medium to resume growth. In all of the lines reported here, the signals are visible in the fluorescence microscope before deconvolution (Supplemental Table III). Indeed, signals in dsRed lines 16:101 and 16:112 are strong enough to be seen under low magnification (160×; data not shown). The primary drawback of these lines is the lower-than-expected frequency of expression of the RP-FP fusion protein. This problem affects all individual lines to varying extents and can be exacerbated in the progeny of some intercrosses (Supplemental Table III). Whether the low frequencies of RP-FP fusion protein expression are more pronounced in root cells than in other cell types remains to be investigated. The contribution of various types of gene silencing to the low signal frequencies is being examined by crossing the tagged transgenic lines with mutants defective in either transcriptional or posttranscriptional gene silencing. In addition, we are examining whether supplying the RP-FP fusion proteins (under the control of different promoters) in trans will alleviate this problem. Even with the frequencies currently attainable, however, these lines provide useful material for analyzing plant interphase chromosomes in their natural state. MATERIALS AND METHODS Constructs and Production of Transgenic Plants Constructs 5 (tetY) and 25 (lacG) and the production of transgenic Arabidopsis (Arabidopsis thaliana) plants (Col-0) using these constructs have been described previously (Matzke et al., 2003). The construct 16 (lacR) was created by replacing EGFP with DsRed2 (purchased from CLONTECH-BD Biosciences) at the pBC stage of the fusion construct using NheI and BsrGI (Fig. 1). At the pBC stage of the fusion construct, the 35S promoter can easily be replaced with another promoter using an XhoI/NheI cleavage (Matzke et al., 2003). Genetic Mapping and Recovery of Flanking DNA Homozygous transgenic Arabidopsis plants (Col-0) were crossed to ecotype Landsberg erecta (Ler) to obtain mapping populations. T-DNA insertions were roughly mapped using a mapping population of around 30 F2 plants and sets of simple sequence length polymorphism (Bell and Ecker, 1994; Lukowitz et al., 2000) and cleaved amplified polymorphic sequence markers (Konieczny and Ausubel, 1993), which detect polymorphisms between the Col-0 and Ler ecotypes (Supplemental Fig. 1). The primer sequence for the simple sequence length polymorphism marker MAC9 on the lower arm of chromosome 5 (developed by W. Aufsatz, personal communication) is 5′-GTC ATG TCAC TGG GGA TAA G-3′; 5′-ATG TGT AAC ACC CAT TGG AC-3′. To recover flanking DNA of the T-DNA inserts, a combination of approaches was used: TAIL-PCR (Weigel and Glazebrook, 2002), lambda cloning using a Lambda FIX II/XhoI partial fill-in vector kit and Gigapack III gold packaging extract (both from Stratagene), and cosmid rescue cloning using the cosmid pWEB::TNC cloning kit (from Epicentre, purchased from Biozym Diagnostics GmbH), which allows direct selection of T-DNA inserts carrying the NPTIIB gene for bacterial selection. Inserts of cosmids were end sequenced using M13 forward and T7 standard primers. In cases where Arabidopsis DNA was detected, we further sequenced using a nopaline synthase (NOS) promoter primer oriented toward the T-DNA border (5′-TTC TGT CAG TTC CAA ACG-3′) and could usually recover the T-DNA-plant DNA junction. If Arabidopsis DNA sequences were not detected by using the NOS promoter primer, we performed primer walking (starting at the Arabidopsis DNA detected by end sequencing) until the T-DNA-plant DNA junction was reached. Primers were designed to allow these T-DNA inserts to be detected in progeny by simple PCR assays (Supplemental Table I). Seedling Growth Conditions Seeds were sterilized and germinated on sterile solid Murashige and Skoog medium in petri dishes. The dishes were placed at 4°C for 2 to 3 d, then moved to a Percival incubator and grown at 23°C in a 16-h-light/8-h-dark cycle. Seedlings used in all experiments were approximately 10 to 20 d old. Fluorescence Microscopy and Image Analysis Whole seedlings were mounted in tap water on indented slides. Leaves were placed in the indented region and the root was stretched out on the regular slide surface. A coverslip placed over the entire seedling was sealed with rubber cement. Photographs and stacks (40 images at 0.2-μm z-axis steps; exposure time for each image, 1 s; total elapsed time for one stack, approximately 2 min) were taken using a Zeiss Axioplan 2 fluorescence microscope (Zeiss) equipped with appropriate filter cubes for YFP, dsRed, and GFP, and with a Quantix CCD camera from Photometrics (purchased from Zeiss) run with MetaView software (Visitron Systems GmbH). Frequency Plots Frequency plots allow visualization of the variability of the signal frequency in interphase nuclei of root cells of a given line: Starting at the root tip, nuclei with signals (fluorescent dots) are counted in consecutive areas that are visible in the microscope using the 63× objective, which equals approximately 0.4-mm distance × 50 sections or about 20 mm (=2 cm) of root. After the root tip area, one can count per section (using DAPI staining of nuclei) about 100 elongated root cells on average (Dolan et al., 1993). Typical frequencies for a given line can be reduced drastically when combined with another locus (Supplemental Table III; see Supplemental Fig. 6 for an example). Distance Measurements Distance measurements were made in three dimensions after deconvolution of the image stacks using AutoDeblure 9.2, configuration WF (wide field), edition gold (AutoQuant Imaging, purchased from Bitplane AG) in MetaMorph (Meta Imaging Series 5.0; Universal Imaging, purchased from Visitron) and/or Imaris 4.1.3 (Bitplane AG). Using the region measurement tool of MetaMorph, the perimeter of each nucleus was manually drawn on the maximal projections following the background fluorescence that fills the nuclei (nuclei were turned in three dimensions to find the maximal area; if odd shaped, the most reasonable estimates were made), light micrographs, or overlays (whatever allowed the most reliable way to follow the nuclear rim), and the nuclear area A was determined. Nuclear diameters were calculated using the equation D = 2(A/pi)″0.5 (Parada et al., 2004). Determination of Height of Tetrahedra in YFP/dsRed Double Homozygous Lines Fluorescent signals (dots) were connected in MetaMorph with white lines using the “Measure XYZ Distance” function. After applying the 3D reconstruction function of MetaMorph, the resulting tetrahedra were turned in three dimensions until three spots formed one line (spanning a triangular plane). Using the caliper function of MetaMorph, the distance of the fourth spot relative to that plane was determined. The minimum height of all possibilities was determined. If this value was less than 15% of the nuclear diameter, the arrangement was considered flat. Monitoring Allelic Distance over Time The distance between dsRed alleles in homozygous line 16:112 was measured as a function of time. Every 5 min a stack (21 images at 0.2-μm z-axis steps) was collected for 80 min. Distance measurements were carried out after deconvolution (AutoDeblur) using the point detection function of Imaris (Supplemental Fig. 9). Fixed material was prepared according to a published procedure (Kato and Lam, 2003). ACKNOWLEDGMENTS We thank Marie-Therese Hauser for suggesting that splitting might be due to separation of sister chromatids in polytene chromosomes. LITERATURE CITED Bell CJ, Ecker JR ( 1994 ) Assignment of 30 microsatellite loci to the linkage map of Arabidopsis. Genomics 19 : 137 –144 Bolzer A, Kreth G, Solovei I, Koehler D, Saracoglu K, Fauth C, Müller S, Eils R, Cremer C, Speicher MR, et al ( 2005 ) Three-dimensional maps of all chromosomes in human male fibroblast nuclei and prometaphase rosettes. PLoS Biol 3 : e157 Bystricky K, Heun P, Gehlen L, Langowski J, Gasser SM ( 2004 ) Long-range compaction and flexibility of interphase chromatin in budding yeast analyzed by high-resolution imaging techniques. 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Chromosoma 113 : 258 –269 Schubert V, Klatte M, Pecinka A, Meister A, Jasencakova Z, Schubert I ( 2005 ) Sister chromatids are often incompletely cohesed in meristematic and endopolyploid interphase nuclei of Arabidopsis thaliana. Genetics doi/10.1534/genetics.105.048363 Spector DL ( 2003 ) The dynamics of chromosome organization and gene regulation. Annu Rev Biochem 72 : 573 –608 Tessadori F, van Driel R, Fransz P ( 2004 ) Cytogenetics as a tool to study gene regulation. Trends Plant Sci 9 : 147 –153 Tumbar T, Sudlow G, Belmont AS ( 1999 ) Large-scale chromatin unfolding and remodelling induced by VP16 acidic activation domain. J Cell Biol 145 : 1341 –1354 Wegel E, Shaw P ( 2005 ) Gene activation and deactivation related changes in the three-dimensional structure of chromatin. Chromosoma doi/10.1007/s00412-005-0015-7 Weigel D, Glazebrook J ( 2002 ) How to isolate a gene defined by a mutation. In Arabidopsis. A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp 143–170 Author notes 1 This work was supported by the Austrian Fonds zur Förderung der Wissenschaftlichen Forschung (grant no. 16545–B12). * Corresponding author; e-mail [email protected]; fax 43–1–4277–29749. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Antonius J.M. Matzke ([email protected]). [W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.105.071068. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Kraft, Edward; Stone, Sophia L.; Ma, Lingeng; Su, Ning; Gao, Ying; Lau, On-Sun; Deng, Xing-Wang; Callis, Judy

doi: 10.1104/pp.105.067983pmid: 16339806

Abstract Attachment of ubiquitin to substrate proteins is catalyzed by the three enzymes E1, E2 (ubiquitin conjugating [UBC]), and E3 (ubiquitin ligase). Forty-one functional proteins with a UBC domain and active-site cysteine are predicted in the Arabidopsis (Arabidopsis thaliana) genome, which includes four that are predicted or shown to function with ubiquitin-like proteins. Only nine were previously characterized biochemically as ubiquitin E2s. We obtained soluble protein for 22 of the 28 uncharacterized UBCs after expression in Escherichia coli and demonstrated that 16 function as ubiquitin E2s. Twelve, plus three previously characterized ubiquitin E2s, were also tested for the ability to catalyze ubiquitination in vitro in the presence of one of 65 really interesting new gene (RING) E3 ligases. UBC22, UBC19-20, and UBC1-6 had variable levels of E3-independent activity. Six UBCs were inactive with all RINGs tested. Closely related UBC8, 10, 11, and 28 were active with the largest number of RING E3s and with all RING types. Expression analysis was performed to determine whether E2s or E3s were expressed in specific organs or under specific environmental conditions. Closely related E2s show unique patterns of expression and most express ubiquitously. Some RING E3s are also ubiquitously expressed; however, others show organ-specific expression. Of all the organs tested, RING mRNAs are most abundant in floral organs. This study demonstrates that E2 diversity includes examples with broad and narrow specificity toward RINGs, and that most ubiquitin E2s are broadly expressed with each having a unique spatial and developmental pattern of expression. Protein ubiquitination is the covalent attachment of the 76-amino acid eukaryotic molecule, ubiquitin, to substrate proteins. The fate of the ubiquitinated substrate depends upon the type of ubiquitin modification and the choice of ubiquitin lysyl residue used to form the attached polyubiquitin chain (Fang and Weissman, 2004; Sun and Chen, 2004) Ligation of a single ubiquitin molecule (monoubiquitination) has been linked to endocytosis and histone modification (Schnell and Hicke, 2003; Umebayashi, 2003). Proteins modified by the attachment of a polyubiquitin chain of four or more ubiquitins linked via ubiquitin lysyl residue 48 are typically targeted for degradation by the 26S proteasome (Thrower et al., 2000). In contrast, modification with a lysyl-63-linked polyubiquitin chain has been implicated in protein activation, stress response, and DNA damage repair (Hoege et al., 2002; Shi and Kehrl, 2003; Zhou et al., 2004). Ubiquitin conjugation is a multistep reaction, sequentially involving three enzymes referred to as an E1 (ubiquitin-activating enzyme [UBA]), an E2 (ubiquitin-conjugating enzyme [UBC]), and an E3 (ubiquitin ligase; Glickman and Ciechanover, 2002). The first event in the cascade is the ATP-dependent formation of a thioester-linked ubiquitin by E1. Thioester-linked ubiquitin is then transferred to a cysteinyl residue of the E2 enzyme. An E3 enzyme facilitates the transfer of ubiquitin to a lysyl group on the substrate. E3 ligases mediate this step either through the formation of a ubiquitin thioester prior to transfer to the substrate or by noncovalent interaction with the E2 carrying thioester-linked ubiquitin. The E3 enzymes are the substrate recognition component and thus are an important determinant of specificity in the ubiquitination pathway. E2s were originally defined as proteins capable of accepting ubiquitin from an E1 through a thioester linkage via a cysteinyl sulfhydryl group (Glickman and Ciechanover, 2002). The cysteinyl residue is found in a conserved region of approximately 140 to 150 amino acids called the UBC domain (Inter-Pro IPR000608). The UBC domain also interacts with the E3 enzyme and, in some cases, it also interacts with the substrate (Kalchman et al., 1996). Other conserved amino acids are also required for the formation of the E2-ubiquitin intermediate (Wu et al., 2003). The yeast (Saccharomyces cerevisiae) genome encodes 11 ubiquitin E2s and there are approximately 50 E2s in the human genome (Bachmair et al., 2001; Jiang and Beaudet, 2004). Previous analysis of the predicted Arabidopsis (Arabidopsis thaliana) proteome identified 37 proteins and one putative pseudogene (At1g35700) that contain a cysteinyl residue within a UBC domain (Bachmair et al., 2001). Additional characterized Arabidopsis proteins with UBC domains include RUB-conjugating enzyme RCE1 (At4g36800), which forms a thioester with the related to ubiquitin-like (RUB) protein RUB1 (At1g31340; Del Pozo and Estelle, 1999) and a closely related protein RCE2 (At2g18600), and SUMO-conjugating enzyme SCE1a (At3g57870) and a truncated protein SCE1b (At5g02240) thought to be a pseudogene (Kurepa et al., 2003). While no functional studies with AtSCE1a have been reported, a Nicotiana benthamiana UBC, 88% identical to AtSCE1a, can activate the ubiquitin-like protein small ubiquitin-related modifier (SUMO1) in vitro and can complement a yeast SUMO E2 mutant (Castillo et al., 2004). Compared to other UBC proteins, RCE1 and SCE1a have higher amino acid identity to human UBC domain-containing proteins, HsUBC12 and HsUBC9, respectively, that form thioester linkages with mammalian orthologs of RUB1 and SUMO. These data suggest that AtRCE1, AtRCE2, and SCE1a do not function in vivo as ubiquitin-conjugating enzymes, but as ubiquitin-like conjugating enzymes (Del Pozo and Estelle, 1999; Kurepa et al., 2003; Castillo et al., 2004). Approximately 1,300 genes of the Arabidopsis genome are predicted to encode for E3 components (Smalle and Vierstra, 2004). The presence of the homology to the E6-AP C terminus (HECT), U-box, or really interesting new gene (RING) domain groups subdivides the Arabidopsis E3 ligases into three classes. These domains have been shown to be essential for E3 ligase function in protein ubiquitination (Lorick et al., 1999; Hatakeyama et al., 2001). The HECT domain class of ubiquitin ligases is unique in that it forms a thioester-linked ubiquitin E3 intermediate prior to transfer of the ubiquitin molecule to the substrate. The remaining classes, RING and U-box proteins, are thought to facilitate ubiquitination by functioning as scaffolds to bring the E2 with thioester-linked ubiquitin and substrate together. The Cys-rich, zinc ion coordinating the RING domain (Freemont et al., 1991; Freemont, 1993) is found in both simple and complex E3 ligases. Complex E3 ligases, such as the well characterized Skp1-Cullin-F-box-type ligase, have the substrate recognition and E2 binding on separate proteins, the F-box protein and the RING-containing protein RBX/ROC/HRT, respectively (Gagne et al., 2002; Kuroda et al., 2002; Lechner et al., 2002; Risseeuw et al., 2003). In contrast, with the simple RING E3 ligases (e.g. ABI3 interacting protein 2 AtAIP2 [Zhang et al., 2005] and SINA of Arabidopsis thaliana 5 AtSINAT5 [Xie et al., 2002]), both substrate and E2 binding occurs within a single protein. The ubiquitination pathway modifies a diverse range of proteins, thus placing protein ubiquitination at the center of numerous cellular processes in all eukaryotic species. With over 5% of the Arabidopsis proteome predicted to be involved in the ubiquitination-26S proteasome pathway, it is not surprising that protein ubiquitination is postulated to be involved in many different aspects of plant growth and development (Smalle and Vierstra, 2004). The large number of potential ubiquitinating enzymes suggests that substrate-specific ubiquitination plays an essential role in cellular regulation in Arabidopsis. However, how the ubiquitination pathway confers specificity and how the pathway is regulated are not well understood in plants. To begin to understand the specificity or selectivity of the pathway, the components involved and how they interact must first be determined. To this end, we carried out an extensive search of the Arabidopsis genome to identify all potential ubiquitin E2 enzymes by both bioinformatics and activity assays. To further define the function of Arabidopsis ubiquitin E2 enzymes, we examined E2-E3 specificity in in vitro ubiquitination assays with the Arabidopsis RING family of E3 ligases. We also compared the expression profiles of UBC- and RING-encoding genes in different Arabidopsis organs and developmental stages using a 70-mer oligo microarray. RESULTS The Arabidopsis Genome Is Predicted to Encode 37 Ubiquitin E2 Proteins Inspection of the Arabidopsis genome for predicted ubiquitin UBC domain-containing proteins identified a previously unannotated predicted protein (At3g24515) in addition to the 36 compiled either from previous activity assays or from predictions of the annotated genome at the time (Bachmair et al., 2001). Only nine of these have been demonstrated to be ubiquitin E2s through their ability to form a ubiquitin thioester or catalyze E1-dependent protein ubiquitination (summarized in Table I Table I. Summary of UBC domain-containing proteins, their nomenclature, and activity as ubiquitin E2 enzymes Previously unnamed UBC domain-containing Arabidopsis proteins were given UBC numbers and grouped together based on sequence similarity (see Fig. 1). InsolubleX, 6×His-tagged protein insoluble when expressed in E. coli; InsolubleY, 6×His-tagged protein insoluble when expressed in either E. coli or insect cells; InsolubleZ, UBC contains predicted transmembrane domain C terminal to UBC domain and protein is insoluble when expressed with predicted transmembrane domain, but soluble when expressed without it. References (as indicated in table by superscript numbers): 1, Sullivan and Vierstra, 1993; 2, Girod et al., 1993; 3, Van Nocker et al., 1996; 4, Bartling et al., 1993; 5, Criqui et al., 2002; 6, Yanagawa et al., 2004; 7, this study; 8, Girod and Vierstra, 1993; and 9, Van Nocker and Vierstra, 1993. Group No. . Gene UBC . AGI Code . Active in TEa . Active in Ubb . Comments . GenBank Accession No. . III 1 At1g14400 Yescd,1,7 Yes1,8,7 E3-ine,7, E3-depf,1,8 DQ027016 III 2 At2g02760 Yes7 Yes7 E3-in7 DQ027017 III 3 At5g62540 ndg Yes7 E3-in7 DQ027018 IV 4 At5g41340 Yesh,8,9 Yes7 E3-in7, E3-dep7 DQ027019 IV 5 At1g63800 nd Yes7 E3-in7 DQ027020 IV 6 At2g46030 nd Yes7 E3-in7 DQ027021 V 7 At5g59300 Yesh,3 nd V 13 At3g46460 Yesh,3 nd DQ027027 V 14 At3g55380 Yesh,3 nd DQ027028 VI 8 At5g41700 Yesh,2,d,7 Yes2,7 E3-dep7 DQ027022 VI 9 At4g27960 Yesd,6 nd DQ027023 VI 10 At5g53300 Yes7 Yes7 E3-dep7 DQ027024 VI 11 At3g08690 Yes7 Yes7 E3-dep7 DQ027025 VI 12 At3g08700 nd nd InsolubleX7 DQ027026 VI 28 At1g64230 Yes7 Yes7 E3-dep7 DQ027041 VI 29 At2g16740 nd Yes7 E3-dep7 DQ027042 VI 30 At5g56150 nd Yes7 E3-dep7 DQ027043 VII 15 At1g45050 Yesi,4 Noj,7 DQ027029 VII 16 At1g75440 nd No7 DQ027030 VII 17 At4g36410 nd No7 DQ027031 VII 18 At5g42990 nd No7 DQ027032 VIII 19 At3g20060 Yesd,5,7 No DQ027033 VIII 20 At1g50490 nd No DQ027034 IX 21 At5g25760 nd nd InsolubleY7 DQ027035 X 22 At5g05080 nd Yes7 E3-in7 DQ027036 XI 23 At2g16920 nd nd Not cloned XI 24 At2g33770 nd nd InsolubleY7 DQ027037 XI 25 At3g15355 nd nd InsolubleY7 DQ027038 XI 26 At1g53020 nd No7 DQ027039 XII 27 At5g50870 Yes7 No7 DQ027040 XIII 31 At1g36340 nd nd InsolubleX7 DQ027044 XIV 32 At3g17000 Yes7 nd InsolubleZ7 DQ027045 XIV 33 At5g50430 nd nd InsolubleZ7 DQ027046 XIV 34 At1g17280 nd Yes7 E3-dep7, InsolubleZ7 DQ027047 XV 35 At1g78870 nd Yes7 E3-dep7 DQ027048 XV 36 At1g16890 Yes7 Yes7 E3-dep7 DQ027049 XVI 37 At3g24515 nd No7 DQ027050 Group No. . Gene UBC . AGI Code . Active in TEa . Active in Ubb . Comments . GenBank Accession No. . III 1 At1g14400 Yescd,1,7 Yes1,8,7 E3-ine,7, E3-depf,1,8 DQ027016 III 2 At2g02760 Yes7 Yes7 E3-in7 DQ027017 III 3 At5g62540 ndg Yes7 E3-in7 DQ027018 IV 4 At5g41340 Yesh,8,9 Yes7 E3-in7, E3-dep7 DQ027019 IV 5 At1g63800 nd Yes7 E3-in7 DQ027020 IV 6 At2g46030 nd Yes7 E3-in7 DQ027021 V 7 At5g59300 Yesh,3 nd V 13 At3g46460 Yesh,3 nd DQ027027 V 14 At3g55380 Yesh,3 nd DQ027028 VI 8 At5g41700 Yesh,2,d,7 Yes2,7 E3-dep7 DQ027022 VI 9 At4g27960 Yesd,6 nd DQ027023 VI 10 At5g53300 Yes7 Yes7 E3-dep7 DQ027024 VI 11 At3g08690 Yes7 Yes7 E3-dep7 DQ027025 VI 12 At3g08700 nd nd InsolubleX7 DQ027026 VI 28 At1g64230 Yes7 Yes7 E3-dep7 DQ027041 VI 29 At2g16740 nd Yes7 E3-dep7 DQ027042 VI 30 At5g56150 nd Yes7 E3-dep7 DQ027043 VII 15 At1g45050 Yesi,4 Noj,7 DQ027029 VII 16 At1g75440 nd No7 DQ027030 VII 17 At4g36410 nd No7 DQ027031 VII 18 At5g42990 nd No7 DQ027032 VIII 19 At3g20060 Yesd,5,7 No DQ027033 VIII 20 At1g50490 nd No DQ027034 IX 21 At5g25760 nd nd InsolubleY7 DQ027035 X 22 At5g05080 nd Yes7 E3-in7 DQ027036 XI 23 At2g16920 nd nd Not cloned XI 24 At2g33770 nd nd InsolubleY7 DQ027037 XI 25 At3g15355 nd nd InsolubleY7 DQ027038 XI 26 At1g53020 nd No7 DQ027039 XII 27 At5g50870 Yes7 No7 DQ027040 XIII 31 At1g36340 nd nd InsolubleX7 DQ027044 XIV 32 At3g17000 Yes7 nd InsolubleZ7 DQ027045 XIV 33 At5g50430 nd nd InsolubleZ7 DQ027046 XIV 34 At1g17280 nd Yes7 E3-dep7, InsolubleZ7 DQ027047 XV 35 At1g78870 nd Yes7 E3-dep7 DQ027048 XV 36 At1g16890 Yes7 Yes7 E3-dep7 DQ027049 XVI 37 At3g24515 nd No7 DQ027050 a A thioester linkage between the E2 and ubiquitin was observed (examples shown in Fig. 2). b Active in ubiquitination and indicates the E2 showed activity in the transfer of ubiquitin to a peptide linkage (see Fig. 4). c Yes, Activity has been demonstrated. d Ub-UBC conjugate eliminated under sulfhydryl-reducing conditions indicating Ub-E2 thioester linkage, as shown in reference. e E3-in, Ubiquitination of the E2 or other proteins that occurs in the absence of added E3 for E3-independent activity (for an E2 with strong activity, see Fig. 3). f E3-dep, Ubiquitination that requires E3 for E3-dependent (for examples, see Fig. 4). g nd, Protein was not assayed for activity. h Ub, E2 conjugate eliminated under sulfhydryl-reducing conditions, indicating Ub-E2 thioester linkage, but data not shown in reference. i Ub-E2 conjugate not eliminated after incubation under sulfhydryl-reducing conditions, but with 100 mm Lys. j No, No activity was detected. Open in new tab Table I. Summary of UBC domain-containing proteins, their nomenclature, and activity as ubiquitin E2 enzymes Previously unnamed UBC domain-containing Arabidopsis proteins were given UBC numbers and grouped together based on sequence similarity (see Fig. 1). InsolubleX, 6×His-tagged protein insoluble when expressed in E. coli; InsolubleY, 6×His-tagged protein insoluble when expressed in either E. coli or insect cells; InsolubleZ, UBC contains predicted transmembrane domain C terminal to UBC domain and protein is insoluble when expressed with predicted transmembrane domain, but soluble when expressed without it. References (as indicated in table by superscript numbers): 1, Sullivan and Vierstra, 1993; 2, Girod et al., 1993; 3, Van Nocker et al., 1996; 4, Bartling et al., 1993; 5, Criqui et al., 2002; 6, Yanagawa et al., 2004; 7, this study; 8, Girod and Vierstra, 1993; and 9, Van Nocker and Vierstra, 1993. Group No. . Gene UBC . AGI Code . Active in TEa . Active in Ubb . Comments . GenBank Accession No. . III 1 At1g14400 Yescd,1,7 Yes1,8,7 E3-ine,7, E3-depf,1,8 DQ027016 III 2 At2g02760 Yes7 Yes7 E3-in7 DQ027017 III 3 At5g62540 ndg Yes7 E3-in7 DQ027018 IV 4 At5g41340 Yesh,8,9 Yes7 E3-in7, E3-dep7 DQ027019 IV 5 At1g63800 nd Yes7 E3-in7 DQ027020 IV 6 At2g46030 nd Yes7 E3-in7 DQ027021 V 7 At5g59300 Yesh,3 nd V 13 At3g46460 Yesh,3 nd DQ027027 V 14 At3g55380 Yesh,3 nd DQ027028 VI 8 At5g41700 Yesh,2,d,7 Yes2,7 E3-dep7 DQ027022 VI 9 At4g27960 Yesd,6 nd DQ027023 VI 10 At5g53300 Yes7 Yes7 E3-dep7 DQ027024 VI 11 At3g08690 Yes7 Yes7 E3-dep7 DQ027025 VI 12 At3g08700 nd nd InsolubleX7 DQ027026 VI 28 At1g64230 Yes7 Yes7 E3-dep7 DQ027041 VI 29 At2g16740 nd Yes7 E3-dep7 DQ027042 VI 30 At5g56150 nd Yes7 E3-dep7 DQ027043 VII 15 At1g45050 Yesi,4 Noj,7 DQ027029 VII 16 At1g75440 nd No7 DQ027030 VII 17 At4g36410 nd No7 DQ027031 VII 18 At5g42990 nd No7 DQ027032 VIII 19 At3g20060 Yesd,5,7 No DQ027033 VIII 20 At1g50490 nd No DQ027034 IX 21 At5g25760 nd nd InsolubleY7 DQ027035 X 22 At5g05080 nd Yes7 E3-in7 DQ027036 XI 23 At2g16920 nd nd Not cloned XI 24 At2g33770 nd nd InsolubleY7 DQ027037 XI 25 At3g15355 nd nd InsolubleY7 DQ027038 XI 26 At1g53020 nd No7 DQ027039 XII 27 At5g50870 Yes7 No7 DQ027040 XIII 31 At1g36340 nd nd InsolubleX7 DQ027044 XIV 32 At3g17000 Yes7 nd InsolubleZ7 DQ027045 XIV 33 At5g50430 nd nd InsolubleZ7 DQ027046 XIV 34 At1g17280 nd Yes7 E3-dep7, InsolubleZ7 DQ027047 XV 35 At1g78870 nd Yes7 E3-dep7 DQ027048 XV 36 At1g16890 Yes7 Yes7 E3-dep7 DQ027049 XVI 37 At3g24515 nd No7 DQ027050 Group No. . Gene UBC . AGI Code . Active in TEa . Active in Ubb . Comments . GenBank Accession No. . III 1 At1g14400 Yescd,1,7 Yes1,8,7 E3-ine,7, E3-depf,1,8 DQ027016 III 2 At2g02760 Yes7 Yes7 E3-in7 DQ027017 III 3 At5g62540 ndg Yes7 E3-in7 DQ027018 IV 4 At5g41340 Yesh,8,9 Yes7 E3-in7, E3-dep7 DQ027019 IV 5 At1g63800 nd Yes7 E3-in7 DQ027020 IV 6 At2g46030 nd Yes7 E3-in7 DQ027021 V 7 At5g59300 Yesh,3 nd V 13 At3g46460 Yesh,3 nd DQ027027 V 14 At3g55380 Yesh,3 nd DQ027028 VI 8 At5g41700 Yesh,2,d,7 Yes2,7 E3-dep7 DQ027022 VI 9 At4g27960 Yesd,6 nd DQ027023 VI 10 At5g53300 Yes7 Yes7 E3-dep7 DQ027024 VI 11 At3g08690 Yes7 Yes7 E3-dep7 DQ027025 VI 12 At3g08700 nd nd InsolubleX7 DQ027026 VI 28 At1g64230 Yes7 Yes7 E3-dep7 DQ027041 VI 29 At2g16740 nd Yes7 E3-dep7 DQ027042 VI 30 At5g56150 nd Yes7 E3-dep7 DQ027043 VII 15 At1g45050 Yesi,4 Noj,7 DQ027029 VII 16 At1g75440 nd No7 DQ027030 VII 17 At4g36410 nd No7 DQ027031 VII 18 At5g42990 nd No7 DQ027032 VIII 19 At3g20060 Yesd,5,7 No DQ027033 VIII 20 At1g50490 nd No DQ027034 IX 21 At5g25760 nd nd InsolubleY7 DQ027035 X 22 At5g05080 nd Yes7 E3-in7 DQ027036 XI 23 At2g16920 nd nd Not cloned XI 24 At2g33770 nd nd InsolubleY7 DQ027037 XI 25 At3g15355 nd nd InsolubleY7 DQ027038 XI 26 At1g53020 nd No7 DQ027039 XII 27 At5g50870 Yes7 No7 DQ027040 XIII 31 At1g36340 nd nd InsolubleX7 DQ027044 XIV 32 At3g17000 Yes7 nd InsolubleZ7 DQ027045 XIV 33 At5g50430 nd nd InsolubleZ7 DQ027046 XIV 34 At1g17280 nd Yes7 E3-dep7, InsolubleZ7 DQ027047 XV 35 At1g78870 nd Yes7 E3-dep7 DQ027048 XV 36 At1g16890 Yes7 Yes7 E3-dep7 DQ027049 XVI 37 At3g24515 nd No7 DQ027050 a A thioester linkage between the E2 and ubiquitin was observed (examples shown in Fig. 2). b Active in ubiquitination and indicates the E2 showed activity in the transfer of ubiquitin to a peptide linkage (see Fig. 4). c Yes, Activity has been demonstrated. d Ub-UBC conjugate eliminated under sulfhydryl-reducing conditions indicating Ub-E2 thioester linkage, as shown in reference. e E3-in, Ubiquitination of the E2 or other proteins that occurs in the absence of added E3 for E3-independent activity (for an E2 with strong activity, see Fig. 3). f E3-dep, Ubiquitination that requires E3 for E3-dependent (for examples, see Fig. 4). g nd, Protein was not assayed for activity. h Ub, E2 conjugate eliminated under sulfhydryl-reducing conditions, indicating Ub-E2 thioester linkage, but data not shown in reference. i Ub-E2 conjugate not eliminated after incubation under sulfhydryl-reducing conditions, but with 100 mm Lys. j No, No activity was detected. Open in new tab and refs. therein). Therefore, one of the purposes of this study was to determine which UBC domain-containing proteins function as bona fide ubiquitin-conjugating (as opposed to ubiquitin-like) enzymes through activity assays. Arabidopsis ubiquitin E2 proteins were initially named according to their identity to yeast UBC domain-containing proteins of which 11 of the 13 function with ubiquitin (Bachmair et al., 2001). Because there cannot be an association of all UBC numbers between the two species given that Arabidopsis has many more UBC domain-containing proteins than yeast, we classified Arabidopsis UBC proteins into subgroups based on their identity to each other. Phylogenetic analysis was performed, including all yeast UBC proteins that function with ubiquitin, RUB, or SUMO, as well as representative human E2s. Also included in this analysis were Arabidopsis ubiquitin-conjugating E2 enzyme variant (UEV) proteins that contain a UBC domain but lack the active-site cysteinyl residue (Sancho et al., 1998). These include constitutive photomorphogenic AtCOP10 and seven other UEV proteins indicated by Arabidopsis Genome Initiative (AGI) numbers (Fig. 1 Figure 1. Open in new tabDownload slide Phylogenetic analysis of Arabidopsis E2s. Phylogenetic tree of the UBC domain from predicted UBC domain-containing proteins of Arabidopsis is shown. Included in the tree are all E2s from budding yeast (Sc, S. cerevisiae) and representative E2s from humans (Hs, Homo sapiens). Phylogenetic tree was generated using a full-heuristic search (PAUP, version 4.0), and Ufc1 from human and Arabidopsis was used as the outgroup (Komatsu et al., 2004). HsCroc-1 (UEV1) and ScSTP22 (VPS23) are ubiquitin-conjugating enzyme variants (Rothofsky and Lin, 1997; Li et al., 1999). Alignment was created from the core UBC domain as designated by Simple Modular Architecture Research Tool (http://smart.embl-heidelberg.de). The SUMO and RUB E2s as well as the Arabidopsis UBC-like UEV proteins are also included for comparison. Arabidopsis UEV proteins are listed by AGI number except for COP10 (Suzuki et al., 2002). The different Roman numeral designations indicate E2 subgroups (see Table I). Bootstrap values from 1,000 replications for each branch are shown. See Supplemental Figure 1 for protein sequence alignment used to generate the tree. Figure 1. Open in new tabDownload slide Phylogenetic analysis of Arabidopsis E2s. Phylogenetic tree of the UBC domain from predicted UBC domain-containing proteins of Arabidopsis is shown. Included in the tree are all E2s from budding yeast (Sc, S. cerevisiae) and representative E2s from humans (Hs, Homo sapiens). Phylogenetic tree was generated using a full-heuristic search (PAUP, version 4.0), and Ufc1 from human and Arabidopsis was used as the outgroup (Komatsu et al., 2004). HsCroc-1 (UEV1) and ScSTP22 (VPS23) are ubiquitin-conjugating enzyme variants (Rothofsky and Lin, 1997; Li et al., 1999). Alignment was created from the core UBC domain as designated by Simple Modular Architecture Research Tool (http://smart.embl-heidelberg.de). The SUMO and RUB E2s as well as the Arabidopsis UBC-like UEV proteins are also included for comparison. Arabidopsis UEV proteins are listed by AGI number except for COP10 (Suzuki et al., 2002). The different Roman numeral designations indicate E2 subgroups (see Table I). Bootstrap values from 1,000 replications for each branch are shown. See Supplemental Figure 1 for protein sequence alignment used to generate the tree. ). Ubiquitin-fold modifier 1-conjugating enzyme (Ufc1) was included to serve as the outgroup. HsUfc1 was recently shown to form a thioester linkage through a conserved cysteinyl residue to the ubiquitin-like protein ubiquitin-fold modifier 1 (Ufm1; Komatsu et al., 2004). Phylogenetic analysis revealed that Arabidopsis UBC domain-containing proteins, excluding the UEV proteins, can be divided into 16 subgroups based on >65% bootstrap support (Fig. 1), with groups I and II functioning in RUB1 and SUMO conjugation pathways, respectively. UEV proteins form several of their own distinct groups, supporting their hypothesized divergence from UBCs with active-site Cys (Fig. 1). Uncharacterized UBC domain-containing proteins, with the exception of the UEV proteins, were given a UBC number based on similarity to previously characterized AtUBCs or to other members of the same subgroup (Table I; Fig. 1). Most Arabidopsis UBC proteins shared highest similarity with another Arabidopsis protein, suggesting duplications; however, two Arabidopsis E2s displayed higher similarity to yeast and human proteins rather than to other Arabidopsis E2s (Bachmair et al., 2001; Fig. 1). AtUBC27 groups with ScUBC1 and HsUBCH1. AtUBC22 groups with a human protein, endemic pemphigus foliaceus (HsEPF5; Liu et al., 1992). Some groups did not have closely related yeast or human E2s. These are Arabidopsis UBC groups V, IX, XIII, and XVI, with Arabidopsis groups IX, XIII, and XVI consisting of a single member (Bachmair et al., 2001; Fig. 1). Most of the Arabidopsis UBCs do not have additional characterized protein-protein interaction domains. The single exception is UBC27, which has a predicted ubiquitin-binding domain called a ubiquitin-associated domain at its C terminus. Instead, several of the putative E2s contain either an acidic extension, a basic extension, or a predicted transmembrane domain (Bachmair et al., 2001). UBC4 to 6 contain acidic C termini, UBC32 to 34 contain a C-terminal-predicted transmembrane domain, and UBC22 contains a basic, Lys-rich C terminus. UBC23 and UBC24 have large N-terminal regions with no defined domains; however, they show significant similarity to a human E2, APOLLON, that contains a baculoviral inverted repeat and was recently shown to function as a chimeric E2-E3 (Hao et al., 2004). The domain structure of UBC26 (At1g53020), which is predicted to contain three UBC domains (Bachmair et al., 2001), was not detected in any other eukaryotic genome. However, multiple cDNA sequences found in GenBank contain only the N-terminal domain, suggesting that the other two domains are not part of the expressed open reading frame (ORF). No product was obtained when The Arabidopsis Information Resource (TAIR) prediction with three domains was used to design primers to isolate a cDNA via reverse transcription (RT)-PCR. Whether these other two UBC domains are expressed remains unresolved. The single expressed UBC domain of UBC26 was used in our studies. The remaining UBC proteins, with the exception of groups VI, IX, and XIV, consist basically of only the UBC domain with short extensions of unknown function. AtUBC8 clades strongly with seven other Arabidopsis E2s: UBC9 to 12 and 28 to 30 (Fig. 1), which form the largest Arabidopsis E2 subgroup, subgroup VI. AtUBC9 to 11 and 28 are very similar to UBC8, with 92% to 96% amino acid identity. UBC29 and 30 show 87% identity to UBC8, while UBC12 shows 78% identity to UBC8 (Fig. 1). These proteins are more similar to human HsUBC5a to c and yeast ScUBC4 and 5 than to other Arabidopsis UBCs (Fig. 1). Most Arabidopsis UBC Domain-Containing Proteins Function as Ubiquitin E2s The ability of seven uncharacterized UBC domain-containing proteins to carry thioester-linked ubiquitin was demonstrated (Fig. 2 Figure 2. Open in new tabDownload slide Thioester formation of select Arabidopsis E2s. A, UBC27 and UBC36 form DTT-sensitive ubiquitin adducts. Immunoblots with anti-ubiquitin and anti-6×His antibodies show the presence of a DTT-sensitive ubiquitin adduct for UBC36 and UBC27. B, Ubiquitin-conjugating enzymes from a variety of subgroups form DTT-sensitive ubiquitin adducts. Reactions were split after 5-min incubation at 37°C and treated with DTT or 8 m urea (−DTT). Reactions were resolved by SDS-PAGE and western blots were performed with indicated antibodies. Figure 2. Open in new tabDownload slide Thioester formation of select Arabidopsis E2s. A, UBC27 and UBC36 form DTT-sensitive ubiquitin adducts. Immunoblots with anti-ubiquitin and anti-6×His antibodies show the presence of a DTT-sensitive ubiquitin adduct for UBC36 and UBC27. B, Ubiquitin-conjugating enzymes from a variety of subgroups form DTT-sensitive ubiquitin adducts. Reactions were split after 5-min incubation at 37°C and treated with DTT or 8 m urea (−DTT). Reactions were resolved by SDS-PAGE and western blots were performed with indicated antibodies. ) and is summarized in Table I. For comparison, AtUBC1 (Sullivan and Vierstra, 1993), AtUBC8 (Girod et al., 1993), and AtUBC19 (Criqui et al., 2002) were included to demonstrate that our expression and purification procedure produced active protein. AtUBC2, 10, 11, 27, 28, 32, and 36 formed adducts with ubiquitin that were lost in the presence of a thiol-reducing agent, indicating that a thioester linkage was formed between ubiquitin and E2 (Fig. 2). These results identify these UBC domain-containing proteins as ubiquitin E2s. Another approach to demonstrate ubiquitin E2 activity is to determine whether a UBC domain-containing protein catalyzes E1-dependent protein ubiquitination. Previously uncharacterized Arabidopsis UBCs were expressed in Escherichia coli and tested for their ability to catalyze transfer of ubiquitin to substrate proteins. We were able to test 25 UBC domain-containing proteins for activity; however, we could not produce sufficient soluble protein to assay five others. UBC12, 21, 24, 25, and 31 were insoluble after expression in E. coli and UBC21, 24, and 25 were also insoluble when expressed in cultured insect cells, precluding any activity assays for these five (Table I). Several UBC proteins were sufficiently active in an E3-independent manner under our in vitro assay conditions to prevent their analyses in E3-dependent ubiquitination assays. UBC1 to 6 (data not shown), 20, and 22 (see below) transferred ubiquitin to proteins dependent upon E1, but independent of an E3 ubiquitin ligase. UBC19 exhibited some E3-independent ubiquitination, but it was low enough such that it would not mask E3-dependent activity (Fig. 4; data not shown). In the E3-dependent ubiquitination assays, the previously uncharacterized UBCs, UBC10, 11, 28, 29, 30, 34, 35, and 36, were active in catalyzing polyubiquitination with one or more RING E3 ligases (Fig. 4; Table II Table II. E3-E2 specificity as determined by in vitro ubiquitination assays Each E3 or GST-RING protein was tested in in vitro ubiquitination assays with a number of Arabidopsis E2s from different subfamilies as described and defined by Stone et al. (2005). Protein ubiquitination was observed (+) for a number of E2-E3 combinations. Two instances occurred where activity was inconsistent (±). For other E2-E3 combinations, protein ubiquitination was not detected (−). E2-E3 combinations that were not determined (nd) are indicated. E3 . RING Type . E2 (UBC) . . . . . . . . . . . . 8 . 10 . 11 . 28 . 29 . 30 . 19 . 34 . 35 . 36 . At1g02860 HCa − − − − − − − − − − At1g12760 H2 + + + + + − − − − − At1g14260 v + + + + − − − − − − At1g18760 D + + + + − − − − − − At1g22500 H2 + + + + + − − − − − At1g50440 v + + + − − ± − − − − At1g61620 HCa − − − − − − − − − − At1g63900 HCa + + + + + − − − − − At1g65430 HCb + + + − − − − − − − At1g68180 H2 + + + − + − − + nd − At1g74410 H2 − − − − − − − − − − At2g18670 H2 − − − − − − − − − − At2g22690 S/T − − − − − − − − − − At2g28530 C2 + − + − − − − − nd − At2g28840 HCa − − − − − − − − − − At2g42360 H2 + + + + + + − − + + At2g44330 H2 + + + + + + − − − − At2g47700 H2 + + + + + − − − + + At3g05250 HCa + + + + − − − − − − At3g05545 H2 + ± + − − − − − − − At3g06330 v + + + + + − − − − − At3g09760 v + + + − − − − − nd − At3g29270 HCa + + + + + − − − − − At3g45555 HCb − − − − − − − − nd − At3g47160 HCa + + + − − − − − − − At3g48070 C2 + + + − − − − − nd − At4g10160 H2 + + + − − − − + nd − At4g14220 H2 + + + + + − − − + + At4g27470 HCa + + + + + − − − − − At5g20910 H2 + + + + + + − − − − At5g37270 H2 + + + + + − − − + + At5g38070 v + + + − − − − − − − At5g42200 H2 + + + + + − − − − − At5g53910 D + + + − − − − − nd − E3 . RING Type . E2 (UBC) . . . . . . . . . . . . 8 . 10 . 11 . 28 . 29 . 30 . 19 . 34 . 35 . 36 . At1g02860 HCa − − − − − − − − − − At1g12760 H2 + + + + + − − − − − At1g14260 v + + + + − − − − − − At1g18760 D + + + + − − − − − − At1g22500 H2 + + + + + − − − − − At1g50440 v + + + − − ± − − − − At1g61620 HCa − − − − − − − − − − At1g63900 HCa + + + + + − − − − − At1g65430 HCb + + + − − − − − − − At1g68180 H2 + + + − + − − + nd − At1g74410 H2 − − − − − − − − − − At2g18670 H2 − − − − − − − − − − At2g22690 S/T − − − − − − − − − − At2g28530 C2 + − + − − − − − nd − At2g28840 HCa − − − − − − − − − − At2g42360 H2 + + + + + + − − + + At2g44330 H2 + + + + + + − − − − At2g47700 H2 + + + + + − − − + + At3g05250 HCa + + + + − − − − − − At3g05545 H2 + ± + − − − − − − − At3g06330 v + + + + + − − − − − At3g09760 v + + + − − − − − nd − At3g29270 HCa + + + + + − − − − − At3g45555 HCb − − − − − − − − nd − At3g47160 HCa + + + − − − − − − − At3g48070 C2 + + + − − − − − nd − At4g10160 H2 + + + − − − − + nd − At4g14220 H2 + + + + + − − − + + At4g27470 HCa + + + + + − − − − − At5g20910 H2 + + + + + + − − − − At5g37270 H2 + + + + + − − − + + At5g38070 v + + + − − − − − − − At5g42200 H2 + + + + + − − − − − At5g53910 D + + + − − − − − nd − Open in new tab Table II. E3-E2 specificity as determined by in vitro ubiquitination assays Each E3 or GST-RING protein was tested in in vitro ubiquitination assays with a number of Arabidopsis E2s from different subfamilies as described and defined by Stone et al. (2005). Protein ubiquitination was observed (+) for a number of E2-E3 combinations. Two instances occurred where activity was inconsistent (±). For other E2-E3 combinations, protein ubiquitination was not detected (−). E2-E3 combinations that were not determined (nd) are indicated. E3 . RING Type . E2 (UBC) . . . . . . . . . . . . 8 . 10 . 11 . 28 . 29 . 30 . 19 . 34 . 35 . 36 . At1g02860 HCa − − − − − − − − − − At1g12760 H2 + + + + + − − − − − At1g14260 v + + + + − − − − − − At1g18760 D + + + + − − − − − − At1g22500 H2 + + + + + − − − − − At1g50440 v + + + − − ± − − − − At1g61620 HCa − − − − − − − − − − At1g63900 HCa + + + + + − − − − − At1g65430 HCb + + + − − − − − − − At1g68180 H2 + + + − + − − + nd − At1g74410 H2 − − − − − − − − − − At2g18670 H2 − − − − − − − − − − At2g22690 S/T − − − − − − − − − − At2g28530 C2 + − + − − − − − nd − At2g28840 HCa − − − − − − − − − − At2g42360 H2 + + + + + + − − + + At2g44330 H2 + + + + + + − − − − At2g47700 H2 + + + + + − − − + + At3g05250 HCa + + + + − − − − − − At3g05545 H2 + ± + − − − − − − − At3g06330 v + + + + + − − − − − At3g09760 v + + + − − − − − nd − At3g29270 HCa + + + + + − − − − − At3g45555 HCb − − − − − − − − nd − At3g47160 HCa + + + − − − − − − − At3g48070 C2 + + + − − − − − nd − At4g10160 H2 + + + − − − − + nd − At4g14220 H2 + + + + + − − − + + At4g27470 HCa + + + + + − − − − − At5g20910 H2 + + + + + + − − − − At5g37270 H2 + + + + + − − − + + At5g38070 v + + + − − − − − − − At5g42200 H2 + + + + + − − − − − At5g53910 D + + + − − − − − nd − E3 . RING Type . E2 (UBC) . . . . . . . . . . . . 8 . 10 . 11 . 28 . 29 . 30 . 19 . 34 . 35 . 36 . At1g02860 HCa − − − − − − − − − − At1g12760 H2 + + + + + − − − − − At1g14260 v + + + + − − − − − − At1g18760 D + + + + − − − − − − At1g22500 H2 + + + + + − − − − − At1g50440 v + + + − − ± − − − − At1g61620 HCa − − − − − − − − − − At1g63900 HCa + + + + + − − − − − At1g65430 HCb + + + − − − − − − − At1g68180 H2 + + + − + − − + nd − At1g74410 H2 − − − − − − − − − − At2g18670 H2 − − − − − − − − − − At2g22690 S/T − − − − − − − − − − At2g28530 C2 + − + − − − − − nd − At2g28840 HCa − − − − − − − − − − At2g42360 H2 + + + + + + − − + + At2g44330 H2 + + + + + + − − − − At2g47700 H2 + + + + + − − − + + At3g05250 HCa + + + + − − − − − − At3g05545 H2 + ± + − − − − − − − At3g06330 v + + + + + − − − − − At3g09760 v + + + − − − − − nd − At3g29270 HCa + + + + + − − − − − At3g45555 HCb − − − − − − − − nd − At3g47160 HCa + + + − − − − − − − At3g48070 C2 + + + − − − − − nd − At4g10160 H2 + + + − − − − + nd − At4g14220 H2 + + + + + − − − + + At4g27470 HCa + + + + + − − − − − At5g20910 H2 + + + + + + − − − − At5g37270 H2 + + + + + − − − + + At5g38070 v + + + − − − − − − − At5g42200 H2 + + + + + − − − − − At5g53910 D + + + − − − − − nd − Open in new tab ; Supplemental Table III), demonstrating that they function as ubiquitin E2s (see below). Three groups of UBC proteins tested were not active under any assay conditions. All four members of group VII, UBC15 to 18, were expressed and soluble, but exhibited no self-ubiquitination, E3-independent, or E3-dependent ubiquitination (Fig. 4; data not shown; Table II). No similar evidence of any activity was obtained for UBC26 and 37. UBC37 was proteolyzed extensively when expressed in bacteria (data not shown). In summary, at least one representative from nine of the 14 UBC protein subgroups demonstrated E2 activity. Three subgroups had one member that produced sufficient soluble protein to test for activity, but did not exhibit activity: UBC26 in subgroup XI, UBC37 in subgroup XVI, and all members of subgroup VII. Two other subgroups could not be tested because they did not produce sufficient protein for ubiquitination assays (UBC21 and UBC31). UBC22 Strongly Promotes Ubiquitination Independent of an E3 Ligase UBC22 consistently showed strong E3-independent activity in in vitro ubiquitination assays when provided with E1 (Fig. 3 Figure 3. Open in new tabDownload slide Biochemical analysis of UBC22 activity. A, UBC22 shows polyubiquitination activity independent of the presence of an E3. The pattern of ubiquitination is identical between reactions lacking an E3 and those with the indicated E3. CIP8 and AGI names (e.g. At5g14420) represent E3 ligases added to the reaction. C (complete) indicates a reaction containing all the necessary components: E1, E2, E3, ubiquitin, and ATP. B, Processive addition of ubiquitin moieties to UBC22. Immunoblot with anti-6×His antibodies shows addition of multiple ubiquitins to UBC22 over times indicated. Figure 3. Open in new tabDownload slide Biochemical analysis of UBC22 activity. A, UBC22 shows polyubiquitination activity independent of the presence of an E3. The pattern of ubiquitination is identical between reactions lacking an E3 and those with the indicated E3. CIP8 and AGI names (e.g. At5g14420) represent E3 ligases added to the reaction. C (complete) indicates a reaction containing all the necessary components: E1, E2, E3, ubiquitin, and ATP. B, Processive addition of ubiquitin moieties to UBC22. Immunoblot with anti-6×His antibodies shows addition of multiple ubiquitins to UBC22 over times indicated. ). The pattern of ubiquitination observed for 6×His-tagged UBC22 (His-UBC22) does not change with the addition of a number of different types of glutathione S-transferase (GST)-RING E3s (Fig. 3). Conversion of the UBC22 catalytic Cys to Ala results in a loss of protein ubiquitination, thus indicating that the observed activity requires an active UBC22 (data not shown). To determine whether His-UBC22 is self-ubiquitinated, a time-course assay was conducted, followed by western blotting with anti-6×His antibodies to visualize accumulation of His-UBC22 linked to one or more ubiquitin moieties. Figure 3B shows the accumulation of a higher Mr UBC22 protein over time, indicating the ubiquitination of His-UBC22. Specificity of E2-E3 Polyubiquitination Previous work in our laboratory tested the ability of recombinant RING E3 ligases expressed as GST fusions to catalyze polyubiquitination in vitro with the 6×His-tagged recombinant AtUBC8 (Stone et al., 2005). While the majority of RING proteins were active in this assay, 19 of the 64 RING proteins tested were inactive. There are several reasons for this inactivity. One possibility is that these RING proteins do not function with AtUBC8, the only E2 tested, but instead with other UBC proteins. Therefore, we determined the ability of RING proteins to catalyze ubiquitination in vitro with other UBC proteins representative of the E2 diversity in Arabidopsis. In addition, we asked whether there was any specificity between the RING domain type and the E2 subgroup in catalyzing polyubiquitination. The 19 GST-RING proteins inactive with AtUBC8 were tested both with other members of the AtUBC8 subgroup as well as with more diverged UBCs. Two GST-RING proteins, At2g15580 and At2g28840, produced variable results with very low levels of higher Mr ubiquitinated species, so no conclusions regarding the activity of these two were made. The remaining 17 were not active with any of the E2s tested (Table II). The RING domain is required for E2 interaction and Arabidopsis encodes three canonical RING domain types, as well as five modified RING domain types (Stone et al., 2005). To determine whether a particular RING domain functions with a specific E2 or E2 group, 46 RING proteins active with AtUBC8 (Stone et al., 2005) representing all RING domain types, including the previously characterized CIP8 (Hardtke et al., 2002), were tested for activity with a spectrum of E2s. Fifteen E2s from six different subgroups were used in the analysis. Representative blots illustrating the results obtained from in vitro ubiquitination assays are shown in Figure 4 Figure 4. Open in new tabDownload slide In vitro ubiquitination assays. Each GST-RING protein (E3) was tested against various E2s in in vitro ubiquitination assays to determine the E2-E3 specificity. Representative anti-ubiquitin blots from the ubiquitination assays are shown. Unconjugated ubiquitin migrates at 5.5 kD and has run off the bottom of the gel. Arabidopsis E2 enzymes used in each assay are indicated above each lane by their UBC number and minus (−) indicates the absence of any E2 enzyme. E2-E3 activity is confirmed by the presence of a smear of ubiquitinated proteins, as indicated by plus (+) below each lane, detected by anti-ubiquitin antibodies. The absence of a smear of ubiquitinated proteins greater than the no-E2 lane indicates that the E2-E3 combination used was not active, as indicated by minus (−) below each lane. Figure 4. Open in new tabDownload slide In vitro ubiquitination assays. Each GST-RING protein (E3) was tested against various E2s in in vitro ubiquitination assays to determine the E2-E3 specificity. Representative anti-ubiquitin blots from the ubiquitination assays are shown. Unconjugated ubiquitin migrates at 5.5 kD and has run off the bottom of the gel. Arabidopsis E2 enzymes used in each assay are indicated above each lane by their UBC number and minus (−) indicates the absence of any E2 enzyme. E2-E3 activity is confirmed by the presence of a smear of ubiquitinated proteins, as indicated by plus (+) below each lane, detected by anti-ubiquitin antibodies. The absence of a smear of ubiquitinated proteins greater than the no-E2 lane indicates that the E2-E3 combination used was not active, as indicated by minus (−) below each lane. and a subset of the data is summarized in Table II. All results obtained from our in vitro assays are shown in Supplemental Table III. Of those E2s that showed activity in these assays, UBC34 (group XIV) had the highest specificity because its activity was only observed with two E3s (At1g68180 and At4g10160), both RING-H2 proteins (Fig. 4; Table II). UBC35 and/or 36 (group XV) displayed activity with only seven of the 45 active RING E3s and were typically both active with the same RING subtype, either RING-HCa or RING-H2 types (Fig. 4; Table II). The most generic E2s were members of the UBC8 group. This group was the only group active with both canonical and modified RING types. The relative order of activity was UBC8 = UBC10 = UBC11 > UBC28 > UBC29 > UBC30 (Supplemental Table III), with UBC30 active with the least number of E3s (eight) and UBC8, 10, and 11 active with the most (46, 44, and 46, respectively). UBC8, 10, and 11 were typically active with the same E3s, although there are a few exceptions. Two E3s active with UBC8 were found to be inactive or have questionable activity with UBC10 (At2g28530 and At3g05545; Table II). Expression of UBCs in Arabidopsis Organs To determine in which organs a particular UBC might function, the relative level of expression of 33 E2s on oligoarrays in selected organs and under different developmental conditions was determined as described previously (Ma et al., 2005). All eight members of the UBC8 family exhibited significant, but varying, levels of expression within each organ and environmental condition examined (Fig. 5 Figure 5. Open in new tabDownload slide Expression analysis of the UBC8 family of ubiquitin-conjugating enzymes. Relative expression level of UBC8 family members in the indicated organs. Determination of relative expression levels of each E2 gene is as described in “Materials and Methods.” Supplemental Table I contains a complete list of Arabidopsis UBC expression levels in all organ types examined. RL, Rosette leaves; RD, roots in dark-grown seedlings; RW, roots in light-grown seedlings; CD, cotyledons in dark-grown seedlings; CW, cotyledons in light-grown seedlings; SE, sepals; PE, petals; SM, stamens; and P1B, pistils 1 d before pollination. Figure 5. Open in new tabDownload slide Expression analysis of the UBC8 family of ubiquitin-conjugating enzymes. Relative expression level of UBC8 family members in the indicated organs. Determination of relative expression levels of each E2 gene is as described in “Materials and Methods.” Supplemental Table I contains a complete list of Arabidopsis UBC expression levels in all organ types examined. RL, Rosette leaves; RD, roots in dark-grown seedlings; RW, roots in light-grown seedlings; CD, cotyledons in dark-grown seedlings; CW, cotyledons in light-grown seedlings; SE, sepals; PE, petals; SM, stamens; and P1B, pistils 1 d before pollination. ). UBC10 was found to have a high level of expression in all organs, with the highest levels in rosette leaves, roots, and petals (Fig. 5, right). Similar to UBC10, UBC8 displayed a consistently high level of expression in most organs, with significantly higher expression in rosette leaves and petals than all other organs and conditions tested (Fig. 5, middle). UBC9, 11, 28, and 29 also display significant levels of expression in all organs examined (Fig. 5, left). The expression levels of UBC12 and 30 are extremely low in all organs. UBC11 is expressed mainly in petals with an approximately 5-fold higher level of expression than observed in rosette leaves. UBC28 is expressed at a consistent level between rosette leaves, stamens, and petals, while UBC29 expression is significantly lower in rosette leaves than in cotyledons, roots, sepals, and petals. These data were compared to that compiled at Genevestigator (Zimmerman et al., 2004; https://www.genevestigator.ethz.ch). The expression patterns of UBC28, 29, and 30 were, in general, consistent with the oligoarray data. Similarly, UBC12 expression was very low. In contrast, UBC8, 9, and 11 differed slightly from the data in this study. While UBC8 showed strong expression in leaf tissue, it did not show equivalently strong expression in petals. UBC9 and 11 showed a more general expression pattern. The expression analyses for the other E2 subgroups collectively show expression in most organ and environmental conditions examined (Supplemental Table I). A predicted pseudogene with a UBC domain, At1g35700, showed a relative expression level that was extremely low in all organs (Supplemental Table I). These data give further support to the conclusion that At1g35700 is a pseudogene and was excluded from the UBC nomenclature. Data obtained in our study for representative UBC genes are shown in Supplemental Figure 2A; corresponding data obtained from Genevestigator are included in Supplemental Figure 2B. Interestingly, none of the members within a subgroup showed coordinated expression, suggesting complex and distinct regulation of UBC mRNA levels. Analysis of stress-response data from Genevestigator (https://www.genevestigator.ethz.ch) shows that several E2 mRNAs were up-regulated at least 3-fold in response to various stimuli. Syringolin, a cell death-inducing chemical, induced UBC3, 11, 13, and 27 (data not shown). Several E2 mRNAs were induced by biotic stresses, including UBC17, 20, and 31. The herbicide isoxaben induced UBC20 and 22. UBC6 was induced in response to senescence. UBC16 was induced in response to 6-benzyl adenine and cycloheximide. UBC24 was induced under low nitrogen conditions. UBC31 was induced in high Glc/Suc conditions. UBC32 was induced in abscisic acid treatment and osmotic stress. Once again, distinct regulation of individual members of each UBC subgroup is seen since none showed coordinated changes in mRNA abundance. Expression Profiling of the Arabidopsis RING-Type E3 Ligases The large number of ubiquitin ligases account for the majority of ubiquitination components encoded by the Arabidopsis genome. The translated Arabidopsis genome is predicted to contain over 450 RING-type E3 ligases that can be grouped into eight subgroups based on the type of RING domain (Stone et al., 2005). To verify that the currently annotated RING genes (Stone et al., 2005) are transcribed, we examined the expression level of the RING E3 family in the same organ samples (Supplemental Table II). Of the 469 genes predicted to encode for RING domain-containing proteins (RING genes), 430 were analyzed on the same microarrays and all 430 could be detected to some extent in at least one of the 17 organs examined. The percentage of RING genes expressed varied from one organ to another, indicating that global RING gene expression was enriched in specific organs (Fig. 6A Figure 6. Open in new tabDownload slide Expression analysis of the Arabidopsis RING genes. A, Relative expression levels of RING genes in each organ as well as in cultured cells. Expression levels of RING genes in roots, cotyledons, and hypocotyls of light- and dark-grown seedlings were also determined. Number as well as percentage of RING genes with expression levels within the indicated range are given. B, Average level of expression of RING genes in each organ. The number of genes with above-average expression levels is shown. Determination of relative expression levels of each RING gene is as described in “Materials and Methods.” C, Average level of UBC expression in each organ type. CL, Cauline leaves; RL, rosette leaves; P1B, pistil 1 d before pollination; P1A, pistil 1 d after pollination; CW, cotyledons in light-grown seedlings; CD, cotyledons in dark-grown seedlings; HW, hypocotyl in light-grown seedlings; HD, hypocotyl in dark-grown seedlings; RW, roots in light-grown seedlings; RD, roots in dark-grown seedlings; SD, seed; PE, petal; SM, stamen; SE, sepals; ST, stem; S8P, siliques 8 d after pollination; S3P, siliques 3 d after pollination; CC, cultured cells. Figure 6. Open in new tabDownload slide Expression analysis of the Arabidopsis RING genes. A, Relative expression levels of RING genes in each organ as well as in cultured cells. Expression levels of RING genes in roots, cotyledons, and hypocotyls of light- and dark-grown seedlings were also determined. Number as well as percentage of RING genes with expression levels within the indicated range are given. B, Average level of expression of RING genes in each organ. The number of genes with above-average expression levels is shown. Determination of relative expression levels of each RING gene is as described in “Materials and Methods.” C, Average level of UBC expression in each organ type. CL, Cauline leaves; RL, rosette leaves; P1B, pistil 1 d before pollination; P1A, pistil 1 d after pollination; CW, cotyledons in light-grown seedlings; CD, cotyledons in dark-grown seedlings; HW, hypocotyl in light-grown seedlings; HD, hypocotyl in dark-grown seedlings; RW, roots in light-grown seedlings; RD, roots in dark-grown seedlings; SD, seed; PE, petal; SM, stamen; SE, sepals; ST, stem; S8P, siliques 8 d after pollination; S3P, siliques 3 d after pollination; CC, cultured cells. ). Eighty-nine percent of RING genes examined were expressed in seeds, while over 99% of the total RING genes on the array were expressed in floral organs such as petals, stamens, and sepals. All 430 genes examined were expressed at some level in roots of dark-grown seedlings, hypocotyls of light-grown seedlings, and cotyledons of both dark- and light-grown seedlings. Over one-half of all RING genes displayed moderate levels of expression in each organ (Fig. 6A). Only seven samples had a small number (1–13) of RING genes expressing in the highest category (>5,000): cauline and rosette leaves, petals, cultured cells, light-grown roots, and light- and dark-grown cotyledons. Organs with greater than 25 RING genes in the next highest level of expression (1,001–5,000) are also rosette leaves, petals, cultured cells, light-grown roots, light- and dark-grown cotyledons, and, additionally, stamens and dark-grown roots (Fig. 6A). Petals had the largest number of RING genes (85) with the two highest categories of expression, while only two RING genes exhibited similar levels of expression in seeds and pistils 1 d after pollination (Fig. 6, A and B). Similarly, additional organs also had a low number (4–12) of highly expressed RING genes, such as hypocotyls, siliques, and sepals. Interestingly, many of the same RING genes account for the high or low levels of expression in almost all organs. For example, At1g71980 has above-average level of expression in all organs, while At1g74370 has a low level of expression in all organs examined (Fig. 7 Figure 7. Open in new tabDownload slide Relative expression levels of representative RING genes. Expression levels of previously characterized RING genes, ARI8, ARI9, and ARI16, are shown. At5g10650 is expressed predominantly in floral organs. This type of organ-specific pattern of expression is observed for a significant number of RING genes. At1g74370 and At1g71980 illustrate, respectively, the low and high levels of expression observed for a number of RING genes. Determination of relative expression levels of each RING gene is as described in “Materials and Methods.” See Supplemental Table II for relative expression levels of all RING genes analyzed. Sample abbreviations are as designated in Figure 6. Figure 7. Open in new tabDownload slide Relative expression levels of representative RING genes. Expression levels of previously characterized RING genes, ARI8, ARI9, and ARI16, are shown. At5g10650 is expressed predominantly in floral organs. This type of organ-specific pattern of expression is observed for a significant number of RING genes. At1g74370 and At1g71980 illustrate, respectively, the low and high levels of expression observed for a number of RING genes. Determination of relative expression levels of each RING gene is as described in “Materials and Methods.” See Supplemental Table II for relative expression levels of all RING genes analyzed. Sample abbreviations are as designated in Figure 6. ). To get some insight into the relative activity of ubiquitin pathway enzymes, the mRNA levels of the UBCs and RING E3s in different organs and developmental stages were compared. The average expression level of the UBC genes reflects that of the RING genes, with similarly high levels of expression observed in organs such as rosette leaves, petals, and roots (compare Fig. 6, B and C). The majority of RING and UBC genes examined display low levels of expression in a number of organs, including seeds, pistils, siliques, hypocotyls, and cauline leaves. This type of expression pattern is illustrated by UBC1, 6, 27, and 35 (Supplemental Fig. 2A) and RING genes At1g71980 and ARI8 (Fig. 7). With the exception of the RING-D genes, this pattern of expression is observed for the majority of RING genes regardless of RING domain type. The RING genes encoding RING-D domain-containing proteins exhibited the lowest average expression levels in all organs and were expressed predominantly in seeds and petals (data not shown). To assist in validating the relative expression levels revealed by our analysis, we determined whether our results correlated with previously published data on the expression levels of known RING genes. To do this, we examined the expression levels of the ARI RING gene family whose mRNA abundance was determined previously (Mladek et al., 2003). The results we obtained correlated with the previously published expression data. For example, ARI8 expression was detected in all organs and developmental conditions (Fig. 7). Similar to the majority of RING genes, ARI8 exhibited a higher level of expression in rosette leaves, stamens, petals, and roots. Previous studies detected very low levels of ARI9 expression below the level required for their quantitation (Mladek et al., 2003). In this study, ARI9 levels were also barely detectable (Fig. 7). ARI16 expression was reported to be specific to siliques (Mladek et al., 2003). A direct comparison to our data is difficult because the developmental stage of this sample is not known. Our analysis showed that there is no silique-specific expression early in silique maturation, but that ARI16 is predominantly expressed in seeds (Fig. 7). This result could be consistent with previous data if ARI16 expression is strong in developing seeds at later stages of silique development and is maintained in the mature seed. DISCUSSION The ubiquitin E1, E2, and E3 ligase enzymes are the enzymatic core of the ubiquitination pathway. The E2-E3 complex interacts with the substrate, catalyzing ubiquitin addition to the substrate. Thus, the expression, activity, localization, and selectivity of ubiquitin E2s and E3s are important parameters that can serve to regulate ubiquitination. The E2-E3 combination used in the ubiquitination reaction can also influence the fate of the substrate through determining the extent and nature of ubiquitin addition. The majority of substrates observed to date are modified by the attachment of a Lys-48-linked ubiquitin chain, which targets the substrate for degradation by the 26S proteasome. Substrates modified with other types of polyubiquitin chains linked via ubiquitin Lys-6, -11, -29, and -63, or modified with a single ubiquitin, face different or unknown fates. For example, the heterodimeric E2 UBC13/methyl methane sulfonate sensitivity 2 functions with the RING E3 TNF receptor-associated factor 6 to catalyze the formation of Lys-63 polyubiquitin chains involved in protein activation (Deng et al., 2000). Monoubiquitination of histone H2B via the UBC2/radiation insensitive 6-Brefeldin A sensitivity 1 E2-E3 pair is required for subsequent methylation of specific Lys on histone H3 (Wood et al., 2003). The function of other ubiquitin linkages in vivo is currently unknown. Specific E2-E3 combinations are also involved in specific functions. In yeast, UBC2/Rad6 along with the RING-type E3s Rad5 and Rad18 are implicated in DNA repair (Jentsch et al., 1987; Bailly et al., 1997). Therefore, identifying and characterizing the components and how they interact in catalyzing ubiquitination is critical for the understanding of E2-E3 combinations required for certain processes. In vitro ubiquitination assays with RING E3 ligases, while operating without a physiological substrate, appear to be accurate reflections of in vivo activity. The requirements for in vivo and in vitro activity, where compared, appear to be identical. For example, mutations in the mammalian RING proteins Mdm2 (Fang et al., 2000) and Praja1 (Sasaki et al., 2003) and the Arabidopsis RING SINAT5 (Xie et al., 2002) that affect in vitro activity also affect in vivo activity. With the aim toward understanding E2-E3 specificity, we have expressed in E. coli multiple UBC domain-containing proteins from Arabidopsis, demonstrated that many function as ubiquitin E2 enzymes, and, using in vitro ubiquitination assays, assessed E2-E3 specificity using a number of different types of RING domain-containing proteins. The E2 enzymes showed little specificity toward different types of RING domain-containing proteins. UBC8, 10, 11, and 28, which show considerable protein sequence similarity to each other, function in vitro with a large number of RING E3 enzymes assayed, including RING E3s with modified RING domains. The high level of expression observed for UBC8, 10, 11, and 28 correlates with the breadth of activity and suggests that these proteins may perform a general ubiquitination function in vivo. Other members of the UBC8 family were not as promiscuous. UBC29 and 30 did not show activity with any of the E3s that contain modified RING domains and only functioned with a fraction of the other RING E3s. UBC29 and 30 are not as similar to UBC8 as other family members and this may account for restricted activity of these E2 enzymes. The changes in the UBC domain of UBC29 and 30 may prevent them from interacting with the modified RING domains (Stone et al., 2005). Alternatively, or in combination with the above suggestion, the spacing variations and amino acid substitutions within the modified RING domains may also impair the interaction with the E2 UBC domain. Greater specificity is observed for UBC35 and 36; both E2s functioned mainly with E3s containing RING-H2 domains. Although UBC35 and 36 are highly similar and functioned with the same set of RING E3s in vitro, their expression patterns are different. Therefore, each E2 may promote protein ubiquitination in different organs or at different times during development. UBC34 functioned with only a few RING-H2 domains. This may be due to the amino acid sequence of the UBC domain that may only allow enhancement of ubiquitin transfer in combination with certain RING domains. In addition, UBC34 shares the greatest identity with E2s known to be endoplasmic reticulum (ER)-associated via their C-terminal transmembrane domain. However, it remains to be determined whether AtUBC34 or the RING proteins it works with in vitro actually localize to the ER and whether it serves a role in ER-associated degradation. The RING-H2 is the most prevalent type of RING protein, so it is not surprising that the majority of E2s function with proteins containing this type of RING domain. Another interesting observation with E2-E3 activity concerns the types of E2s we found to be active in our study. E2s of group VI and XV showed activity with the greatest number of E3s in this study. Members of these groups consist only of the core catalytic UBC domain. The remainder of the E2s of Arabidopsis, with the exception of UBC21, contains extensions of varying lengths outside the UBC domain or within the catalytic core itself, as in the case of group V members (for review, see Bachmair et al., 2001). These extra sequences outside the UBC domain may serve as a regulatory mechanism to preclude promiscuous activity with inappropriate E3s. Alternatively, these sequences could allow additional protein-protein interactions with other components required for ubiquitin transfer. Previous work with UBCH10 has shown that the E2 associates with the cullin subunit anaphase-promoting complex (APC2) and not with the RING protein (APC11) and requires APC2 in addition to APC11 to facilitate ubiquitination (Tang et al., 2001). This suggests that additional elements outside the RING domain-UBC domain interaction may be required for ubiquitination to occur. However, the region of UBCH10 required for this direct interaction remains to be determined. Limited studies of E2 activity with other types of E3 ligases have been performed. Mudgil et al. (2004) did not find a clear subgroup of E2s that showed preference for the U-box proteins tested. One U-box protein, AtPUB38, was active with AtUBC8, but not with AtUBC7, while another U-box protein, AtPUB18, had the opposite activity (Mudgil et al., 2004). This finding is also supported by analysis of mammalian U-box proteins where, of the five tested, three were active with human E2s, hUBC4 and hUBC5c, but two others were active with other UBCs (Hatakeyama et al., 2001). Arabidopsis HECT E3 ligases have been shown to be active with members of the Arabidopsis UBC8 subgroup (Bates and Vierstra, 1999), although the testing of other E2s was not extensive. The E2 enzymes that did not function in our in vitro assays with RING E3s can be divided into two classes, those that showed some E3-independent activity and those that did not. No E2 activity at all was observed for the latter class and the possible reasons for this are multiple. These proteins may not fold properly after expression in E. coli; they may require another type of E3 other than the RING types tested here; they may function with only a specific RING protein not yet tested; or they may be unable to transfer ubiquitin to nonphysiological substrates. Another possibility is that they may require cofactors such as UEVs that are not present in the in vitro assay to promote protein ubiquitination. The fact that these E2s may not conjugate ubiquitin, but instead conjugate a ubiquitin-like protein and require a different E1, should also be considered. For the former class, the E3-independent conjugation observed for a number of E2s suggests that these enzymes are capable of E1-dependent ubiquitin thioester formation. However, the inability of these E2s to catalyze E3-dependent polyubiquitination could be because they may not be able to function with the RING E3s tested, may not be able to transfer to a nonphysiological substrate, or may require a cofactor or UEV for E3-dependent activity. Self-ubiquitination of one E2, AtUBC22, was easily detected and occurred both in the presence and absence of a RING E3. Whether self-ubiquitination of AtUBC22 or other E2s represents an additional level of regulation in vivo in Arabidopsis is not known. However, there are a few examples from other organisms that suggest self-ubiquitination may serve an in vivo function. Yeast Cdc34p self-ubiquitination is thought to regulate its own levels (Skowyra et al., 1999), and the Drosophila E2 Vihar E2-C of APC is degraded during mitosis to slow cyclin B degradation (Mathe et al., 2004). To examine whether ubiquitin pathway enzyme expression at the mRNA level was coordinated, we analyzed E2 expression patterns along with that of over 400 RING E3 genes. Overall, specific UBC genes show a generic expression pattern with no UBC gene showing a 3-fold higher level of expression in a single specific organ based on our microarray analysis or data available through Genevestigator (Zimmerman et al., 2004). RING genes, in contrast, contained many examples whose expression was very specific and limited. At5g01520 and At1g23980 are expressed at over 3-fold greater levels in flowers and seeds, respectively, than any other organ examined. Surprisingly, expression of the UBC and RING genes in pistils, siliques, and seeds was reduced compared to other organs. The low expression of the ubiquitination enzymes may reflect a tighter regulation of UBC and RING genes in these organs. Another possibility is that the E2 and/or E3 enzymes utilized in these organs may have broad specificity. Alternatively, the total level of protein ubiquitination that occurs within seeds, for example, may be reduced. Therefore, the requirement for ubiquitinating enzymes would be reduced. The expression we observed for both UBC and RING genes generally correlates with previously reported expression data (Thoma et al., 1996; Jensen et al., 1998; Mladek et al., 2003). The ubiquitination system is hierarchical in that few E2 enzymes (dozens) exist in comparison to the large number of E3s (hundreds). In the case of Arabidopsis, two E1s and as many as 34 to 37 ubiquitin E2 enzymes are utilized by hundreds of different E3 enzymes to facilitate ubiquitination of their cognate target proteins. Therefore, it would not be surprising to find a requirement for additional proteins to help guide the specificity of E2-E3 interactions. MATERIALS AND METHODS Plant Material Seeds from Arabidopsis (Arabidopsis thaliana) ecotype Columbia (Col-0) were either sown on soil and grown under photoperiodic cycles of 16 h light and 8 h dark at 16°C with 50% relative humidity or seeds surface sterilized with 30% (v/v) bleach and 0.1% (v/v) Triton X-100 were grown on 1% (w/v) agar with 1× Murashige and Skoog and 1% (w/v) Suc under continuous light. Identification of Arabidopsis E2 Enzymes The UBC domain of Arabidopsis UBC8 was used in BLAST searches against the complete nonredundant Arabidopsis genome (TAIR, April 16, 2003; http://www.arabidopsis.org). The Simple Modular Architecture Research Tool database was used to analyze retrieved sequences (version 4.0, May 28, 2004; http://smart.embl-heidelberg.de) followed by manual inspection to confirm the presence of the complete UBC domain. The previously named UBC12 (Girod et al., 1993) corresponded to a cDNA for UBC10, so UBC12 here refers to a previously uncharacterized ORF that is closely related to UBC8. Sequence Analysis The ClustalX program was used to generate an alignment of the UBC protein sequence. The alignment was generated using a PAM350 protein matrix, with gap opening and gap extension penalty parameters of 35.0 and 0.75, respectively, in pairwise alignment and 15.0 and 0.3, respectively, in the multiple alignments (Thompson et al., 1997). MacClade sequence editor (Sinauer Associates) was used to manually edit the alignment (Supplemental Fig. 1). The rooted phylogenetic trees were created by PAUP* (Phylogenetic Analysis Using Parsimony, version 4.0; Sinauer Associates) using the heuristic search method with 1,000 bootstrap replicates. Microarray Hybridization and Expression Analysis The dataset described in Ma et al. (2005) was used in the analyses presented here. Data are represented as mean normalized intensity value after subtraction of the value from a set of 192 negative control oligos contained on the same slide (Ma et al., 2005). Relative expression was classified as low (<50), moderate (50–1,000), high (1,001–5,000), and highest (>5,000). Cloning of Arabidopsis UBCs and RING E3s Arabidopsis E2 cDNAs were cloned by RT reactions followed by PCR to amplify the predicted ORF for each UBC. RNA isolated from either Arabidopsis ecotype Col-0 10-d-old seedlings or floral tissue from 6- to 7-week-old plants was used. The Qiagen RNeasy plant RNA extraction kit was used to isolate total RNA, according to the manufacturer's instructions. Coding regions for 35 UBCs were first introduced into the Gateway entry vector, pDONR (Invitrogen), and the DNA sequence was determined. Attempts to isolate cDNA for UBC23 failed. Sequences of each UBC cDNA were compared to the predicted ORF available on TAIR (http://www.arabidopsis.org), the Arabidopsis genome annotation database. Sequences obtained for UBC7 were different from TAIR predictions, as previously reported (Bachmair et al., 2001). UBC37 has an incorrect nucleotide prediction listed in TAIR. This results in a codon change from Ser to Leu at nucleotide position 1,193. UBC26 was isolated based on cDNAs predicting only the N-terminal UBC domain to be expressed. TAIR suggests the presence of three UBC domains encoded by the computer-predicted cDNA. All other UBCs isolated matched TAIR-predicted ORFs and the sequences obtained are available from GenBank (Table I). cDNAs determined to be correct were introduced into the Gateway-compatible pDEST17 vector to allow for recombinant 6×His-tagged protein to be produced in and purified from Escherichia coli. Truncations of UBC32, 33, and 34 were made to remove C-terminal transmembrane domains to improve solubility. For mutation of the conserved Cys of UBC22, site-directed mutagenesis (Stratagene) was used to change the Cys codon to an Ala codon. pDONR vector containing the UBC22 cDNA was used as a PCR template. After verification by DNA sequencing, the mutated UBC22 cDNA was then cloned, via the Gateway system, into the pDEST17 vector. The RING E3 cDNA GST expression vectors were as described in Stone et al. (2005). Protein Expression, Purification, in Vitro Ubiquitination Assays, and Western-Blot Analysis 6×His UBC fusions were expressed in E. coli strain BL21 AI or BL21-pLysS. Transformed cells were grown at 37°C for 2 to 3 h or to an OD600 of 0.4 to 0.6 before induction with 0.2% Ara or 0.5 to 1.0 mm isopropylthio-β-galactoside, respectively, for 2 to 3 h at 37°C. Cells were harvested by centrifugation and lysed in lysis buffer containing 25 mm Tris-HCl, pH 7.5, 500 mm NaCl, 0.01% Triton X-100, and 5 mm imidazole. For purification, nickel-nitrilotriacetic acid agarose (Qiagen) was added to cleared lysates and incubated for 1 h at 4°C. Beads were then washed four times with wash buffer containing 25 mm Tris-HCl, pH 7.5, 300 mm NaCl, 0.01% Triton X-100, and 5 mm imidazole. 6×His fusion proteins were eluted with elution buffer containing 25 mm Tris-HCl, pH 7.5, 150 mm NaCl, and 0.01% Triton X-100 supplemented with 300 mm imidazole. Glycerol was added to the eluted protein to a final concentration of 40%. Proteins were stored at −80°C until needed. GST-RING fusions were expressed and purified from bacterial extracts as described in Stone et al. (2005). SDS-PAGE electrophoresis followed by Coomassie Blue staining and Bradford assays (Bio-Rad) were used to quantify purified proteins. Ubiquitination assays were carried out as previously described (Stone et al., 2005). Thioester Assay Thioester assays were performed in a total reaction volume of 30 μL, consisting of 50 mm Tris-HCl, pH 7.4, 10 mm MgCl2, 10 mm ATP, 100 ng rabbit E1 (Boston Biochem), 500 ng of recombinant E2s (2 μg for AtUBC32), and 10 μg ubiquitin (Boston Biochem). Reactions were split after incubation for 5 min at 37°C and terminated by SDS sample buffer with dithiothreitol (DTT) or 8 m urea sample buffer without DTT. Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers DQ027017 to DQ027050 (see Table I). ACKNOWLEDGMENTS We would like to thank Christina Tan for assistance with the ubiquitination assays and Mandy Hsia for recombinant E1; Andy Troy and Michael Kerber for their assistance with E2 analysis; and Jemma Jowett and Kate Dreher for comments on the manuscript, as well as other members of the Callis laboratory for helpful discussions. 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Cullman Jr. fellowship and L.M. was a long-term postdoctoral fellow of the Human Frontier Science Program. 2 These authors contributed equally to the paper. * Corresponding author; e-mail [email protected]; fax 530–752–3085. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Judy Callis ([email protected]). [W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.105.067983. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Haberer, Georg; Young, Sarah; Bharti, Arvind K.; Gundlach, Heidrun; Raymond, Christina; Fuks, Galina; Butler, Ed; Wing, Rod A.; Rounsley, Steve; Birren, Bruce; Nusbaum, Chad; Mayer, Klaus F.X.; Messing, Joachim

doi:

Linebarger, Carla R. Lyerly; Boehlein, Susan K.; Sewell, Aileen K.; Shaw, Janine; Hannah, L. Curtis

doi: 10.1104/pp.105.067637pmid: 16299180

Abstract ADP-glucose pyrophosphorylase (AGPase) is a key regulatory enzyme in starch biosynthesis. However, plant AGPases differ in several parameters, including spatial and temporal expression, allosteric regulation, and heat stability. AGPases of cereal endosperms are heat labile, while those in other tissues, such as the potato (Solanum tuberosum) tuber, are heat stable. Sequence comparisons of heat-stable and heat-labile AGPases identified an N-terminal motif unique to heat-stable enzymes. Insertion of this motif into recombinant maize (Zea mays) endosperm AGPase increased the half-life at 58°C more than 70-fold. Km values for physiological substrates were unaffected, although Kcat was doubled. A cysteine within the inserted motif gives rise to small subunit homodimers not found in the wild-type maize enzyme. Placement of this N-terminal motif into a mosaic small subunit containing the N terminus from maize endosperm and the C terminus from potato tuber AGPase increases heat stability more than 300-fold. ADP-Glc pyrophosphorylase (AGPase) is ubiquitous in starch-synthesizing plant tissues and plays an essential role in glycogen biosynthesis in bacteria (for review, see Preiss and Romeo, 1994; Preiss and Sivak, 1996). AGPase catalyzes the initial step in the starch biosynthetic pathway by converting ATP and α-Glc-1-P (G-1-P) to ADP-Glc and pyrophosphate (for review, see Hannah, 1997). Seminal evidence for the crucial role AGPase plays in starch synthesis was derived from transgenic experiments in which altered AGPases were expressed in plants. An increase in starch synthesis was found in potato (Solanum tuberosum) tubers (Stark et al., 1992) and seeds of wheat (Triticum aestivum; Smidansky et al., 2002), rice (Oryza sativa; Smidansky et al., 2003), and maize (Zea mays; Giroux et al., 1996; T.W. Greene and L.C. Hannah, unpublished data). Accordingly, the stability and regulatory properties of AGPase have been subjects of intense study. Plant AGPases are heterotetramers composed of two small subunits and two large subunits. While the two subunits are not interchangeable, both evolved from a common progenitor (Bae et al., 1990; Bhave et al., 1990) and each exhibits a low level of activity when expressed alone in an Escherichia coli expression system (Iglesias et al., 1993; Burger et al., 2003). Mutations affecting catalytic and allosteric properties of AGPase map to both subunits (Cross et al., 2004, 2005; Hwang et al., 2005). Modulation of plant AGPase activity can involve several mechanisms: allosteric regulation by small effecter molecules, thermal inactivation, and reductive activation. AGPase is activated by 3-phosphoglyceric acid (3-PGA) and inhibited by inorganic phosphate (Pi) in many plant tissues. 3-PGA also can inhibit activity at high concentrations (Hwang et al., 2005). Several AGPases, such as the potato tuber enzyme, are fully stable at 70°C (Sowokinos and Preiss, 1982; Okita et al., 1990), whereas others are quite labile. The maize endosperm AGPase loses 96% of its activity when heated at 57°C for 5 min (Hannah et al., 1980). Temperature extremes are responsible for reduced grain yield in many cereal crops of worldwide importance, such as maize, wheat, and rice (Singletary et al., 1994). AGPase is one of the enzymes most profoundly affected by elevated temperature (Singletary et al., 1993, 1994). Since AGPase is rate limiting in starch biosynthesis, high temperatures adversely affect starch production and, in turn, yield. Heat stability and reductive activation of the potato tuber AGPase involve a small subunit N-terminal Cys (Ballicora et al., 1995). This Cys enhances stability of the potato tuber AGPase at high temperatures, presumably by formation of a disulfide bridge between the two small subunits (Fu et al., 1998; Ballicora et al., 1999). The presence of this disulfide bridge was also observed in the crystal structure of an allosterically inhibited potato small subunit homotetramer (Jin et al., 2005). Although the disulfide bridge conveys enzyme stability, it can have negative effects on enzyme activity. A two-step process has been proposed for maximal enzyme activity levels in the absence of 3-PGA. First, the disulfide bridge must be reduced and then an ADP-Glc-induced conformational change must occur (Fu et al., 1998; Tiessen et al., 2002). However, the specific activity of the reductively activated enzyme is 13-fold lower in the ADP-Glc synthesis reaction than that observed in the presence of 3-PGA (Fu et al., 1998). Comparison of heat-stable and heat-labile AGPases (Hannah et al., 2001) identified a conserved amino acid motif in the N terminus of the small subunit of heat-stable enzymes, designated QTCL (containing Gln, Thr, Cys, and Leu). This motif contains the Cys residue described above. This motif is absent in the heat-labile AGPases of the maize and barley (Hordeum vulgare) endosperms as well as other cereal seeds. Here, the QTCL motif of heat-stable AGPases and variants thereof were placed into the maize endosperm AGPase expressed in E. coli. The Cys causes formation of an intermolecular small subunit disulfide bond and greatly enhances heat stability apparently by stabilizing heterotetramers. The presence of the QTCL motif in the maize small subunit does not alter binding of physiological substrates or the physiological inhibitor, Pi; however, it does double Kcat and increases sensitivity to the activator, 3-PGA. Placement of the heat-stable motif into a recently described (Boehlein et al., 2005), Pi-insensitive mosaic subunit derived from maize and potato revealed complex interactions within the small subunit. While incorporation of the QTCL motif gave rise to an AGPase 300 times more heat stable than wild type, the Pi insensitivity of the maize/potato mosaic was nullified by the QTCL motif. RESULTS Identification of Motifs Important in Heat Stability Sequence alignments of heat-stable and heat-labile AGPase small subunits revealed pronounced sequence conservation (Hannah et al., 2001). However, significant sequence variation occurs in both termini. Of particular interest was the N-terminal motif QTCL (Fig. 1A Figure 1. Open in new tabDownload slide Alignment of small subunits and created mutations. A, Position of the first amino acid shown is given in parentheses following the small subunit origin. Sequence numbering is based on the maize endosperm small subunit. Maize and barley endosperm are heat sensitive while maize embryo, maize leaf, and potato tuber are heat stable. Amino acids examined here are in bold and boxed. Shaded areas indicated highly conserved amino acids. B, Changes in the N termini of the various mutants are listed. Inserted amino acids are in bold. Figure 1. Open in new tabDownload slide Alignment of small subunits and created mutations. A, Position of the first amino acid shown is given in parentheses following the small subunit origin. Sequence numbering is based on the maize endosperm small subunit. Maize and barley endosperm are heat sensitive while maize embryo, maize leaf, and potato tuber are heat stable. Amino acids examined here are in bold and boxed. Shaded areas indicated highly conserved amino acids. B, Changes in the N termini of the various mutants are listed. Inserted amino acids are in bold. ). This sequence motif occurs only in the heat-stable AGPases shown in Figure 1 and Hannah et al. (2001). The Cys residue in this motif has been implicated in conferring heat stability to the potato tuber AGPase (Ballicora et al., 1999). To establish whether this motif bestows heat stability in the otherwise heat-labile maize endosperm AGPase, four variants were created in the N terminus of the maize endosperm small subunit (Fig. 1B). The mutation STCL replaces the native endosperm Tyr residue with a Cys. The mutation termed QTCL adds a Gln residue and changes the Tyr residue to a Cys. To evaluate any alteration caused by the Gln insertion, two additional constructs were created, ETCL and QTYL. Each recombinant AGPase small subunit plus a wild-type maize endosperm large subunit on a separate, compatible vector was expressed in an E. coli mutant harboring an inactive AGPase. Functional AGPase encoded by the plasmids complements the mutant E. coli, resulting in glycogen production. This is easily detected by brown staining of colonies following exposure to iodine vapors. All variant small subunits were functional in E. coli (Fig. 2 Figure 2. Open in new tabDownload slide Staining of mutations. Cells were grown overnight and then exposed to iodine vapors for 1 min. Complementation is observed by the production of brown staining. The positive control (WT) is wild-type maize endosperm AGPase, and the negative control (NEG) is cells containing empty vectors. Figure 2. Open in new tabDownload slide Staining of mutations. Cells were grown overnight and then exposed to iodine vapors for 1 min. Complementation is observed by the production of brown staining. The positive control (WT) is wild-type maize endosperm AGPase, and the negative control (NEG) is cells containing empty vectors. ). In agreement with the in vivo-produced glycogen, AGPase activity levels of the variants were comparable to that of the recombinant maize enzyme (Table I Table I. Activity of small subunit mutations compared to the wild-type maize endosperm AGPase in crude preparationsa Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 100 100 ETCL 123 ± 25 130 ± 5 QTCL 170 ± 18 165 ± 7 STCL 135 ± 5 150 ± 5 QTYL 120 ± 5 135 ± 21 Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 100 100 ETCL 123 ± 25 130 ± 5 QTCL 170 ± 18 165 ± 7 STCL 135 ± 5 150 ± 5 QTYL 120 ± 5 135 ± 21 a Results are the averages of three independent experiments for each genotype. Assays within each experiment were performed in triplicate. The forward activity was measured in the direction of ADP-Glc synthesis in the presence of 10 mm 3-PGA. The nonradioactive reverse assay was used (see “Materials and Methods”). Open in new tab Table I. Activity of small subunit mutations compared to the wild-type maize endosperm AGPase in crude preparationsa Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 100 100 ETCL 123 ± 25 130 ± 5 QTCL 170 ± 18 165 ± 7 STCL 135 ± 5 150 ± 5 QTYL 120 ± 5 135 ± 21 Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 100 100 ETCL 123 ± 25 130 ± 5 QTCL 170 ± 18 165 ± 7 STCL 135 ± 5 150 ± 5 QTYL 120 ± 5 135 ± 21 a Results are the averages of three independent experiments for each genotype. Assays within each experiment were performed in triplicate. The forward activity was measured in the direction of ADP-Glc synthesis in the presence of 10 mm 3-PGA. The nonradioactive reverse assay was used (see “Materials and Methods”). Open in new tab ). AGPase activity from whole cell extracts was measured in both the forward and backward direction. Activity levels of the variants were moderately increased compared to the recombinant wild-type enzyme. To determine whether any of the mutations conferred heat stability, whole cell extracts were incubated for 6 min at 58°C, and the percentage of AGPase activity remaining was determined. All constructs containing the added Cys (ETCL, QTCL, and STCL) exhibited significant increases in heat stability. Cys-containing variants retained 30% to 50% of AGPase activity (Table II Table II. Percent heat stability of small subunit mutations in crude preparationa Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 2.4% ± 0.8 0.9% ± 1.2 ETCL 31.7% ± 2.5 44.5% ± 17.7 QTCL 50.0% ± 7.2 68.5% ± 0.7 STCL 44.3% ± 1.5 55.0% ± 5.7 QTYL 1.7% ± 1.1 1.4% ± 2.0 Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 2.4% ± 0.8 0.9% ± 1.2 ETCL 31.7% ± 2.5 44.5% ± 17.7 QTCL 50.0% ± 7.2 68.5% ± 0.7 STCL 44.3% ± 1.5 55.0% ± 5.7 QTYL 1.7% ± 1.1 1.4% ± 2.0 a Results are the averages of at least two independent experiments. Enzyme preparations were placed at 58°C for 6 min then submerged in ice prior to assay. Percentage heat stability is activity remaining after heat treatment divided by activity before heating. Open in new tab Table II. Percent heat stability of small subunit mutations in crude preparationa Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 2.4% ± 0.8 0.9% ± 1.2 ETCL 31.7% ± 2.5 44.5% ± 17.7 QTCL 50.0% ± 7.2 68.5% ± 0.7 STCL 44.3% ± 1.5 55.0% ± 5.7 QTYL 1.7% ± 1.1 1.4% ± 2.0 Sample . Assay A (Forward) . Assay B (Reverse) . STYL (wild type) 2.4% ± 0.8 0.9% ± 1.2 ETCL 31.7% ± 2.5 44.5% ± 17.7 QTCL 50.0% ± 7.2 68.5% ± 0.7 STCL 44.3% ± 1.5 55.0% ± 5.7 QTYL 1.7% ± 1.1 1.4% ± 2.0 a Results are the averages of at least two independent experiments. Enzyme preparations were placed at 58°C for 6 min then submerged in ice prior to assay. Percentage heat stability is activity remaining after heat treatment divided by activity before heating. Open in new tab ), compared to only 2% for those without the Cys (wild type and QTYL). In addition, the Gln within this motif may aid in enhancing heat stability of mosaic subunits containing the added Cys. Kinetic Analysis Because of its increased activity and heat stability, detailed kinetic analysis was performed on the QTCL variant. Recombinant QTCL and wild-type enzymes were purified as described previously (Boehlein et al., 2005). Km and Vmax values for ATP and for G-1-P were determined for both enzymes in the presence of 3-PGA (Table III Table III. Kinetic parameters of purified QTCL and wild-type AGPases Reactions were run with 10 mm 3-PGA. Mutant . ATP . . . G-1-P . . . . Km . Vmax . Kcat/Km . Km . Vmax . Kcat/Km . mm μmolmin−1mg−1 mm μmolmin−1mg−1 Wild typea 0.12 ± 0.003 26.7 ± 0.22 0.85 × 106 0.058 ± 0.001 23.2 ± 0.15 1.45 × 106 QTCL 0.10 ± 0.009 48.6 ± 1.18 1.82 × 106 0.042 ± 0.003 43.3 ± 0.65 3.77 × 106 MPa 0.10 ± 0.006 37.3 ± 0.65 1.41 × 106 0.050 ± 0.002 31.7 ± 0.34 2.3 × 106 MP-QTCL 0.09 ± 0.003 49.1 ± 0.52 1.91 × 106 0.051 ± 0.003 43.9 ± 0.67 3.15 × 106 Mutant . ATP . . . G-1-P . . . . Km . Vmax . Kcat/Km . Km . Vmax . Kcat/Km . mm μmolmin−1mg−1 mm μmolmin−1mg−1 Wild typea 0.12 ± 0.003 26.7 ± 0.22 0.85 × 106 0.058 ± 0.001 23.2 ± 0.15 1.45 × 106 QTCL 0.10 ± 0.009 48.6 ± 1.18 1.82 × 106 0.042 ± 0.003 43.3 ± 0.65 3.77 × 106 MPa 0.10 ± 0.006 37.3 ± 0.65 1.41 × 106 0.050 ± 0.002 31.7 ± 0.34 2.3 × 106 MP-QTCL 0.09 ± 0.003 49.1 ± 0.52 1.91 × 106 0.051 ± 0.003 43.9 ± 0.67 3.15 × 106 a Data were taken from Boehlein et al. (2005) and are included here for comparison. Open in new tab Table III. Kinetic parameters of purified QTCL and wild-type AGPases Reactions were run with 10 mm 3-PGA. Mutant . ATP . . . G-1-P . . . . Km . Vmax . Kcat/Km . Km . Vmax . Kcat/Km . mm μmolmin−1mg−1 mm μmolmin−1mg−1 Wild typea 0.12 ± 0.003 26.7 ± 0.22 0.85 × 106 0.058 ± 0.001 23.2 ± 0.15 1.45 × 106 QTCL 0.10 ± 0.009 48.6 ± 1.18 1.82 × 106 0.042 ± 0.003 43.3 ± 0.65 3.77 × 106 MPa 0.10 ± 0.006 37.3 ± 0.65 1.41 × 106 0.050 ± 0.002 31.7 ± 0.34 2.3 × 106 MP-QTCL 0.09 ± 0.003 49.1 ± 0.52 1.91 × 106 0.051 ± 0.003 43.9 ± 0.67 3.15 × 106 Mutant . ATP . . . G-1-P . . . . Km . Vmax . Kcat/Km . Km . Vmax . Kcat/Km . mm μmolmin−1mg−1 mm μmolmin−1mg−1 Wild typea 0.12 ± 0.003 26.7 ± 0.22 0.85 × 106 0.058 ± 0.001 23.2 ± 0.15 1.45 × 106 QTCL 0.10 ± 0.009 48.6 ± 1.18 1.82 × 106 0.042 ± 0.003 43.3 ± 0.65 3.77 × 106 MPa 0.10 ± 0.006 37.3 ± 0.65 1.41 × 106 0.050 ± 0.002 31.7 ± 0.34 2.3 × 106 MP-QTCL 0.09 ± 0.003 49.1 ± 0.52 1.91 × 106 0.051 ± 0.003 43.9 ± 0.67 3.15 × 106 a Data were taken from Boehlein et al. (2005) and are included here for comparison. Open in new tab , first two entries). An inspection of the Km values shows that the QTCL alteration has little to no effect on these parameters, indicating that this mutation does not have a direct role in substrate binding or catalysis. The alteration does affect catalytic efficiency as evidenced by the doubling of Vmax of the purified enzyme. An increased Vmax without changes in Km values was also noted in the absence of 3-PGA (data not shown). Allosteric properties of QTCL were also examined. Since the maize endosperm AGPase is activated by 3-PGA and deactivated by Pi, the Ka value for 3-PGA and the Ki value for Pi were determined. Resulting data show that the QTCL motif causes a small, but significant, increase in 3-PGA sensitivity (Table IV Table IV. Activation and inhibition of QTCL and wild type Assays were in the forward direction (assay C) using standard reaction conditions Mutant . 3-PGA . Pi . . Ka . Ki . Wild typea 0.17 ± 0.015 2.96 ± 0.12 QTCL 0.11 ± 0.013 2.62 ± 0.10 MPa 0.03 ± 0.008 12.28 ± 0.73 MP-QTCL 0.10 ± 0.014 2.87 ± 0.08 Mutant . 3-PGA . Pi . . Ka . Ki . Wild typea 0.17 ± 0.015 2.96 ± 0.12 QTCL 0.11 ± 0.013 2.62 ± 0.10 MPa 0.03 ± 0.008 12.28 ± 0.73 MP-QTCL 0.10 ± 0.014 2.87 ± 0.08 a Data were previously published in Boehlein et al. (2005). Ki for phosphate was determined in the presence of 2.5 mm 3-PGA. Open in new tab Table IV. Activation and inhibition of QTCL and wild type Assays were in the forward direction (assay C) using standard reaction conditions Mutant . 3-PGA . Pi . . Ka . Ki . Wild typea 0.17 ± 0.015 2.96 ± 0.12 QTCL 0.11 ± 0.013 2.62 ± 0.10 MPa 0.03 ± 0.008 12.28 ± 0.73 MP-QTCL 0.10 ± 0.014 2.87 ± 0.08 Mutant . 3-PGA . Pi . . Ka . Ki . Wild typea 0.17 ± 0.015 2.96 ± 0.12 QTCL 0.11 ± 0.013 2.62 ± 0.10 MPa 0.03 ± 0.008 12.28 ± 0.73 MP-QTCL 0.10 ± 0.014 2.87 ± 0.08 a Data were previously published in Boehlein et al. (2005). Ki for phosphate was determined in the presence of 2.5 mm 3-PGA. Open in new tab ). The extent of phosphate modulation was determined in the presence of 3-PGA. Comparable Ki values for QTCL and wild-type recombinant AGPase were detected. QTCL Disulfide Bridge Formation The Cys of QTCL has been shown to be involved in disulfide bridge formation between the two small subunits (Jin et al., 2005). Because the wild-type maize endosperm lacks this Cys and the QTCL variant contains it, both were monitored for disulfide bond formation. Purified recombinant wild-type and QTCL AGPases were subjected to a nondenaturing SDS-PAGE analysis, blotted, and probed with antibodies against the large or small AGPase subunit. Proteins recognized by these antibodies are approximately 50 and 100 kD, the sizes of monomers and dimers of the two subunits, respectively (Fig. 3 Figure 3. Open in new tabDownload slide Disulfide bridge formation. A nonreducing 10% SDS-PAGE gel followed by a western transfer was performed to investigate the formation of the disulfide bridge. A, Western blot developed with polyclonal small subunit. B, Western blot probed with a polyclonal large subunit antibody. Both antibodies recognize a dimer in QTCL and wild-type AGPase that depolymerizes in the presence of DTT. The small subunit dimer of QTCL is substantially more intense than that of wild type. Figure 3. Open in new tabDownload slide Disulfide bridge formation. A nonreducing 10% SDS-PAGE gel followed by a western transfer was performed to investigate the formation of the disulfide bridge. A, Western blot developed with polyclonal small subunit. B, Western blot probed with a polyclonal large subunit antibody. Both antibodies recognize a dimer in QTCL and wild-type AGPase that depolymerizes in the presence of DTT. The small subunit dimer of QTCL is substantially more intense than that of wild type. ). Exposure to dithiothreitol (DTT) prior to electrophoresis abolishes the 100-kD proteins and intensifies the 50-kD band. We conclude that disulfide bridges maintain dimers not only of the small subunit, but also of the large subunit. Heterotetrameric AGPase was not observed in the presence or absence of DTT on the SDS gels. The presence of QTCL in the maize small subunit does not alter the size or amounts of proteins recognized by the large subunit antibody (Fig. 3B); however, its presence dramatically enhances the amount of the small subunit in the 100-kD protein at the expense of the 50-kD protein. Because the dimer involving the small subunit is abolished by the reducing agent and is enriched by the addition of the Cys in the N terminus, a disulfide bridge involving the small subunit is involved in dimerization. The low-intensity 100-kD band detected with the small subunit polyclonal antibody in wild type was shown to be nonspecific binding. This band is absent when an identical blot is developed with a monoclonal small subunit antibody (data not shown). We suspect that the cross-reaction of the polyclonal small subunit antibody with the large subunit is due to sequence conservation between the subunits. Incorporation of the QTCL motif into the small subunit promotes dimer formation between small subunits but not between small and large subunits. Perusal of the two blots in Figure 3 shows that QTCL enhances the amount of the dimer detected with the small subunit antibody but not the dimer detected with large subunit antibody. Were heterodimers enhanced by the QTCL mutation, an increased intensity of both dimers would have been seen. We also identified a disulfide bond involving the large subunit in our purified extracts. Activation of QTCL by DTT in the Absence of 3-PGA Reduction of the Cys residue in the QTCL motif may represent a form of physiological control of the potato tuber AGPase (Ballicora et al., 2000; Tiessen et al., 2002). In the absence of 3-PGA, potato tuber AGPase is activated by preincubation with DTT and the substrate ADP-Glc (Fu et al., 1998). Accordingly, DTT and substrate modulation of recombinant maize endosperm AGPase containing or lacking the QTCL small subunit motif was investigated. Activities of the two AGPases were measured after preincubation with 3 mm DTT and/or 2.0 mm ADP-Glc (Fig. 4A Figure 4. Open in new tabDownload slide Reduction of the disulfide bridge. Reactions were incubated for 15 min at room temperature in 20 μL of a mixture containing 100 mm HEPES, pH 7.4, 0.2 mg/mL BSA, 5 mm MgCl2, and the specified treatment listed. Following preincubation, reactions were performed using assay B with the following modification. The ADP-Glc concentration was adjusted from 2.0 mm so that the final concentration in all assays was 1.4 mm. The concentration of DTT used was 3.0 mm. Each assay was performed from 0 to 6 min and the slope of the velocity versus time plot was used to calculate the specific activity. A, Activity (μmol min−1 mg−1) in the absence of 3-PGA. Black bars represent wild-type and cross-hatching QTCL. B, Activity (μmol min−1 mg−1) of QTCL in the absence (white bars) or presence (gray bars) of 10 mm 3-PGA. Figure 4. Open in new tabDownload slide Reduction of the disulfide bridge. Reactions were incubated for 15 min at room temperature in 20 μL of a mixture containing 100 mm HEPES, pH 7.4, 0.2 mg/mL BSA, 5 mm MgCl2, and the specified treatment listed. Following preincubation, reactions were performed using assay B with the following modification. The ADP-Glc concentration was adjusted from 2.0 mm so that the final concentration in all assays was 1.4 mm. The concentration of DTT used was 3.0 mm. Each assay was performed from 0 to 6 min and the slope of the velocity versus time plot was used to calculate the specific activity. A, Activity (μmol min−1 mg−1) in the absence of 3-PGA. Black bars represent wild-type and cross-hatching QTCL. B, Activity (μmol min−1 mg−1) of QTCL in the absence (white bars) or presence (gray bars) of 10 mm 3-PGA. ). Preincubation of the QTCL variant with DTT and/or ADP-Glc enhanced activity almost 3-fold; however, wild-type AGPase was only slightly affected. While the greatest enhancement of QTCL occurred when both DTT and ADP-Glc were present, the modulating effects of the two are not additive (Fig. 4A). Interestingly, activation by DTT and/or ADP-Glc is not obligatory for maximal activity since the presence of saturating concentrations of the activator 3-PGA during the reaction more than compensates for preincubation activation (Fig. 4B). QTCL Heterotetramer Formation Addition of QTCL to the N terminus of the small subunit confers substantial heat stability to the recombinant maize endosperm AGPase and enhances homodimer formation, as shown above. Next, the aggregation state of purified AGPase was monitored at various stages of heat inactivation (Fig. 5 Figure 5. Open in new tabDownload slide Heat stability of purified QTCL at 42°C. Enzymes were placed in a water bath at 42°C for varying times and then cooled on ice. Each enzyme was assayed for 10 min in the forward direction in the presence of 10 mm 3-PGA. Reactions were started with 0.065 μg of enzyme. Data were plotted as log% activity versus time (min), and the inactivation constant t1/2 was calculated as follows: slope = −k/(2.3). t1/2 is calculated from the equation k = 0.693/t1/2. A, Inactivation time course at 42°C. Symbols are as follows: ▪, QTCL mutant; and ▴, wild-type enzyme. B, BN-PAGE gel transferred and probed with small subunit antibody at the time points from A. T, The approximately 220-kD tetramer; D, the approximately 100-kD dimer; and M, the approximately 50-kD monomer. Figure 5. Open in new tabDownload slide Heat stability of purified QTCL at 42°C. Enzymes were placed in a water bath at 42°C for varying times and then cooled on ice. Each enzyme was assayed for 10 min in the forward direction in the presence of 10 mm 3-PGA. Reactions were started with 0.065 μg of enzyme. Data were plotted as log% activity versus time (min), and the inactivation constant t1/2 was calculated as follows: slope = −k/(2.3). t1/2 is calculated from the equation k = 0.693/t1/2. A, Inactivation time course at 42°C. Symbols are as follows: ▪, QTCL mutant; and ▴, wild-type enzyme. B, BN-PAGE gel transferred and probed with small subunit antibody at the time points from A. T, The approximately 220-kD tetramer; D, the approximately 100-kD dimer; and M, the approximately 50-kD monomer. ). Purified recombinant AGPases were incubated at 42°C for varying times and the levels of activity and their aggregation states were determined. The half-life of AGPase activity was increased approximately 10-fold at this temperature by addition of the QTCL motif, from 1.50 min to 13.0 min (Fig. 5A). Blue native (BN)-PAGE gels were used to investigate the aggregation state of the enzyme as a function of inactivation (Fig. 5B). Both AGPases exist primarily in the heterotetrameric state before exposure to elevated temperature. However, associated with the rapid loss of activity in wild type is the production of monomers and dimers. The heterotetrameric band is virtually abolished by 20 min, and the only proteins detected after this time are high-Mr aggregates. In contrast, the QTCL enzyme remains predominantly as a heterotetramer, even after a 30-min heat treatment. Note that bovine serum albumin (BSA) was not added to these samples to stabilize the activity of the enzymes; therefore, the half-life of QTCL at 42°C appears to be less than when incubated at 58°C. When this experiment was repeated with BSA, the half-life for the wild-type enzyme was approximately 4.5 min and the half-life for the QTCL mutant could not be determined due to its long stability over the course of the 45-min experiment (data not shown). Further Small Subunit Modifications Boehlein et al. (2005) recently reported an AGPase small subunit mosaic that contains amino acids 1 to 199 of the maize small subunit and 200 to 475 from the potato small subunit (MPss). Compared to the recombinant wild-type maize endosperm AGPase, this mosaic was recalcitrant to phosphate inhibition and exhibited increased heat stability. The half-life was 18 min at 42°C or 4 times greater than wild type. To determine whether there was an additive effect in heat stability, the QTCL mutation was placed into MPss. This new variant, termed QTCL-MP, was kinetically indistinguishable from the QTCL mutation (Table III). Interestingly, the increased 3-PGA sensitivity and decreased Pi inhibition caused by substituting the terminal 275 potato tuber small subunit amino acids for their counterparts of maize were nullified by the N-terminal QTCL mutation (Table IV). The QTCL-MP mutation is approximately 5 times more heat stable than QTCL and over 300 times more stable than the recombinant wild-type maize endosperm enzyme (Fig. 6 Figure 6. Open in new tabDownload slide Heat stability of purified QTCL, wild type, and QTCL-MP at 55°C. Reactions were carried out as described in Figure 5, except temperature was raised to 55°C. ▪, Wild type; ⋄, MP; ▵, QTCL; ∇, QTCL-MP. Figure 6. Open in new tabDownload slide Heat stability of purified QTCL, wild type, and QTCL-MP at 55°C. Reactions were carried out as described in Figure 5, except temperature was raised to 55°C. ▪, Wild type; ⋄, MP; ▵, QTCL; ∇, QTCL-MP. ). DISCUSSION Here, we report recombinant maize endosperm AGPases with greatly enhanced heat stability. Substitution of a motif that we term QTCL from small subunits of heat-stable AGPases into the otherwise heat-labile maize endosperm AGPase causes a 70-fold increase in heat stability at 55°C. In addition, this substitution conditioned two other changes that may be beneficial in the cereal endosperm. The turnover number of the enzyme was doubled and sensitivity to the activator 3-PGA was increased. Significantly, inhibition caused by phosphate was not altered, nor were Km values for the physiologically important substrates, G-1-P and ATP. Further enhancement of maize endosperm AGPase heat stability was accomplished by placement of the QTCL motif into a modified small subunit described recently by Boehlein et al. (2005). This mosaic subunit, termed MPss, contains amino acids 1 to 199 from the maize endosperm small subunit and amino acids 200 to 475 from the potato tuber subunit. Whereas substitution of MPss for the maize small subunit increases heat stability 4-fold, placement of QTCL into MPss increases heat stability over 300-fold. Previous work with the potato tuber AGPase (Fu et al., 1998) identified this Cys as being important in enzyme stability. The Cys residue in the potato tuber AGPase is also involved in the reductive activation of the enzyme. This is a two-part mechanism involving substrates and a reducing agent (Fu et al., 1998). Here we show that placement of this Cys into the recombinant maize endosperm AGPase gives rise to activation kinetics similar, although not identical, to those described for the potato AGPase. In the absence of 3-PGA, preincubation of the QTCL mutant with the substrate ADP-Glc and DTT gives rise to a 3.75-fold increase in activity. This does not occur with the wild-type maize endosperm enzyme, nor does it occur with either enzyme in the presence of the activator 3-PGA. One notable difference does distinguish activation of the maize AGPase from the reported potato tuber observations. Whereas preincubation of the potato enzyme with DTT reduces activity, the Cys-containing maize endosperm enzyme is activated by DTT preincubation at room temperature. We exploited a BN gel system recently modified by us for AGPases (Boehlein et al., 2005) to monitor changes in aggregation states of AGPase caused by the Cys substitution, as well as changes induced by high temperatures. In the absence of SDS and reducing agents, different polymers of AGPase ranging from monomers to higher order polymers can be detected. Polymers involving disulfide bridges are detected in the absence of DTT on nonreducing SDS gels, whereas AGPase monomeric subunits are seen when samples contained DTT. This disulfide bridge does not form in the wild-type maize small subunit, which lacks the QTCL motif. The formation of the disulfide bridge is interesting since potato and maize bear little similarity in the extreme N terminus, with maize having a 25-amino acid extension. A fundamental difference distinguishes the maize endosperm and the potato tuber AGPases. Whereas disulfide bridges form only between small subunits in the wild-type potato tuber AGPase (Fu et al., 1998), here we show that disulfide bridge formation occurs only between large subunits in the recombinant wild-type maize endosperm enzyme. In view of this, it is noteworthy that hybrid AGPases involving maize and potato subunits exhibit vastly different properties. Whereas the hybrid involving the potato small subunit with the maize large subunit (Pss/Mls) is more active than either parental AGPase, the hybrid involving the maize small subunit with the potato large subunit (Mss/Pls) exhibits barely detectable activity as judged by glycogen production in E. coli (Cross et al., 2004, 2005). The Pss/Mls hybrid could potentially form disulfide bridges both between the small subunits and between the large subunits, whereas Mss/Pls lacks the ability to form these disulfide bridges. In addition, it is interesting to note that a variant of the Mss/Pls enzyme that contains 55 potato-derived amino acids substituted into maize [Mos(1-321,377-457)] produces copious amounts of glycogen in E. coli. However, the AGPase produced by this variant dissociates rapidly following extraction (Cross et al., 2005). These observations suggest that the ability to form at least one disulfide bridge has been selected over evolutionary time. Due to the interchangeability of the subunits early in evolution, the ability to form a disulfide bridge was selected in the large subunit of the maize endosperm (and likely other cereal seed AGPases), whereas the small subunit in potato and likely other AGPases harbor the Cys residues involved in bond formation. Mutations in the small subunit of maize AGPase are not the only alterations that lead to increased heat stability. Greene and Hannah (1998), through mutation analysis, identified a large subunit heat-stable variant of the maize endosperm. This heat-stable mutation, termed HS33, functions by strengthening subunit interactions forming the heterotetramer. While HS33 enhances heat stability when combined with the wild-type small subunit, the enzyme containing HS33 is much less heat stable than the variants reported here. A detailed analysis of heat-induced activity loss and enzyme aggregation state shows that the QTCL variant not only enhances enzyme stability, but also aids in maintenance of the tetrameric state. A comparison of heat-induced changes in the aggregation state of wild type and QTCL shows that the QTCL enzyme remains as a heterotetramer much longer than does the wild-type enzyme. In addition, it appears that activity is lost in both wild type and QTCL before loss of the heterotetramer. Several roles for the N-terminal QTCL motif of the small subunit have now been identified. In addition to its role in disulfide bridge formation and enhanced heat stability, the data presented here show that it plays a significant role in Pi sensitivity. We (Boehlein et al., 2005) reported earlier that substitution of potato sequences into the carboxyl-terminal region of the small subunit effectively removed Pi inhibition. Here we show that incorporation of the QTCL motif into the Pi-insensitive enzyme restores Pi sensitivity. Hence, the QTCL motif plays an important role in modulating AGPase activity. The lack of this important motif in cereal endosperms of worldwide importance is an interesting observation. As noted previously (Geigenberger et al., 2005), the absence of this Cys indicates redox insensitivity. Here we show that its presence in the maize endosperm AGPase greatly enhances heat stability. It is possible that the QTCL motif has been under negative selection pressure in cereal seeds. Earlier, Greene and Hannah (1998) speculated that, under adverse temperature conditions, the heat lability of the endosperm AGPase might favor greater carbon flow into the embryo rather than the endosperm. This possibly enhances seed viability under adverse conditions. It is also interesting to note that expression in Arabidopsis leaves of the wild-type small subunit protein conjoined to a chloroplast transit peptide functionally substitutes for the endogenous small subunit gene (J. Zou, K. Folta, and L.C. Hannah, unpublished data). Hence, because the Arabidopsis small subunit contains the QTCL, this motif and the associated heat stability and reductive activation are not obligatory for normal starch synthesis in the Arabidopsis leaf. Starch production in cereals relies heavily on the function of AGPase. Thus, modifications of this enzyme can lead to enhancement or reduction in starch content (Stark et al., 1992; Giroux and Hannah, 1994; Giroux et al., 1996; Smidansky et al., 2002, 2003; T.W. Greene and L.C. Hannah, unpublished data). Since yield loss is associated with temperature stress (Singletary et al., 1993, 1994), a superior AGPase with increased heat stability would be beneficial to cereal crop production. Therefore, further testing of QTCL or QTCL-MP variants in transgenic plants will determine whether the enhanced catalytic activity and heat stability seen in E. coli translates to increases in starch synthesis. MATERIALS AND METHODS Site-Directed Mutagenesis Mutations in the maize (Zea mays) small subunit were created essentially as described by Horton et al. (1993). PCR-mutagenized DNA fragments of the maize endosperm small subunit were digested with NcoI and KpnI. These were used to replace the equivalent wild-type region of small subunit in an expression vector. Mutations were verified by sequencing. The vector was transformed into the Escherichia coli strain AC70R1-504, which also contained the wild-type large subunit coding region on a compatible expression vector (Giroux et al., 1996). The AC70RI-504 cell line contains a mutation that renders the strain incapable of producing bacterial AGPase (Iglesias et al., 1993). Growth and Purification of Maize AGPase from E. coli Protein inductions were as described by Burger et al. (2003), except that induction was performed for 3 h at room temperature. Following cell harvest by centrifugation, cell pellets were stored at −80°C. Enzyme Purification Purification of the wild type and the QTCL variant AGPase was done as described by Boehlein et al. (2005). For crude extracts, the bacterial pellets were resuspended in 1.0 mL of extraction buffer (50 mm HEPES, pH 7.5, 200 mm KCl, 10 mm MgCl2, 2.5 mm EDTA, and 5% Suc) containing 20% ammonium sulfate, 50 μg/mL lysozyme, 1 μg/mL pepstatin, 1 μg/mL leupeptin, 1 mm phenylmethylsulfonyl fluoride, 10 μg/mL chymostatin, and 1 mm benzamidine. The lysate was maintained on ice and sonicated three times for 10 s each. The sample was centrifuged for 5 min at 12,500 rpm at 4°C and the supernatant was conserved on ice. Solid ammonium sulfate was added to 45% saturation and the sample was centrifuged for 5 min at 12,500 rpm at 4°C. The pellet was resuspended in extraction buffer containing protease inhibitors and stored on ice. The concentration of the crude protein extract was determined using the Bio-Rad protein assay using BSA as a standard. Assay A (Forward Direction, Radioactive) For heat-stability measurements, the enzymes were diluted to 1.0 μg/μL and divided into two tubes. A single tube remained on ice while the second tube was placed at 58°C for 6 min with occasional gentle agitation. AGPase activity of the crude extracts was determined in the direction of ADP-Glc synthesis as described in Burger et al. (2003) with a reaction time of 5 min. Reactions were started by the addition of the enzyme. The value reported within an experiment is the average from triplicate samples. Assay B (Reverse Direction, Nonradioactive) A nonradioactive endpoint assay was used to measure G-1-P produced by coupling synthesis to NADH production using phosphoglucomutase and Glc-6-P dehydrogenase as described by Boehlein et al. (2005). The temperature of all the assays was 37°C unless otherwise specified. Standard reaction mixtures contained 100 mm MOPS HCl, pH 7.4, 0.4 mg/mL BSA, 5 mm MgCl2, 1 mm ADP-Glc, 20 mm 3-PGA, 1 mm sodium pyrophosphate, and enzyme in 100-μL reaction volume. Reactions were incubated for 5 min and terminated by boiling in a water bath for 1 min. After termination, 330 μL of water were added, followed by 70 μL of a development mixture containing a final concentration of 100 mm MOPS HCl, pH 7.4, 0.1 mg/mL BSA, 7 mm MgCl2, 0.6 mm NAD, 1 unit Glc-6-P dehydrogenase, and 1 unit phosphoglucomutase. Reactions were centrifuged for 5 min and then the absorbance read at 340 nm. Amount of G-1-P produced was calculated from a standard curve using freshly prepared G-1-P instead of enzyme. Assay tubes were prewarmed to 37°C prior to assaying. All assays were initiated by the addition of enzyme. Specific activity is defined as a unit/mg protein. Purification was always monitored using the reverse assay. Assay C (Forward Reaction, Nonradioactive) A nonradioactive endpoint assay was used to determine the amount of inorganic pyrophospate produced by coupling it to a decrease in NADH using a pyrophosphate reagent (Sigma P-7275) as previously described (Boehlein et al., 2005). Standard reaction mixtures contained 50 mm HEPES, pH 7.4, 15 mm MgCl2, 4.0 mm ATP, and 4.0 mm G-1-P in 200 μL. 3-PGA was added at varying amounts, as specified. Reactions were terminated after 10 min by boiling in a water bath for 1 min. The reactions were developed by adding 300 μL of pyrophosphate reagent (one bottle diluted with 22.5 mL water) to each assay and then the absorbance read at 340 nm. Change in absorbance between the blank and the reaction was used to calculate the amount to inorganic pyrophospate produced. All reactions were linear with time and enzyme concentration. Assay tubes were prewarmed to 37°C prior to assay and were initiated by enzyme addition. Enzyme Kinetics Determination of Km and Vmax Reactions were performed in the presence of 10 mm 3-PGA using assay C. The reactions were performed in triplicate and started by the addition of 0.043 μg of purified wild-type AGPase and 0.022 μg of QTCL or QTCL-MP enzyme. The amount of ATP or G-1-P varied from 0 to 3 mm. Determination of Ka and Ki To determine the activation constant for the recombinant maize wild-type AGPase (0.043 μg), QTCL (0.0217 μg) and QTCL-MP (0.0217 μg) of purified AGPase was incubated for 10 min using assay C. 3-PGA concentrations ranged from 0 to 5.0 mm. The value of the Ki for Pi was calculated in the presence of 2.5 mm 3-PGA and varied from 0 to 15 mm. Curves were fit using Graph Pad Prism nonlinear or linear regression. Activity was not detectable in the absence of 3-PGA at these protein concentrations. Heat Stability The half-life of wild type and QTCL at 42°C was determined by desalting enzyme in 50 mm HEPES, pH 6.5, 5.0 mm MgCl2, 0.5 mm EDTA. Heat was applied to desalted enzyme (0.065 mg/mL) and, at the appropriate time, enzyme was withdrawn from the tube and placed on ice. This enzyme was used for activity assays and BN gels. All reactions were carried out in triplicate using assay C with the addition of 10 mm 3-PGA. Both SDS-PAGE and BN-PAGE gels were prepared as outlined (Boehlein et al., 2005). Briefly, the gradient for the BN-PAGE was 5% to 18%. The buffers used are according to Schagger et al. (1994) with minor modifications. Two types of cathode buffer were prepared, one contained 0.002% Coomassie G and the other lacked Coomassie G. Aminocaproic acid was not used in the gel buffer. The gels were run at 4°C for 20 min at 100 V in cathode buffer containing Coomassie G. The voltage then was increased to 200 V for an additional 20 min. Finally, the gel was transferred to cathode buffer without Coomassie and run at 200 V until the dye front was off the gel. The gel was equilibrated in cold 1× transfer buffer (25 mm Tris base, 192 mm Gly, and 20% methanol) plus 1% SDS. A standard western transfer procedure was performed using polyvinylidene difluoride membrane with a 0.2 μm pore size (Bio-Rad). The molecular mass markers were as follows: thyroglobulin, 669,000 kD; ferritin, 440,000 kD; catalase, 232,000 kD; lactate dehydrogenase, 140,000 kD; and BSA, 67,000 kD (Amersham-Pharmacia Biotech electrophoresis calibration kit). Each lane contained approximately 0.87 μg of purified enzyme. All BN-PAGE gel western blots were developed with polyclonal antibodies against the maize AGPase small subunit. Heat stability experiments at 55°C were performed as described above, except each enzyme was desalted into 50 mm HEPES, pH 7.4, 5 mm MgCl2, 0.5 mm EDTA followed by the addition of BSA (0.5 mg/mL). Data were plotted as log% activity versus time, and the inactivation constant t1/2 was calculated as follows: slope = −k/(2.3). t1/2 is calculated from the equation k = 0.693/t1/2. Nonreducing 10% SDS-PAGE gels were prepared according to standard procedures (Sambrook et al., 1989). Approximately 0.43 μg of purified enzyme was loaded in each lane, and western blots were developed using polyclonal antibodies raised against the small and large subunits of the maize AGPase. Disulfide Bridge Reduction Enzymes were desalted into 50 mm HEPES, pH 7.4, 5 mm MgCl2, and 0.5 mm EDTA. Protein concentrations were determined and BSA was added to a final concentration of 0.5 mg/mL to stabilize the desalted enzyme. Enzymes were incubated for 15 min at room temperature in a mixture containing 100 mm HEPES, pH 7.4, 0.2 mg/mL BSA, 5 mm MgCl2, and the appropriate treatment in a 20-μL volume. Following this incubation, reactions were performed using assay B with the following modification. The final concentration in samples with DTT was 3.0 mm and the ADP-Glc concentration was adjusted so that the final concentration in the assay was always 1.4 mm. Each assay was performed for 2, 3, 4, 5, and 6 min and the slope of the plot of velocity versus time was used to calculate the specific activity. ACKNOWLEDGMENTS We thank Dr. Jon Stewart, Dr. Thomas Greene, and members of the Hannah Laboratory for many useful comments and discussions. LITERATURE CITED Bae J, Giroux M, Hannah LC ( 1990 ) Cloning and molecular characterization of the brittle-2 gene of maize. Maydica 35 : 317 –322 Ballicora MA, Frueauf JB, Fu Y, Schurmann P, Preiss J ( 2000 ) Activation of the potato tuber ADP-glucose pyrophosphorylase by thioredoxin. J Biol Chem 275 : 1315 –1320 Ballicora MA, Fu Y, Frueauf JB, Preiss J ( 1999 ) Heat stability of the potato tuber ADP-glucose pyrophosphorylase: role of Cys residue 12 in the small subunit. Biochem Biophys Res Commun 257 : 782 –786 Ballicora MA, Laughlin MJ, Fu Y, Okita TW, Barry GF, Preiss J ( 1995 ) Adenosine 5′-diphosphate-glucose pyrophosphorylase from potato tuber. Significance of the N terminus of the small subunit for catalytic properties and heat stability. Plant Physiol 109 : 245 –251 Bhave M, Lawrence S, Barton C, Hannah LC ( 1990 ) Identification and molecular characterization of shrunken-2 cDNA clones of maize. Plant Cell 2 : 581 –588 Boehlein SK, Sewell AK, Cross J, Stewart JD, Hannah LC ( 2005 ) Purification and characterization of adenosine diphosphate glucose pyrophosphorylase from maize/potato mosaics. Plant Physiol 138 : 1552 –1562 Burger BT, Cross JM, Shaw JR, Caren JR, Greene TW, Okita TW, Hannah LC ( 2003 ) Relative turnover numbers of maize endosperm and potato tuber ADP-glucose pyrophosphorylases in the absence and presence of 3-phosphoglyceric acid. Planta 217 : 449 –456 Cross JM, Clancy M, Shaw JR, Boehlein SK, Greene TW, Schmidt RR, Okita TW, Hannah LC ( 2005 ) A polymorphic motif in the small subunit of ADP-glucose pyrophosphorylase modulates interactions between the small and large subunits. Plant J 41 : 501 –511 Cross JM, Clancy M, Shaw JR, Greene TW, Schmidt RR, Okita TW, Hannah LC ( 2004 ) Both subunits of ADP-glucose pyrophosphorylase are regulatory. Plant Physiol 135 : 137 –144 Fu Y, Ballicora MA, Leykam JF, Preiss J ( 1998 ) Mechanism of reductive activation of potato tuber ADP-glucose pyrophosphorylase. J Biol Chem 273 : 25045 –25052 Geigenberger P, Kolbe A, Tiessen A ( 2005 ) Redox regulation of carbon storage and partitioning in response to light and sugars. J Exp Bot 56 : 1469 –1479 Giroux MJ, Hannah LC ( 1994 ) ADP-glucose pyrophosphorylase in shrunken-2 and brittle-2 mutants of maize. Mol Gen Genet 243 : 400 –408 Giroux MJ, Shaw J, Barry G, Cobb BG, Greene T, Okita T, Hannah LC ( 1996 ) A single mutation that increases maize seed weight. Proc Natl Acad Sci USA 93 : 5824 –5829 Greene TW, Hannah LC ( 1998 ) Enhanced stability of maize endosperm ADP-glucose pyrophosphorylase is gained through mutants that alter subunit interactions. Proc Natl Acad Sci USA 95 : 13342 –13347 Hannah L, Tuschall D, Mans R ( 1980 ) Multiple forms of maize endosperm ADP-glucose pyrophosphorylase and their control by Shrunken-2 and Brittle-2. Genetics 95 : 961 –970 Hannah LC ( 1997 ) Starch synthesis in the maize endosperm. In BA Larkins, IK Vasil, eds, Advances in Cellular and Molecular Biology of Plants, Vol 4. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp 375–405 Hannah LC, Shaw JR, Giroux MJ, Reyss A, Prioul JL, Bae JM, Lee JY ( 2001 ) Maize genes encoding the small subunit of ADP-glucose pyrophosphorylase. Plant Physiol 127 : 173 –183 Horton R, Steffan N, Pullen J, Hunt H, Cai Z, Pease L ( 1993 ) Gene splicing by overlap extension. In J Abelson, M Simon, eds, Methods in Enzymology. Academic Press, New York, pp 270–280 Hwang SK, Salamone PR, Okita TW ( 2005 ) Allosteric regulation of the higher plant ADP-glucose pyrophosphorylase is a product of synergy between the two subunits. FEBS Lett 579 : 983 –990 Iglesias AA, Barry GF, Meyer C, Bloksberg L, Nakata PA, Greene T, Laughlin MJ, Okita TW, Kishore GM, Preiss J ( 1993 ) Expression of the potato tuber ADP-glucose pyrophosphorylase in Escherichia coli. J Biol Chem 268 : 1081 –1086 Jin X, Ballicora MA, Preiss J, Geiger JH ( 2005 ) Crystal structure of potato tuber ADP-glucose pyrophosphorylase. EMBO J 24 : 1 –11 Okita T, Nakata P, Anderson J, Sowokinos J, Morell M, Preiss J ( 1990 ) The subunit structure of potato tuber ADPglucose pyrophosphorylase. Plant Physiol 93 : 785 –790 Preiss J, Romeo T ( 1994 ) Molecular biology and regulatory aspects of glycogen biosynthesis in bacteria. Prog Nucleic Acid Res Mol Biol 47 : 299 –329 Preiss J, Sivak M ( 1996 ) Starch synthesis in sinks and sources. In E Zamski, ed, Photoassimilate Distribution in Plants and Crops: Source-Sink Relationships. Marcel Dekker, New York, pp 139–168 Sambrook J, Fritsch E, Maniatis T ( 1989 ) Molecular Cloning: A Laboratory Manual, Ed 2. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY Schagger H, Cramer W, von Jagow G ( 1994 ) Analysis of molecular masses and oligomeric states of protein complexes by blue native electrophoresis and isolation of membrane protein complexes by two-dimensional native electrophoresis. Anal Biochem 217 : 220 –230 Singletary G, Banisadr R, Keeling P ( 1993 ) Decreased starch synthesis in heat stressed maize kernels results from reduced ADPG-pyrophosphorylase and starch synthase activities. Plant Physiol (Suppl) 102 : 6 Singletary G, Banisadr R, Keeling P ( 1994 ) Heat stress during grain filling in maize: effects of carbohydrate storage and metabolism. Aust J Plant Physiol 21 : 829 –841 Smidansky ED, Clancy M, Meyer FD, Lanning SP, Blake NK, Talbert LE, Giroux MJ ( 2002 ) Enhanced ADP-glucose pyrophosphorylase activity in wheat endosperm increases seed yield. Proc Natl Acad Sci USA 99 : 1724 –1729 Smidansky ED, Martin JM, Hannah LC, Fischer AM, Giroux MJ ( 2003 ) Seed yield and plant biomass increases in rice are conferred by deregulation of endosperm ADP-glucose pyrophosphorylase. Planta 216 : 656 –664 Sowokinos J, Preiss J ( 1982 ) Pyrophosphorylases in Solanum tuberosum. III. Purification, physical, and catalytic properties of ADPglucose pyrophosphorylase in potatoes. Plant Physiol 69 : 1459 –1466 Stark D, Timmerman K, Barry G, Preiss J, Kishore G ( 1992 ) Regulation of the amount of starch in plant tissues by ADP glucose pyrophosphorylase. Science 258 : 287 –291 Tiessen A, Hendriks JH, Stitt M, Branscheid A, Gibon Y, Farre EM, Geigenberger P ( 2002 ) Starch synthesis in potato tubers is regulated by post-translational redox modification of ADP-glucose pyrophosphorylase: a novel regulatory mechanism linking starch synthesis to the sucrose supply. Plant Cell 14 : 2191 –2213 Author notes 1 This work was supported by the National Science Foundation (IBN–9316887, IBN–960416, IBN–9982626, IBN–0444031, and MCB–9420422), the U.S. Department of Agriculture Competitive Grants Program (94–37300–453, 9500836, 95–37301–2080, 9701964, 97–36306–4461, 98–01006, and 2000–01488), and the Florida Agricultural Experiment Station (Journal Series no. R–10889). * Corresponding author; e-mail [email protected]; fax 352–392–6957. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: L. Curtis Hannah ([email protected]). Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.105.067637. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Keskitalo, Johanna; Bergquist, Gustaf; Gardeström, Per; Jansson, Stefan

doi: 10.1104/pp.105.066845pmid: 16299183

Abstract We have studied autumn leaf senescence in a free-growing aspen (Populus tremula) by following changes in pigment, metabolite and nutrient content, photosynthesis, and cell and organelle integrity. The senescence process started on September 11, 2003, apparently initiated solely by the photoperiod, and progressed steadily without any obvious influence of other environmental signals. For example, after this date, senescing leaves accumulated anthocyanins in response to conditions inducing photooxidative stress, but at the beginning of September the leaves did not. Degradation of leaf constituents took place over an 18-d period, and, although the cells in each leaf did not all senesce in parallel, senescence in the tree as a whole was synchronous. Lutein and β-carotene were degraded in parallel with chlorophyll, whereas neoxanthin and the xanthophyll cycle pigments were retained longer. Chloroplasts in each cell were rapidly converted to gerontoplasts and many, although not all, cells died. From September 19, when chlorophyll levels had dropped by 50%, mitochondrial respiration provided the energy for nutrient remobilization. Remobilization seemed to stop on September 29, probably due to the cessation of phloem transport, but, up to abscission of the last leaves (over 1 week later), some cells were metabolically active and had chlorophyll-containing gerontoplasts. About 80% of the nitrogen and phosphorus was remobilized, and on September 29 a sudden change occurred in the δ15n of the cellular content, indicating that volatile compounds may have been released. Autumnal senescence may attract more attention from the public, but less from scientists, than any other plant developmental process. Every autumn deciduous trees need to shed their leaves and prepare for the winter. Autumnal senescence is spectacular in many trees, when leaves change color from green to yellow and/or red (Lee et al., 2003). The different colors arise from the preferential degradation of chlorophylls over carotenoids and the synthesis of red-colored pigments like anthocyanins (Goodwin, 1958; Lichtenthaler, 1987). Autumn senescence, like other forms of leaf senescence, is a type of programmed cell death, i.e. the leaf cells die in an organized, predetermined way controlled by the nucleus. On the other hand, there are fundamental differences between the leaf senescence program and apoptotic processes, which have been studied in great detail in animal systems and more recently in plants (Noodén et al., 1997; Kuriyama and Fukuda, 2002). In fact, it has even been speculated that the processes are mutually antagonistic (Ougham et al., 2005) since an appropriately executed senescence program avoids the pathological consequences of cell death (Hörtensteiner, 2004). Senescence has adaptive value because of the associated remobilization of nutrients, especially nitrogen, and, to a lesser extent, phosphorus, sulfur, and other elements (Himelblau and Amasino, 2001; Hörtensteiner and Feller, 2002). The timing of autumn senescence can be regarded as the result of a trade-off between the conflicting requirements for optimizing the nitrogen and carbon status of the plant. Trees entering senescence early will efficiently remobilize nitrogen at the expense of photosynthetic yield, while trees entering senescence late will gain more photosynthates, but in some years their leaves will die before the remobilization of their nutrients is complete. Nitrogen status influences the onset of autumn senescence. For instance, alders (Alnus subsp.) that host a nitrogen-fixing symbiont (Frankia) shed their leaves while they are still green, and gardeners in temperate regions know that nitrogen-rich fertilization should be avoided since it delays both autumn senescence and the development of hardiness, thereby compromising winter survival (especially of woody perennials). Autumn senescence has not been well characterized at the cellular and molecular levels, but more information is available regarding leaf senescence in annuals (Buchanan-Wollaston, 1997; Quirino et al., 2000). The chloroplasts, present in green leaves, differentiate into gerontoplasts that lack stacked thylakoid membranes but are rich in electron-dense lipid bodies, plastoglobuli, which have an especially high content of carotenoids and carotenoid esters (Tevini and Steinmuller, 1985). Nutrient remobilization requires the breakdown of macromolecules and conversion of the breakdown products into transportable compounds. Many genes coding for catabolic enzymes (proteases, lipases, nucleases, etc.) are among those that are induced during senescence (Bhalerao et al., 2003; Buchanan-Wollaston et al., 2003; Andersson et al., 2004; Guo et al., 2004). When the photosynthetic capacity of the leaves is lost, the energy required for the remobilization must be provided by mitochondrial respiration and, consequently, nuclear genes coding for components of the mitochondrial electron transport chain are not down-regulated during autumn senescence, in contrast to genes coding for components of the photosynthetic apparatus (Buchanan-Wollaston, 1997; Andersson et al., 2004). Although the leaves do not perform photosynthesis, phloem transport out of the leaf is needed to export the nutrients so, from a transport perspective, senescing leaves are still source leaves. Eventually, however, phloem transport stops, a protective layer is formed on the inner side of the petiole, a separation or abscission layer is formed proximal to it, and when the cell walls in the separation layer are gradually loosened, the leaf eventually falls (Roberts et al., 2002). It is well known that autumn senescence in most trees is triggered by reductions in the photoperiod, a signal that is more reliable than temperature as a harbinger of the first strong frost, allowing autumn senescence calendars to be constructed that are quite constant over the years (e.g. www.great-lakes.net/tourism/fallcolor.html; www.arkansas.com/calendar/fall_foliage_pg1.asp). The pivotal role of phytochromes in the process has also been well documented (Olsen et al., 1997; Chen et al., 2002), but the extent to which low temperatures, perhaps coupled to light intensities inducing photooxidative stress, modulate the process is not clear. It is common knowledge that cold autumns result in intense leaf coloration, but the physiological basis of this color change is not well understood. We have started a project to elucidate the genetic basis of autumn senescence in aspen (Populus tremula), and as part of the project we have identified many genes that are expressed in autumn leaves (Bhalerao et al., 2003) and constructed a transcriptional timetable for autumn senescence (Andersson et al., 2004). However, studies of gene expression alone do not always provide sufficient information for understanding the cellular events that occur during the senescence process, especially since not only anabolism, but also gene expression, ceases during the later stages of autumn senescence (Andersson et al., 2004). Consequently, catabolic processes leading to the degradation of cellular components and, eventually, cell death should perhaps be studied at the cell biological level rather than the transcriptional level. Many fundamental questions concerning cellular processes in autumn leaves in aspen, and trees in general, have not yet been answered, or in some cases even addressed. For instance, is the conversion of chloroplasts to gerontoplasts manifested by the appearance of autumn colors driven only by developmental factors, or do environmental stress factors like photooxidation also contribute? When and to what extent are the nutrients of the leaf remobilized? When has photosynthesis decreased to the point that mitochondrial respiration has to provide the energy required for the degradation process and, after this point, do the plastids still play a role in the remobilization of nutrients? When, and how, are the gerontoplasts, nuclei, and mitochondria degraded, and when does the cell eventually undergo cell death? Finally, what are the signals that trigger the onset of autumn senescence? To address these questions, we have monitored the timing of autumn senescence events in a free-growing aspen, and here present a cellular timetable for them. RESULTS Autumn Senescence Progressed through Four Phases As reported for the same tree in the autumn of 1999 (Bhalerao et al., 2003), autumn senescence in 2003 largely progressed in a synchronized fashion. Autumn colors appeared earlier in some leaves than in others, and patches of each leaf turned yellow before others, but there were no apparent differences between the different parts of the tree in these respects (Fig. 1, A and B Figure 1. Open in new tabDownload slide A, Autumn senescence in a free-growing aspen. The images were taken at the time of leaf sampling (11:30 am) during the autumn of 2003. B, Senescing aspen leaf (September 25). C, Weather conditions during the sampling period. Gray bars represent the amount of sunlight (W/m2) per day and black bars represent the millimeters of precipitation per day. The lines correspond to the maximum and minimum temperatures for each day. sep, September; oct, October. Figure 1. Open in new tabDownload slide A, Autumn senescence in a free-growing aspen. The images were taken at the time of leaf sampling (11:30 am) during the autumn of 2003. B, Senescing aspen leaf (September 25). C, Weather conditions during the sampling period. Gray bars represent the amount of sunlight (W/m2) per day and black bars represent the millimeters of precipitation per day. The lines correspond to the maximum and minimum temperatures for each day. sep, September; oct, October. ). This is typical for aspen trees, but not for all other tree species. Over the years, we have visually examined thousands of aspens, but we have found no cases in which one part of the tree had progressed further into autumn senescence than others. In contrast, maples (Acer subsp.), for example, frequently have autumn coloration in the part of the crown that is most exposed to sunlight, while the leaves are still green in other parts. Yet other species may have green shoot tips, while the inner parts are yellow. Such differences, together with differences in leaf color, often allow tree species to be recognized from long distances (Chapman et al., 2000). This feature makes aspen a convenient model system for studies of autumn senescence since sampling and pooling randomly selected leaves are likely to give results that are representative for the whole tree. The weather conditions during the experimental period, recorded by a weather station about 200 m from the tree, are displayed in Figure 1B. After a few cold days with bright sun (August 31 to September 2), the conditions were very stable, with relatively mild, clear days up to September 19, when the temperature fell sharply. The chlorophyll content of the leaves (Fig. 2A Figure 2. Open in new tabDownload slide A, Chlorophyll concentrations in aspen autumn leaves. White and black circles represent total chlorophyll (a + b) for 2003 and the chlorophyll a/b ratio for 2003, respectively. Each point is the mean of three measurements (±sd). The gray background line represents total chlorophyll (a + b) for 1999 (as trend comparison, not actual values). B, Leaf weight (dry). Five leaves were pooled, ground, and weighed, and then dry leaf weight was calculated from the weight loss (%) after drying a small amount of the material. Figure 2. Open in new tabDownload slide A, Chlorophyll concentrations in aspen autumn leaves. White and black circles represent total chlorophyll (a + b) for 2003 and the chlorophyll a/b ratio for 2003, respectively. Each point is the mean of three measurements (±sd). The gray background line represents total chlorophyll (a + b) for 1999 (as trend comparison, not actual values). B, Leaf weight (dry). Five leaves were pooled, ground, and weighed, and then dry leaf weight was calculated from the weight loss (%) after drying a small amount of the material. ), expressed on a dry-weight basis, was more or less stable up to September 11. Concentrations of both chlorophyll a and b started to decrease on September 11 (Fig. 2A). The chlorophyll a/b ratio was constant until September 24, after which it decreased for about 3 d, stabilized again, and finally decreased in phase 4. The minor variations observed in chlorophyll concentration before September 11 could have been due to inhomogeneity in the sampled leaf material. In a free-growing tree, fungal infections, and to some extent insect herbivory, induce patches of senescence in the leaf. These are stochastic events; note that the standard deviation of the measurements of chlorophyll concentration are larger up to the point where chlorophyll degradation due to autumn senescence started, apparently overruling these leaf-to-leaf differences. However, it is possible that the dip in chlorophyll concentration observed around September 1 was a consequence of the cold, sunny weather at that time, inducing photooxidative stress and some photobleaching. If so, the recovery of chlorophyll levels in the first week of September indicates that the leaves were still not fully prepared to undergo autumn senescence. Whether or not this is true, the time up to September 11, when chlorophyll levels were stable, clearly represents a presenescent stage, which we designated phase 1. After September 11, chlorophyll levels started to decrease (phase 2) at a constant rate until the last days of September, when chlorophyll concentration (on a dry-weight basis) had decreased to about 10% of phase 1 levels. In phase 3, little or no further degradation of chlorophyll occurred, but a few days before abscission, starting on about October 6, there may have been a small further decrease in chlorophyll content (phase 4). Degradation of leaf components leads, of course, to the loss of leaf biomass. In order to monitor the changes in leaf biomass over the period, we calculated leaf dry weights of five randomly chosen leaves each day during the studied period. Although differences in the size of the five leaves resulted in fluctuations in the dry-weight data (Fig. 2B), there was an overall tendency for average leaf weight to start declining, from an initial value of around 150 mg, at about the same time as chlorophyll levels (about September 11), reaching a final value of about 75 mg. On average, leaf dry weight fell by 40% to 50%, showing that roughly 50% of the dry mass had been remobilized from the leaves. This resulted in chlorophyll levels in phase 3, per leaf, being about 5% of phase 1 values. It also seems that some chlorophyll degradation occurred in phase 3, since the average leaf dry weight decreased slowly, while the amount of chlorophyll per unit dry leaf weight was constant. In order to compare the progression of senescence in 2003 with the transcriptional timetable of autumn senescence that we generated for the same tree for the autumn of 1999, the chlorophyll concentrations in the leaves used for RNA preparation in 1999 were calculated and are presented in Figure 2A. The dates of the transitions between phases 1, 2, and 3 were almost identical in the two years. Our separation of the autumn senescence process into four phases provides a framework for the following examination of the cellular timetable for autumn senescence. The fact that chlorophyll degradation was initiated on the same date in both years indicates that the photoperiod was the dominant trigger for the transition into phase 2, since the weather conditions in the 10 d preceding September 11 were very stable and mild in both 2003 (Fig. 1B) and 1999 (data not shown). However, in both years, the days around September 1 were unusually cold but sunny, and these conditions could have triggered entry into phase 2, following a 10-d delay. To prove or disprove this hypothesis, we also examined chorophyll concentrations in the leaves sampled in 2001 and correlated them with the weather conditions (data not shown). In this year, the days around September 1 were not unusually cold, but chlorophyll degradation was nevertheless initiated around September 11, indicating that the tree initiated autumn senescence in response solely to changes in the photoperiod. Dynamic Changes in Carotenoid and Anthocyanin Metabolism To establish whether the yellow autumn color in these leaves depends on synthesis of new carotenoids or their slower degradation, we also measured their carotenoid content and composition throughout the period using HPLC. Figure 3A Figure 3. Open in new tabDownload slide Carotenoid concentrations in aspen autumn leaves. A, Total carotenoid content. Symbols refer to neoxanthin (circles), xanthophylls (triangles), lutein (diamonds), and β-carotene (squares). Each point is the mean value of two measurements (±sd). B, Epoxidation state of xanthophyll cycle pigments. C, Accumulation of carotenoid degradation products. Lines correspond to the different peaks in Table I. Figure 3. Open in new tabDownload slide Carotenoid concentrations in aspen autumn leaves. A, Total carotenoid content. Symbols refer to neoxanthin (circles), xanthophylls (triangles), lutein (diamonds), and β-carotene (squares). Each point is the mean value of two measurements (±sd). B, Epoxidation state of xanthophyll cycle pigments. C, Accumulation of carotenoid degradation products. Lines correspond to the different peaks in Table I. shows the levels of neoxanthin, lutein, β-carotene, and the xanthophyll cycle pigments. The total carotenoid concentration started to decrease at the same time that chlorophyll degradation started, during the transition from phase 1 to phase 2. However, the carotenoid degradation rate was slower, resulting in the color shift of the leaves from green to yellow. At the time when the leaves appeared yellow, the chlorophyll levels had decreased by 75%, but carotenoid levels only by about 50%. In phase 3, carotenoid levels were quite stable, but in phase 4 they again started to decrease. There were significant differences in degradation rates between the different carotenoids. β-Carotene was degraded largely in parallel to chlorophyll; the degradation rate for lutein also followed the four-phase pattern, although the levels did not decrease as much as for chlorophyll. Neoxanthin levels roughly followed chlorophyll levels throughout phase 2, but also continued to decrease in phases 3 and 4. The total concentration of the xanthophyll cycle pigments (violaxanthin, antheraxanthin, and zeaxanthin) were more stable and decreased only by about 50% on a dry-weight basis (75% on a per leaf basis) during the studied period. The xanthophyll cycle pigments can be interconverted in the xanthophyll cycle, and in green leaves the dynamics of the xanthophyll pools are determined by environmental factors. Violaxanthin is converted to zeaxanthin via antheraxanthin in a reaction catalyzed by the enzyme violaxanthin de-epoxidase when the lumenal pH decreases as a consequence of the light reactions, producing a higher proton gradient than can be utilized in CO2 fixation (Demmig-Adams, 1990). Such weather-dependent changes in the xanthophyll pools were clearly observed until the end of phase 2 (Fig. 3B). However, even throughout phase 3, such changes occurred, although the amplitude was greatly reduced, and the changes observed were perhaps not significant. From the middle of phase 2, new peaks started to appear in the chromatograms (Suzuki and Shioi, 2004). We could distinguish nine pigments in addition to the normal photosynthetic pigments, which accumulated during autumn senescence (Fig. 3C). These carotenoids/carotenoid derivatives were tentatively identified by recording their absorbance spectra and by exposing the extracts to high pH (saponification), a treatment that breaks the ester bond of carotenoid esters and releases the carotenoid moieties (Granado et al., 2001). To our knowledge, these pigments have not been thoroughly characterized before, and we did not attempt to rigorously do so here; thus, the assignments are only tentative (Table I Table I. Photosynthetic pigments and pigment catabolites in aspen autumn leaves, separated by HPLC Several of the peak assignments are tentative (see “Materials and Methods” for details). Peak . Retention Time . Compounds . min 1 1.97 trans-Neoxanthin 2 2.14 9-cis-Neoxanthin 3 2.59 Violaxanthin 4 3.71 Antheraxanthin 5 5.15 Lutein 6 5.44 Zeaxanthin 7 6.19 Carotenoid X (CX) 8 6.51 Carotenoid Y (CY) 9 7.71 Chlorophyll b 10 8.00 Chlorophyll a 11 8.18 Neoxanthin ester 1 (NE1) 12 8.30 Neoxanthin ester 2 (NE2) 13 8.50 Violaxanthin ester 1 (VE1) 14 8.86 Neoxanthin ester 3 (NE3) 15 9.40 Neoxanthin ester 4 (NE4) 16 9.64 Violaxanthin ester 2 (VE2) 17 10.00 Violaxanthin X (VX) 18 10.73 β-Carotene Peak . Retention Time . Compounds . min 1 1.97 trans-Neoxanthin 2 2.14 9-cis-Neoxanthin 3 2.59 Violaxanthin 4 3.71 Antheraxanthin 5 5.15 Lutein 6 5.44 Zeaxanthin 7 6.19 Carotenoid X (CX) 8 6.51 Carotenoid Y (CY) 9 7.71 Chlorophyll b 10 8.00 Chlorophyll a 11 8.18 Neoxanthin ester 1 (NE1) 12 8.30 Neoxanthin ester 2 (NE2) 13 8.50 Violaxanthin ester 1 (VE1) 14 8.86 Neoxanthin ester 3 (NE3) 15 9.40 Neoxanthin ester 4 (NE4) 16 9.64 Violaxanthin ester 2 (VE2) 17 10.00 Violaxanthin X (VX) 18 10.73 β-Carotene Open in new tab Table I. Photosynthetic pigments and pigment catabolites in aspen autumn leaves, separated by HPLC Several of the peak assignments are tentative (see “Materials and Methods” for details). Peak . Retention Time . Compounds . min 1 1.97 trans-Neoxanthin 2 2.14 9-cis-Neoxanthin 3 2.59 Violaxanthin 4 3.71 Antheraxanthin 5 5.15 Lutein 6 5.44 Zeaxanthin 7 6.19 Carotenoid X (CX) 8 6.51 Carotenoid Y (CY) 9 7.71 Chlorophyll b 10 8.00 Chlorophyll a 11 8.18 Neoxanthin ester 1 (NE1) 12 8.30 Neoxanthin ester 2 (NE2) 13 8.50 Violaxanthin ester 1 (VE1) 14 8.86 Neoxanthin ester 3 (NE3) 15 9.40 Neoxanthin ester 4 (NE4) 16 9.64 Violaxanthin ester 2 (VE2) 17 10.00 Violaxanthin X (VX) 18 10.73 β-Carotene Peak . Retention Time . Compounds . min 1 1.97 trans-Neoxanthin 2 2.14 9-cis-Neoxanthin 3 2.59 Violaxanthin 4 3.71 Antheraxanthin 5 5.15 Lutein 6 5.44 Zeaxanthin 7 6.19 Carotenoid X (CX) 8 6.51 Carotenoid Y (CY) 9 7.71 Chlorophyll b 10 8.00 Chlorophyll a 11 8.18 Neoxanthin ester 1 (NE1) 12 8.30 Neoxanthin ester 2 (NE2) 13 8.50 Violaxanthin ester 1 (VE1) 14 8.86 Neoxanthin ester 3 (NE3) 15 9.40 Neoxanthin ester 4 (NE4) 16 9.64 Violaxanthin ester 2 (VE2) 17 10.00 Violaxanthin X (VX) 18 10.73 β-Carotene Open in new tab ; for details of the tentative identification, see “Materials and Methods”). Although some of these pigments (CX, NE2, NE3, and VE1) were already present in small amounts in phase 1, accumulation started in phase 2 (on September 12, 15, and 21 for VX, NE4, and VE2, respectively). The pattern for CY and NE1 was not clear, but accumulation seemed to start around September 20. Formation of the first, as-yet uncharacterized carotenoid catabolites appeared to coincide with the onset of chlorophyll breakdown, but several of them did not appear until later stages of senescence when chlorophyll levels had decreased by 50% or more. During phase 4, these carotenoid catabolites decreased in parallel with the decrease in other carotenoids. Although these pigments accumulated strongly during autumn senescence, none of them seemed to become the dominant pigments in the autumn leaves. We do not know their exact extinction coefficients so they could not be accurately quantified, but since saponification appeared to convert them quantitatively into neoxanthin, violaxanthin, and antheraxanthin, the increases in the concentrations of these pigments after saponification is likely to represent the amounts of the corresponding carotenoid esters. On October 3, neoxanthin, violaxanthin, and antheraxanthin collectively accounted for about 25% of the total carotenoid pool (including lutein, zeaxanthin, and β-carotene), and since the amount of these pigments increased by about 50% upon saponification, we estimate that the total amount of esterified carotenoids corresponded to 10% to 15% of the total carotenoid pool. Carotenoids are not the only pigments responsible for autumn colors; anthocyanins often accumulate (Lee et al., 2003), giving rise to dark-reddish leaf colors, and have been proposed to function as light protectants during the nutrient recycling of senescing leaves (Feild et al., 2001; Hoch et al., 2003). Therefore, we quantified anthocyanin levels to see whether they changed during autumn senescence in aspen leaves. Spectroscopic measurements of the total anthocyanin concentration in extracts of the leaves showed that they did indeed increase during autumn senescence (Fig. 4 Figure 4. Open in new tabDownload slide Anthocyanin concentrations in aspen autumn leaves. Each point is the mean value of two measurements (±sd). Figure 4. Open in new tabDownload slide Anthocyanin concentrations in aspen autumn leaves. Each point is the mean value of two measurements (±sd). ). The levels were low and constant up to about 5 d into phase 2 (September 15), at which point chlorophyll levels had decreased by about 25%. The onset of anthocyanin accumulation was not apparently correlated to photooxidative stress, since the 12 d preceding September 15 were all sunny but very mild. However, once anthocyanin accumulation had been initiated, further accumulation seemed to be strongly dependent on excess light conditions, i.e. cold and sunny weather. The clear, cold days on September 19 and 20 induced a huge accumulation of anthocyanins, more than one-half of which disappeared in the milder and rainy week that followed, while frost and sun on September 30 resulted in a new accumulation, followed by degradation during the mild and rainy days at the beginning of October. Finally, from October 4 onward, the weather was clear and cold, resulting in a very high accumulation of anthocyanins. Leaves Lose Photosynthesis Activity and Energy Requirements Are Then Met by Mitochondria When the chlorophyll starts to be degraded and the chlorophyll-binding proteins are remobilized, the photosystems are affected and photosynthesis declines. Although chlorophyll levels reflect the overall amounts of chlorophyll-binding proteins, they provide no direct indication of whether or not the photosynthetic apparatus is functional, so we wanted to directly estimate the photosynthetic activity in autumn leaves. Performing accurate gas exchange measurements on attached branches directly in the field was not feasible, and detached branches lost their photosynthetic capacity quite rapidly, presumably due to stomatal closure. The most convenient way is to measure chlorophyll fluorescence, which provides a sensitive tool for studying photosynthetically active PSII centers in vivo. In healthy, unstressed leaves, the Fv/Fm ratio is 0.80 to 0.85, whereas in stressed leaves, the ratio can decrease to 0.70 or less (Öquist and Wass, 1988). However, when interpreting the Fv/Fm ratio data, it must be remembered that they are relative measures that reflect the status of the PSII centers present and do not provide accurate quantifications of the number of centers. We followed four leaves during the autumn and measured Fv/Fm twice a week (Fig. 5 Figure 5. Open in new tabDownload slide Photochemical efficiency of PSII (Fv/Fm) in aspen autumn leaves. Four leaves were followed during the autumn. Figure 5. Open in new tabDownload slide Photochemical efficiency of PSII (Fv/Fm) in aspen autumn leaves. Four leaves were followed during the autumn. ). One leaf appeared to be significantly stressed at the beginning of the measurement series and showed Fv/Fm values below 0.6. However, this leaf recovered and its values became similar to those of the other leaves, showing that at least up to the first week of September, the PSII repair capacity was sufficient to restore damaged PSII centers. Somewhat surprisingly, Fv/Fm ratios stayed high and more or less constant until September 26, when chlorophyll concentrations had decreased by about 80%, i.e. throughout almost all of phase 2. Apparently, therefore, the remaining PSII centers were photosynthetically active even very late in the senescence program. Since a wide variety of stresses that affect either the chloroplast or the rest of the cell can influence Fv/Fm, this finding shows that degradation of the photosynthetic apparatus was tightly regulated. At the very end of phase 2 and later in phase 3, Fv/Fm declined in all leaves, but a measurable Fv/Fm ratio was present in one of the leaves even on October 3, indicative of active PSII centers. When the photosynthetic capacity decreases, mitochondria must presumably take over as the main energy sources when the chloroplasts and other cell components are degraded. To examine this assumption, we measured ATP and ADP levels in leaves that were instantly frozen in liquid nitrogen in the light (Fig. 6A Figure 6. Open in new tabDownload slide Changes in energy status and sugar depletion in aspen autumn leaves. Each point is the mean value of three measurements. A, Adenylate content (ATP + ADP) and ATP/ADP ratio. B, Concentrations of soluble sugars and starch. Figure 6. Open in new tabDownload slide Changes in energy status and sugar depletion in aspen autumn leaves. Each point is the mean value of three measurements. A, Adenylate content (ATP + ADP) and ATP/ADP ratio. B, Concentrations of soluble sugars and starch. ). The adenylate content (ATP + ADP) was constant during phase 1 and the ATP/ADP ratio was initially relatively low (around 2), but gradually increased to almost 4. During phase 2, the content of adenylates decreased in a very similar way to chlorophyll and, during the first part of phase 2, the ATP/ADP ratio rapidly increased from 4 to about 8. The decrease in adenylate content may reflect degradation of chloroplasts, which contain most of the cellular adenylate pool (Gardeström and Wigge, 1988). Also, the chloroplast compartment typically has a lower ATP/ADP ratio than the rest of the cell (Gardeström and Wigge, 1988; Bykova et al., 2005). Thus, the observed increases in ATP/ADP ratios are consistent with degradation of the chloroplast pool. Taken together, this evidence shows that chloroplasts are the major energy sources for the leaves in phase 1. In phase 2, the degradation of the photosynthetic apparatus creates a gradual shift toward an increasing dependence on mitochondrial respiration, and from the middle of phase 2 (September 19) and throughout phase 3, mitochondria seem to provide most of the autumnal leaves' energy. In phase 4, the drop in ATP/ADP ratios and the total adenylate pool also indicates that remaining mitochondria are being degraded. To get further insight into this transition from an organ dependent on photosynthesis to an organ dependent on respiration, we also wanted to determine the time at which the cells stopped accumulating sugars and starch. Sugars (Suc, Glc, and Fru) and starch were measured in a combined assay from the pool of leaves that was also used for adenylate analysis. During phase 1, Suc and starch were the major components, and very low amounts of Glc and Fru were detected. In phase 2, Glc and Fru levels initially increased, while Suc and starch were still abundant. Consequently, the highest content of total carbohydrates was observed on September 15, when chlorophyll had already started to decrease. After that, Suc and starch rapidly decreased so that, after September 22, Glc and Fru were the dominating sugars, staying at high levels throughout phase 2, before decreasing during phases 3 and 4. Nitrogen and Phosphorus Are Remobilized in Aspen Autumn Leaves To obtain data on net fluxes of nutrients from the autumn leaves, we subjected leaves to elementary analysis, measuring δ13C and δ15n and levels of carbon, nitrogen, potassium, sulfur, phosphorus, and iron in the leaves during the senescence process. The 13C/12C ratio did not change significantly (data not shown) but, as shown in Figure 7A Figure 7. Open in new tabDownload slide Elemental composition of aspen autumn leaves. A, Changes in relative contents (%) of carbon and nitrogen and in δ15n values (‰). B, Changes in relative content (%) of potassium, sulfur, phosphorus, and iron. Each point is the mean value of three measurements (±sd × 2). Figure 7. Open in new tabDownload slide Elemental composition of aspen autumn leaves. A, Changes in relative contents (%) of carbon and nitrogen and in δ15n values (‰). B, Changes in relative content (%) of potassium, sulfur, phosphorus, and iron. Each point is the mean value of three measurements (±sd × 2). , the relative content of carbon and nitrogen and the δ15n value changed substantially during the autumn. On a dry-weight basis, the carbon content was constant, which is not surprising since carbohydrates account for the vast majority of leaf biomass. Since the average weight of the leaf decreased by about 50% over the time period, it seems that about 50% of the leaf carbon was lost. Nitrogen content, on the other hand, perhaps decreased slightly even in phase 1, and very markedly during phase 2. In phases 3 and 4, nitrogen appeared to be remobilized slowly or not at all since the nitrogen content on a dry-weight basis only changed marginally. In total, about 80% of total leaf nitrogen (60% on a dry-weight basis) was withdrawn during autumn senescence. Phosphorus content also decreased by about 80% on a per leaf basis (60% on a dry-weight basis) during the autumn, although the phosphorus content started to decrease in phase 1 even before September 11 (Fig. 7B) and leveled out in phases 3 and 4. The remobilization of 80% of phosphorus could be compared with the breakdown of adenylates, which was 90% on leaf basis. Not surprisingly, phosphates bound to other complexes are less readily remobilized than those of the adenylate pool. Sulfur remobilization was less efficient, but still significant: the sulfur content decreased in parallel with the decrease in leaf dry weight; and the level on a dry-weight basis decreased by about 10%, showing that about 50% of the total leaf sulfur was remobilized. Potassium was also remobilized, mainly during a short time interval immediately before entry into phase 2. In phases 2 and 3, potassium levels did not change significantly on a dry-weight basis. On a dry-weight basis, the iron levels increased by about 60%. This corresponds to only about 20% of the leaf iron being remobilized. Overall, remobilization of leaf nutrients, in particular nitrogen, was efficient. Unexpectedly, we noticed a rapid change in the δ15N values of cellular content, which decreased slowly throughout phase I, showing that macromolecules with high δ15N values had a slightly higher catabolic rate than the average nitrogenous molecules. This was probably because different fractions of leaf proteins had slightly different δ15N values, due to differences in their biosynthetic history, and the degradation of the different fractions was not equally efficient. However, there was a very dramatic change in δ15N between September 28 and September 30 (transition to phase 3), and they decreased slowly but stayed high throughout phases 3 and 4. Chloroplasts Are Degraded and Dissolved, But a Few Plastids Remain Intact and Retain Their Chlorophyll In order to follow the autumn senescence process in the cells ultrastructurally, we studied the leaf cells and organelles using transmission electron microscopy (TEM; Fig. 8A Figure 8. Open in new tabDownload slide Changes in the ultrastructure of mesophyll cells during the autumn. Structural changes in cells (A, C, E, G, I) and chloroplasts (B, D, F, H, J) at five time points of autumn senescence (see Fig. 10). Size bars correspond to 10 μm (G and I), 5 μm (A and E), 2 μm (C), 1 μm (D, F, H, and J), and 0.5 μm (B). sep, September; oct, October. Figure 8. Open in new tabDownload slide Changes in the ultrastructure of mesophyll cells during the autumn. Structural changes in cells (A, C, E, G, I) and chloroplasts (B, D, F, H, J) at five time points of autumn senescence (see Fig. 10). Size bars correspond to 10 μm (G and I), 5 μm (A and E), 2 μm (C), 1 μm (D, F, H, and J), and 0.5 μm (B). sep, September; oct, October. ). Representative electron micrographs from five dates were chosen to illustrate the cellular changes during senescence, depicting the ultrastructure of both whole cells (left) and chloroplasts (right). At the end of phase 1 (September 8), the mesophyll cells had a normal shape, were lined with chloroplasts and mitochondria, and both the cytoplasm and vacuole were intact (Fig. 8A). The chloroplasts had a defined shape and structure (Fig. 8B), with plenty of starch granules and visible grana stacks. However, even at this stage the chloroplasts contained many plastoglobuli. In the middle of phase 2 (September 18), the chlorophyll concentration had decreased by 50%. Many cells (Fig. 8C) and chloroplasts (Fig. 8D) appeared almost unchanged, but the amount of electron-dense material in the cells had decreased, on average. The tonoplast was less defined and unknown structures (small vesicles) could be seen. In some cells, the chloroplasts had lost their internal membrane structure, whereas in other cells they appeared active with starch grains. There were clear differences between different sections of the leaf, which was not surprising since patchiness was also apparent at the macroscopic level at this stage, many leaves having a yellow-green mosaic pattern (Fig. 1A). At the end of phase 2 (September 25, when 25% of the initial chlorophyll remained), many cells had lost most of their electron-dense material (Fig. 8E). Chloroplast deterioration started to be frequent (Fig. 8F), manifested as a loss of starch granules, increases in the number and size of plastoglobuli, and loss of normal thylakoid membrane structure (replaced by swellings of the lamellae). Many small vesicle-like structures were present (Fig. 8F), apparently not located in the cytoplasm but in the vacuolar space, which gradually became less structured but more electron dense than in phase 1. Cells at this stage seemed to contain a lower number of chloroplasts and mitochondria and the cytoplasm was confined to small areas along the walls. Some cells appeared almost empty, sometimes situated adjacent to cells containing chloroplasts. No cells seemed to contain plastids in different developmental stages, i.e. both chloroplasts and gerontoplasts, but every cell seemed to be fully synchronized in terms of plastid development. In late phase 3, on October 3 (when less than 5% of the initial chlorophyll remained), the cells in the electron micrographs looked even less structured, and any remaining cytoplasm and plastids were aggregating in the corners of some cells, whereas others appeared empty (Fig. 8G). The plastids found were swollen gerontoplasts, consisting mainly of large plastoglobuli, but some mitochondria and a few nuclei still looked intact (Fig. 8H). Some plastoglobuli were also starting to extrude in blobs from the gerontoplasts, perhaps corresponding to the mass exodus that has been reported from soybean gerontoplasts (Guiamet et al., 1999), and the cytoplasmic regions were heterogeneous and contained vesicle-like structures. In phase 4, finally, most cells appeared empty and dead (Fig. 8I), although some still contained gerontoplasts filled with plastoglobuli, and some mitochondria-like structures were found (Fig. 8J). We were intrigued by the findings that the chlorophyll content of phase 3 leaves was about 5% of that of phase 1 leaves (although they appeared yellow), and that some leaves yielded a detectable Fv/Fm ratio. To get a better overview of the chlorophyll compartmentalization in late phase 3 leaves, we scanned leaves sampled on October 6 using confocal microscopy and recorded chlorophyll autofluorescence. Figure 9 Figure 9. Open in new tabDownload slide Chlorophyll autofluorescence (red) measured using confocal microscopy in a phase 4 leaf (October 6). A, Mesophyll tissue. B, Leaf segment. Size bars correspond to 50 μm (top) and 100 μm (bottom). ep, Epidermis; pal, palisade; mes, mesophyll. Figure 9. Open in new tabDownload slide Chlorophyll autofluorescence (red) measured using confocal microscopy in a phase 4 leaf (October 6). A, Mesophyll tissue. B, Leaf segment. Size bars correspond to 50 μm (top) and 100 μm (bottom). ep, Epidermis; pal, palisade; mes, mesophyll. shows a sectioned leaf at two different magnifications. The remaining chlorophyll was found in plastids, or at least organelle-like structures, throughout the entire mesophyll (both palisade cells and spongy mesophyll), but more frequently close to the veins. Some cells were clearly devoid of plastids, but some contained one or a few chlorophyll-containing plastids, sometimes located in opposite corners of the cell. This and the TEM micrographs show that, even at this very late stage of senescence, some cells contained chlorophyll-containing plastids, perhaps performing a metabolic function. DISCUSSION The tree we are studying has several million leaves. Each leaf contains about 30 million cells and each cell on average around 40 chloroplasts. The magnificent appearance of autumn senescence is the result of the synchronized and apparently tightly controlled conversion of these ≈1015 chloroplasts to gerontoplasts and the subsequent degradation of the majority of the cell organelles, including the gerontoplasts. We have followed this process by measuring pigments, key metabolites like ATP and Suc, macronutrients, photosynthesis, and cell and organelle integrity throughout the whole process. The data generated can be compiled to create a cellular timetable of autumn senescence (Fig. 10 Figure 10. Open in new tabDownload slide A cellular timetable of Populus tremula autumn senescence, compiled from the data presented in this paper. “Loss of cytoplasm” denotes the combination of two events, cytoplasm degradation and vacuolar burst. MES, Main energy source. Figure 10. Open in new tabDownload slide A cellular timetable of Populus tremula autumn senescence, compiled from the data presented in this paper. “Loss of cytoplasm” denotes the combination of two events, cytoplasm degradation and vacuolar burst. MES, Main energy source. ) in which the senescence process is divided into four temporal phases to emphasize key events, the first detailed description of the cellular events that occur during autumn senescence. This is, of course, not an absolute and generic timetable, since different tree species follow different patterns and even within a single species there is significant variation in, for example, the onset of the process. Factors like nutrient status also affect the initiation of the process. Nevertheless, we believe that studies of a single tree in a single year have general significance and are informative about the order of events involved. Reliable data on the sequence of events are required for a mechanistic understanding of the process and to allow detailed studies to address variation between species and individuals. By constructing the cellular timetable of autumn senescence for an aspen tree, we believe that we have obtained novel insights into some of the major unresolved questions concerning autumn senescence. We have shown that the entry into phase 2 was triggered by the photoperiod and that temperature (and light) had little or no effect on the onset of senescence. The autumn senescence process, once initiated by environmental factors, seems to be a tightly controlled developmental program that is not significantly altered by environmental factors. Chlorophyll was degraded at a fairly constant rate, although weather conditions fluctuated (and differed between years). If photooxidation of pigments had been a major determinant, the degradation rate would have been increased by cold clear days (like September 19 and 20), for example, which was not the case (Kukavica and Jovanovic, 2004). Chlorophyll was degraded at a constant rate, and carotenoid catabolites started to appear after September 12, before gerontoplast formation was observed. Photooxidative stress, on the other hand, seemed to be the major determinant of anthocyanin accumulation, and anthocyanin levels changed dynamically depending on the weather. The stimulation of autumn colors by cold weather seemed therefore to be solely due to effects on anthocyanin accumulation. Anthocyanin accumulation seemed to correlate better with excess light (high light and cold) than with either high light or cold alone, consistent with the hypothesis that anthocyanins have a photoprotective role. The large fluctuations in anthocyanin content showed that anthocyanins were rapidly metabolized right up to leaf abscission. Anthocyanin catabolism may be light regulated or, alternatively, regulated anabolism in aspen leaves could compete with a seemingly unregulated catabolism, similar to the regulation of the xanthophyll cycle. Interestingly, leaves in phase 1 did not seem to be as competent to respond to the stimuli that later led to anthocyanin accumulation. September 1 and 2 were as cold as September 19, and the light was brighter, but no anthocyanin accumulation was induced at this time. These observations, together with the fact that damaged PSII centers were also repaired during the first week of September, demonstrate that the leaves at this point were not yet ready to enter the senescence process. This is consistent with our previous suggestion that a peak in transcriptional activity in the first week of September coincides with a developmental switch reprogramming the leaf to senescence (Andersson et al., 2004). On September 11, the degradation process started (phase 2). This date for entry into phase 2 was the same over several years, with different weather conditions, and is therefore likely to be controlled by the photoperiod. A senescence signal triggered the cells to degrade their photosystems (and chlorophyll), thereby releasing carotenoids, some of which (lutein and β-carotene) were degraded, whereas others (particularly neoxanthin and violaxanthin) were esterified and accumulated in the plastoglobuli. This process seemed, within each plastid, to be very rapid, since the Fv/Fm ratios stayed high and constant until very late in the process. If large proportions of PSII centers present in the chloroplasts had been undergoing degradation processes, this would probably have been manifested by reductions in the overall Fv/Fm values. Apparently, therefore, although total photosynthetic capacity decreases along with the chlorophyll degradation, the photosynthetic parameters of the remaining PSII centers stay constant until very late in the process. We interpret these data as indicating that the transition from a photosynthetically fully active chloroplast to a gerontoplast is very rapid, and these data are consistent with a model in which all of the chloroplasts in a given cell undergo the transition simultaneously, while a neighboring cell could be in a different developmental stage. Changes in the chlorophyll a/b ratio showed that the degradation of antenna chlorophylls at the end of phase 2 is slower than that of chlorophyll a in the reaction centers. This resembles somewhat the situation in some stay-green mutants, where the remaining chlorophyll in otherwise senescing leaves is bound to light-harvesting complex II (Hilditch et al., 1986). The adaptive benefit of keeping the antenna and dismantling the reaction centers is not clear, but has been demonstrated during leaf senescence in other species (Wolf, 1956; Lichtenthaler, 1987). In parallel with the chloroplast degeneration, expression of the photosynthetic genes was strongly repressed (Andersson et al., 2004). After entry into phase 2, the cytoplasm started to be degraded, eventually making it difficult to distinguish cytoplasm from the vacuole. We could detect small vesicles, perhaps similar to the vesicles reported by Otegui et al. (2005), nuclei gradually disappeared, and the cells seem to largely stop synthesizing protein at this point; at least we have not been able to extract high-quality RNA from the leaves after September 24 (Andersson et al., 2004). During the leaf senescence process, most of the nitrogen and phosphorus was retrieved. The relative content of carbon was not changed (Fig. 7A), but since the leaf dry weight decreased (Fig. 2B) some carbon may also be retained. This massive remobilization requires, of course, the involvement of many catabolic enzymes, and genes encoding various types of proteases, lipases, and enzymes involved in nucleotide metabolism, glyconeogenesis, and nitrogen remobilization (inter alia) are either induced, or continue to be expressed, in the autumn leaves (Bhalerao et al., 2003; Andersson et al., 2004; Terce-Laforgue et al., 2004). Senescence is an energy-requiring process in which ATP is needed for degradation, recycling, and transporting nutrients out of the cell. As photosynthesis declines in phase 2, mitochondrial respiration needs to take over to provide the cell with the energy required for these processes. Thus, a significant fraction of the carbon lost from the leaf is likely to be due to respiration and not to be retrieved. We did not measure respiration during the process, but it is clear that mitochondria stay active throughout senescence since intact mitochondria could be seen in electron microscopy images late into the process. Genes encoding components of mitochondrial electron transport also continue to be expressed throughout the period (Andersson et al., 2004). By monitoring changes in the levels of key metabolites (ATP/ADP ratios and pool size, starch, and various soluble sugars), we found evidence indicating that a shift in the main energy sources of the cell from chloroplasts to mitochondria probably occurred around September 19. On September 30, the degradation process was largely complete and the leaves had entered phase 3. It is likely that this transition coincides with the formation of the protection and separation layers in the petiole, which blocks further phloem transport out of the leaf. The changes after this date were minor, and since leaves started to be shed and individual leaves in which the process was somewhat retarded were more likely to be withheld, this may explain the weakness of some trends in the data and suggests that our separation between phase 3 and phase 4 may be artificial. Some cells seemed at this stage to be empty (and probably dead), whereas many contained nuclei, mitochondria, and a small number of gerontoplasts. Vacuoles and cytoplasm could not be distinguished from each other. The dynamic changes in variables, such as the levels of xanthophyll cycle pigments and anthocyanins, and the presence of a few active photosystems during phase 3 suggest that a significant fraction of the cells were still alive and metabolically active. Many of these cells contained chlorophyll-containing plastids, and the residual chlorophyll level was approximately 5% of the phase 1 levels, but only a small fraction of this remaining chlorophyll seemed to be photosynthetically active, and this activity was lost during phase 3. The residual chlorophyll is likely to be localized in the plastoglobuli, where the high amounts of carotenoids could serve as efficient quenchers of excess excitation energy. The sudden shift in the nitrogen isotope ratio at the very end of phase 2 (September 29) was an unexpected finding. Changes in the nitrogen isotope ratios of senescing pine needles have previously been reported, and three hypotheses have been put forward to explain them (Näsholm, 1994). Since two of the hypotheses (discrimination against δ15N in the process of nitrogen translocation and the possibility that nitrogen pools that are not degraded, like cell wall proteins, have relatively high δ15N values) are incompatible with such a dramatic shift in the ratio; the third hypothesis, the emission of nitrogen-containing volatile compounds (low in δ15N), may be the best explanation for our findings. Emission of volatiles from senescing leaves has been reported (Husted et al., 1996), and our finding that the sudden change in isotope ratio coincided with the transition from phase 2 to phase 3, and probably with the termination of phloem transport out of the leaves, is compatible with a hypothesis that nitrogen-containing compounds may accumulate in the leaf to toxic levels, since protein catabolism probably continues so ammonia may be released to reduce the amount of excess nitrogen. It is possible that this sudden change in δ15N is a signature of the end of the remobilization process in autumn leaves. There are two possible explanations for the retention of some cells and organelles throughout phase 3. We believe that, by the end of phase 2, the tree has gone through the essential steps in the senescence process, enabling nutrient retrieval to occur, and that a separation layer has formed, so very little further remobilization is possible. However, if there are leaves that are still able to export some material, catabolic activities will be required for the remobilization. The accumulation of plastids, probably an indicator of living cells, closer to the veins is compatible with the hypothesis that, during phase 3 (and 4), the cells located far away from the veins may die, but that cells closer to the veins could participate in the degradation of macromolecules. The tree may be prepared for the winter by the end of phase 2, but if no frost hard enough to kill the cells occurs, the tree gets a bonus phase in which a small fraction of the remaining nutrients could perhaps be remobilized. The second possible explanation is that if the leaf cells have functionally separated from the trunk, their fate will not influence the rest of the tree so they may live or die without any particular adaptive benefit. In any case, these data are compatible with the concept that leaf senescence and cell death are to some extent antagonistic processes. Our data show that anthocyanin accumulation is likely to have a role in photoprotection, as suggested by several authors (Feild et al., 2001; Hoch et al., 2003). In the ecological literature, there has recently been speculation about a possible connection between autumn senescence and insect herbivory (Archetti, 2000; Hagen et al., 2003), assuming that autumn colors may be an honest, or dishonest, signal to insects about the status of the tree. Strong arguments against this, from a physiological perspective, have been put forward by Ougham et al. (2005) and, given the fact that accumulation and degradation of pigments in autumn leaves can be explained by obvious physiological and developmental factors, we think that there is unlikely to be a direct link between autumn colors and herbivore preference, and instead the phenomena are highly correlated because both are highly dependent on the nitrogen status of the tree. Is there a point of no return in autumn senescence of aspen leaves? To our knowledge, this question has not been experimentally addressed, but considering the events that we have followed here, one could speculate that the reprogramming of gene expression late in phase 1 could be an irreversible process or, alternatively, that the leaves could re-green if the whole tree is exposed to a longer photoperiod. However, it is doubtful that such a capability would have adaptive significance in nature, since trees will never experience an increase in the photoperiod during the autumn. However, we now have the tools to also address this question. By detailed studies of a tree during autumn senescence, we believe that we have obtained novel insights into the fundamental questions posed in the introduction. The conversion of chloroplasts to gerontoplasts is driven by developmental factors, the nutrients are largely remobilized over a period of 18 d (Sept 11 to Sept 29, phase 2), and, in the middle of this phase, mitochondria take over as the main providers of energy for the process. During this phase, plastids are converted to gerontoplasts and many cells die, but some survive and retain their gerontoplasts, mitochondria, and perhaps also nuclei until the leaves abscise, up to 1 week later. Finally, the photoperiod seems to be the sole trigger for the onset of autumn senescence in aspen, and the signal transduction chain between the photoreceptor phytochrome and the downstream genes that execute the senescence program can now be investigated with molecular and genomic tools. MATERIALS AND METHODS Leaf Material Leaves were sampled every day at 11 to 11:30 am between August 25 and October 9, 2003, from a free-growing aspen tree (Populus tremula) on the Umeå University campus (see Bhalerao et al., 2003). Five leaves were picked at each time point, pooled, and ground in liquid nitrogen. Once or twice a week, two single leaves were picked for fixation, embedding, and TEM. Fresh weight and dry weight were measured and mean leaf weight was calculated. Spectroscopic Measurements Chlorophyll Triplicates of about 30 mg of ground leaf material (a mixture of five leaves for each date) were extracted in 80% aqueous acetone solution buffered with 25 mm HEPES. The extract was analyzed for absorbance at wavelengths of 646.6, 663.6, and 750.0 nm using a Biochrom 4060 spectrophotometer (Pharmacia LKB Biochrom). The concentrations of chlorophyll a and chlorophyll b were then calculated using the equations of Porra et al. (1989). Anthocyanins Duplicates of about 40 mg of ground leaf material (a mixture of five leaves for each date) were extracted with 85% aqueous acetone and 1% HCl (85:15) at 4°C. After phase separation with diethylether to remove chlorophylls, the aqueous extract was analyzed for absorbance at wavelengths of 535.0, 650.0, and 750.0 nm using a Biochrom 4060 spectrophotometer (Pharmacia LKB Biochrom). Anthocyanin concentrations were then calculated using the extinction coefficient (⁠ \(E_{1\mathrm{cm}}^{1\%}{=}98.2\) at λ = 535 nm) according to Do and Cormier (1991). Pigment Analysis by HPLC Carotenoids were extracted with 96% ethanol and quantified according to Król et al. (1995). Pigments were detected by measuring the absorbance at 440 nm. Retention times and response factors for chlorophyll a, chlorophyll b, lutein, β-carotene, neoxanthin, violaxanthin, antheraxanthin, and zeaxanthin were previously determined (Król et al., 1995). Analysis was performed using the chromatograph's software (System Gold; Beckham Instruments). To minimize instrumental bias, samples were run in random order. The epoxidation state of xanthophyll cycle pigments was calculated according to the formula (V + 1/2A)/(V + A + Z). Several unidentified peaks were found in the chromatograms in extracts from the later stages of senescence. We assumed that some of these pigments may have been carotenoid esters, so we saponified extracts from late phase 3 (October 3) by adding 25 μL of 20% KOH to the extracts, then incubating the mixtures in darkness on ice for 18 h. Saponified extracts were filtered again before the HPLC analysis. The concentrations of neoxanthin, violaxanthin, and antheraxanthin increased by about 50% in the saponified extracts, indicating that these carotenoids were released from esters upon saponification. The amounts of zeaxanthin, lutein, β-carotene, chlorophyll a, and chlorophyll b changed only marginally. Thus, the results suggest that esters of neoxanthin, violaxanthin, and antheraxanthin were present in the autumn leaves. Two of the pigments (with retention times of 6.2 and 6.5 min, respectively) were not affected by saponification; the first pigment only marginally accumulated during senescence and is sometimes present in leaves of other plants (data not shown), while the second was not detected until September 19 and accumulated thereafter. The first pigment had a similar absorbance spectrum to violaxanthin (peaks at 443 and 472 nm in our system), and the second pigment had peaks at 448 and 473 nm. These are denoted carotenoids X and Y (CX and CY). Two pigments (retention times 8.2 and 8.3 min, respectively) had absorbance peaks at 438 and 467 nm (similar to neoxanthin) and disappeared upon saponification. These pigments were tentatively identified as neoxanthin esters (NE1 and NE2). Another pigment with a retention time of 8.5 min also disappeared upon saponification but had an absorbance spectrum resembling that of violaxanthin, with peaks at 443 and 471 nm, so this pigment could represent a violaxanthin ester (VE1). The peak with a retention time of 8.9 min was a nonresolved doublet that disappeared in the saponified extract. The absorbance spectrum of this peak most resembled that of neoxanthin, so we denoted it NE3, despite the fact that it most likely corresponds to two different carotenoid esters. A doublet appeared at retention times of 9.4 and 9.6. These pigments disappeared upon saponification. The absorbance spectra of the leading and trailing peaks in the doublet resembled those of neoxanthin and violaxanthin, respectively. These peaks were denoted NE4 and VE2, respectively. Finally, the peak with the largest area (retention time 10 min) had a spectrum resembling that of violaxanthin, but this pigment was stable during saponification. We tentatively denoted this pigment VX. These pigments have not been rigorously identified, so the assignments were made purely for practical reasons and are only tentative. For example, the increase in the amount of antheraxanthin after saponification indicates that antheraxanthin esters were present, but none of the peaks have been assigned as antheraxanthin esters. In addition to the pigments discussed here, several minor unassigned peaks (with typical carotenoid absorbance spectra) also appeared in the chromatograms. ATP and Sugar Measurements About 20 mg of ground leaf material (a mixture of five leaves for each date) was extracted in 3% TCA. ATP was determined by the firefly luciferase method (Gardeström and Wigge, 1988). ADP was measured after pyruvate kinase-catalyzed conversion to ATP (Roche Diagnostics). Soluble sugars (Suc, Glc, and Fru) and starch were measured in the soluble and residual fractions of ethanol-water extracts according to Stitt et al. (1989). TEM Ten pieces, each with a surface area of about 1.5 mm2, were cut from each leaf and infiltrated under mild vacuum for 15 min in a 3% (w/v) glutaraldehyde/0.1 m phosphate buffer (pH 7.2), then incubated for 4 h at room temperature (primary fixation), washed with phosphate buffer, and changed to 2% (w/v) osmium tetroxide for 2 h (secondary fixation). After washing in distilled water, the samples were dehydrated in a graded ethanol series, transferred to propyleneoxide for 3 × 10 min, and then epoxyresin (TAAB 812; TAAB Laboratories) was added dropwise every 10 min until the resin concentration reached approximately 10% (v/v). The samples were then left on a rotator overnight. The next day, the resin was sequentially changed every 4 h to 25%, 50%, and finally 75%, then incubated at 75% overnight, after which they were kept in 100% resin for one-half day and then embedded on silicon frames and incubated at 60°C for 1 d. After embedding, the samples were cut with a diamond knife and transferred to Formvar-coated 200 mesh copper grids. The grids were stained with 3% aqueous uranyl acetate for 30 min and with lead citrate for 10 min before being examined under a JEOL JEM 1230 transmission electron microscope operated at 80 kV. Elemental Analysis N%, C%, δ13C, and δ15N values of the pooled leaf samples were measured according to Ohlsson and Wallmark (1999), using a continuous flow isotope ratio mass spectrometer (20–20 Stable Isotope Analyzer; Europa Scientific) interfaced with an elemental analyzer unit (ANCA-NT system, solid/liquid preparation module; Europa Scientific). Iron, sulfur, phosphorus, and potassium concentrations of the pooled leaf samples were measured according to Emteryd (2003), using an inductively coupled plasma mass spectrometer (Elan 6100; Perkin-Elmer). 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Planta 163 : 91 –96 Wolf FT ( 1956 ) Changes in chlorophylls a and b in autumn leaves. Am J Bot 43 : 714 –718 Author notes 1 This work was supported by the Swedish Research Council and the Swedish Research Council for the Environment, Agricultural Sciences, and Spatial Planning. * Corresponding author; e-mail [email protected]; fax 46–786–66–76. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Johanna Keskitalo ([email protected]). Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.105.066845. © 2005 American Society of Plant Biologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Suh, Mi Chung; Samuels, A. Lacey; Jetter, Reinhard; Kunst, Ljerka; Pollard, Mike; Ohlrogge, John; Beisson, Fred

doi: 10.1104/pp.105.070805pmid: 16299169

Abstract All vascular plants are protected from the environment by a cuticle, a lipophilic layer synthesized by epidermal cells and composed of a cutin polymer matrix and waxes. The mechanism by which epidermal cells accumulate and assemble cuticle components in rapidly expanding organs is largely unknown. We have begun to address this question by analyzing the lipid compositional variance, the surface micromorphology, and the transcriptome of epidermal cells in elongating Arabidopsis (Arabidopsis thaliana) stems. The rate of cell elongation is maximal near the apical meristem and decreases steeply toward the middle of the stem, where it is 10 times slower. During and after this elongation, the cuticular wax load and composition remain remarkably constant (32 μg/cm2), indicating that the biosynthetic flux into waxes is closely matched to surface area expansion. By contrast, the load of polyester monomers per unit surface area decreases more than 2-fold from the upper (8 μg/cm2) to the lower (3 μg/cm2) portion of the stem, although the compositional variance is minor. To aid identification of proteins involved in the biosynthesis of waxes and cutin, we have isolated epidermal peels from Arabidopsis stems and determined transcript profiles in both rapidly expanding and nonexpanding cells. This transcriptome analysis was validated by the correct classification of known epidermis-specific genes. The 15% transcripts preferentially expressed in the epidermis were enriched in genes encoding proteins predicted to be membrane associated and involved in lipid metabolism. An analysis of the lipid-related subset is presented. The plant cuticle is a continuous lipophilic layer covering the surface of all epidermal cell types (Esau, 1977) and is one of their distinctive characteristics (Holloway, 1982; Jeffree, 1996; Kunst et al., 2005). It forms a vital hydrophobic barrier over the aerial surfaces of land plants during primary stages of development, limiting nonstomatal water loss and gaseous exchanges, controlling the absorption of lipophilic compounds, providing mechanical strength and viscoelastic properties (Baker et al., 1982; Hoffmann-Benning and Kende, 1994; Riederer and Schreiber, 2001), preventing organ fusion during development (Lolle et al., 1998; Sieber et al., 2000), as well as protecting the plant from nonbiotic and biotic stressors from the environment (Schweizer et al., 1996). The cuticle is composed of cutin (Kolattukudy, 2001), a polymer of fatty acid derivatives, which abuts the cell wall and is embedded in and covered with a mixture of ubiquitous aliphatic compounds (mainly C24-C34 alkanes, alcohols, and ketones) called cuticular waxes (Kunst and Samuels, 2003), as well as minor, extremely diverse, compounds (Jetter, 2000; Jetter et al., 2002; Vermeer et al., 2003). Due to their close physical association, it is difficult to distinguish between the relative contribution of the cutin matrix and cuticular waxes to the physical properties and the biological functions of the cuticle. However, experimentally, the cutin and cuticular waxes are usually considered and analyzed separately, as the waxes are easily extracted in organic solvents while the cutin polymer remains insoluble. The cutin matrix is composed mainly of C16 and C18 hydroxy and epoxy fatty acid monomers. Minor amounts of aromatic compounds are also present (Kolattukudy, 1980; Nawrath, 2002; Heredia, 2003). It has long been believed that cutin is formed by interesterification of fatty acids only, but it is now clear that glycerol is present and also esterified to the fatty acid monomers (Graça et al., 2002). The size and the exact structure of the polymer (dendrimeric, cross-linked, etc.) are still a matter of uncertainty. In Arabidopsis (Arabidopsis thaliana) epidermis, the major monomers released by polyester depolymerization of delipidated residues are not ω-hydroxy fatty acids, but dicarboxylic acids (Bonaventure et al., 2004). Thus it is not clear whether the rigid classification of cutin as highly enriched in ω-hydroxy fatty acids needs to be modified or whether the dicarboxylic acids represent a novel polyester domain. Knowledge on the biosynthesis of cutin is based largely on a few early studies, mostly done with broad bean (Vicia faba) leaves. These studies indicated that cutin biosynthesis involves ω-hydroxylation followed by midchain hydroxylation and incorporation into the polymer (Kolattukudy and Walton, 1972), and that hydroxy fatty acids can be incorporated into cutin by a reaction requiring CoA and ATP (Croteau and Kolattukudy, 1974). Genetic approaches to cutin biosynthesis are relatively undeveloped. In the Arabidopsis fatb-ko line, where a disruption in an acyl-carrier protein (ACP) thioesterase gene results in reduced general availability of cytosolic palmitate (Bonaventure et al., 2003), there is an 80% loss of C16 monomers and a compensatory increase in C18 monomers in epidermal polyesters (Bonaventure et al., 2004). The first mutant demonstrated by chemical analysis to be specifically affected in cutin metabolism has been reported only very recently (Xiao et al., 2004). The ethyl methanesulfonate-induced mutant att1 shows enhanced disease severity to a virulent strain of Pseudomonas syringae, a 70% decrease in cutin amount, a loose cuticle ultrastructure, and increased permeability to water vapor. ATT1 (At4g00360) encodes a P450 monooxygenase belonging to the CYP86A family (Xiao et al., 2004) and has been demonstrated to catalyze the ω-hydroxylation of fatty acids (Duan and Schuler, 2005). ATT1 is thus probably responsible for the synthesis of ω-hydroxy fatty acid and/or dicarboxylic acid monomers found in Arabidopsis epidermal polyesters. In maize (Zea mays) leaves, a peroxygenase has been shown to be involved in the synthesis of C18 epoxy monomers of cutin (Lequeu et al., 2003). This enzyme, associated with a cytochrome P450 monooxygenase (Pinot et al., 1999) and a membrane-bound epoxide hydrolase, can catalyze in vitro the formation of these monomers (Blée and Schuber, 1993). The characterization of the Arabidopsis lacs2 mutant indicates that LACS2, an epidermis-specific long-chain acyl-CoA synthetase, is involved in leaf cuticle formation and barrier function (Schnurr et al., 2004) and suggests the likely involvement of specific acyl-CoA pools in cutin biosynthesis. Whether the polymerization reactions of the cutin monomers occur inside or outside the epidermal cells, which enzymes are involved, and how cutin synthesis is coordinated with the deposition of waxes is still completely unknown and a major challenge in the study of plant epidermis. Cuticular waxes of the Arabidopsis shoot are found both above the cutin matrix (epicuticular) as well as embedded in the cutin (intracuticular). The chemical compositions of the Arabidopsis stem and leaf waxes have been investigated in both wild-type and eceriferum (glossy) plant lines (Hannoufa et al., 1993; Jenks et al., 1995; Rashotte et al., 2001, 2004). The predominant wax components of Arabidopsis stems are C29 alkane, ketone, and secondary alcohols, together with smaller quantities of C28 primary alcohol and C30 aldehyde, while leaves have C31 alkane but lack the abundant ketones and secondary alcohols of the stem (for review, see Jenks et al., 2001). Stems also have much higher wax loads than leaves (more than 30 times higher in Arabidopsis ecotype Columbia [Col]-0). These saturated aliphatic wax components are synthesized as C16-C18 fatty acids in the plastid, followed by fatty acid elongation associated with the endoplasmic reticulum to form very-long-chain fatty acids (VLCFA; Post-Beittenmiller, 1996; Kunst and Samuels, 2003). The function of one of the condensing enzymes of the elongase complex was demonstrated by the cer6 knockdown, which has a highly glossy stem phenotype and 7% of wild-type wax (Millar et al., 1999). Subsequent modifications occur to the VLCFAs, but the mechanisms and gene products involved are less well characterized. The mechanism of transport of waxes onto the epidermal surface is better understood as a result of the recent discovery of CER5, an ATP-binding cassette (ABC) transporter involved in the export of cuticular waxes to the stem surface (Pighin et al., 2004). The cuticle is known to be synthesized at very early stages of embryo and organ development (Szczuka and Szczuka, 2003). In rapidly growing internodes of deepwater rice (Oryza sativa), it has been observed that the cuticle thickness remains constant (Hoffmann-Benning and Kende, 1994). However, whether the secretions of its cutin and wax components occur at the same time and whether cuticles have a constant composition during the dynamic process of rapid expansion of young organs has not been documented, although it is clearly an important question regarding the mechanism of cuticle formation. To investigate the timing of wax and cutin deposition, we have performed structural studies and lipid analyses on the epidermis of different parts of rapidly elongating Arabidopsis stems. In addition, to gain insights into the identity of the enzyme families and isoforms involved in the deposition of wax and cutin, we have undertaken a gene expression study of the epidermal cells of the stem, which is also the first genome-wide study of the transcriptome of the aerial plant epidermis. We report here on the results of these combined approaches and discuss the expression of a subset of genes involved in lipid metabolism in the light of the surface structure and the lipid composition of the stem cuticle. RESULTS The Length of Epidermal Cells Increases More Than 80-Fold during Stem Elongation The bolting inflorescence stem of Arabidopsis provides an opportunity to study the biosynthesis of cuticular lipids by the epidermis during cell expansion. When growth was measured over a 24-h period for 9- to 11-cm-long bolting stems, the total length of the stems increased by 3.7 cm ± 0.3 (mean ± se, n = 6). The elongation of apical, but not basal, 0.5-cm segments was observed (Fig. 1 Figure 1. Open in new tabDownload slide Size of Arabidopsis stem epidermal cells and stem elongation within 0.5-cm-long segments. A to C, Confocal scanning laser microscopy with propidium iodide staining of the cell wall of regions near the shoot apex (A), in the middle of the stem (B), and at the bottom of the stem (C). D, Thin marks were made with black ink along 9- to 11-cm stems, starting at the apex and defining 0.5-cm initial segments. The position of each mark relative to the base of the stem was measured after 24 h and elongation of each segment calculated. Values are means ± se (n = 6). Figure 1. Open in new tabDownload slide Size of Arabidopsis stem epidermal cells and stem elongation within 0.5-cm-long segments. A to C, Confocal scanning laser microscopy with propidium iodide staining of the cell wall of regions near the shoot apex (A), in the middle of the stem (B), and at the bottom of the stem (C). D, Thin marks were made with black ink along 9- to 11-cm stems, starting at the apex and defining 0.5-cm initial segments. The position of each mark relative to the base of the stem was measured after 24 h and elongation of each segment calculated. Values are means ± se (n = 6). ). Most of the increase in total length (about 85%) occurred in the top 3-cm segment of the stem, 15% in the middle part, while no increase was detected below 7 cm from the top. The elongation rate was greater nearer to the apex and maximal in the first 0.5-cm portion. Given typical mean diameters of the stem segments, this elongation corresponds to an increase in epidermis surface area of 60 mm2/24 h in the top 3 cm of the stem and 8 mm2/24 h in the middle 3-cm segment. Growth of epidermal cells was followed with cryo-scanning electron microscopy (SEM) and confocal scanning laser microscopy with propidium iodide staining of the cell wall. Near the shoot apex (Fig. 1A), the cells of the epidermis were nearly isodiametric, but they soon underwent anisotropic growth to elongate greatly in the axial dimension (Fig. 1B). At the base of a 10-cm stem, the cells were elongated and trichomes were seen (Figs. 1C and 2C Figure 2. Open in new tabDownload slide Cryo-SEM of developing inflorescence stem of Arabidopsis (Col-0). A to C, Low magnification view of top, within 1 cm of the apex (A), middle, between 4 to 5 cm from the apex (B), and base, between 8 to 9 cm (C). D to F, Higher magnification view of epicuticular crystals on the surface of the top (D), middle (E), and base (F). Figure 2. Open in new tabDownload slide Cryo-SEM of developing inflorescence stem of Arabidopsis (Col-0). A to C, Low magnification view of top, within 1 cm of the apex (A), middle, between 4 to 5 cm from the apex (B), and base, between 8 to 9 cm (C). D to F, Higher magnification view of epicuticular crystals on the surface of the top (D), middle (E), and base (F). ). No trichomes were present in the middle and top segments (Fig. 2, A and B). When the cell dimensions were quantified using cryo-SEM, the length of the cells increased from 6 μm ± 0.3 (mean ± se, n = 24) to 532 ± 44 μm (mean ± se, n = 14), an increase of 87-fold. In contrast, cell widths did not increase significantly. The Load and Composition of Waxes Remain Constant along the Stem To test whether the biosynthesis of cuticular waxes is coordinated with the rapid expansion of the stem epidermal cells, cryo-SEM and gas chromatography (GC) with mass spectroscopy (MS) and flame ionization detection (FID) were used. Cryo-SEM provided information about the epicuticular wax crystals, while the GC-FID and GC-MS provided quantitative wax load and composition information. Epicuticular wax crystals were found over the entire stem surface, both vertical rods, tubes, longitudinal bundles of rodlets, and horizontal, reticulate plates (Fig. 2, D–F). In the region of most rapid elongation, the cuticle showed striations and crystals distorted along the axis of elongation. The cuticle around stomata did not have epicuticular crystals, only a smooth epicuticular film (Fig. 2, E and F). Qualitative and quantitative chemical analyses were performed to compare the cuticular wax in the top, middle, and basal 3-cm segments of the bolting inflorescence stem. The wax load, expressed on a per-unit-area basis, was constant between these segments of the stem (Fig. 3 Figure 3. Open in new tabDownload slide Epidermal loads of wax, polyester, and other acyl lipids along the stem. Fatty acid derivatives were analyzed using sections of whole stems for wax and polyesters and epidermal peels for other acyl lipids. The loads are expressed in micrograms of fatty acid derivatives per square centimeter of epidermis. Difference in polyester load between the middle (or top) and the base of stems was significant (Student's t test, P < 0.05). Other differences were not significant. Values are means ± se (n = 5). Figure 3. Open in new tabDownload slide Epidermal loads of wax, polyester, and other acyl lipids along the stem. Fatty acid derivatives were analyzed using sections of whole stems for wax and polyesters and epidermal peels for other acyl lipids. The loads are expressed in micrograms of fatty acid derivatives per square centimeter of epidermis. Difference in polyester load between the middle (or top) and the base of stems was significant (Student's t test, P < 0.05). Other differences were not significant. Values are means ± se (n = 5). ). The composition along the stem did not vary significantly (Fig. 4a Figure 4. Open in new tabDownload slide Relative composition of fatty acid derivatives in waxes and polyesters along the stem. Waxes (a) and polyesters (b) were analyzed using sections of whole stems. Values are means ± se (n = 5). Figure 4. Open in new tabDownload slide Relative composition of fatty acid derivatives in waxes and polyesters along the stem. Waxes (a) and polyesters (b) were analyzed using sections of whole stems. Values are means ± se (n = 5). ), with C29 alkane, ketone, and secondary alcohols predominating, and the overall composition is similar to that reported previously (Hannoufa et al., 1993; Jenks et al., 1995; Rashotte et al., 2001). In addition, primary alcohols of chain lengths C26-C30 and alkyl esters were observed in all segments of the stem at very similar percentages. To increase the spatiotemporal resolution of our analysis and gain insights into the timing of wax deposition in the region of the stem elongating most rapidly, the top 3-cm segment of the stem was further divided into 1-cm portions and the waxes analyzed. No significant differences were found in the wax load or wax composition between the three portions of the top segment (data not shown). Polyester Composition Is Constant along the Stem, But the Load Decreases at the Base To quantify the polyester monomers in the stem, the method bypassing cuticle isolation we have reported earlier (Bonaventure et al., 2004) was used with minor modifications. Because our previous report has demonstrated that stem polyester monomers were located exclusively in the epidermis, analyses were done on whole-stem segments. The aliphatic monomer polyester composition in the 9- to 11-cm elongating stems was largely unsaturated aliphatic dicarboxylates and similar to the composition reported earlier for 20-cm stems of the same ecotype (Bonaventure et al., 2004; Fig. 4b). The variations in monomer composition observed in the three stem segments were minor. By contrast, the total load in aliphatic polyester monomers per surface area unit decreased substantially (about 2-fold) and significantly between the middle and lower portions of the stems, but not significantly between the upper and the middle portions (Fig. 3). Neither the load nor the composition of stem polyesters was significantly changed in the three stem segments by increasing the depolymerization time from 48 to 72 h (data not shown). This decrease in the amount of polyester monomers at the base of the stem could indicate a change in structure giving rise to a polymer less or not susceptible to depolymerization (e.g. cutan). A more refined analysis of the polyesters in portions of the apical segment as performed for waxes was hampered by the low amount of polyesters. Such an analysis may require the use of a plant species with larger stems. In Stem Epidermis There Is a Considerable Flux of Lipids Exported onto the Surface We also determined the fatty acid content of the intracellular acyl lipids (membrane and storage lipids) of epidermal peels. When compared to the amounts of fatty acid derivatives measured in waxes and polyesters, the intracellular lipids represented less than one-half of the total cell lipids (Fig. 3). The fatty acid composition of the stem epidermis was found to be similar to that of the whole stem (Supplemental Fig. 1). In all stem segments, the ratio of surface lipids to epidermal intracellular acyl lipids was about 3:2 in mass (Fig. 3). This means that the epidermal cells of the elongating stem top need to have machinery capable of exporting onto the surface more than one-half of the approximately 75 μg of fatty acids produced per square centimeter of epidermis. Given an average rate of expansion of 0.6 cm2/24 h for the stem top, the accumulation of total fatty acids in the corresponding epidermis can be estimated to be around 1.9 μg cm−2 h−1 and the net rate of export 1.1 μg cm−2 h−1 (assuming that fatty acid accumulation commences when cell elongation starts). This represents a considerable flux of hydrophobic compounds, possibly partially polymerized, that must go through the plasma membrane and the aqueous cell wall. Since this specialized machinery of synthesis and export of surface lipids is specific to the epidermis, it is reasonable to assume that most of the genes encoding the enzymes, transporters, and other proteins that are part of the machinery will be transcriptionally up-regulated in the epidermis compared to the other tissues of the stem. Expression of these genes could be stronger in the elongating top epidermis or restricted to it. The above rationale provided the basis for the microarray analysis described below. Genome-Wide Microarray Analysis Identified about 1,900 Genes Up-Regulated in the Stem Epidermis To investigate the mRNA levels of genes expressed in the epidermis of the apical and basal segments of the stems, we used the GeneChip Arabidopsis ATH1 Genome Array (Affymetrix) that represents 22,748 probe sets covering approximately 23,750 Arabidopsis genes (Redman et al., 2004). Epidermis manually peeled from stem segments (Fig. 5 Figure 5. Open in new tabDownload slide Peeling of epidermis from Arabidopsis stems. A, Transverse section of stems viewed under a light microscope after staining with phloroglucinol. The epidermis is the transparent outermost single cell layer. B, Using manual dissection, epidermis can be isolated from the stem as a transparent film. Figure 5. Open in new tabDownload slide Peeling of epidermis from Arabidopsis stems. A, Transverse section of stems viewed under a light microscope after staining with phloroglucinol. The epidermis is the transparent outermost single cell layer. B, Using manual dissection, epidermis can be isolated from the stem as a transparent film. ) was used as the source of RNAs. The small amounts of chlorophyll detected in the epidermis peels (on average 0.05 mg chlorophyll/g fresh weight versus 0.6 mg chlorophyll/g fresh weight in total stems), indicated that contamination by nonepidermal tissues was minor. In fact, the green pigmentation in the peels may arise largely from epidermal plastids, which in fluorescence microscopy show a low, but detectable, amount of red autofluorescence usually attributed to chlorophyll in chloroplasts (data not shown). Enrichment of epidermal mRNAs in the epidermal peels was confirmed by the epidermis-to-stem gene expression ratios measured for genes of known epidermal, extraepidermal, or ubiquitous expression (Table I Table I. Genes with known preferential expression in epidermis or in vascular tissues were correctly identified by the microarray experiments using epidermal peels from stems List of genes whose preferential or specific site of expression is epidermis (top of the list) or vascular tissues (bottom) as shown by techniques other than microarrays. The organ investigated and the reference is given. For known housekeeping genes (middle), only a few randomly selected examples are given. Molecular function of the proteins encoded by the genes: ACR4, putative receptor kinase; ATML1 and PDF2, homeodomain protein; AtPIN1, putative auxin efflux carrier protein; AtPPT1 (=cue1), phosphoenolpyruvate/phosphate translocator; CER2, unknown protein; CER5, ABC transporter; CBP20 and CBP80, nuclear cap binding protein; CER6 (=CUT1) and FDH (FIDDLEHEAD), β-ketoacyl-CoA synthase; LACERATA, fatty acid ω-hydroxylase; LACS2, long-chain acyl-CoA synthetase; L23a, 60S ribosomal protein; PPX1 and PPX2, protein phosphatase; UBQ1, ubiquitin extension protein; XCP1 and XCP2, cysteine peptidase; WAX2 (=YORE-YORE), unknown. Gene Locus . Gene Name . Expression Ratioa . . Preferential Expression . Reference . . . Top . Base . . . Epidermal At3g59420 ACR4 24.8 19.1     Epidermis (all organs) Tanaka et al. (2002) At4g21750 ATML1 6.2 5.5     L1 layer of shoot apex Sessions et al. (1999) At4g04890 PDF2 6.0 5.9     L1 layer of shoot apex Abe et al. (2003) At4g24510 CER2 4.1 6.0     Epidermis Xia et al. (1997) At2g26250 FDH 4.0 3.9     Epidermis Yephremov et al. (1999); Pruitt et al. (2000) At1g51500 CER5 3.5 3.1     Epidermis (stem) Pighin et al. (2004) At1g68530 CER6 3.1 3.1     Shoot epidermis Millar et al. (1999) At2g45970 LACERATA 3.1 2.2     Epidermis-specific Wellesen et al. (2001) At5g57800 WAX2 2.6 2.2     L1 layer of shoot apex Chen et al. (2003); Kurata et al. (2003) At1g49430 LACS2 2.5 1.2     Epidermis (leaf) Schnurr et al. (2004) Ubiquitous At5g55260 PPX2 1.6 1.6     All tissues Pujol et al. (2000) At3g52590 UBQ1 1.1 1.6     Most tissues Holtorf et al. (1995) At4g26720 PPX1 1.0 1.2     All tissues Pujol et al. (2000) At3g18780 Actin 2 0.9 0.7     Vegetative tissues An et al. (1996) At2g13540 CBP80 0.8 0.8     Most tissues Kmieciak et al. (2002) At5g44200 CBP20 0.7 1.0     Most tissues Kmieciak et al. (2002) At1g49240 Actin 8 0.6 0.7     Vegetative tissues An et al. (1996) At3g55280 L23a 0.6 1.4     Most tissues Volkov et al. (2003) Vascular At5g33320 AtPPT1 0.4 1.0     Stem vasculature Knappe et al. (2003) At1g73590 AtPIN1 0.2 0.2     Xylem and cambium Gälweiler et al. (1998) At4g35350 XCP1 0.1 0.5     Xylem Funk et al. (2002) At1g20850 XCP2 0.1 0.3     Xylem Zhao et al. (2000) Gene Locus . Gene Name . Expression Ratioa . . Preferential Expression . Reference . . . Top . Base . . . Epidermal At3g59420 ACR4 24.8 19.1     Epidermis (all organs) Tanaka et al. (2002) At4g21750 ATML1 6.2 5.5     L1 layer of shoot apex Sessions et al. (1999) At4g04890 PDF2 6.0 5.9     L1 layer of shoot apex Abe et al. (2003) At4g24510 CER2 4.1 6.0     Epidermis Xia et al. (1997) At2g26250 FDH 4.0 3.9     Epidermis Yephremov et al. (1999); Pruitt et al. (2000) At1g51500 CER5 3.5 3.1     Epidermis (stem) Pighin et al. (2004) At1g68530 CER6 3.1 3.1     Shoot epidermis Millar et al. (1999) At2g45970 LACERATA 3.1 2.2     Epidermis-specific Wellesen et al. (2001) At5g57800 WAX2 2.6 2.2     L1 layer of shoot apex Chen et al. (2003); Kurata et al. (2003) At1g49430 LACS2 2.5 1.2     Epidermis (leaf) Schnurr et al. (2004) Ubiquitous At5g55260 PPX2 1.6 1.6     All tissues Pujol et al. (2000) At3g52590 UBQ1 1.1 1.6     Most tissues Holtorf et al. (1995) At4g26720 PPX1 1.0 1.2     All tissues Pujol et al. (2000) At3g18780 Actin 2 0.9 0.7     Vegetative tissues An et al. (1996) At2g13540 CBP80 0.8 0.8     Most tissues Kmieciak et al. (2002) At5g44200 CBP20 0.7 1.0     Most tissues Kmieciak et al. (2002) At1g49240 Actin 8 0.6 0.7     Vegetative tissues An et al. (1996) At3g55280 L23a 0.6 1.4     Most tissues Volkov et al. (2003) Vascular At5g33320 AtPPT1 0.4 1.0     Stem vasculature Knappe et al. (2003) At1g73590 AtPIN1 0.2 0.2     Xylem and cambium Gälweiler et al. (1998) At4g35350 XCP1 0.1 0.5     Xylem Funk et al. (2002) At1g20850 XCP2 0.1 0.3     Xylem Zhao et al. (2000) a Mean ratio calculated from four epidermis-to-stem gene expression ratios, i.e. ratio of transcripts in epidermis versus transcripts in stems, determined by microarray experiments (Affymetrix ATH1 GeneChip) using the epidermis of a stem section (top or base) and the whole-stem section (top or base) as a reference. Open in new tab Table I. Genes with known preferential expression in epidermis or in vascular tissues were correctly identified by the microarray experiments using epidermal peels from stems List of genes whose preferential or specific site of expression is epidermis (top of the list) or vascular tissues (bottom) as shown by techniques other than microarrays. The organ investigated and the reference is given. For known housekeeping genes (middle), only a few randomly selected examples are given. Molecular function of the proteins encoded by the genes: ACR4, putative receptor kinase; ATML1 and PDF2, homeodomain protein; AtPIN1, putative auxin efflux carrier protein; AtPPT1 (=cue1), phosphoenolpyruvate/phosphate translocator; CER2, unknown protein; CER5, ABC transporter; CBP20 and CBP80, nuclear cap binding protein; CER6 (=CUT1) and FDH (FIDDLEHEAD), β-ketoacyl-CoA synthase; LACERATA, fatty acid ω-hydroxylase; LACS2, long-chain acyl-CoA synthetase; L23a, 60S ribosomal protein; PPX1 and PPX2, protein phosphatase; UBQ1, ubiquitin extension protein; XCP1 and XCP2, cysteine peptidase; WAX2 (=YORE-YORE), unknown. Gene Locus . Gene Name . Expression Ratioa . . Preferential Expression . Reference . . . Top . Base . . . Epidermal At3g59420 ACR4 24.8 19.1     Epidermis (all organs) Tanaka et al. (2002) At4g21750 ATML1 6.2 5.5     L1 layer of shoot apex Sessions et al. (1999) At4g04890 PDF2 6.0 5.9     L1 layer of shoot apex Abe et al. (2003) At4g24510 CER2 4.1 6.0     Epidermis Xia et al. (1997) At2g26250 FDH 4.0 3.9     Epidermis Yephremov et al. (1999); Pruitt et al. (2000) At1g51500 CER5 3.5 3.1     Epidermis (stem) Pighin et al. (2004) At1g68530 CER6 3.1 3.1     Shoot epidermis Millar et al. (1999) At2g45970 LACERATA 3.1 2.2     Epidermis-specific Wellesen et al. (2001) At5g57800 WAX2 2.6 2.2     L1 layer of shoot apex Chen et al. (2003); Kurata et al. (2003) At1g49430 LACS2 2.5 1.2     Epidermis (leaf) Schnurr et al. (2004) Ubiquitous At5g55260 PPX2 1.6 1.6     All tissues Pujol et al. (2000) At3g52590 UBQ1 1.1 1.6     Most tissues Holtorf et al. (1995) At4g26720 PPX1 1.0 1.2     All tissues Pujol et al. (2000) At3g18780 Actin 2 0.9 0.7     Vegetative tissues An et al. (1996) At2g13540 CBP80 0.8 0.8     Most tissues Kmieciak et al. (2002) At5g44200 CBP20 0.7 1.0     Most tissues Kmieciak et al. (2002) At1g49240 Actin 8 0.6 0.7     Vegetative tissues An et al. (1996) At3g55280 L23a 0.6 1.4     Most tissues Volkov et al. (2003) Vascular At5g33320 AtPPT1 0.4 1.0     Stem vasculature Knappe et al. (2003) At1g73590 AtPIN1 0.2 0.2     Xylem and cambium Gälweiler et al. (1998) At4g35350 XCP1 0.1 0.5     Xylem Funk et al. (2002) At1g20850 XCP2 0.1 0.3     Xylem Zhao et al. (2000) Gene Locus . Gene Name . Expression Ratioa . . Preferential Expression . Reference . . . Top . Base . . . Epidermal At3g59420 ACR4 24.8 19.1     Epidermis (all organs) Tanaka et al. (2002) At4g21750 ATML1 6.2 5.5     L1 layer of shoot apex Sessions et al. (1999) At4g04890 PDF2 6.0 5.9     L1 layer of shoot apex Abe et al. (2003) At4g24510 CER2 4.1 6.0     Epidermis Xia et al. (1997) At2g26250 FDH 4.0 3.9     Epidermis Yephremov et al. (1999); Pruitt et al. (2000) At1g51500 CER5 3.5 3.1     Epidermis (stem) Pighin et al. (2004) At1g68530 CER6 3.1 3.1     Shoot epidermis Millar et al. (1999) At2g45970 LACERATA 3.1 2.2     Epidermis-specific Wellesen et al. (2001) At5g57800 WAX2 2.6 2.2     L1 layer of shoot apex Chen et al. (2003); Kurata et al. (2003) At1g49430 LACS2 2.5 1.2     Epidermis (leaf) Schnurr et al. (2004) Ubiquitous At5g55260 PPX2 1.6 1.6     All tissues Pujol et al. (2000) At3g52590 UBQ1 1.1 1.6     Most tissues Holtorf et al. (1995) At4g26720 PPX1 1.0 1.2     All tissues Pujol et al. (2000) At3g18780 Actin 2 0.9 0.7     Vegetative tissues An et al. (1996) At2g13540 CBP80 0.8 0.8     Most tissues Kmieciak et al. (2002) At5g44200 CBP20 0.7 1.0     Most tissues Kmieciak et al. (2002) At1g49240 Actin 8 0.6 0.7     Vegetative tissues An et al. (1996) At3g55280 L23a 0.6 1.4     Most tissues Volkov et al. (2003) Vascular At5g33320 AtPPT1 0.4 1.0     Stem vasculature Knappe et al. (2003) At1g73590 AtPIN1 0.2 0.2     Xylem and cambium Gälweiler et al. (1998) At4g35350 XCP1 0.1 0.5     Xylem Funk et al. (2002) At1g20850 XCP2 0.1 0.3     Xylem Zhao et al. (2000) a Mean ratio calculated from four epidermis-to-stem gene expression ratios, i.e. ratio of transcripts in epidermis versus transcripts in stems, determined by microarray experiments (Affymetrix ATH1 GeneChip) using the epidermis of a stem section (top or base) and the whole-stem section (top or base) as a reference. Open in new tab ). In all cases, the genes known to have epidermal expression by techniques other than microarrays were found to have a top epidermis-to-top stem mean gene expression ratio of 2.5 or above, while housekeeping genes had ratios between 1.6 and 0.7 and extraepidermal-expressed genes showed ratios of 1 or below. A global analysis of transcripts found to be up-regulated in the stem epidermis compared to the whole stem is presented below and in the supplemental data, with a particular emphasis on genes known or suspected of being involved in lipid metabolism. Transcripts from a total of about 13,000 genes were detected in the stem segments (Supplemental Table I), which is very close to the value of 60% of expressed genes in the stem reported recently (Ma et al., 2005). The Affymetrix Microarray Suite 5.0 (MAS 5.0) software identified more than 3,000 genes whose expression was increased in the epidermis compared to the whole stem (data not shown). Using some statistical parameters calculated by MAS 5.0 and a conservative cutoff value of 2.0 for the lower estimate of the epidermis-to-stem gene expression ratio (see “Materials and Methods”), we identified about 1,900 genes up-regulated in the stem epidermis. This list breaks down into three subsets of about 600 each (Supplemental Table I): up-regulated in the epidermis of stem segments (top and base), in top segments alone (elongating), and in basal segments alone (nonelongating). These three subsets of transcripts (see Supplemental Table III for the identity) are likely to be enriched in genes related to various aspects of epidermal biology, such as trichome formation for the basal-only subset or cell elongation for the apical-only subset. A list of the 40 most up-regulated, highly expressed transcripts in the apical-only subset is given in Table II Table II. List of the 40 most up-regulated highly expressed genes in top stem epidermis classified by functional category Putative Functional Category and Locus . Description of Encoded Protein . Expression Ratioa (Top) . Expression Ratioa (Base) . Lipid Metabolism     At1g07720 β-Ketoacyl-CoA synthase family 9.9 2.7     At1g64400 Long-chain-acyl-CoA synthetase (LACS3) 9.4 1.7     At2g39400 Similar to lipase 13.4 4.8 Transport     At5g40780 Putative Lys and His transporter 16.9 3.3     At3g55130 ABC transporter family 12.3 3.1     At5g55930 Oligopeptide transporter OPT family protein 8.2 9.0 Cell Wall Metabolism     At4g30280 Putative xyloglucan:xyloglucosyl transferase 33.7 27.9     At4g02330 Similar to pectinesterase 29.7 211     At3g10720 Similar to pectinesterase 8.8 57.4 Disease Resistance     At1g78860 Lectin family protein (mannose-binding) 33.8 19     At2g33050 Similar to disease resistance protein Hcr2 10.2 4.4     At1g75040 Pathogenesis-related protein 5 (PR-5) 10.0 1.0 Signal Transduction     At1g21270 Wall-associated kinase 2 18.2 2.9     At5g20050 Protein kinase family protein 10.0 5.9     At1g11350 Ser-Thr kinase related 9.9 7.1     At1g21250 Wall-associated kinase 1 9.9 1.1     At2g13790 Leu-rich-repeat protein kinase family 9.3 3.4     At1g51805 Leu-rich-repeat protein kinase family 8.6 3.4     At4g31000 Calmodulin-binding protein 8.6 5.1     At4g23180 Ser-Thr kinase related 8.3 5.5     At2g01890 Putative purple acid phosphatase 8.3 1.8     At4g38550 Phospholipase-like protein 7.9 3.4 Transcription     At4g31800 WRKY family transcription factor 11 3.8     At2g38470 WRKY family transcription factor 8.9 4.2     At2g02450 No apical meristem (NAM) family protein 8.2 2.8 Unknown     At1g22890 Expressed protein 31.9 14.9     At1g78460 Expressed protein 25.2 9.2     At2g28570 Expressed protein 20.6 3.2     At3g28290 Integrin-related protein 14a 18.9 91.1     At5g20230 Plastocyanin-like domain-containing protein 14.9 9.1     At3g26200 Cytochrome P450-like 13.6 10.1     At2g05540 Gly-rich protein 11.2 80.2     At1g56150 Auxin-responsive family protein 10.5 6.1     At3g11600 Expressed protein 9.5 7.8     At5g05440 Expressed protein 9.2 2.3     At2g20950 Expressed protein 8.8 3.3     At4g36500 Expressed protein 7.9 3.4     At1g33600 Leu-rich-repeat family protein 7.9 8.4     At1g29430 Auxin-responsive family protein 7.8 9.6     At5g44020 Similar to soybean stem glycoprotein 7.4 36.2 Putative Functional Category and Locus . Description of Encoded Protein . Expression Ratioa (Top) . Expression Ratioa (Base) . Lipid Metabolism     At1g07720 β-Ketoacyl-CoA synthase family 9.9 2.7     At1g64400 Long-chain-acyl-CoA synthetase (LACS3) 9.4 1.7     At2g39400 Similar to lipase 13.4 4.8 Transport     At5g40780 Putative Lys and His transporter 16.9 3.3     At3g55130 ABC transporter family 12.3 3.1     At5g55930 Oligopeptide transporter OPT family protein 8.2 9.0 Cell Wall Metabolism     At4g30280 Putative xyloglucan:xyloglucosyl transferase 33.7 27.9     At4g02330 Similar to pectinesterase 29.7 211     At3g10720 Similar to pectinesterase 8.8 57.4 Disease Resistance     At1g78860 Lectin family protein (mannose-binding) 33.8 19     At2g33050 Similar to disease resistance protein Hcr2 10.2 4.4     At1g75040 Pathogenesis-related protein 5 (PR-5) 10.0 1.0 Signal Transduction     At1g21270 Wall-associated kinase 2 18.2 2.9     At5g20050 Protein kinase family protein 10.0 5.9     At1g11350 Ser-Thr kinase related 9.9 7.1     At1g21250 Wall-associated kinase 1 9.9 1.1     At2g13790 Leu-rich-repeat protein kinase family 9.3 3.4     At1g51805 Leu-rich-repeat protein kinase family 8.6 3.4     At4g31000 Calmodulin-binding protein 8.6 5.1     At4g23180 Ser-Thr kinase related 8.3 5.5     At2g01890 Putative purple acid phosphatase 8.3 1.8     At4g38550 Phospholipase-like protein 7.9 3.4 Transcription     At4g31800 WRKY family transcription factor 11 3.8     At2g38470 WRKY family transcription factor 8.9 4.2     At2g02450 No apical meristem (NAM) family protein 8.2 2.8 Unknown     At1g22890 Expressed protein 31.9 14.9     At1g78460 Expressed protein 25.2 9.2     At2g28570 Expressed protein 20.6 3.2     At3g28290 Integrin-related protein 14a 18.9 91.1     At5g20230 Plastocyanin-like domain-containing protein 14.9 9.1     At3g26200 Cytochrome P450-like 13.6 10.1     At2g05540 Gly-rich protein 11.2 80.2     At1g56150 Auxin-responsive family protein 10.5 6.1     At3g11600 Expressed protein 9.5 7.8     At5g05440 Expressed protein 9.2 2.3     At2g20950 Expressed protein 8.8 3.3     At4g36500 Expressed protein 7.9 3.4     At1g33600 Leu-rich-repeat family protein 7.9 8.4     At1g29430 Auxin-responsive family protein 7.8 9.6     At5g44020 Similar to soybean stem glycoprotein 7.4 36.2 a Mean ratio of epidermis-to-stem gene expression. Gene expressions were determined as in Table I. Among genes up-regulated in top stem epidermis, only those showing a signal intensity >1,000 in top epidermis are shown. Descriptions of proteins are from The Institute for Genomic Research, Munich Information Center for Protein Sequences, or Swiss-Prot. Open in new tab Table II. List of the 40 most up-regulated highly expressed genes in top stem epidermis classified by functional category Putative Functional Category and Locus . Description of Encoded Protein . Expression Ratioa (Top) . Expression Ratioa (Base) . Lipid Metabolism     At1g07720 β-Ketoacyl-CoA synthase family 9.9 2.7     At1g64400 Long-chain-acyl-CoA synthetase (LACS3) 9.4 1.7     At2g39400 Similar to lipase 13.4 4.8 Transport     At5g40780 Putative Lys and His transporter 16.9 3.3     At3g55130 ABC transporter family 12.3 3.1     At5g55930 Oligopeptide transporter OPT family protein 8.2 9.0 Cell Wall Metabolism     At4g30280 Putative xyloglucan:xyloglucosyl transferase 33.7 27.9     At4g02330 Similar to pectinesterase 29.7 211     At3g10720 Similar to pectinesterase 8.8 57.4 Disease Resistance     At1g78860 Lectin family protein (mannose-binding) 33.8 19     At2g33050 Similar to disease resistance protein Hcr2 10.2 4.4     At1g75040 Pathogenesis-related protein 5 (PR-5) 10.0 1.0 Signal Transduction     At1g21270 Wall-associated kinase 2 18.2 2.9     At5g20050 Protein kinase family protein 10.0 5.9     At1g11350 Ser-Thr kinase related 9.9 7.1     At1g21250 Wall-associated kinase 1 9.9 1.1     At2g13790 Leu-rich-repeat protein kinase family 9.3 3.4     At1g51805 Leu-rich-repeat protein kinase family 8.6 3.4     At4g31000 Calmodulin-binding protein 8.6 5.1     At4g23180 Ser-Thr kinase related 8.3 5.5     At2g01890 Putative purple acid phosphatase 8.3 1.8     At4g38550 Phospholipase-like protein 7.9 3.4 Transcription     At4g31800 WRKY family transcription factor 11 3.8     At2g38470 WRKY family transcription factor 8.9 4.2     At2g02450 No apical meristem (NAM) family protein 8.2 2.8 Unknown     At1g22890 Expressed protein 31.9 14.9     At1g78460 Expressed protein 25.2 9.2     At2g28570 Expressed protein 20.6 3.2     At3g28290 Integrin-related protein 14a 18.9 91.1     At5g20230 Plastocyanin-like domain-containing protein 14.9 9.1     At3g26200 Cytochrome P450-like 13.6 10.1     At2g05540 Gly-rich protein 11.2 80.2     At1g56150 Auxin-responsive family protein 10.5 6.1     At3g11600 Expressed protein 9.5 7.8     At5g05440 Expressed protein 9.2 2.3     At2g20950 Expressed protein 8.8 3.3     At4g36500 Expressed protein 7.9 3.4     At1g33600 Leu-rich-repeat family protein 7.9 8.4     At1g29430 Auxin-responsive family protein 7.8 9.6     At5g44020 Similar to soybean stem glycoprotein 7.4 36.2 Putative Functional Category and Locus . Description of Encoded Protein . Expression Ratioa (Top) . Expression Ratioa (Base) . Lipid Metabolism     At1g07720 β-Ketoacyl-CoA synthase family 9.9 2.7     At1g64400 Long-chain-acyl-CoA synthetase (LACS3) 9.4 1.7     At2g39400 Similar to lipase 13.4 4.8 Transport     At5g40780 Putative Lys and His transporter 16.9 3.3     At3g55130 ABC transporter family 12.3 3.1     At5g55930 Oligopeptide transporter OPT family protein 8.2 9.0 Cell Wall Metabolism     At4g30280 Putative xyloglucan:xyloglucosyl transferase 33.7 27.9     At4g02330 Similar to pectinesterase 29.7 211     At3g10720 Similar to pectinesterase 8.8 57.4 Disease Resistance     At1g78860 Lectin family protein (mannose-binding) 33.8 19     At2g33050 Similar to disease resistance protein Hcr2 10.2 4.4     At1g75040 Pathogenesis-related protein 5 (PR-5) 10.0 1.0 Signal Transduction     At1g21270 Wall-associated kinase 2 18.2 2.9     At5g20050 Protein kinase family protein 10.0 5.9     At1g11350 Ser-Thr kinase related 9.9 7.1     At1g21250 Wall-associated kinase 1 9.9 1.1     At2g13790 Leu-rich-repeat protein kinase family 9.3 3.4     At1g51805 Leu-rich-repeat protein kinase family 8.6 3.4     At4g31000 Calmodulin-binding protein 8.6 5.1     At4g23180 Ser-Thr kinase related 8.3 5.5     At2g01890 Putative purple acid phosphatase 8.3 1.8     At4g38550 Phospholipase-like protein 7.9 3.4 Transcription     At4g31800 WRKY family transcription factor 11 3.8     At2g38470 WRKY family transcription factor 8.9 4.2     At2g02450 No apical meristem (NAM) family protein 8.2 2.8 Unknown     At1g22890 Expressed protein 31.9 14.9     At1g78460 Expressed protein 25.2 9.2     At2g28570 Expressed protein 20.6 3.2     At3g28290 Integrin-related protein 14a 18.9 91.1     At5g20230 Plastocyanin-like domain-containing protein 14.9 9.1     At3g26200 Cytochrome P450-like 13.6 10.1     At2g05540 Gly-rich protein 11.2 80.2     At1g56150 Auxin-responsive family protein 10.5 6.1     At3g11600 Expressed protein 9.5 7.8     At5g05440 Expressed protein 9.2 2.3     At2g20950 Expressed protein 8.8 3.3     At4g36500 Expressed protein 7.9 3.4     At1g33600 Leu-rich-repeat family protein 7.9 8.4     At1g29430 Auxin-responsive family protein 7.8 9.6     At5g44020 Similar to soybean stem glycoprotein 7.4 36.2 a Mean ratio of epidermis-to-stem gene expression. Gene expressions were determined as in Table I. Among genes up-regulated in top stem epidermis, only those showing a signal intensity >1,000 in top epidermis are shown. Descriptions of proteins are from The Institute for Genomic Research, Munich Information Center for Protein Sequences, or Swiss-Prot. Open in new tab as an example. Among these, several encode proteins belonging to expected categories, such as enzymes of cuticle biosynthesis or defense against pathogens. Interestingly, almost one-third of the proteins have putative regulatory functions, such as the seven putative protein kinases, a group highly represented in the epidermis (Fig. 6 Figure 6. Open in new tabDownload slide Transcripts up-regulated in stem epidermis are enriched in sequences encoding proteins from several functional categories. For each of three sets of transcript sequences (epidermis expressed, epidermis up-regulated, and present on the ATH1 GeneChip), the figure presents the percent of corresponding predicted proteins that are assigned to a selected functional category. Percentages refer to a functional category within each of four types of annotation. Functional categories extracellular, cell wall, and membranes refer to the type of annotation: cellular components from Gene Ontology (GO); secreted, targeting signals predicted by targetP software (http://www.cbs.dtu.dk/services/TargetP); kinase activity, molecular functions from GO; responses to stress/stimulus and lipid metabolism, biological processes from GO to which lipid metabolism was added. Functional categorization was done using annotations from The Arabidopsis Information Resource (http://www.arabidopsis.org) and the Arabidopsis Lipid Gene Database (Beisson et al., 2003). Expressed genes, called present in epidermis of stem tops and stem bottoms by Affymetrix MAS 5.0 software; upregulated genes, show an epidermis-to-stem gene expression ratio of ≥2.0 in stem tops and stem bases (90% confidence level). The complete charts showing all functional categories for each type of annotation are available from Supplemental Figure 2. Figure 6. Open in new tabDownload slide Transcripts up-regulated in stem epidermis are enriched in sequences encoding proteins from several functional categories. For each of three sets of transcript sequences (epidermis expressed, epidermis up-regulated, and present on the ATH1 GeneChip), the figure presents the percent of corresponding predicted proteins that are assigned to a selected functional category. Percentages refer to a functional category within each of four types of annotation. Functional categories extracellular, cell wall, and membranes refer to the type of annotation: cellular components from Gene Ontology (GO); secreted, targeting signals predicted by targetP software (http://www.cbs.dtu.dk/services/TargetP); kinase activity, molecular functions from GO; responses to stress/stimulus and lipid metabolism, biological processes from GO to which lipid metabolism was added. Functional categorization was done using annotations from The Arabidopsis Information Resource (http://www.arabidopsis.org) and the Arabidopsis Lipid Gene Database (Beisson et al., 2003). Expressed genes, called present in epidermis of stem tops and stem bottoms by Affymetrix MAS 5.0 software; upregulated genes, show an epidermis-to-stem gene expression ratio of ≥2.0 in stem tops and stem bases (90% confidence level). The complete charts showing all functional categories for each type of annotation are available from Supplemental Figure 2. ). This list is thus likely to yield good candidates for the cellular signaling pathways involved in the division, differentiation, or elongation of apical epidermal cells. Another important group representing about 40% of the total in Table II is proteins from families completely uncharacterized or whose function is so far unclear. The identification of these proteins as epidermis up-regulated should provide helpful clues in the determination of their function in the cell. Finally, the list contains a few proteins related to cell wall metabolism that could indicate the existence in the epidermis of a specific composition or structure of the external cell wall to which the cuticle may be anchored and through which surface lipids must be transported during cuticle formation. The Epidermis Up-Regulated Set of Genes Is Enriched in Candidates Encoding Proteins Predicted to Be Membrane Associated, Extracellular, or Related to Stress/Stimulus or Lipid Metabolism The subcellular location of the synthesis of waxes and cutin monomers and the mechanism of secretion are mostly unknown. However, due to the hydrophobicity of these molecules, it is likely to involve membrane-associated enzymes and (at least in the case of waxes) transporters like CER5 (Pighin et al., 2004). Also, the mechanism by which waxes and cutin monomers, oligomers, or polymers reach their final destination through the cell wall is likely to involve extracellular proteins. Assembly of cutin monomers or oligomers might even take place outside the cell. For all these reasons, the epidermis up-regulated genes encoding proteins predicted to be (1) membrane associated, (2) secreted (especially extracellular), (3) related to transport, and/or (4) related to lipid metabolism are groups likely to contain strong candidates for the metabolism of surface lipids. A clear enrichment in the genes of three of four of these categories was in fact observed in the set up-regulated in the epidermis of the elongating top part of the stem, which is consistent with the lipid secretory function of epidermal cells. The enrichments observed in proteins predicted to be membrane associated, extracellular, secreted, and lipid related were between 1.5- and 3-fold (Fig. 6). A clear enrichment (1.5- to 2-fold) in proteins likely to play a role in the interaction between the plant epidermis and the environment, such as proteins involved in response to abiotic/biotic stimulus and response to stress, was also observed (Fig. 6). A Subset of 85 Genes Related to Lipid Metabolism and Up-Regulated in the Epidermis Provided Strong Candidates for Wax and Cutin Synthesis Among the 620 genes known or thought to be involved in acyl lipid metabolism in Arabidopsis (Beisson et al., 2003), a subset of about 85 was found to be up-regulated in the epidermis of the top stem (Table III Table III. Genes known or suspected of being involved in acyl lipid metabolism that are up-regulated in the epidermis of top stems Putative or Known Function of Protein . Locus Code . Ratioa (Top) . Ratioa (Base) . Signalb (Top) . Signalb (Base) . Acyl-ACP thioesterase FatB At1g08510 2.4 1.4 9,644 6,072 Stearoyl-ACP desaturase At5g16240 5.5 2.1 810 853 Acyl-CoA desaturase-like At1g06350 3.1 41.3 11,859 1,360 Dihydroxyacetone phosphate reductase At2g41540 5.1 11.3 3,889 787 Plastidial phosphatidic acid phosphatase At2g01180 2.4 6.4 1,198 2,054 Choline kinase At1g71697 4.2 2.9 770 1,011 Fatty acid alcohol oxidase At3g23410 2.8 2.2 718 1,260 Monoacylglycerol lipase At2g39400 13.4 4.8 4,426 5,573 Monoacylglycerol lipase At2g47630 6 3.8 282 37 Monoacylglycerol lipase At5g11650 4.1 1 535 934 Lipid acylhydrolase-like At5g14930 2.7 4.2 236 337 Cytosolic homomeric acyl-CoA carboxylase At1g36160 4.2 2.3 6,025 2,174 Ketoacyl-CoA synthase At1g07720 9.9 2.7 2,866 2,121 Ketoacyl-CoA synthase At5g04530 8.7 41.3 254 62 Ketoacyl-CoA synthase At1g04220 8.1 3.6 6,599 1,867 Ketoacyl-CoA synthase At2g15090 6.6 2.2 296 740 Ketoacyl-CoA synthase (CER60) At1g25450 5.8 10.9 1,527 699 Ketoacyl-CoA synthase At2g28630 5.5 7.4 3,857 2,420 Ketoacyl-CoA synthase At5g43760 4.8 4.6 9,032 4,370 Ketoacyl-CoA synthase (KCS1) At1g01120 4.7 4.2 21,884 11,052 Ketoacyl-CoA synthase (FIDDLEHEAD) At2g26250 4 4 9,547 3,618 Ketoacyl-CoA synthase (CUT1=CER6) At1g68530 3.1 2.8 19,397 13,537 Ketoacyl-CoA synthase At2g16280 3.1 3.6 9,337 4,767 Ketoacyl-CoA reductase At1g24470 5.6 3.5 1,219 854 Ketoacyl-CoA reductase At1g67730 2.7 2.1 13,441 4,209 Fatty acyl-CoA reductase At3g56700 31.8 2.3 491 4,471 Fatty acyl-CoA reductase (CER4) At4g33790 2.6 4.7 14,665 5,669 WAX2 protein (=YORE-YORE) At5g57800 2.7 2.2 25,019 19,380 CER1 protein At1g02205 2.2 2.2 30,389 18,784 CER2 protein At4g24510 4.1 6 8,305 2,540 Putative oxidoreductase (HOTHEAD) At1g72970 4.1 4.3 1,888 296 Fatty acid ω-hydroxylase (CYP86A2=ATT1) At4g00360 5.4 3.5 8,455 5,994 Fatty acid ω-hydroxylase (CYP86A8=LACERATA) At2g45970 3.1 2.2 529 175 Fatty acid ω-hydroxylase (CYP86A7) At1g63710 2.7 6.5 794 48 Fatty acid ω-hydroxylase (CYP86A4) At1g01600 2.5 18.1 1,979 195 Fatty acid ω-hydroxylase (CYP94B3) At3g48520 3.4 6.8 246 1,484 Acyl-activating enzyme (AAE16) At3g23790 2.9 1.6 380 345 Long-chain acyl-CoA synthetase (LACS3) At1g64400 9.5 1.7 5,753 3,511 Long-chain acyl-CoA synthetase (LACS1) At2g47240 4 8.6 8,888 2,647 Long-chain acyl-CoA synthetase (LACS2) At1g49430 2.5 1.2 2,521 398 Glycerol-3-P acyltransferase (GPAT8) At4g00400 4 9.5 4,352 1,494 Glycerol-3-P acyltransferase (GPAT4) At1g01610 3.5 12.3 6,183 1,568 Glycerol-3-P acyltransferase (GPAT2) At1g02390 2.5 5.9 118 366 1-Acylglycerol-P acyltransferase (LPAT5) At3g18850 2.3 2.2 6,123 1,214 Bifunctional wax ester synthase/DAGAT At3g49210 7.8 3.1 805 304 Bifunctional wax ester synthase/DAGAT At5g37300 4.2 16.6 6,431 1,258 Bifunctional wax ester synthase/DAGAT At5g12420 3.6 11.2 123 537 Bifunctional wax ester synthase/DAGAT At1g72110 2.9 12 1,960 83 Wax synthase At3g51970 4.8 2.8 116 121 ABC transporter (WBC12=CER5) At1g51500 3.5 3.1 14,194 4,110 ABC transporter (WBC1) At2g39350 2.3 14.4 140 162 ABC transporter (WBC11) At1g17840 3.6 4.2 10,137 4,644 ABC transporter (WBC18) At3g55110 3.0 2.3 2,915 686 ABC transporter (WBC19) At3g55130 12.3 3.1 3,172 4,884 Translocase At1g72700 2.1 1.3 1,240 1,185 Translocase At1g13210 6.9 4.1 1,810 1,815 Lipid transfer protein type 1 (LTP2) At2g38530 19.5 24.5 904 827 Lipid transfer protein type 1 (LTP8) At2g15050 4.3 3.4 18,270 10,531 Lipid transfer protein type 1 (LTP5) At3g51600 2.8 3.6 28,335 19,619 Lipid transfer protein type 5 At1g27950 3.8 8.7 11,827 4,481 Lipid transfer protein type 5 At3g43720 3.2 4.8 13,127 6,167 Lipid transfer protein type 5 At1g55260 2.6 10.6 7,506 1,076 Lipid transfer protein type 5 At1g62790 2.5 1 4,199 2,033 Diacylglycerol kinase At4g30340 2.9 1.4 1,929 704 DAD1-like acylhydrolase At1g06800 4.6 6 297 203 DAD1-like acylhydrolase At2g30550 4.1 0.8 2,585 1,462 Plastidial lipoxygenase At1g72520 10.1 13.4 681 1,925 Plastidial lipoxygenase At1g67560 2.0 1.8 2,333 1,541 Plastidial lipoxygenase At1g17420 6.6 10 265 1,668 Cytosolic lipoxygenase At1g55020 3.6 1.2 511 345 Oxo-phytodienoic acid reductase At2g06050 5 1.9 2,015 5,873 Oxo-phytodienoic acid reductase At1g76690 2.6 1.8 1,451 2,094 Oxo-phytodienoic acid reductase At1g17990 2.3 1.4 1,801 3,781 Type II phosphoinositide 5-phosphatase At1g05630 3.3 2.1 763 212 Patatin-like acyl-hydrolase At3g63200 7.2 2 1,280 1,191 Patatin-like acyl-hydrolase At3g54950 5.1 0.6 258 309 Phosphatidylinositol-4-kinase-γ At1g26270 3.6 1.5 1,246 1,114 Phosphatidylinositol phosphate kinase type I/II A At4g01190 3.2 1.8 195 140 Phosphatidylinositol phosphate kinase type III At3g14270 3.5 2.1 1,181 1,452 Phosphatidylinositol phosphate kinase type III At1g34260 2.5 1.2 498 607 PI-specific phospholipase C At5g58670 3.3 1.2 417 1,018 Phospholipase D-γ At4g11830 2.7 2.0 931 727 Phospholipase D-γ At4g11850 2.2 2.5 109 216 Lysophospholipase At1g64670 4.0 10.6 2,110 495 Lysophospholipase At5g17780 2.0 1.7 141 92 PPT1-like thioesterase At4g17480 10.4 0.9 200 12 PPT1-like thioesterase At4g17440 36.9 184.4 136 518 PPT1-like thioesterase At5g47330 8.8 1.5 838 973 Putative or Known Function of Protein . Locus Code . Ratioa (Top) . Ratioa (Base) . Signalb (Top) . Signalb (Base) . Acyl-ACP thioesterase FatB At1g08510 2.4 1.4 9,644 6,072 Stearoyl-ACP desaturase At5g16240 5.5 2.1 810 853 Acyl-CoA desaturase-like At1g06350 3.1 41.3 11,859 1,360 Dihydroxyacetone phosphate reductase At2g41540 5.1 11.3 3,889 787 Plastidial phosphatidic acid phosphatase At2g01180 2.4 6.4 1,198 2,054 Choline kinase At1g71697 4.2 2.9 770 1,011 Fatty acid alcohol oxidase At3g23410 2.8 2.2 718 1,260 Monoacylglycerol lipase At2g39400 13.4 4.8 4,426 5,573 Monoacylglycerol lipase At2g47630 6 3.8 282 37 Monoacylglycerol lipase At5g11650 4.1 1 535 934 Lipid acylhydrolase-like At5g14930 2.7 4.2 236 337 Cytosolic homomeric acyl-CoA carboxylase At1g36160 4.2 2.3 6,025 2,174 Ketoacyl-CoA synthase At1g07720 9.9 2.7 2,866 2,121 Ketoacyl-CoA synthase At5g04530 8.7 41.3 254 62 Ketoacyl-CoA synthase At1g04220 8.1 3.6 6,599 1,867 Ketoacyl-CoA synthase At2g15090 6.6 2.2 296 740 Ketoacyl-CoA synthase (CER60) At1g25450 5.8 10.9 1,527 699 Ketoacyl-CoA synthase At2g28630 5.5 7.4 3,857 2,420 Ketoacyl-CoA synthase At5g43760 4.8 4.6 9,032 4,370 Ketoacyl-CoA synthase (KCS1) At1g01120 4.7 4.2 21,884 11,052 Ketoacyl-CoA synthase (FIDDLEHEAD) At2g26250 4 4 9,547 3,618 Ketoacyl-CoA synthase (CUT1=CER6) At1g68530 3.1 2.8 19,397 13,537 Ketoacyl-CoA synthase At2g16280 3.1 3.6 9,337 4,767 Ketoacyl-CoA reductase At1g24470 5.6 3.5 1,219 854 Ketoacyl-CoA reductase At1g67730 2.7 2.1 13,441 4,209 Fatty acyl-CoA reductase At3g56700 31.8 2.3 491 4,471 Fatty acyl-CoA reductase (CER4) At4g33790 2.6 4.7 14,665 5,669 WAX2 protein (=YORE-YORE) At5g57800 2.7 2.2 25,019 19,380 CER1 protein At1g02205 2.2 2.2 30,389 18,784 CER2 protein At4g24510 4.1 6 8,305 2,540 Putative oxidoreductase (HOTHEAD) At1g72970 4.1 4.3 1,888 296 Fatty acid ω-hydroxylase (CYP86A2=ATT1) At4g00360 5.4 3.5 8,455 5,994 Fatty acid ω-hydroxylase (CYP86A8=LACERATA) At2g45970 3.1 2.2 529 175 Fatty acid ω-hydroxylase (CYP86A7) At1g63710 2.7 6.5 794 48 Fatty acid ω-hydroxylase (CYP86A4) At1g01600 2.5 18.1 1,979 195 Fatty acid ω-hydroxylase (CYP94B3) At3g48520 3.4 6.8 246 1,484 Acyl-activating enzyme (AAE16) At3g23790 2.9 1.6 380 345 Long-chain acyl-CoA synthetase (LACS3) At1g64400 9.5 1.7 5,753 3,511 Long-chain acyl-CoA synthetase (LACS1) At2g47240 4 8.6 8,888 2,647 Long-chain acyl-CoA synthetase (LACS2) At1g49430 2.5 1.2 2,521 398 Glycerol-3-P acyltransferase (GPAT8) At4g00400 4 9.5 4,352 1,494 Glycerol-3-P acyltransferase (GPAT4) At1g01610 3.5 12.3 6,183 1,568 Glycerol-3-P acyltransferase (GPAT2) At1g02390 2.5 5.9 118 366 1-Acylglycerol-P acyltransferase (LPAT5) At3g18850 2.3 2.2 6,123 1,214 Bifunctional wax ester synthase/DAGAT At3g49210 7.8 3.1 805 304 Bifunctional wax ester synthase/DAGAT At5g37300 4.2 16.6 6,431 1,258 Bifunctional wax ester synthase/DAGAT At5g12420 3.6 11.2 123 537 Bifunctional wax ester synthase/DAGAT At1g72110 2.9 12 1,960 83 Wax synthase At3g51970 4.8 2.8 116 121 ABC transporter (WBC12=CER5) At1g51500 3.5 3.1 14,194 4,110 ABC transporter (WBC1) At2g39350 2.3 14.4 140 162 ABC transporter (WBC11) At1g17840 3.6 4.2 10,137 4,644 ABC transporter (WBC18) At3g55110 3.0 2.3 2,915 686 ABC transporter (WBC19) At3g55130 12.3 3.1 3,172 4,884 Translocase At1g72700 2.1 1.3 1,240 1,185 Translocase At1g13210 6.9 4.1 1,810 1,815 Lipid transfer protein type 1 (LTP2) At2g38530 19.5 24.5 904 827 Lipid transfer protein type 1 (LTP8) At2g15050 4.3 3.4 18,270 10,531 Lipid transfer protein type 1 (LTP5) At3g51600 2.8 3.6 28,335 19,619 Lipid transfer protein type 5 At1g27950 3.8 8.7 11,827 4,481 Lipid transfer protein type 5 At3g43720 3.2 4.8 13,127 6,167 Lipid transfer protein type 5 At1g55260 2.6 10.6 7,506 1,076 Lipid transfer protein type 5 At1g62790 2.5 1 4,199 2,033 Diacylglycerol kinase At4g30340 2.9 1.4 1,929 704 DAD1-like acylhydrolase At1g06800 4.6 6 297 203 DAD1-like acylhydrolase At2g30550 4.1 0.8 2,585 1,462 Plastidial lipoxygenase At1g72520 10.1 13.4 681 1,925 Plastidial lipoxygenase At1g67560 2.0 1.8 2,333 1,541 Plastidial lipoxygenase At1g17420 6.6 10 265 1,668 Cytosolic lipoxygenase At1g55020 3.6 1.2 511 345 Oxo-phytodienoic acid reductase At2g06050 5 1.9 2,015 5,873 Oxo-phytodienoic acid reductase At1g76690 2.6 1.8 1,451 2,094 Oxo-phytodienoic acid reductase At1g17990 2.3 1.4 1,801 3,781 Type II phosphoinositide 5-phosphatase At1g05630 3.3 2.1 763 212 Patatin-like acyl-hydrolase At3g63200 7.2 2 1,280 1,191 Patatin-like acyl-hydrolase At3g54950 5.1 0.6 258 309 Phosphatidylinositol-4-kinase-γ At1g26270 3.6 1.5 1,246 1,114 Phosphatidylinositol phosphate kinase type I/II A At4g01190 3.2 1.8 195 140 Phosphatidylinositol phosphate kinase type III At3g14270 3.5 2.1 1,181 1,452 Phosphatidylinositol phosphate kinase type III At1g34260 2.5 1.2 498 607 PI-specific phospholipase C At5g58670 3.3 1.2 417 1,018 Phospholipase D-γ At4g11830 2.7 2.0 931 727 Phospholipase D-γ At4g11850 2.2 2.5 109 216 Lysophospholipase At1g64670 4.0 10.6 2,110 495 Lysophospholipase At5g17780 2.0 1.7 141 92 PPT1-like thioesterase At4g17480 10.4 0.9 200 12 PPT1-like thioesterase At4g17440 36.9 184.4 136 518 PPT1-like thioesterase At5g47330 8.8 1.5 838 973 a Mean epidermis-to-stem gene expression ratio obtained with the top or base of stems. Gene expressions were determined as in Table I. b Mean signal intensities are indicated to give an estimate of relative expression levels of the genes in the epidermis of top stems as well as in the epidermis of the base of stems. Open in new tab Table III. Genes known or suspected of being involved in acyl lipid metabolism that are up-regulated in the epidermis of top stems Putative or Known Function of Protein . Locus Code . Ratioa (Top) . Ratioa (Base) . Signalb (Top) . Signalb (Base) . Acyl-ACP thioesterase FatB At1g08510 2.4 1.4 9,644 6,072 Stearoyl-ACP desaturase At5g16240 5.5 2.1 810 853 Acyl-CoA desaturase-like At1g06350 3.1 41.3 11,859 1,360 Dihydroxyacetone phosphate reductase At2g41540 5.1 11.3 3,889 787 Plastidial phosphatidic acid phosphatase At2g01180 2.4 6.4 1,198 2,054 Choline kinase At1g71697 4.2 2.9 770 1,011 Fatty acid alcohol oxidase At3g23410 2.8 2.2 718 1,260 Monoacylglycerol lipase At2g39400 13.4 4.8 4,426 5,573 Monoacylglycerol lipase At2g47630 6 3.8 282 37 Monoacylglycerol lipase At5g11650 4.1 1 535 934 Lipid acylhydrolase-like At5g14930 2.7 4.2 236 337 Cytosolic homomeric acyl-CoA carboxylase At1g36160 4.2 2.3 6,025 2,174 Ketoacyl-CoA synthase At1g07720 9.9 2.7 2,866 2,121 Ketoacyl-CoA synthase At5g04530 8.7 41.3 254 62 Ketoacyl-CoA synthase At1g04220 8.1 3.6 6,599 1,867 Ketoacyl-CoA synthase At2g15090 6.6 2.2 296 740 Ketoacyl-CoA synthase (CER60) At1g25450 5.8 10.9 1,527 699 Ketoacyl-CoA synthase At2g28630 5.5 7.4 3,857 2,420 Ketoacyl-CoA synthase At5g43760 4.8 4.6 9,032 4,370 Ketoacyl-CoA synthase (KCS1) At1g01120 4.7 4.2 21,884 11,052 Ketoacyl-CoA synthase (FIDDLEHEAD) At2g26250 4 4 9,547 3,618 Ketoacyl-CoA synthase (CUT1=CER6) At1g68530 3.1 2.8 19,397 13,537 Ketoacyl-CoA synthase At2g16280 3.1 3.6 9,337 4,767 Ketoacyl-CoA reductase At1g24470 5.6 3.5 1,219 854 Ketoacyl-CoA reductase At1g67730 2.7 2.1 13,441 4,209 Fatty acyl-CoA reductase At3g56700 31.8 2.3 491 4,471 Fatty acyl-CoA reductase (CER4) At4g33790 2.6 4.7 14,665 5,669 WAX2 protein (=YORE-YORE) At5g57800 2.7 2.2 25,019 19,380 CER1 protein At1g02205 2.2 2.2 30,389 18,784 CER2 protein At4g24510 4.1 6 8,305 2,540 Putative oxidoreductase (HOTHEAD) At1g72970 4.1 4.3 1,888 296 Fatty acid ω-hydroxylase (CYP86A2=ATT1) At4g00360 5.4 3.5 8,455 5,994 Fatty acid ω-hydroxylase (CYP86A8=LACERATA) At2g45970 3.1 2.2 529 175 Fatty acid ω-hydroxylase (CYP86A7) At1g63710 2.7 6.5 794 48 Fatty acid ω-hydroxylase (CYP86A4) At1g01600 2.5 18.1 1,979 195 Fatty acid ω-hydroxylase (CYP94B3) At3g48520 3.4 6.8 246 1,484 Acyl-activating enzyme (AAE16) At3g23790 2.9 1.6 380 345 Long-chain acyl-CoA synthetase (LACS3) At1g64400 9.5 1.7 5,753 3,511 Long-chain acyl-CoA synthetase (LACS1) At2g47240 4 8.6 8,888 2,647 Long-chain acyl-CoA synthetase (LACS2) At1g49430 2.5 1.2 2,521 398 Glycerol-3-P acyltransferase (GPAT8) At4g00400 4 9.5 4,352 1,494 Glycerol-3-P acyltransferase (GPAT4) At1g01610 3.5 12.3 6,183 1,568 Glycerol-3-P acyltransferase (GPAT2) At1g02390 2.5 5.9 118 366 1-Acylglycerol-P acyltransferase (LPAT5) At3g18850 2.3 2.2 6,123 1,214 Bifunctional wax ester synthase/DAGAT At3g49210 7.8 3.1 805 304 Bifunctional wax ester synthase/DAGAT At5g37300 4.2 16.6 6,431 1,258 Bifunctional wax ester synthase/DAGAT At5g12420 3.6 11.2 123 537 Bifunctional wax ester synthase/DAGAT At1g72110 2.9 12 1,960 83 Wax synthase At3g51970 4.8 2.8 116 121 ABC transporter (WBC12=CER5) At1g51500 3.5 3.1 14,194 4,110 ABC transporter (WBC1) At2g39350 2.3 14.4 140 162 ABC transporter (WBC11) At1g17840 3.6 4.2 10,137 4,644 ABC transporter (WBC18) At3g55110 3.0 2.3 2,915 686 ABC transporter (WBC19) At3g55130 12.3 3.1 3,172 4,884 Translocase At1g72700 2.1 1.3 1,240 1,185 Translocase At1g13210 6.9 4.1 1,810 1,815 Lipid transfer protein type 1 (LTP2) At2g38530 19.5 24.5 904 827 Lipid transfer protein type 1 (LTP8) At2g15050 4.3 3.4 18,270 10,531 Lipid transfer protein type 1 (LTP5) At3g51600 2.8 3.6 28,335 19,619 Lipid transfer protein type 5 At1g27950 3.8 8.7 11,827 4,481 Lipid transfer protein type 5 At3g43720 3.2 4.8 13,127 6,167 Lipid transfer protein type 5 At1g55260 2.6 10.6 7,506 1,076 Lipid transfer protein type 5 At1g62790 2.5 1 4,199 2,033 Diacylglycerol kinase At4g30340 2.9 1.4 1,929 704 DAD1-like acylhydrolase At1g06800 4.6 6 297 203 DAD1-like acylhydrolase At2g30550 4.1 0.8 2,585 1,462 Plastidial lipoxygenase At1g72520 10.1 13.4 681 1,925 Plastidial lipoxygenase At1g67560 2.0 1.8 2,333 1,541 Plastidial lipoxygenase At1g17420 6.6 10 265 1,668 Cytosolic lipoxygenase At1g55020 3.6 1.2 511 345 Oxo-phytodienoic acid reductase At2g06050 5 1.9 2,015 5,873 Oxo-phytodienoic acid reductase At1g76690 2.6 1.8 1,451 2,094 Oxo-phytodienoic acid reductase At1g17990 2.3 1.4 1,801 3,781 Type II phosphoinositide 5-phosphatase At1g05630 3.3 2.1 763 212 Patatin-like acyl-hydrolase At3g63200 7.2 2 1,280 1,191 Patatin-like acyl-hydrolase At3g54950 5.1 0.6 258 309 Phosphatidylinositol-4-kinase-γ At1g26270 3.6 1.5 1,246 1,114 Phosphatidylinositol phosphate kinase type I/II A At4g01190 3.2 1.8 195 140 Phosphatidylinositol phosphate kinase type III At3g14270 3.5 2.1 1,181 1,452 Phosphatidylinositol phosphate kinase type III At1g34260 2.5 1.2 498 607 PI-specific phospholipase C At5g58670 3.3 1.2 417 1,018 Phospholipase D-γ At4g11830 2.7 2.0 931 727 Phospholipase D-γ At4g11850 2.2 2.5 109 216 Lysophospholipase At1g64670 4.0 10.6 2,110 495 Lysophospholipase At5g17780 2.0 1.7 141 92 PPT1-like thioesterase At4g17480 10.4 0.9 200 12 PPT1-like thioesterase At4g17440 36.9 184.4 136 518 PPT1-like thioesterase At5g47330 8.8 1.5 838 973 Putative or Known Function of Protein . Locus Code . Ratioa (Top) . Ratioa (Base) . Signalb (Top) . Signalb (Base) . Acyl-ACP thioesterase FatB At1g08510 2.4 1.4 9,644 6,072 Stearoyl-ACP desaturase At5g16240 5.5 2.1 810 853 Acyl-CoA desaturase-like At1g06350 3.1 41.3 11,859 1,360 Dihydroxyacetone phosphate reductase At2g41540 5.1 11.3 3,889 787 Plastidial phosphatidic acid phosphatase At2g01180 2.4 6.4 1,198 2,054 Choline kinase At1g71697 4.2 2.9 770 1,011 Fatty acid alcohol oxidase At3g23410 2.8 2.2 718 1,260 Monoacylglycerol lipase At2g39400 13.4 4.8 4,426 5,573 Monoacylglycerol lipase At2g47630 6 3.8 282 37 Monoacylglycerol lipase At5g11650 4.1 1 535 934 Lipid acylhydrolase-like At5g14930 2.7 4.2 236 337 Cytosolic homomeric acyl-CoA carboxylase At1g36160 4.2 2.3 6,025 2,174 Ketoacyl-CoA synthase At1g07720 9.9 2.7 2,866 2,121 Ketoacyl-CoA synthase At5g04530 8.7 41.3 254 62 Ketoacyl-CoA synthase At1g04220 8.1 3.6 6,599 1,867 Ketoacyl-CoA synthase At2g15090 6.6 2.2 296 740 Ketoacyl-CoA synthase (CER60) At1g25450 5.8 10.9 1,527 699 Ketoacyl-CoA synthase At2g28630 5.5 7.4 3,857 2,420 Ketoacyl-CoA synthase At5g43760 4.8 4.6 9,032 4,370 Ketoacyl-CoA synthase (KCS1) At1g01120 4.7 4.2 21,884 11,052 Ketoacyl-CoA synthase (FIDDLEHEAD) At2g26250 4 4 9,547 3,618 Ketoacyl-CoA synthase (CUT1=CER6) At1g68530 3.1 2.8 19,397 13,537 Ketoacyl-CoA synthase At2g16280 3.1 3.6 9,337 4,767 Ketoacyl-CoA reductase At1g24470 5.6 3.5 1,219 854 Ketoacyl-CoA reductase At1g67730 2.7 2.1 13,441 4,209 Fatty acyl-CoA reductase At3g56700 31.8 2.3 491 4,471 Fatty acyl-CoA reductase (CER4) At4g33790 2.6 4.7 14,665 5,669 WAX2 protein (=YORE-YORE) At5g57800 2.7 2.2 25,019 19,380 CER1 protein At1g02205 2.2 2.2 30,389 18,784 CER2 protein At4g24510 4.1 6 8,305 2,540 Putative oxidoreductase (HOTHEAD) At1g72970 4.1 4.3 1,888 296 Fatty acid ω-hydroxylase (CYP86A2=ATT1) At4g00360 5.4 3.5 8,455 5,994 Fatty acid ω-hydroxylase (CYP86A8=LACERATA) At2g45970 3.1 2.2 529 175 Fatty acid ω-hydroxylase (CYP86A7) At1g63710 2.7 6.5 794 48 Fatty acid ω-hydroxylase (CYP86A4) At1g01600 2.5 18.1 1,979 195 Fatty acid ω-hydroxylase (CYP94B3) At3g48520 3.4 6.8 246 1,484 Acyl-activating enzyme (AAE16) At3g23790 2.9 1.6 380 345 Long-chain acyl-CoA synthetase (LACS3) At1g64400 9.5 1.7 5,753 3,511 Long-chain acyl-CoA synthetase (LACS1) At2g47240 4 8.6 8,888 2,647 Long-chain acyl-CoA synthetase (LACS2) At1g49430 2.5 1.2 2,521 398 Glycerol-3-P acyltransferase (GPAT8) At4g00400 4 9.5 4,352 1,494 Glycerol-3-P acyltransferase (GPAT4) At1g01610 3.5 12.3 6,183 1,568 Glycerol-3-P acyltransferase (GPAT2) At1g02390 2.5 5.9 118 366 1-Acylglycerol-P acyltransferase (LPAT5) At3g18850 2.3 2.2 6,123 1,214 Bifunctional wax ester synthase/DAGAT At3g49210 7.8 3.1 805 304 Bifunctional wax ester synthase/DAGAT At5g37300 4.2 16.6 6,431 1,258 Bifunctional wax ester synthase/DAGAT At5g12420 3.6 11.2 123 537 Bifunctional wax ester synthase/DAGAT At1g72110 2.9 12 1,960 83 Wax synthase At3g51970 4.8 2.8 116 121 ABC transporter (WBC12=CER5) At1g51500 3.5 3.1 14,194 4,110 ABC transporter (WBC1) At2g39350 2.3 14.4 140 162 ABC transporter (WBC11) At1g17840 3.6 4.2 10,137 4,644 ABC transporter (WBC18) At3g55110 3.0 2.3 2,915 686 ABC transporter (WBC19) At3g55130 12.3 3.1 3,172 4,884 Translocase At1g72700 2.1 1.3 1,240 1,185 Translocase At1g13210 6.9 4.1 1,810 1,815 Lipid transfer protein type 1 (LTP2) At2g38530 19.5 24.5 904 827 Lipid transfer protein type 1 (LTP8) At2g15050 4.3 3.4 18,270 10,531 Lipid transfer protein type 1 (LTP5) At3g51600 2.8 3.6 28,335 19,619 Lipid transfer protein type 5 At1g27950 3.8 8.7 11,827 4,481 Lipid transfer protein type 5 At3g43720 3.2 4.8 13,127 6,167 Lipid transfer protein type 5 At1g55260 2.6 10.6 7,506 1,076 Lipid transfer protein type 5 At1g62790 2.5 1 4,199 2,033 Diacylglycerol kinase At4g30340 2.9 1.4 1,929 704 DAD1-like acylhydrolase At1g06800 4.6 6 297 203 DAD1-like acylhydrolase At2g30550 4.1 0.8 2,585 1,462 Plastidial lipoxygenase At1g72520 10.1 13.4 681 1,925 Plastidial lipoxygenase At1g67560 2.0 1.8 2,333 1,541 Plastidial lipoxygenase At1g17420 6.6 10 265 1,668 Cytosolic lipoxygenase At1g55020 3.6 1.2 511 345 Oxo-phytodienoic acid reductase At2g06050 5 1.9 2,015 5,873 Oxo-phytodienoic acid reductase At1g76690 2.6 1.8 1,451 2,094 Oxo-phytodienoic acid reductase At1g17990 2.3 1.4 1,801 3,781 Type II phosphoinositide 5-phosphatase At1g05630 3.3 2.1 763 212 Patatin-like acyl-hydrolase At3g63200 7.2 2 1,280 1,191 Patatin-like acyl-hydrolase At3g54950 5.1 0.6 258 309 Phosphatidylinositol-4-kinase-γ At1g26270 3.6 1.5 1,246 1,114 Phosphatidylinositol phosphate kinase type I/II A At4g01190 3.2 1.8 195 140 Phosphatidylinositol phosphate kinase type III At3g14270 3.5 2.1 1,181 1,452 Phosphatidylinositol phosphate kinase type III At1g34260 2.5 1.2 498 607 PI-specific phospholipase C At5g58670 3.3 1.2 417 1,018 Phospholipase D-γ At4g11830 2.7 2.0 931 727 Phospholipase D-γ At4g11850 2.2 2.5 109 216 Lysophospholipase At1g64670 4.0 10.6 2,110 495 Lysophospholipase At5g17780 2.0 1.7 141 92 PPT1-like thioesterase At4g17480 10.4 0.9 200 12 PPT1-like thioesterase At4g17440 36.9 184.4 136 518 PPT1-like thioesterase At5g47330 8.8 1.5 838 973 a Mean epidermis-to-stem gene expression ratio obtained with the top or base of stems. Gene expressions were determined as in Table I. b Mean signal intensities are indicated to give an estimate of relative expression levels of the genes in the epidermis of top stems as well as in the epidermis of the base of stems. Open in new tab ). Among this subset are several genes previously identified in forward-genetics screens for mutants with altered wax (WAX2, CER5, CER6, etc.) as well as genes reported to be involved in cutin biosynthesis (ATT1, LACS2, FATB). This concordance of the microarray data with numerous results of forward genetics clearly validates this epidermal microarray approach as suitable for identification of additional gene candidates likely to be involved in wax and cutin biosynthesis. For example, the new candidates for wax synthesis include specific members of multigenic families for which some members have already been previously shown to be involved in wax synthesis, e.g. the ketoacyl-CoA synthase (KCS) family. Figure 7 Figure 7. Open in new tabDownload slide Relative gene expression levels of the members of the KCS gene family in epidermis of stem tops and in total stems. Figure 7. Open in new tabDownload slide Relative gene expression levels of the members of the KCS gene family in epidermis of stem tops and in total stems. summarizes the level of gene expression and the epidermis-to-stem gene expression ratios found for the 20 members of the KCS family (out of 21) present on the ATH1 array. It can be clearly seen that there are five uncharacterized KCS genes showing an epidermal up-regulation higher than the CER60 gene, which is known to affect wax synthesis. These genes are thus good candidates for the putative specific elongases thought to be responsible for each of the multiple elongation steps of wax synthesis. In addition, and as expected, the KCS FAE1 isoform (At4g34520) that is responsible for the elongation of fatty acids for storage lipids in seeds is not among the epidermis up-regulated KCS genes. DISCUSSION Using rapidly elongating Arabidopsis stems, we have performed measurements of the elongation of epidermal cells in conjunction with quantitative analyses of the cuticular lipids. In brief, the results indicate that polyester and waxes are deposited in the elongating apical part of the stem at a rate that allows the cuticle to keep a constant load and composition of surface lipids, despite the fast rate of elongation of the epidermal cells. A major metabolic function of these epidermal cells is extracellular lipid synthesis as illustrated by the fact that more lipid is transported out of the cell than remains in membrane glycerolipids (Fig. 3) and by the enriched expression of transcripts for many enzymes of lipid metabolism (Fig. 6). These isoforms preferentially expressed in the epidermis are thus strong candidates for roles in wax and cutin synthesis (Table III). Deposition of Waxes The wax analysis shows that the amounts of individual wax constituent do not vary significantly along the stem (Figs. 2–434) and within the apical stem segment (data not shown). The wax load on the total stem, determined in control experiments, confirms the findings for stem segments. These results for total stem wax match previous reports on the Col ecotype (Pighin et al., 2004), and are dramatically higher than the wax load of 0.75 μg/cm2 on leaves (data not shown). The distribution of waxes on the surface of various stem segments is not likely to be influenced by diffusional transport within the cuticle because, clearly, the polyester components are not mobile and diffusion coefficients measured for fatty acids and alkanes in reconstituted cuticular waxes (Schreiber et al., 1996) imply that lateral wax diffusion will be a very slow process, less than 0.01 mm/d. Because surface area expansion is greatest and unequal in the top segment (Fig. 1), the constant wax load and composition found both along the stem and within the top segment implies that the rate of net deposition of wax is strictly synchronized with epidermal cell expansion and largely limited to the top zone of the stem, i.e. early on during development. All wax constituents are therefore formed at similar rates and deposited at the same time in the expanding epidermal cells. Expression of all proteins involved in wax biosynthesis is likely synchronized and must be highly up-regulated during rapid epidermal cell expansion either transcriptionally or posttranscriptionally. Constant wax loads on the lower segments of the stems might reflect a dynamic equilibrium between wax accumulation and wax loss by erosion or back transport (Jetter and Schaffer, 2001). But if wax turnover occurred, then it would certainly be at much slower rates than the accumulation in the top zone. Deposition of the Polyester Matrix The fact that the highest polyester loads were observed in the youngest part of the stem is consistent with previous observations that the major cutin synthesis occurs in young tissues (Kolattukudy, 1970). Palmitate applied exogenously to broad bean leaves was most highly incorporated into cutin in the youngest leaves, but incorporation was not detectable in the oldest leaves. Assuming that palmitate uptake was not limiting, this implies that cutin synthesis indeed occurs primarily in young organs. The 2-fold polyester reduction seen at the base of the Arabidopsis stems as compared to the middle of the stems (Fig. 3) is not likely to be caused by surface area expansion and dilution of the initial polyester load accumulated in the top zone (i.e. early during the elongation of epidermal cells). Indeed, there is very little or no more elongation in the stem bases (Fig. 1) and, in the zone where cell elongation is maximal (between the apex and the middle of the stems), the average polyester load does not vary significantly (Fig. 2). The extent to which the decrease in polyester load seen in the base is caused by a turnover of the components of the polyester matrix with a slow rate of resynthesis, or more likely by conversion of cutin (which can be depolymerized) to cutan (which cannot be depolymerized), remains to be determined. Monomers containing epoxy groups have been shown to be present only in young leaves in Clivia miniata (Jeffree, 1996) and could thus be involved in cutan formation in older organs. However, due to the difficulty of polyester analysis in Arabidopsis, as well as very low amounts, the putative Arabidopsis epoxy monomers (probably among the C18 in-chain substituted fatty acids in Fig. 4) were not fully identified. Dicarboxylic acids are major polyester monomers in Arabidopsis and no evidence was uncovered that indicated there is a significant difference in timing of their deposition as compared to ω-hydroxy fatty acids. These results are consistent with the idea that dicarboxylic acids are also part of the cutin polymer, or part of an epidermal polymer that is functionally closely related to cutin, and that together they form the matrix of the cuticle. Wax and Polyester Loads The wax load of the leaf is about 2 times lower than the polyester load: 0.75 μg/cm2 (data not shown) versus 1.5 μg/cm2 (this study). Since the average polyester load of the stem is 6 μg/cm2, one would expect about 3 μg/cm2 of waxes to be present on the stem surface (assuming the wax-to-polyester ratio of the leaf is the same). However, a much higher wax load was found in the stem: on average, 32 μg/cm2 (this study). The difference is thus likely to come from the abundant epicuticular waxes of the stem, as evidenced by the wax crystals seen in the ultrastructure of the stem surface (Fig. 2), but not in that of the leaf surface (data not shown). It can therefore be estimated that epicuticular wax crystals represent about 90% of the total wax load in the stem. Finally, it should be stressed that the actual ratio of wax to matrix in the cuticle could be different from the wax-to-polyester ratio because the amount of cuticle lipid that cannot be depolymerized is unknown. Gene Expression in Epidermal Cells About 1,900 genes (15% of the genes detected in the stem) were identified as preferentially expressed in the epidermis of top and/or basal segments of stems. The majority of these genes (70%) had a transcript epidermis-to-stem ratio that fell in the 2.0 to 4.0 range, while 30% had a ratio greater than 4.0 (Supplemental Table II). This is consistent with what has been observed in maize for transcripts accumulating to at least 2-fold higher levels in epidermal cells than in vascular tissues (Nakazono et al., 2003). Moreover, more than 70% of the unique genes (or genes belonging to small gene families) identified as preferentially up-regulated in the maize epidermis had their best Arabidopsis hit (BLASTX search), showing also a clear up-regulation in the epidermis of Arabidopsis stems (data not shown). These include, for example, genes encoding the acyl-ACP thioesterase FatB or the cytosolic acetyl-CoA carboxylase (Table III). We have found that a subset of about 600 genes (about 5% of genes detected in stems) was specifically up-regulated in the epidermis of the elongating top segments (Supplemental Tables I and III). Interestingly, this is similar to the size of the subset of genes (672) that were found to be preferentially expressed in the epidermis of roots at a stage of longitudinal cell elongation and that represented about 6% of the genes detected in roots (Birnbaum et al., 2003). These observations and the correct classification of known epidermis-specific genes (Table I) suggest that many of the genes identified in this study as epidermis up-regulated are true hits and are thus likely to play a role in the biological processes occurring specifically in the epidermis, e.g. the synthesis of the cuticle. The epidermal peel approach used here is based on the harvest of all epidermal cell types and thus transcripts specifically expressed in less abundant cell types, like guard cells, will be diluted by more abundant cell types. The candidate genes obtained here are therefore more likely to be genes expressed in all epidermal cells of a particular stem region. However, the example of the guard cell-specific MYB transcription factor At1g08810 (Cominelli et al., 2005) that is found to be 3.4-fold up-regulated in the epidermis in our dataset (Supplemental Table II) shows that cell type-specific proteins could also be included in the list of candidates. The genes that are up-regulated to the same extent in both the top and the basal epidermis are presumably related to functions or processes specific to the whole epidermis, whereas genes up-regulated only in the top or the basal epidermis are more likely to be involved, for example, in epidermal cell division, elongation, or cuticle formation for the top, and in trichome differentiation for the base. To identify gene candidates for wax and cutin synthesis, we have focused mostly on the epidermis-to-stem gene expression ratio for the top stem. Even if there is a turnover of surface lipids in the epidermal cells of the nonelongating stem base, the rapid elongation and the high wax-polyester content observed in the stem top indicate that this machinery must be more active in the epidermis of the top than in the epidermis of the base. In the case of transcriptional regulation, the transcripts of the corresponding genes are thus expected to be more abundant in the epidermis of the top segment than in the epidermis of the basal segment of the stem. However, it is striking that many genes involved in wax synthesis, such as the major elongase-condensing enzyme (CER6) that supplies alkane and alkane-derived (secondary alcohols and ketones) waxes, are still highly expressed or even up-regulated in the lower stem epidermis (Table III), where we would expect very low net flux into these components. It is possible that there is a turnover of the wax and/or cutin components in the nonelongating parts of the stems, requiring continued synthesis to maintain constant loads. An alternative hypothesis is that many of the genes involved in surface lipid metabolism could be expressed in the epidermis of all stem segments, but strongly controlled at the posttranscriptional level, depending on the elongation rate of the cell. This control might, for example, include a system that senses the wax-cutin loads to provide feedback to biosynthesis. Lipid-related genes of Table III may not all be involved in surface lipid metabolism. Some are clearly related to cellular signaling, possibly playing a role in epidermal cell differentiation, elongation, and/or interaction with the environment. It is not likely that the signaling genes up-regulated in the epidermis are induced by wounding during the harvest of epidermal peels because peels were frozen in liquid nitrogen immediately and also several essential and well characterized genes of wound response (Delessert et al., 2004), such as At4g15440 (hydroperoxide lyase) and At5g42650 (allene oxide synthase), were not found to be significantly up-regulated in the epidermal peels as compared to the nonpeeled stem segments used as a reference (see Supplemental Table II). Candidate Genes for Wax and Cutin Biosynthesis Many reactions in the pathways for the synthesis of wax components remain obscure and some proteins identified by mapping of wax mutants cannot be assigned a clear molecular function (Kunst and Samuels, 2003). The only protein that has been demonstrated to play a specific role in cutin biosynthesis by a chemical analysis of cutin (att1 Arabidopsis mutant) is a fatty acid hydroxylase (Xiao et al., 2004). Important enzymes, like putative acyltransferases, that might be responsible for the assembly of cutin polyester chains are clearly lacking in the current list of candidates (Yephremov and Schreiber, 2005), and therefore more gene candidate searches are needed. Analysis of cutin components is time consuming and not easy to use in a high-throughput screening of a forward-genetics approach, while identification of cutin mutants by indirect assays, such as organ fusion phenotype, chlorophyll leaching, or dye uptake, might yield other mutants not primarily affected in cutin synthesis. The analysis of the polyesters of insertion lines in selected candidate genes might therefore prove useful to discover members of large protein families or new proteins specifically involved in cutin as well as wax metabolism. In addition to the KCS family (Fig. 7), another example of the utility of Table III for large gene families is provided by analysis of the expression of the subfamilies of putative lipid transfer proteins (LTPs) that have been suggested to be involved in wax-polyester monomer transport through the cell wall. In Arabidopsis, there are 72 putative LTPs that were classified into eight types based on the conserved Cys pattern (Beisson et al., 2003). Interestingly, all the LTPs up-regulated in the growing epidermis were found to be from type 1, the original group of LTPs shown to have in vitro lipid-binding properties, and from the large uncharacterized type 5 group. No LTP from the type 3 group was found, although this group is as large (11 genes represented on the ATH1 array) as the type 1 group. These results point toward type 1 and 5 groups as candidates for a function in cuticle synthesis, whereas the type 3 group might be related to other functions, such as signaling in the apoplast as suggested for one LTP of type 3 (Maldonado et al., 2002). Also potentially related to wax secretion, several ABC transporters from the same white-brown complex (WBC) subfamily as CER5 were up-regulated in the epidermis. Our list of candidates for wax synthesis (Table III) includes uncharacterized members of the KCS family, such as At5g16280 or At2g43760, that have been recently identified as highly expressed in the aerial parts of 15-d-old expanding seedlings (Costaglioli et al., 2005). Our results thus confirm that they are likely to play an important role in wax synthesis. But other KCS genes, like At1g04220 or At1g07720, were also found to be clearly up-regulated in stem epidermis, although they are low expressed in 15-d-old seedlings. The same is true for LTP2 (At2g38530), which has a low global expression level when measured in seedlings, but is among the few epidermis up-regulated LTPs identified in the elongating stem. These genes exemplify the strength of our approach using only epidermis material in the discovery of gene candidates for cuticle synthesis. These examples could also indicate that, regarding candidate profiling for cuticle synthesis, it might be difficult to extend the results obtained on young seedlings to older plants and, conversely, because specific isoforms might be expressed in the shoot epidermis at different stages of development. Concerning cutin biosynthesis, putative fatty acid hydroxylases and acyltransferases are proteins of special interest. The protein affected in the Arabidopsis att1 cutin mutant (Xiao et al., 2004) is CYP86A2 (At4g00360 locus), a fatty acid hydroxylase belonging to the same cytochrome P450 monooxygenase subfamily as CYP86A8, which has also been implicated in cuticle formation in the Arabidopsis lacerata mutant (Wellesen et al., 2001). Not surprisingly, there are other members of the same subfamily that are also up-regulated in the epidermis (Table III). The other Arabidopsis cytochrome P450 monooxygenase subfamily thought to encode fatty acid hydroxylases is the CYP94B family (Duan and Schuler, 2005). It is therefore likely that the CYP94B3 member listed in Table III is also related to oxidation of surface lipids or epidermal oxylipins. Since this gene is more expressed and up-regulated in the epidermis of the base of the stem than of the top of the stem, it is, for instance, a good candidate for the polyester modifications (e.g. cutan synthesis) that might occur at the base of the stem. Arabidopsis cytochrome P450 monooxygenases belonging to other subfamilies that are uncharacterized or involved in the oxidation of substrates not related to fatty acids were not listed in Table III. The polyester synthases responsible for the assembly of cutin chains remain completely unknown and the acyltransferases listed in Table III are therefore potential candidates for this function. It is not likely that the epidermis up-regulated acyltransferases listed in Table III (1-acylglycerol-3-P acyltransferase [LPAT5] and glycerol-3-P acyltransferase [GPAT4], etc.) are involved in housekeeping membrane biogenesis because the cells of the tissues underlying the epidermis are also elongating and need to synthesize membrane lipids at about the same rate as epidermal cells. Thus, it is more likely that the acyltransferases of Table III are somehow involved in surface lipid synthesis. Indeed, LPAT activity was not detected for the recombinant LPAT5 (Kim et al., 2005), and no GPAT activity was found for the GPAT2 and GPAT3 isoforms (Zheng et al., 2003). Taken together, these observations raise the possibility that at least some of the GPAT acyltransferases isoforms, as well as LPAT5, act on intermediates of cutin and/or wax biosynthesis and are not involved in the synthesis of the glycerolipids composing the membranes or storage-oil bodies of the cell. Preliminary analysis of T-DNA insertion lines in some of these genes supports this hypothesis (data not shown). CONCLUSION The accumulation of cuticular lipids by the plant epidermis has been studied by a combined quantitative and qualitative analysis of the surface lipids and of the transcripts present in expanding epidermal cells during the elongation of Arabidopsis stems. The high loads and constant composition of polyesters and waxes found in the elongating top and middle zone of stems indicated that the synthesis and the secretion of these two components of the cuticle are highly coregulated to keep up with the pace of rapid epidermal cell expansion. Our microarray approach identified a subset of the genome (about 15% of the genes detected in the stem) that is preferentially expressed in the epidermis and provided a list of potential gene candidates for the machinery of assembly, secretion, and synthesis of surface lipids that can be tested by reverse-genetics approaches. This dataset should accelerate progress toward elucidation of the wax biosynthesis pathways and aid in unraveling the biosynthesis of cutin. MATERIALS AND METHODS Plant Material and Growth Conditions The wild-type ecotype Col-0 of Arabidopsis (Arabidopsis thaliana) was used for all experiments. Seeds were stratified for 3 to 4 d at 4°C and plants were grown on a mixture of soil:vermiculite:perlite (1:1:1) under white fluorescent light (80–100 μE m−2 s−1) in an 18-h-light/6-h-dark photoperiod. The temperature was set at 20°C to 22°C and the relative humidity at 50% to 70%. Stem Harvest and Stem Diameter Measurements Each harvested 10- to 11-cm-long primary stem was cut into 3-cm-long segments (3 cm from the base of the stem, 3 cm from the base of the inflorescence, and 3 cm in the middle of the stem). The cauline leaves or siliques were cut off and the stem segments were kept in liquid nitrogen until use. Diameters were measured using transverse hand sections of stems and light microscopy and, in addition, diameters of intact stems were measured with cryo-SEM. Typical diameters were around 0.6 mm for the basal and middle segments of the stems and around 0.4 mm for the apical segment. The surface areas of the segments were calculated using an average diameter for each segment and assuming cylinder geometry. In order to detect any significant departure from these standard diameters in the samples used for routine lipid analysis, the stem segments used were photocopied prior to chemical analysis and diameters were estimated from magnified copies. Cryo-SEM Segments from the apical 1 cm of stem were mounted onto cryo-SEM stubs with 25% dextran and plunged into liquid nitrogen. Frozen samples were transferred into an Emitech K1250 cryo-system, where water was sublimed for 30 min at −77°C and subsequently sputter coated with gold for 2.5 min at 35 mA. The coated samples were viewed with a Hitachi S4700 field emission SEM using an accelerating voltage of 2 kV and a working distance of 12 mm. Propidium Iodide Staining and Confocal Laser Scanning Microscopy Stem segments of Arabidopsis plants were immersed in a solution of propidium iodide (100 μg/mL; Sigma) and examined with a Radiance 2000 confocal laser-scanning microscope (Bio-Rad). The excitation wavelength was 568 nm with the emission filter set at 580 to 600 nm. All confocal images obtained were processed with ImageJ (http://rsb.info.nih.gov/ij) and Photoshop 5.0 (Adobe Systems) software. Polyester Analysis Thirty to 100 3-cm-long segments from 10- to 11-cm-long primary stems were used for each replicate. The analysis of the polyesters was conducted according to Bonaventure et al. (2004) with a slight modification (the use of centrifugation in 50-mL glass tubes at 800g for 20 min instead of filtration steps). Internal standards were methyl heptadecanoate and ω-pentadecalactone (Sigma). Polyesters from dried solvent-extracted residues of stems were depolymerized by hydrogenolysis with LiAlH4 or methanolysis with NaOCH3. The products recovered after hydrogenolysis were separated and quantified by GC as described previously (Bonaventure et al., 2004). For the products corresponding to several possible fatty acid derivatives, the proportions of diacids, diols, and ω-hydroxyl fatty acids were estimated based on the methanolysis data. Wax Analysis The cuticular waxes were extracted by immersing whole inflorescence stems or 3-cm-long segments of stems two times for 30 s into 5 mL of chloroform (CHCl3) at room temperature. Both CHCl3 solutions were combined and n-tetracosane was added as internal standard. The solvent was removed under a gentle stream of nitrogen gas, and the remaining wax mixture was redissolved in 1 mL of CHCl3 and stored at 4°C until used. The extracted area was determined by measuring the height and diameter of the stems assuming cylinder geometry. Prior to GC analysis, chloroform was evaporated from the samples under a gentle stream of nitrogen gas while heating to 50°C. Then the wax mixtures were treated with bis-N,N-(trimethylsilyl) trifluoroacetamide (BSTFA; Sigma) in pyridine (30 min at 70°C) to transform all hydroxyl-containing compounds into the corresponding trimethylsilyl derivatives. The qualitative composition was studied with capillary GC (5890 N; column 30 m Hewlett-Packard-1, 0.32-mm i.d., film thickness = 0.1 μm; Agilent) with He carrier gas inlet pressure regulated for constant flow of 1.4 mL min−1 and a MS detector (5973 N; Agilent). GC was carried out with temperature-programmed injection in a 50°C oven, 2 min at 50°C, raised by 40°C min−1 to 200°C, held for 2 min at 200°C, raised by 3°C min−1 to 320°C, and held for 30 min at 320°C. The quantitative composition of the mixtures was studied using capillary GC with FID under the same GC conditions as above, but H2 carrier gas inlet pressure regulated for constant flow of 2 mL min−1. Single compounds were quantified against the internal standard by automatically integrating peak areas. Isolation of Epidermal Peels Epidermal peels were manually dissected as a thin transparent film using thin forceps under a dissecting microscope (Fig. 5). For each freshly cut 3-cm-long stem segment of 10- to 11-cm-long primary stems, peels were collected, immediately frozen in liquid nitrogen, and stored at −80°C for RNA isolation or fatty acid analysis. Fatty Acid Analysis Epidermal peels from about two to three stem segments were heated at 85°C for 1.5 h in 1 mL of methanol containing 5% H2SO4 (v/v) with triheptadecanoylglycerol and ω-hydroxy-pentadecanoic acid as controls. Alternatively, the epidermal peels were quenched 10 min in isopropanol at 85°C and the lipids were extracted with hexane (3:2 [v/v] hexane:isopropanol final) before transmethylation. Fatty acid methyl esters (FAMEs) were extracted two times with 2 mL hexane and the solvent was evaporated under nitrogen gas. FAMEs were redissolved in 100 μL pyridine; 100 μL of acetic anhydride was added and the mixture was vortexed and heated for 1 h at 60°C. After evaporation of the solvents under nitrogen gas, the FAMEs were redissolved in hexane and analyzed by GC with FID on a DB-23 capillary column (J&W Scientific) under conditions allowing the separation of regular and VLCFAs. Gene Expression Analysis Total RNAs were isolated from epidermal peels or whole-stem segments using the RNeasy kit (Qiagen). Double-stranded cDNA was synthesized from approximately 15 μg of total RNA by using a SuperScript double-stranded cDNA synthesis kit (Invitrogen) with oligo d(T) primer containing a T7 RNA polymerase promoter sequence at its 5′ end (GGCCAGTGAATTGTAATACGACTCACTATAGGGAGGC-GG-(dT)24-3′; Genset). After synthesis of the second-strand cDNA, the cDNA was extracted with phenol-chloroform-isoamylalcohol, precipitated with ethanol, and resuspended in ribonuclease-free water. Labeled cRNA was generated from cDNA by in vitro transcription using a bioarray high-yield RNA transcript-labeling kit (Enzo Diagnostics) following the manufacturer's instructions and incorporating biotinylated CTP and UTP. After purification of biotin-labeled cRNA using an RNeasy column, 15 μg of the labeled cRNA were fragmented to a size of 35 to 200 bases by incubating at 94°C for 35 min in fragmentation buffer (40 mm Tris-acetate, pH 8.1, 100 mm potassium acetate, and 30 mm magnesium acetate). The fragmented cRNA was used for hybridization of Arabidopsis ATH1 gene chips (Affymetrix), which was performed in the Research Technology Support Facility at Michigan State University. Hybridization, washing, and detection of labeled cRNA were done as recommended by Affymetrix. Image acquisition and global data scaling were performed with Affymetrix MAS 5.0 software. The signal intensity values were scaled to a mean of 500 for each chip. The signal intensities indicated in the “Results” section are the mean from two biological replicates. The ratios of gene expression are the mean of four ratios of epidermis versus whole-stem segments calculated using multiple pairwise comparisons of epidermis versus stem. For each gene, a 90% confidence interval was calculated for the epidermis-to-stem ratio using a T-score and a degree of freedom of 3. Genes were considered as up-regulated in the epidermis if they were called present by the MAS 5.0 software in both biological replicates, and if the lower bound of the 90% confidence interval for the epidermis-to-stem gene expression ratio was ≥2.0. Indeed, for all epidermis-specific genes (Table I), the epidermis-to-stem ratio was found to lie in a 90% confidence interval whose lower bound was 2.0 or above. These results thus provided an empirical basis for the use of 2.0 as a cutoff value for the lower estimates of the ratios to identify epidermis-specific genes in our dataset. Other studies have indicated that robust gene expression changes can be consistently identified by selecting transcripts with a gene expression ratio >2.0 (mean ratio) for increases (Wodicka et al., 1997). Hence, a value of 2.0 for the lower estimate of the ratio is clearly conservative. 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Plant Cell 15 : 1872 –1887 Author notes 1 This work was supported by the Dow Chemical Company, Dow AgroSciences, a U.S. Department of Agriculture National Research Initiative grant (grant no. 05–35318–15419 to M.P. and F.B.), the Canadian National Science and Engineering Research Council of Canada Special Research Opportunity Grant (grant no. 305360–04 to A.L.S., R.J., and L.K.), and the Plant Signaling Network Research Center and the Agricultural Plant Stress Research Center Grant of the Korea Science and Engineering Foundation (grant no. R11–2001–0920301–0 to M.C.S.). * Corresponding author; e-mail [email protected]; fax 517–353–1926. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Fred Beisson ([email protected]). [W] The online version of this article contains Web-only data. 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