Transcriptional Control of SET DOMAIN GROUP 8 and ... - Cell Press

Molecular Plant

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Volume 3

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Number 1

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Pages 174–191

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January 2010

RESEARCH ARTICLE

Transcriptional Control of SET DOMAIN GROUP 8 and CAROTENOID ISOMERASE during Arabidopsis Development Christopher I. Cazzonelli, Andrea C. Roberts, Melanie E. Carmody and Barry J. Pogson1 Australian Research Council Centre of Excellence in Plant Energy Biology, Research School of Biology, The Australian National University, Canberra, ACT 0200, Australia

ABSTRACT Carotenoids are pigments required for photosynthesis, photoprotection and the production of carotenoidderived hormones such as ABA and strigolactones. The carotenoid biosynthetic pathway bifurcates after lycopene to produce epsilon- and beta-carotenoids and this branch is critical for determining carotenoid composition. Here, we show how the branch point can be regulated by the chromatin-modifying histone methyltransferase, Set Domain Group 8 (SDG8) targeting the carotenoid isomerase (CRTISO). SDG8 is required to maintain permissive expression of CRTISO during seedling development, in leaves, shoot apex, and some floral organs. The CRTISO and SDG8 promoters show overlapping tissue-specific patterns of reporter gene activity. Interestingly, CRTISO showed atypical reporter gene expression in terms of greater variability between different lines compared to the Cauliflower Mosaic Virus 35S promoter (CaMV35s) and eLCY promoters, potentially due to chromosomal position effects. Regulation of the CRTISO promoter was dependent in part upon the presence or absence of SDG8. Knockouts of SDG8 (carotenoid and chloroplast regulation (ccr1)) and CRTISO (ccr2) result in altered carotenoid composition and this could be restored in ccr2 using the CaMV35s or CRTISO promoters. In contrast, varying degrees of GUS expression and carotenoid complementation by CRTISO overexpression using CaMV35S or CRTISO promoters in the ccr1 background demonstrated that both the CRTISO promoter and open reading frame are necessary for SDG8-mediated expression of CRTISO. Key words: Photosynthesis; secondary metabolism—terpenoids, isoprenoids, and carotenoids; chloroplast biology; epigenetics; gene expression; gene regulation.

INTRODUCTION Carotenoids are the second most abundant pigment in the natural world, ranging from the yellow, orange, and red of many fruits, vegetables, flowers, and autumn leaves through to the colors of butterflies and crayfish. The bright colors of carotenoid pigments attract insects for pollination and dispersal of seeds. Fruits and vegetables provide essential human dietary sources of carotenoids for proVitamin A activity, antioxidants, and limiting age-related macular degeneration of the eye. Carotenoids are important for photosynthetic organisms, where they play crucial roles in photosystem assembly, lightharvesting, and photoprotection, and thus their function and biosynthesis have been reviewed extensively (Niyogi, 1999; Cuttriss and Pogson, 2004; DellaPenna and Pogson, 2006; Lu and Li, 2008). The proportion of different xanthophylls greatly affects plant viability and photoprotection. Typically, leaf tissues accumulate one e-carotenoid, lutein, and three b-carotenoids—b-carotene, violaxanthin, and

neoxanthin—with changes in this profile altering photosynthesis, antenna assembly, and photoprotection (DemmigAdams and Adams, 2006; Pogson et al., 2006). Thus, to avoid extensive photo-oxidative damage, the synthesis of carotenoids, chlorophylls, and their subsequent binding to pigment-binding proteins must be precisely balanced to meet the appropriate photosynthetic demands that plants are exposed to on a daily and seasonal basis. Consequently, carotenoid biosynthesis appears to be tightly regulated throughout the lifecycle, with dynamic changes in composition matched to developmental requirements including germination,

1 To whom correspondence should be addressed. E-mail barry.pogson@ anu.edu.au, tel. 011-61-2-61252663.

ª The Author 2009. Published by the Molecular Plant Shanghai Editorial Office in association with Oxford University Press on behalf of CSPP and IPPE, SIBS, CAS. doi: 10.1093/mp/ssp092, Advance Access publication 17 November 2009 Received 3 July 2009; accepted 22 September 2009

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photomorphogenesis, and fruit development (Welsch et al., 2000; Hirschberg, 2001; Fraser et al., 2008). Carotenoid pigments also provide substrate precursors for the biosynthesis of phytohormones such as abscisic acid (ABA) and strigolactones (Figure 1) (Nambara and Marion-Poll, 2005; DellaPenna and Pogson, 2006; Howitt and Pogson, 2006; Gomez-Roldan et al., 2008; Umehara et al., 2008). The strigo-

Figure 1. A Model Showing Key Steps in the Regulation of Carotenoid Biosynthesis in Arabidopsis. Arabidopsis mutations ccr1, ccr2, lut1, lut2, lut5, aba1, and npq1 are shown in gray italics. The first committed step in the synthesis of phytoene from geranylgeranyl pyrophosphate is catalyzed by the light-responsive enzyme, PSY. The branch point is a key regulatory step in carotenoid biosynthesis and involves the isomerization of tetra-cis-lycopene to lycopene by CRTISO and light (e.g. photoisomerization). Epigenetic regulation by a chromatin modifying histone methyltransferase, SET DOMAIN GROUP8 (SDG8), and metabolic feedback mechanisms control rate-limiting enzymes, CRTISO and eLCY, respectively. Lycopene undergoes further modifications by eLCY and b-LCY to produce a- and b-carotene, respectively, which serve as substrates for the production of lutein, phytohormones (abscisic acid and strigolactones), and an unknown signaling compound mediated by BYPASS (BPS). bLCY, b-cyclase; bOH, b-hydroxylase; CCD, carotenoid cleavage dioxygenase; CRTISO, carotenoid isomerase; eLCY, e-cyclase; eOH, e-hydroxylase; NCED, 9-cis-epoxycarotenoid dioxygenase; NXS, neoxanthin synthase; PDS, phytoene desaturase; PSY, phytoene synthase; VDE, violaxanthin de-epoxidase; ZDS, f-carotene desaturase; ZE, zeaxanthin epoxidase; Z-ISO, f-carotene isomerase.

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lactone class of carotenoid metabolites are graft transmissible and inhibit shoot branching in Arabidopsis, pea, petunia, and rice, while also functioning to stimulate a symbiotic relationship with fungi in the rhizosphere (Gomez-Roldan et al., 2008; Pichersky, 2008; Umehara et al., 2008). ABA is required to maintain shoot and root growth (LeNoble et al., 2004; Rodrigues et al., 2009). A novel graft transmissible b-carotene-derived signaling compound was also shown to be required for normal root and shoot development in Arabidopsis (Figure 1) (Van Norman and Sieburth, 2007). Given the role of roots as sources of ABA and strigolactones, it is perhaps surprising that the essentially colorless root tissues have not been investigated in detail, although trace levels of neoxanthin and violaxanthin were reported for Arabidopsis, pea, and tobacco (Parry and Horgan, 1992). Uncovering the molecular nature of regulatory mechanisms and the processes by which carotenoid composition is controlled has been a major challenge in understanding carotenoid metabolism (Lu and Li, 2008). Despite knowledge that flux through the branch point in the carotenoid pathway can affect plant development in response to environmental stimuli, the major regulators of this pathway have remained rather elusive. Two steps in the primary pathway are considered to be major targets for regulation of carotenoid composition. The first step, phytoene synthase (PSY) that converts geranylgeranyl diphosphate to phytoene, has been described as the bottleneck and a number of recent studies have focused on the transcriptional regulation of PSY (Figure 1) (Welsch et al., 2007, 2008). In greening seedlings, PSY is strongly light-induced (Welsch et al., 2000) and the transcription factor, RAP2.2 (AP2/EREBP family), has been shown to bind to the PSY promoter (Welsch et al., 2007). However, modulating RAP2.2 levels resulted in only small pigment alterations in Arabidopsis root calli (Welsch et al., 2007). The second major step in the pathway can be considered the branch point between flux towards epsilon-carotenoids (e.g. lutein) or beta-carotenoids (e.g. violaxanthin), determined in part by two rate-limiting enzymes—epsilon cyclase (eLCY) and carotenoid isomerase, which provides substrates for beta and epsilon cyclases (Figure 1) (Cuttriss et al., 2007). CRTISO somerases cis-carotenes, such as tetra-cis-lycopene to all-trans-lycopene, which is the substrate for the cyclases (Beyer, 1989; Schnurr et al., 1996; Bartley et al., 1999; Giuliano et al., 2002; Isaacson et al., 2002; Park et al., 2002; Isaacson et al., 2004; Breitenbach and Sandmann, 2005; Li et al., 2007). Mutations in crtiso (ccr2) alter carotenoid composition in Arabidopsis, rice, and tomato (Isaacson et al., 2002; Park et al., 2002; Fang et al., 2008) and, in particular, the most abundant carotenoid, lutein, is reduced in light-grown tissues (Park et al., 2002). The branch point of pathway has been shown to be regulated by a chromatin modifying enzyme, SET DOMAIN GROUP 8 (SDG8), which also regulates flowering time, as well as shoot and root branching (Kim et al., 2005; Zhao et al., 2005; Dong et al., 2008; Xu et al., 2008; Cazzonelli et al., 2009a, 2009c). The

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question is: does this indicate a link between carotenoid biosynthesis and key developmental processes (Cazzonelli et al., 2009b)? The absence of SDG8 in ccr1 alters the methylation of chromatin surrounding the carotenoid isomerase gene, impairs lutein biosynthesis, alters root branching, and increases shoot branching (Cazzonelli et al., 2009a, 2009c). The enhanced shoot branching displayed by ccr1 is partly explained through limiting biosynthesis of the carotenoidderived branching hormone strigolactone (Cazzonelli et al., 2009a), as well as by controlling the methylation status around target shoot branching genes (Dong et al., 2008). So it is becoming increasingly clear that carotenoid biosynthesis is required for the production of graft transmissible signaling compounds. Regulation at the branch point of the biosynthetic pathway may therefore represent a mechanism for influencing root and shoot branching in addition to carotenoid composition. This investigation looks at the transcriptional regulation of CRTISO and SDG8. In particular, we investigated when and where SDG8 regulates CRTISO expression during Arabidopsis development. CRTISO expression requires SDG8 activity during seedling development and expression of their promoters overlap in specific tissues such as the hypocotyl, shoot apical meristem region, leaf vasculature, and pollen. Sequences located in both the CRTISO promoter and open reading frame are necessary for SDG8 to promote expression driven by the CRTISO promoter.

cDNA sequence data obtained from Genbank (–8 bp; BX816901 and –12 bp; NM_100559). The actual CRTISO transcription start site was determined by RNA ligase-mediated and oligo-capping rapid PCR amplification of cDNA 5# ends and products were cloned and sequenced. Seven potential transcription start sites spanning –53 bp upstream from the ATG start codon were identified (Supplemental Figure 1A). Six of the potential start sites were represented by either one (–8 and –53 bp), two (–29, –36, and –48 bp), or three (–40 bp) clones. An adenine residue positioned –20 bp upstream from the ATG was confirmed by six sequence clones to be the most likely start of transcription, and matched the eukaryotic consensus sequence (CA) (Bucher and Trifonov, 1986). In silico analysis of the SDG8 promoter (–837 bp) did not uncover a discernible TATA, CAAT, or transcription start site in the core domain (Supplemental Figure 1B). The upstream regulatory domain shared two of the cis-elements identified in CRTISO, that is it contained a gibberellin-responsive element (GARE) (Sutoh and Yamauchi, 2003) and W-box motifs (Eulgem et al., 1999) (Supplemental Figure 1B). A 12-bp DE-1 cis-acting element involved in phytochrome down-regulation of gene expression was identified (Inaba et al., 2000). Both the SDG8 and CRTISO promoters contain a 12-bp sequence fragment in opposite orientations and the function of this element, if any, remains unknown (Supplemental Figure 1A and 1B).

The Promoters of CRTISO and SDG8 Display Unusual Variability in Expression

RESULTS Bioinformatic Identification of Core and Upstream Regulatory Elements of CRTISO Regulatory domains of the CRTISO (–570 bp) promoters were analyzed in silico for sequence elements that may function in basal transcription, direct tissue-specific expression, or mediate an environmental and/or developmental signal (Supplemental Figure 1A). A search for core promoter elements (e.g. TATA and CAAT boxes) that are consistent with other published core promoter consensus sequences (approximately –30 bp upstream from TSS) did not uncover any clear candidates (Yamaguchi et al., 1998; Molina and Grotewold, 2005; Yamamoto et al., 2007). In silico database searches (Lescot et al., 2002) for regulatory sequences within the CRTISO upstream regulatory domain (–570 bp) identified a number of putative cis-elements that have been demonstrated to be involved in responses to gibberellins, a direct repeat of a gibberellin response element (GARE) (Sutoh and Yamauchi, 2003); auxin, namely NDE (McClure et al., 1989) and TGA (Dhadi et al., 2009); and wounding elicitation, W-box (Eulgem et al., 1999). In addition, a number of direct repeat elements were identified (Cazzonelli and Velten, 2008) (Supplemental Figure 1A). Analysis of the CRTISO core promoter sequence identified two possible transcription start sites based upon full-length

The strength of promoter regulatory regions controlling CRTISO and SDG8 gene expression were tested in wild-type and/or the ccr1 mutant using an in vivo luciferase reporter gene activity assay (Velten et al., 2008). SDG8 maintains permissive gene expression of a small number of target genes presumably by facilitating an open configuration of chromatin surrounding the transcription start site (Cazzonelli et al., 2009b). Independent transgenic lines harboring a promoter– reporter gene fusion can show considerable variation in the strength and tissue-specific pattern of expression due to a combination of distal cis-acting enhancer elements and other effects associated with regulatory state of chromatin neighboring the site of T-DNA insertion. Therefore, a large number of transgenic lines (13–70) harboring test promoter– reporter fusions (pTPromoter:intron modified Firefly (Photinus pyralis) luciferase gene (FiLUC); Figure 2) were generated in order to determine representative patterns of tissue specific expression and select individual lines for further characterization. Individual transformation events were analyzed for luciferase activity across multiple promoter–luciferase constructs (Figure 3). Both the full-length CRTISO promoter (pTCRTISO1F:FILUC; –1977 bp) and a deletion fragment (pTCRTISO5F:FiLUC; –570 bp) enable near background to low levels of luciferase activity (,1000 Relative Light Units (RLU)) in

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Figure 2. Schematic Diagram Showing Overexpression and Promoter–Reporter Gene Constructs. (A) Representative design of all pT* series of vectors showing a promoter:gene:nopaline synthase terminator (Nost) fusion adjacent to the right border of the binary vector, pPZP200 (Cazzonelli and Velten, 2008). (B) Overexpression constructs contain a CRTISO or CaMV35s promoter fragment fused adjacent to the wild-type CRTISO open reading frame and terminator (Cazzonelli et al., 2009a). Promoters used to drive CRTISO overexpression include –46 bp of the minimal CaMV35s (pTm35:CRTISO), –768 bp of a double CaMV35s enhancer (pMDC32:CRTISO), –1977 bp (pTCPCG-1F), and –570 bp (pTCPCG-5F) from the CRTISO upstream regulatory domain. (C, D) Regulation of CRTISO (–1977and –570 bp), SDG8 (–1787and–873 bp),and eLCY(–470 bp) promoteractivitywas analyzedusing luciferase (pT*-FiLUC) and/or GUSi (pT*-GUSi) reporter gene vectors. The CaMV35s and SYNPRO3 strong constitutive promoters were included as positive controls for either luciferase (pT35enh-FiLUC) or GUS (pSYNPRO3-GUSi) reporter gene analysis (Cazzonelli and Velten, 2008).

wild-type leaf tissues: 132 and 134 RLU, respectively (Figure 3A and 3C). Stronger luminescence was recorded in leaf tissues from the eLCY (pTeLCY-4F:FiLUC; –450 bp) and SDG8

(pTSDG8-H3:FiLUC –1787 bp and pTSDG8-1F:FiLUC; –837 bp) promoters in comparison to the CRTISO promoter, emitting, on average, 3511 and 7177 RLU above background,

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respectively (Figure 3E–3G). All three promoters are several hundred-fold weaker in comparison to the strong, constitutive CaMV35s promoter (Figure 3H; ,221 000 RLU). The full-length and the deletion fragment of the CRTISO promoter enabled similar luciferase activity levels in wild-type and ccr1 leaf tissues (Figure 3A–3D). Among the independent lines, there was an unusual variability in luciferase activity and the activity spectrum increased in an exponential manner. That is, the average of luciferase activity among the independent lines was off-center and shifted more towards the lines showing strongest luciferase activity. The exponential activity spec-

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trum was more evident for the –570-bp CRTISO promoter deletion (Figure 3C) and was slightly less pronounced in ccr1 tissues (compare Figure 3A with 3B and 3C with 3D). The –1787 and –837-bp fragments of the SDG8 promoter also displayed an exponential increase in luciferase activity (Figure 3E and 3F). In contrast, the luciferase activity directed by the eLCY promoter (Figure 3G) in leaf tissues increased in a linear nature similar to that of strong, constitutive CaMV35s promoter (Figure 3H). Therefore, it appears that both the CRTISO and SDG8 promoters result in an unusual and variable reporter gene activity spectrum among the different transgenic lines.


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Figure 3. Luciferase Activity Spectrum for CRTISO, eLCY, and SDG8 Promoter Lines in Wild-Type and ccr1 Leaf Tissues. Multiple independent transgenic lines harboring pTCRTISO1F:FiLUC (A, B), pTCRTISO5F-FiLUC (C, D), pTm35enh-FiLUC (E), pTeLCY4F-FiLUC (F, G), pTSDG81F-FiLUC (H), pTSDG8H3-FiLUC (I), and pTSYNPRO3-FiLUC (J) promoter–luciferase fusions were analyzed for luciferase activity and presented in the order of increasing activity along the x-axis. All promoter constructs were tested in a wild-type Col background and, in addition, the CRTISO and eLCY promoters were each tested in the ccr1 mutant (B, D). Select lines chosen for further analysis of SDG8:FiLUC regulation after germination are displayed. The luciferase activity of mature leaf tissues (at least two leaves per plant) from independent transgenic lines (R0 plants) were measured using the in vivo leaf-disk assay (at least two leaf discs per leaf).

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SDG8 Is Required for Permissive Expression of CRTISO during Seedling Development SDG8 gene expression was shown to be regulated during seedling development where transcript levels increased 8 d after germination (DAG) (Kim et al., 2005). A selection of independent transgenic lines (Figure 3A and 3E) harboring pTCRTISO1F:FiLUC (#C6, A3, and A11) and pTSDG8-1F:FiLUC (#N1, G2, G3, D3, K1, D2, J1, and A1) promoter–reporter fusions were analyzed during seedling development. The SDG8 promoter lines were grouped into four categories based upon their luminescence levels at day 5 relative to day 9. Group 1 were always highly expressed, group 2 displayed a strong switch from low basal to high levels, group 3 switched from undetectable to low basal levels, and group 4 showed no luminescence (Figure 4). A CaMV35s control promoter was included in all experiments to confirm efficient uptake of luciferin (Figure 4). The SDG8 promoter showed stronger luminescence in the cotyledons, shoot apex, and hypocotyl tissues at 9 DAG; however, the intensity of luminescence was variable, depending not only on the transgenic line, but also among siblings of

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a hemizygous population (Figure 4). The intensity of luminescence ranged from strong (A1 and J1) to moderate (D2 and K1) to weak (D3 and G3), depending upon the line. This pattern correlates with the unusual variability between lines observed in the activity assays (compare Figures 3E and 4). In vivo imaging of CRTISO:LUC activity in seedling tissues showed very low levels of luminescence in the shoot apex, if any, and therefore made it difficult to determine the regulatory pattern of the CRTISO promoter (data not shown), which is consistent with the low levels of luciferase activity measured in mature leaf tissues (Figure 3A). Due to low levels of CRTISO promoter–luciferase activity, real-time quantitative PCR (RT–qPCR) was used to quantify CRTISO and SDG8 transcript levels in 2–10-day-old seedling tissues (minus the roots) growing on sucrose containing MS agar. SDG8 and CRTISO mRNA were detected at each stage during seedling development in two experimental replicates. In one experiment, there was a slight increase in relative SDG8 and CRTISO mRNA levels at 10 DAG in comparison to 5 DAG (Figure 5A). However, in a second experiment, the SDG8 and CRTISO mRNA levels showed a significant decrease during seedling development (Figure 5B). One possible explanation for the variation was that the plants in the first experiment showed anthocyanin accumulation—a sign of stress—and this was not apparent in the second experiment. Regardless, the overall abundance of CRTISO and SDG8 mRNA transcript levels was similar in both experiments. In ccr1 mutant seedlings, the relative expression of CRTISO was reduced by greater than 80% throughout seedling development (compare 2, 4, and 9 DAG) and demonstrates that SDG8 is required to maintain CRTISO gene expression during seedling development (Figure 5D) in addition to mature leaves (Cazzonelli et al., 2009a).

CRTISO and SDG8 Promoters Drive Tissue-Specific and Overlapping GUS Expression

Figure 4. Regulation of the SDG8 Promoter during Seedling Development. In vivo luciferase expression in seedling tissues from independent lines harboring SDG8 (pSDG8-1F:FiLUC) or CaMV35s (pT35enh: FiLUC) promoters. Bioluminescence was assayed 5 and 9 d after germination. Images were adjusted by false colorization using Image J software (version 1.41, National Institutes of Health, USA) and the threshold was adjusted such that black (700) indicates no luminescence, blue reflects low levels of luminescence, and white (1000) reflects higher levels of luminescence. Representative lines are arranged in a descending order of light intensity (group 1 = strongly active; group 2 = strong switch; group 3 = weak switch; group 4 = no expression; and CaMV35s control = strong constitutive promoter).

The expression patterns driven by the CRTISO and SDG8 promoters were further characterized using GUS as a reporter for analyzing tissue-specific gene expression. Selected lines were grown on sucrose containing MS agar (1–2 DAG) and/ or soil (4–50 DAG) and GUS histochemical assays were used to visualize GUS activity in tissues from multiple hemizygous (F2) or homozygous (F3) plants. A very thorough analysis of GUS tissue-specific staining was performed during various stages of growth (seed germination: day 1, seedling development: days 2–9, mature vegetative development: days 13–25, and flowering: days 35–50) in multiple experiments, using 37–90 transgenic lines per construct. Mature tissues from transgenic lines harboring pTCRTISO-1F:GUS or pTSDG8-1F:GUS were analyzed for promoter activity. A GUS intensity score was determined for individual CRTISO and SDG8 lines by calculating an average score of multiple tissues and experiments by qualitatively scoring the intensity of GUS staining relative to the strong constitutive Synthetic Promoter (SYNPRO)3 (Figure 6A and 6B). The expression of promoter:GUS activity

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Figure 5. Relative Transcript Abundance of SDG8 and CRTISO. Quantification of SDG8 and CRTISO transcript abundance during seedling development was performed using (A) whole seedlings showing some symptoms of stress, such as anthocyanins, and (B) whole seedlings with no obvious signs of stress. Wild-type and ccr1.1 were grown on MS media and whole seedlings minus the roots harvested at 2, 5, or 9 DAG. Seedling tissues were assayed in triplicate and normalized to the earliest day examined. (C) Analysis of SDG8 and CRTISO transcript levels in floral organs. Transcript levels were measured in leaf, stamen (anther and filament), and petal tissues using RT–qPCR and the average of duplicate samples normalized to leaf tissues. (D) qPCR analysis of CRTISO transcript levels in seedling tissues from ccr1. RT–qPCR was used to quantify gene expression levels in three biological replicate per seedling stage, and transcript levels normalized to day 2 wild-type seedlings. (E) CRTISO transcript abundance in ccr1 rosette leaf, shoot apical meristem region, and flowering tissues relative to wild-type leaves. Data represent two biological repeats of pooled tissues. CRTISO mRNA expression levels were measured in leaf (day 23), shoot apical meristem region (day 23), and open flowers (day 29) from mature plants grown on soil. A housekeeping gene (TIP41) was used to normalize differences in biological variation among different tissues, and a secondary housekeeping gene (CYCLO), as well as a synthetic technical control (IC), reveal biological and technical variation. Standard error bars are displayed (n = 2–3). Abbreviations are in Supplemental Table 2.

in specific tissues was calculated as an average of the staining intensity in 5–16 lines during various stages of growth and in the roots, stem (hypocotyl), cotyledons, shoot apical meristem region, leaf veins (vasculature), and true leaves (Figure 6C and 6D). Based upon the overall GUS staining scores during various stages of development and associated tissue-specific patterns, representative lines with a stronger GUS staining pattern of expression were identified and labeled CRTISO:GUS lines (#D14, C1, B1) and SDG8:GUS lines (#I102, H63, C3, and E7), respectively (Figure 6A and 6B). Several SDG8:GUS lines displayed strong and consistent GUS staining in the hypocotyl (stem), cotyledon and shoot apex, and, to a lesser degree, in the leaf vasculature and older true leaves (Figure 7I–7P). GUS expression was also strongly apparent in germinating seeds and in the mature pollen from anthers (Figure 7I and 7O). Representative CRTISO:GUS lines (#D14, C1, B1) mimicked the SDG8:GUS expression patterns and overlapped considerably in the shoot apical meristem, leaf vasculature, and pollen, as well as, to a lesser degree, in the hypocotyl and cotyledons (compare Figure 7A–7H with 7I– 7P). CRTISO:GUS activity was visible in immature seeds from developing siliques (Figure 7G); however, in contrast to the SDG8 promoter lines, GUS expression was not visible in germinating seeds (Figure 7A and 7I). Interestingly, the GUS lines, like the

luciferase lines (Figure 4), could be grouped into four categories based on their staining patterns between 2 and 9 DAG, although segregation into each of these groups was less clear in GUS lines. Weak and variable GUS staining driven by the SDG8 and CRTISO promoters was observed in the roots, older rosette leaves, leaf mesophyll, and floral stems in some experiments; however, the expression was far too inconsistent to draw strong conclusions (data not shown). The CRTISO and SDG8 promoters did not show visible GUS expression in flower tissues such as the petals, sepals, or maternal organs, such as the stigma, style, and ovary. The relative abundance of CRTISO mRNA transcripts in the leaves, shoot apical meristem region, and open flowers from ccr1.4 relative to wild-type plants was quantified by RT–qPCR. Both SDG8 and CRTISO transcripts were detected in wild-type tissues (data not shown), which correlates with GUS assay data, suggesting that these two genes are co-expressed in the shoot meristem, male anthers, and leaf vasculature (Figures 6 and 7). Also, CRTISO transcript levels were severely reduced in all ccr1 tissues examined (.90%) (Figure 5E). Bioinformatics analysis of microarray data using Genevestigator showed that SDG8 and CRTISO transcript levels are detectable throughout plant development and in most tissues (Supplemental Figure 2A and 2B). Most interestingly, both SDG8 and CRTISO are highly

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Figure 6. Qualitative Analysis of CRTISO and SDG8 Promoter Expression in Multiple Lines. GUS staining in tissues was qualitatively scored on a scale of 1 to 3 (1 = weak, 2 = moderate, and 3 = strong) relative to strong, constitutive SYNPRO3 and CaMV35s promoters. (A, B) GUS staining for multiple wild-type lines harboring pTCRTISO-1F:GUS (n = 37) or pTSDG8-1F:GUS (n = 38), respectively. (C, D) GUS staining in select tissues from multiple wild-type lines harboring pTCRTISO-1F:GUS or pTSDG8-1F:GUS, respectively. The average GUS staining score of various tissues (roots, hypocotyls, cotyledons, leaf mesophyll, leaf veins, shoot apical meristem, and flowers) were averaged from multiple assays performed throughout development. Each score represents the average of multiple samples in at least two independent experiments.

expressed in flowers and SDG8 expression was even more pronounced in the stamen and pollen (Supplemental Figure 2B). Quantification of CRTISO mRNA abundance in wild-type stamens (anther and filament) and petals revealed transcript levels that were only 33 and 43%, respectively, of transcript levels quantified from leaf tissues (Figure 5C). SDG8 mRNA levels were almost six-fold higher in the stamens when compared with leaves and petals (Figure 5C). Both SDG8 and CRTISO transcripts were detectable in the petals, which is inconsistent with the promoter assays that did not reveal any visible GUS staining. Due to the proximity of pollen to petals, perhaps the CRTISO and SDG8 transcripts that were detected resulted from contamination of petals with pollen material. In summary, transcriptional regulation of CRTISO and SDG8 appears to

overlap in a tissue-specific manner, especially in the shoot apical meristem and pollen.

Regulation of the CRTISO Promoter Is Partially Impaired in ccr1 The regulation of the CRTISO promoter was tested in the SDG8 mutant, ccr1, in order to determine whether SDG8 recognizes and targets specific control sequences contained with the upstream regulatory region of CRTISO (Figure 8). GUS staining scores were determined as described above. The average score of multiple tissues from independent experiments revealed that a third of the 15 ccr1 transgenic lines harboring pTCRTISO-1F:GUS displayed a GUS staining score similar to that of wild-type lines (Figure 8A). Further analysis of select lines

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Figure 7. GUS Histochemical Analysis of CRTISO and SDG8 Promoter Expression in Wild-Type Plants. GUS assays were performed throughout development using multiple plants (5–30) from pTCRTISO-1F:GUS (A–F) and pTSDG8-1F:GUS (G–L) transgenics. Independent lines (SDG8:GUS#C3, E7, H63, and I102 and CRTISO:GUS#B1, C1, and D14) displaying strong GUS activity in representative tissues from the analysis of 15–30 lines are displayed. Stages of development analyzed include seed germination (day 1; A and G), seedling development (day 2; B and H; day 4; C and I), true leaf development (day 9; D and J), mature plantlets (days 12–14; E and K), and flowering tissues (day 50; F and L).

growing on soil (C2, E2, E1, G1, and J1) and or sucrose containing MS agar (A1, A2, A3, A4, A5, C1, F1, G2, and J2) showed a tissue-specific pattern of GUS expression (Figure 8B) similar to that observed for wild-type lines (Figure 6D) during various stages of development. Even though GUS staining was visible in the shoot apex, pollen, leaf veins, and more variably in the hypocotyl, cotyledon, and true leaves, the intensity of staining appeared considerably lower in ccr1 than wild-type (Figure 8C). The majority of ccr1 lines did not show any GUS staining in the shoot apical meristem or hypocotyl, although minor GUS stain-

ing was more usually apparent in the leaf veins during later stages of seedling development (day 9) and in the pollen (Figure 8C). The strong constitutive SYNPRO3 promoter was used as a control to rule out artifacts associated with the histochemical analysis of GUS expression in all tissues tested (Figure 8D). In general, the activity of the CRTISO promoter was partially impaired in the ccr1 mutant. The variability among independent lines might be attributed to the chromatin environment surrounding the T-DNA insertion (Lam et al., 2009; Luo et al., 2009).

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Figure 8. GUS Analysis of the CRTISO Promoter in ccr1. GUS staining was qualitatively scored on a scale of 1 to 3 (1 = weak, 2 = moderate, and 3 = strong) in comparison to strong, constitutive promoters such as SYNPRO3 (SP) and CaMV35s (35S). (A) Relative GUS staining scores of multiple ccr1.1 lines harboring pTCRTISO-1F:GUS. Scores represent the average of multiple tissues analyzed from eight independent experiments. ccr1 lines crossed to wild-type Columbia are indicated by a star. (B) Relative GUS staining scores for roots, hypocotyls, cotyledons, leaf mesophyll, leaf veins, shoot meristem region, and flowers. Scores represent the average of tissues assayed from multiple ccr1.1 lines during several stages of development. (C, D) GUS histochemical analysis of representative lines harboring pTCRTISO-1F:GUS or pTSYNPRO3-GUS in ccr1 and wild-type plants, respectively. GUS histochemical assays were performed during several stages of development as indicated. A control for penetration artifacts associated with the GUS histochemical assay was the strong and constitutive SYNPRO3 promoter line (pTSYNPRO3-GUS#B4).

It was hypothesized that ccr1 lines displaying an impaired pattern of CRTISO:GUS expression would display a wild-type GUS expression pattern if SDG8 activity was restored. Therefore, select lines (C2, G1, J1; Figure 8A) showing low GUS staining during early stages of development and minor GUS expression in the leaf veins were crossed to wild-type plants in order to restore SDG8 activity. Seed from successful crosses was subsequently grown to maturity and self-fertilized. At least two lines displaying a wild-type phenotype and homozygosity for the promoter–reporter transgene (C2#8A and 1E,

G1#18B and 22A, J1#20B and 21A) were grown on soil and/ or MS agar and GUS expression analyzed during development. While GUS staining was observed in mature tissues such as the leaf vasculature and pollen, GUS expression was not restored in the shoot apical meristem region or hypocotyl.

SDG8 Targets Sequences Contained within the CRTISO Open Reading Frame A range of promoters were fused to the CRTISO gene (Figure 2B) in order to test the requirement of the CRTISO promoter

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and gene sequences to enable SDG8 activity and maintain a wild-type carotenoid composition. Mutants of sdg8 (ccr1-1, ccr1-4) and crtiso (ccr2) were transformed with a genomic fragment (including the 3’ untranslated region) of the CAROTENOID ISOMERASE gene driven by either the CaMV35s, CRTISO-1F, CRTISO-5F, or minimal CaMV35s promoters. The carotenoid composition of leaves from hemizygous plants were quantified by HPLC. Relative to wild-type plants, the carotenoid profiles of ccr1 and ccr2 (Table 1A) show an increase in violaxanthin, antheraxanthin, and b-carotene, and a substantial decrease in lutein (,23%), all of which is consistent with previous findings due to the lack of CRTISO activity (Park et al., 2002). The minimal CaMV35s promoter, which enables basal levels of reporter gene expression in tobacco leaf, cotyledon, and stem tissues (Cazzonelli and Velten, 2008), was insufficient to restore a wild-type carotenoid profile in ccr2 or ccr1 lines (Table 1A). However, the strong, constitutive CaMV35s promoter did restore wild-type levels of lutein, violaxanthin, antheraxanthin, and b-carotene. Interestingly, the full-length (–1977 bp) and deletion fragment (–570 bp) of the CRTISO promoter were sufficient to restore carotenoid content in ccr2, but not ccr1 (Table 1A). TheabundanceofCRTISOmRNAinmature leaftissues(day32, growth in soil) from duplicate ccr2, ccr1, and wild-type lines harboring either the CaMV35s or CRTISO-1F promoter–genefusions was quantified by RT–qPCR. The CaMV35s promoter enabled a 36–64-fold increase in CRTISO mRNA, irrespective of the genotype. In contrast, the CRTISO-1F construct failed to restore wildtype levels of CRTISO mRNA in ccr1 (0.1–0.14-fold) despite overexpression in ccr2 (2–32-fold) and wild-type (7–14-fold) plants. Three independent crosses between a complemented ccr2.5 line harboring CRTISO-1F:CRTISO (pTCPCG-1F) and ccr1.1 were undertaken in order to rule out any position effects surround-

ing the T-DNA insertion on the regulation of the CRTISO promoter–gene fusion. The F2 progeny were phenotyped (early flowering, increased shoot branching, and lutein content) to distinguish ccr1 from ccr2, and the presence of the transgene confirmed by herbicide selection. Three progeny confirmed as ccr1 (phenotype = branchy, early flowering, and pale green) and containing the CRTISO transgene showed reduced lutein levels (29–37%), whereas genetic and HPLC analyses of plants with wild-type flowering and branching habit showed co-segregation of the restoration of lutein levels and the presence of the transgene in both wild-type and ccr2 genotypes. This confirms that the CRTISO promoter–CRTISO gene fusion was unable to restore lutein levels in the ccr1 mutant, despite complementation of carotenoids in ccr2.

DISCUSSION SDG8 Maintains CRTISO Expression during Seedling Development SDG8 is required to maintain active transcription of CRTISO (Cazzonelli et al., 2009a). In order to determine where and when the regulation of CRTISO is under permissive control by SDG8, the transcriptional regulation of both SDG8 and CRTISO was investigated throughout development. The SDG8 promoter:GUS and luciferase lines displayed a tissue-specific pattern of reporter expression during germination and seedling development (Figures 4 and 7), although there was greater variability in the strength of luciferase expression (Figure 3E). In general, SDG8 expression was observed 1 DAG within the seed and emerging root (Figure 7I) and young seedlings showed intense GUS staining in the hypocotyl, shoot apical meristem region, cotyledons, vasculature, and more variably in the primary root (Figure 7I–7M).

Table 1. The CRTISO Promoter–Gene Fusion Restores Carotenoid Composition in ccr2, But Not ccr1. Percentage Carotenoid Composition Genotype

Promoter

Gene

Wild Type

Control

CRTISO

ccr2

ccr1

Neoxanthin

Violaxanthin

Antheraxanthin

10

13 (60.6)

0.6 (60.1)

Lutein 49 (60.4)

Zeaxanthin

b-carotene

Transgenic Lines

0.5

27 (60.5)

20

CaMV35s

CRTISO

12

13 (60.2)

0.1 (60.0)

49 (60.4)

0.5

25 (60.5)

53

CRTISO-1.9kb

CRTISO

10

16 (61.4)

0.7 (60.1)

45 (62.8)

0.4

27 (61.7)

17

CRTISO-0.5kb

CRTISO

11

15 (61.5)

0.5 (60.1)

47 (60.4)

0.2

29 (60.4)

7

Control

CRTISO

10

31 (61.7)

4.1 (60.6)

18 (62.6)

1.1

36 (60.9)

13

Tmin35

CRTISO

12

31 (60.6)

4.6 (60.4)

17 (60.8)

1.0

35 (60.7)

24

CaMV35s

CRTISO

9

14 (61.2)

0.4 (60.1)

46 (62.0)

1.3

29 (61.0)

23

CRTISO-1.9kb

CRTISO

11

17 (60.8)

0.9 (60.1)

43(61.2)

0.5

30 (60.4)

41

CRTISO-0.5kb

CRTISO

11

14 (60.8)

0.8 (60.2)

44 (61.2)

0.4

29 (60.4)

30 25

Control

CRTISO

10

29 (61.1)

1.7 (60.3)

23 (61.0)

1.0

36 (60.4)

Tmin35

CRTISO

12

25 (62.2)

2.5 (60.3)

26 (62.8)

0.2

34 (60.6)

5

CaMV35s

CRTISO

11

16 (60.5)

0.3 (60.0)

44 (60.7)

0.5

28 (60.4)

63

CRTISO-1.9kb

CRTISO

10

22 (61.3)

1.6 (60.2)

34 (61.7)

0.4

33 (60.4)

37

CRTISO-0.5kb

CRTISO

10

27 (62.2)

3.4 (60.5)

27 (62.1)

0.9

35 (60.6)

14

2.2

0.6

2.8

0.2

1.7

Maximum Standard Error

1.2

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The analysis of SDG8:FiLUC lines revealed that SDG8 may be up-regulated during seedling development (Figure 4). Representative lines were placed into four groups based upon the intensity of luminescence; group 2 and 3 lines switched from low basal luminescence at 5 DAG to stronger luciferase expression at 9 DAG (Figure 4B and 4C). This pattern was also observed in a small subset of SDG8:GUS lines (data not shown). However, a large number of SDG8 promoter lines did not show an up-regulation of GUS expression during seedling development (group 1, Figure 4). The increase in reporter expression enabled by the SDG8 promoter in select lines agrees in part with previous findings that SDG8 transcript levels were low immediately following germination and increased over the first 8 d of seedling development (Kim et al., 2005). We conducted one experiment using slightly stressed seedling tissues showing a small increase in SDG8 and CRTISO transcript accumulation (Figure 5A). However, in a second experiment using healthy seedling tissues, CRTISO and SDG8 transcript levels decreased slightly during development (Figure 5B). This, together with the variability in GUS across different lines, indicates SDG8 can fluctuate during seedling development, perhaps in response to other developmental, biotic, and/or abiotic factors. There was striking overlap in tissue-specific reporter activities enabled by SDG8 and CRTISO promoters, especially in the cotyledons and hypocotyl (Figure 7B and 7J). However, the expression of SDG8 in dormant and germinating seeds did not universally coincide with the detection of CRTISO. Where SDG8 showed visible GUS staining in germinating seeds, CRTISO did not (Figure 7A and 7I). In summary, the co-regulation of CRTISO and SDG8 gene expression (Figure 5A and 5B) and the substantial reduction in CRTISO mRNA levels in ccr1 seedlings (Figure 5D) demonstrate that SDG8 is required to confer transcriptional expression of CRTISO during seedling development.

Organ-Specific Expression of SDG8 and CRTISO Overall, the tissue-specific GUS staining patterns regulated by the CRTISO promoter lines displayed striking overlap with those of SDG8. Most notable was the strong expression observed in the shoot apical meristem, hypocotyl (stem), cotyledons, and leaf vasculature, while more variable and somewhat weaker GUS staining was displayed in mesophyll cells (Figure 7). Why the expression of a carotenoid biosynthetic gene:GUS fusion is stronger in vasculature, seedlings, and shoot apical meristem region than photosynthetic mesophyll cells is interesting, but not without precedent. Indeed, many genes, including a number of high light inducible genes, such as ASCORBATE PEROXIDISE 2 (APX2) and ZAT10, are strongly expressed in the bundle sheath cells, but not in mesophyll (Fryer et al., 2003; Ball et al., 2004; Rossel et al., 2006, 2007). Also, in rapidly growing (e.g. seedlings) and dividing (e.g. shoot apical meristem region) tissues, there may be a greater requirement for carotenoid biosynthesis and photosystem assembly. Also, carotenoid biosynthesis is necessary for

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the production of ABA and strigolactones, which are key phytohormones in plant development and stress response systems. Perhaps the link between the tissue-specificity of SDG8 and CRTISO and their regulation of key phytohormones provides insight into additional functions for carotenoid biosynthesis that have yet to be investigated. One such function is the role of carotenoid biosynthesis in root development. In this study, the promoters of CRTISO and SDG8 enabled weak, yet variable, GUS expression in the primary root, but rarely in the root tip (Figure 7C, 7D, 7K, and 7L). SDG8 has previously been shown to be necessary for regulation of CRTISO gene expression in root tissues (Cazzonelli et al., 2009a) and both CRTISO and SDG8 transcripts are also detectable in the roots (Supplemental Figure 2B). The production of carotenoid-derived signaling hormones such as abscisic acid and strigolactones, as well as the novel graft transmissible b-carotene-derived signaling compound (BYPASS), in root tissues implies that there is a link between the regulation of the carotenoid biosynthetic branch point and normal root and shoot development in Arabidopsis (Figure 1). Given the fact that SDG8 affects root branching (Cazzonelli et al., 2009c) and that roots are a proven source for ABA and strigolactones (Nambara and Marion-Poll, 2005; Van Norman and Sieburth, 2007; Gomez-Roldan et al., 2008; Umehara et al., 2008), future studies will address the role of carotenoid biosynthesis in the near colorless root. Another potential function for carotenoid biosynthesis may be in pollen development and epigenetic programming. The strong co-expression of CRTISO and SDG8 in male reproductive organs was most intriguing. Flowers were dissected into stamens and petals, and relative transcript levels compared to those from leaf tissues (Figure 5C and Supplemental Figure 2B). The strong SDG8 mRNA expression and SDG8:GUS activity in pollen compared to that in leaf and petal tissues is consistent with the partial male sterility and impaired pollen dehiscence of ccr1 mutants, suggesting that SDG8 is both expressed in and required for normal pollen development (Soppe et al., 1999; Xu et al., 2008; Cazzonelli et al., 2009a). This may have important implications for understanding the role of histone methyltransferases in coordinating the memory of epigenetic regulatory events in meristematic tissues such as the apical meristem, as well as the resetting of epigenetic processes, such as vernalization (Sheldon et al., 2008). However, the primary gene targets of SDG8 in pollen are currently unknown, as is the extent to which SDG8 responds to stimuli and contributes to epigenetic reprogramming. There were some similarities and substantial differences in the tissue-specific patterns of expression driven by our SDG8 promoter–reporter gene fusion in comparison to previous reports in which the GUS gene was inserted into the BspEI restriction site of the 14th exon of an SDG8 genomic clone (EFS:GUS) (Kim et al., 2005; Choi et al., 2009). As annotated in TAIR9, the 14th exon of SDG8 can undergo splice variation, that is in some cDNAs, the GUS gene would have been intronic and spliced out. Our analysis of sequenced cDNA clones confirms

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that SDG8 shows splice variants in the 14th exon (present in EU014690, NM_106379 cDNAs and excised in DQ340869 and NM_001084367 cDNAs, see NCBI). It was reported that the EFS:GUS fusion was strongly expressed in the shoot and root apex, as well as the carpels and ovules, but not the stamens (Kim et al., 2005; Choi et al., 2009). Our analysis did not uncover any clear GUS staining in the root tips, carpels, and/or ovules, but we observed strong GUS staining in the pollen (Figure 7). We therefore attribute these discrepancies to be associated with variation in splicing at the 14th exon of the EFS:GUS fusion and/or that there are regulatory cis-elements located outside the promoter region (–1787 bp) characterized.

SDG8, Chromatin, and Regulation of CRTISO CRTISO mRNA, GUS, and luciferase activity were only ever detected in tissues in which SDG8 was also co-expressed. Furthermore, if SDG8 was knocked out, as in ccr1, the expression of CRTISO was almost undetectable in the shoot apex and flower tissues (Figure 5E). Similarly, in developing seedlings, CRTISO transcript levels were almost completely downregulated (.90%) in ccr1 (Figure 5D). This demonstrates that SDG8 is required for permissive regulation of CRTISO in tissues that undergo rapid cell division and differentiation, and highlights the need for SDG8 to maintain transcriptional regulation of CRTISO. The behavior of SDG8 and CRTISO promoter lines in wildtype Columbia was quite unusual in comparison to the CaMV35s and eLCY promoters (Figure 3). While there was large variation in the strength of reporter gene activity observed among lines and in segregating progeny, the tissue-specific patterns driven by multiple lines harboring both the CRTISO and SDG8 promoter–reporter fusions were nonetheless consistent. It was not surprising to observe such variation among the CRTISO promoter lines, as permissive regulation of this gene requires SDG8 activity (Cazzonelli et al., 2009a) and the chromatin state surrounding the site of T-DNA insertion is therefore likely to affect regulation of the CRTISO promoter. It is not readily clear, however, why SDG8 promoter–reporter lines also displayed an unusual variation in luciferase activity levels (Figure 3E and 3F). Perhaps the regulation of SDG8 is also influenced by the chromatin state in which it resides. Analysis of histone-3 lysine-4 methylation patterns in the Arabidopsis epigenome has uncovered significant enhancement of mono-methylation throughout the SDG8 coding regions and localized tri-methylation to the transcriptional start site (Zhang et al., 2009), indicating that SDG8 may also be affected by the chromatin state in which it resides. Future studies investigating epigenetic marks associated with the site of T-DNA integration may uncouple this unusual regulatory behavior displayed by the CRTISO and SDG8 promoters. The CRTISO promoter:GUS activity was determined to be reduced, albeit variably, in the ccr1 mutant compared to wild-type (Figure 8). That is, in comparison to wild-type, the majority of ccr1 lines harboring the CRTISO-1F:GUS fusion displayed weaker GUS staining overall (Figure 8A) and specifically

in the shoot apical meristem region and leaf vasculature (compare Figures 6D and 8B). However, when select lines were crossed to wild-type Columbia, thereby restoring SDG8 function, GUS activity remained unchanged in the shoot apical meristem region and leaf vasculature. In general, the small differences we observed in GUS staining patterns between ccr1 and wild-type plants indicate that target sequences required to enable SDG8 activity are likely to reside not only within, but also outside the region of the CRTISO promoter tested. We tested several promoter–CRTISO gene fusions in an attempt to uncover the molecular nature by which SDG8 maintains transcriptional regulation of CRTISO. As anticipated, the strong and constitutive CaMV35s promoter was sufficient to up-regulate CRTISO transcription by .36-fold and completely restore carotenoid composition in ccr1 and ccr2. In contrast, the minimal CaMV35s core regulatory domain, which can enable basal transcription (Cazzonelli and Velten, 2006, 2008), failed to restore carotenoid levels in both mutants. Interestingly, the CRTISO promoter–gene fusion was insufficient to restore wild-type CRTISO mRNA levels and only partially restored carotenoid composition in ccr1, demonstrating that sequences contained within the CRTISO promoter and gene are necessary for SDG8 activity. In summary, sequence domains required for interacting with SDG8 and/or recruiting other regulatory proteins are likely to be present within –570 bp of the CRTISO promoter and the CRTISO open reading frame. CRTISO functions at the branch point of carotenoid biosynthesis and is emerging as a key regulatory step in the pathway. Evidence is accumulating that it may modulate biosynthesis of signaling compounds and phytohormones. Where sequences within the gene and open reading frame of CRTISO are necessary for SDG8 action, SDG8 also regulates CRTISO expression in a tissue-specific manner. These include rapidly dividing and growing tissues, as well as in photosynthetic and floral male reproductive tissues. Thus, these two genes are expressed in many sites essential for defining plant architecture and development.

METHODS Plant Growth and Mutants Soil-grown plants were incubated at 4C for 2–3 d in the dark before transferring to 16 h of illumination (100–150 lE) and temperature maintained at 21C. Media-grown plants were prepared by sterilizing seeds with 3% HCl/bleach for 3 h, plated onto Murashige and Skoog media (4.4 g L 1 MS salts, 1 MS salts, 0.5 g L 1 sucrose, 0.8% agar) and incubated in the dark for 3 d at 4C before transferring plates to a growth chamber maintained at 21C and illuminated at 150 lE of light for 16 h. All germplasm are in the Arabidopsis thaliana ecotype Columbia (Col-0) background and mutants used in this study include ccr2-1, ccr2-3, and ccr2-5, which are null alleles of CRTISO (Park et al., 2002), as well as ccr1-1 and ccr1-4, which contain lesions in SDG8 (Cazzonelli et al., 2009a).

Cazzonelli et al.

Construction of CRTISO Overexpression and Promoter– Reporter Gene Constructs Plasmid constructs were prepared using standard cloning techniques (Sambrook and Russell, 2001) and appropriate DNA segments sequenced to confirm the final structure. Promoter–reporter gene constructs were designed to test promoter function using stable and/or transient gene expression systems. The length of a promoter sequence is defined from the sequence position upstream from the start codon. Promoter sequences were amplified by PCR using specific oligonucleotides (Supplemental Table 1) and DYNAzyme or Phusion Polymerases (FINNZYMES) according to the manufacturer’s instructions. The forward PCR primer contained a 5’ XbaI restriction enzyme site and the reverse primer generated an NcoI restriction site surrounding the ATG start codon (Supplemental Table 1 and Figure 2A). Promoter fragments for CRTISO-1F (–1977 bp), CRTISO-5F (–570 bp), eLCY-4F (–460 bp), and SDG8-1F (–1787 bp) were PCR amplified and cloned into intermediate vectors, such as pGEM-T (PROMEGA) and the ZERO Blunt TOPO PCR cloning kit (INVITROGEN), following the manufacturer’s instructions. Vectors used during this study are functionally grouped into CRTISO overexpression, promoter–luciferase, and promoter–GUS test constructs (Figure 2B–2D). The CRTISO overexpression constructs, pMDC32:CRTISO and pTCRTISOPromoter1FCRTISOGene (pTCPCG-1F), were constructed as previously described (Cuttriss et al., 2007; Cazzonelli et al., 2009a) (Figure 2B). The minimal CaMV35s promoter (–46 bp) was amplified from pTm35:FiLUC (pPZP200 binary vector harboring an intron containing Firefly luciferase reporter gene under control by the minimal CaMV35S promoter) (Cazzonelli and Velten, 2008; Velten et al., 2008) using TMCS-cc1F/TMCS-cc1R primers, digested and cloned into pTCPCG-1F (digested with XbaI/NcoI) to create pTmin35PromoterCRTISOGene (pTm35-CRTISO) (Figure 2B). pTCRTISOPromoter5FCRTISOGene (pTCPCG-5F) was constructed by amplifying –570 bp of the CRTISO promoter (using CRTISOProm-5F/CRTISOProm-1R), cloning the PCR product into an intermediate vector, and subsequently transferring the promoter sequence to pTCPCG-1F (digested with XbaI/NcoI) (Supplemental Figure 2B). A promoter–luciferase reporter vector (pTm35:FiLUC) was previously designed to quantify promoter enhancer strength in stably transformed plant tissues (Cazzonelli et al., 2005; Velten et al., 2008). pTCRTISO-1F:FiLUC and pTm35enh:FiLUC were constructed as previously described (Cazzonelli and Velten, 2008; Cazzonelli et al., 2009a) (Figure 2D). The CRTISO, eLYC, and SDG8 promoter sequences were transferred from their intermediate vectors into pTm35:FiLUC (digested with XbaI/NcoI to remove the minimal CaMV35s promoter) to create pTCRTISO5F:FiLUC, pTeLCY-4F:FiLUC, and pTSDG8-1F:FiLUC (Supplemental Figure 2DM). pTSDG8-H3:FiLUC was created by digesting pSDG8-1F:FiLUC with HindIII, removing the upstream (–837 to –1787 bp) portion of the SDG8 promoter by QIAEXII agarose gel extraction (QIAGEN) and re-ligating in order to create pTSDG8-H3:FiLUC (–837 bp) (Figure 2D).

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A new promoter–GUS reporter vector was designed to test tissue-specific activity of the SDG8 and CRTISO promoters. The pSYNPRO3–intron-modified b-glucuronidase (GUSi) vector was previously described (Cazzonelli and Velten, 2008) (Figure 2C). pTmin35:GUSi was created by removing the GUSi reporter from pSYNPRO3–GUSi using NcoI/AflII and ligated into pTmin35-FiLUC (digested with NcoI/AflII). pTCRTISO-1F:GUS and pTSDG8-1F:GUS were then created by excising the corresponding promoters from their intermediate vectors using XbaI/NcoI and ligating into pTmin35:GUSi (digested with XbaI/NcoI to remove the min35 fragment) (Figure 2C).

Plant Transformation and Selection The binary vectors were subsequently transformed into Agrobacterium tumefaciens strain LBA4404 by electroporation followed by selection on media containing 50 lg mL 1 kanamycin (pMDC32:CRTISO) or 100 lg mL 1 spectinomycin (pT* family of vectors). Agrobacterium-mediated transformation of wild-type and mutant (ccr1 and ccr2 alleles) Arabidopsis plants was performed according to the floral-dip method (Clough and Bent, 1998). Transformants harboring pMDC32:CaMV35S-CRTISO were selected by plating sterilized seeds on Murashige and Skoog media (4.4 g L 1 MS salts, 1 MS salts, 0.5 g L 1 sucrose, 0.8% agar) containing 50 lg mL hygromycin (Invitrogen) and heterozygous lines were transferred to soil for self-fertilization. Independent transgenic lines transformed with the pT* family of binary vectors (Supplemental Figure 2) were selected by spraying soil-grown seedlings with 50 mg L 1 BASTA (glufosinate-ammonium salt, Sigma-Aldrich).

Determination of Transcription Start Site The CRTISO transcription start site was determined using the GeneRacer kit (Invitrogen) according to the manufacturer’s instructions. PCR was performed using DYNAzyme EXT DNA Polymerase according to the manufacturer’s instructions. The CRTISO 5’ untranslated leader region was amplified using 1 lL cDNA and GeneRacer 5’ Primer/CRTISO-ch4R primer pair (Supplemental Table 1), which was further amplified using a nested primer pair (GeneRacer 5’ Nested Primer/CRTISOch3R) and subsequently cloned into pGEM-T (PROMEGA) and sequenced.

Carotenoid Assays Carotenoid measurements were performed as previously described (Park et al., 2002).

Real-Time Quantitative PCR Total RNA was extracted using the QIAGEN RNeasy or SIGMAaldrich Spectrum kits and included an on-column DNase treatment step using the QIAGEN RNase-Free DNase Set following the manufacturer’s instructions. First-strand cDNA synthesis was performed using Oligo dT primer and SuperScript III Reverse Transcriptase (Invitrogen) according to the manufacturer’s instructions. The relative transcript abundance was

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quantified using LightCycler 480 SYBR Green I Master and three technical replicates for each of one to three biological replicates were performed using the Light Cycler 480 (ROCHE, Australia). All PCR mixtures contained 5 lL of Roche SYBRgreen master mix, 2 lL of each primer (2 lM), and 1 lL of diluted RT product (1–5 ng) in a 384-well optical reaction plate. The take-off point was determined using relative quantification {Target Eff Ct(Wt–target)/Reference Eff Ct(Wt–target)} and fit point analysis (Pfaffl, 2001). Cyclophilin (At2g29960), Protein Phosphatase 2A (At1g13320), TIP41 (At4g34270), and a synthetic internal control (GQ215228) (kindly donated by Dr Jeff Velten) were included as housekeeper reference control genes (Czechowski et al., 2005). Real-time SYBR-green dissociation curves showed one species of amplicon for each primer combination. Primer sequences are listed in Supplemental Table 2. A synthetic internal was utilized to confirm the efficiency of reverse transcription among biological treatments. The internal control (IC) DNA sequence was developed by Dr Jeff Velten (GQ215228) and does not contain sequence homology to any transcribed regions of the Arabidopsis genome. The IC sequence was developed by ligation mediated-PCR using a series of overlapping oligonucleotides, which was then cloned into pUC57 to create pUC57-IC2. IC DNA was PCR amplified using IC2r-T7 (TAATACGACTCACTATAGGGAGATGGTCAGCCTCTAATGGCTCG) and IC2r-pA (AAAAAAAAAAAAAAAAAAAACAAGGAGATCACTGCTTTCG) as primers and TaqF1 DNA Polymerase (Fisher-Biotec) according to the manufacturer’s instructions. In vitro transcription of the IC DNA fragment by T7 RNA polymerase was achieved by using Ambion’s MEGAscript Kit as per instructions, DNA removed by DNase treatment, and mRNA purified using Qiagen’s RNeasy spin column. IC mRNA was added prior to reverse transcription at a final concentration of 1 ng lL 1.

b-Glucuronidase (GUS) Reporter Gene Assays GUS histochemical assays were performed as previously described, with minor modifications (Jefferson et al., 1987). Plant tissues or intact seedlings were vacuum infiltrated for 5–20 min in a reaction mixture containing 50 mM Na2HPO4 pH 7.0, 0.5% Triton X-100, 10 mM EDTA, 0.5 mM K4F3(CN)6, 0.5 mM K3Fe3(CN)6:H2O, and 2 mM X-Gluc dissolved in DMSO. Infiltrated tissues were incubated in the GUS reaction media for 24 h at 37C or until the blue indigo dye precipitate was observed. Immediately following the GUS assay, the tissues were cleared by washing in a series of ethanol steps and stored in 70% ethanol. Tissue sections were imaged using an Olympus SZX9 microscope and photographed with an Olympus DP70 camera.

Luciferase Assays Quantitative measurement of luciferase activity in leaf tissues from transgenic lines was achieved using the in vivo floating leaf-disk assay (Cazzonelli and Velten, 2006). Multiple independent lines were assayed (two leaf disks per hemizygous or homozygous plant, average of two to five plants per line).

Individual disks were floated on Luciferase Assay Media (LAM: 1 mM luciferin, 50 mM 2-(N-Morpholino)ethanesulfonic acid (MES) (pH 5.6), 0.5% glucose, 2 mM NaPO4, 0.5% v/v DMSO) in separate wells of a 96-well, white-walled, microtiter plate and light production measured over time using a FLUOstar Optima luminometer from BMG Lab Technologies Inc. The peak of light emission over a 90-min time course was determined for each sample and referred to as the relative light units (RLU) emitted (Velten et al., 2008). RLU values were usually 10–100 000-fold over background (disks from wild-type Arabidopsis leaves). Semi-quantitative in planta bioluminescence assays were performed using a luminescence CCD camera (Model DV435 by Andor Technologies, Japan) and ImagePro Plus 4.5.1 software (Media Cybernatics, CA, USA). In order to minimize substrate uptake artifacts associated with the spraying of luciferin on mature plants, individual tissues such as rosette/cauline leaves, floral stems, flowers, and roots from multiple plants of an independent line were combined and submerged in an aqueous solution containing 0.4 mM luciferin and 100 mM sodium citrate (pH ; 5.6) as previously described (Velten et al., 2008). Photon counting (exposures of 300 and 600 sec) was performed after 2 min of pre-incubation (at ambient temperature in the dark). Under these conditions, control untransformed Arabidopsis plants display no detectable bioluminescence. Image J software (version 1.32) was used to visually enhance different light intensities using a false color display (threshold settings are displayed with each image).

Accession Numbers Arabidopsis Genome Initiative locus identifiers for the genes mentioned in this article are as follows: At1g77300 (SDG8), At1g06810 (CYCLOPHILIN), At4g34270 (TIP41-like protein), At1g13320 (PROTEIN PHOSPHATASE 2A), At1g06820 (CRTISO), At5g57030 (eLCY).

SUPPLEMENTARY DATA Supplementary Data are available at Molecular Plant Online.

FUNDING We were supported by the Australian Research Council Centre of Excellence in Plant Energy Biology (CE0561495).

ACKNOWLEDGMENTS We thank Jean Finnegan (CSIRO, PI) for providing the GeneRacer Kit to determine the CRTISO transcription start site. We thank Dr Jeff Velten (USDA-ARS) for kindly providing pUC57-IC2 (RT–qPCR synthetic internal housekeeper gene). Many thanks to Derek Collinge, Kuide Yin, Stephanie Webb, Alice Burgess, and Peter Crisp for their valuable contributions. No conflict of interest declared.

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