Journal of Experimental Botany, Vol. 53, No. 373, pp. 1397-1409,
June 2002
© 2002 Oxford University Press
Original Papers |
Expression of the grape dihydroflavonol reductase gene and analysis of its promoter region1
Department of Fruit Tree Breeding and Molecular Genetics, ARO, The Volcani Center, PO Box 6, Bet-Dagan 50250, Israel
Received 6 July 2001; Accepted 22 January 2002
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Abstract |
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Dihydroflavonol reductase (DFR) is a key enzyme involved in anthocyanin biosynthesis and proanthocyanidin synthesis in grape. DFR catalyses the reduction of dihydroflavonols to leucoanthocyanidins in the anthocyanin pathway. The DFR products, the leucoanthocyanidins, are substrates for the next step in the anthocyanin pathway and are also the substrates for the proanthocyanidin pathway. In the present study the promoter of the grape dfr gene was cloned. Analysis of the dfr promoter sequence revealed the existence of several putative DNA binding motifs. The dfr promoter was fused to the uidA gene and the control of this fusion and the endogenous dfr gene expression, was studied in transformed plants and in red cell suspension originated from fruits. The dfr promoter–uidA gene fusion was expressed in leaves, roots and stems. Deletions of the dfr promoter influenced the specificity of the expression of the GUS gene fusion in plantlet roots and the level of expression in plants and in the red cell suspension originated from fruits. The deletion analysis of the dfr promoter suggests that a specific sequence located between -725 to -233 might be involved in expression of the dfr gene in fruits. Light, calcium and sucrose induced the dfr gene expression. In the transformed suspension cultures, expression of both the endogenous dfr gene and the dfr promoter–uidA gene fusions was induced by white light. The induction by both light and calcium suggests the possible involvement of a UV receptors signal transduction pathway in the induction of the dfr gene. The induction of the dfr gene and the dfr promoter–uidA gene fusions by light and sucrose indicates a close interaction between sucrose and light signalling pathways.
Key words: Developmental expression, dfr, grape, light signalling, sucrose signalling.
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Introduction |
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Flavonoids are involved in many plant functions, such as pigmentation, UV protection and defence against pathogens (Koes et al., 1994
). The anthocyanins, which are flavonoids, are important plant pigments, responsible for most of the brick red, red and blue colours in plants (Holton and Cornish, 1995). The synthesis of anthocyanin biosynthesis pathway enzymes is under developmental and tissue-specific control (Van Tunen et al., 1988). The genes in this pathway are induced by environmental factors such as UV light (Christie and Jenkins, 1996; Frohnmeyer et al., 1997), red light (Bowler and Chua, 1994) and pathogens (Lamb et al., 1989). Other factors, such as salts and carbon source, also influence anthocyanin biosynthesis (Seitz and Hinderer, 1988). High sucrose and low phosphate and nitrate concentrations induced anthocyanin formation in Vitis cell suspension cultures (Yamakawa et al., 1983).
Light controls a wide range of aspects of plant growth, development and gene expression. Three different photoreceptor systems mediate the effects of light: the phytochromes, blue/UV-A photoreceptors and UV-B photoreceptors (Khurana et al., 1998). Phytochrome A (PHY A) controls anthocyanin synthesis and chloroplast development. In the PHY A signal transduction pathway anthocyanin synthesis and chalcone synthase (chs) expression have been shown to be positively regulated by cGMP and negatively by calcium and calmodulin (Bowler et al., 1994a, b). The blue/UV-A and UV-B photoreceptors also induce chs expression. Calcium and calmodulin, but not cGMP, have been shown to be involved in this induction (Christie and Jenkins, 1996; Frohnmeyer et al., 1997).
The colour of red and black grapes results from the accumulation of anthocyanins that are usually located in the berry skin (Boss et al., 1996a). Anthocyanin biosynthesis commences when berry ripening begins and sugar accumulation has just begun (Boss et al., 1996b). The major genes for the enzymes in the anthocyanin biosynthesis pathway have been cloned in grape (Sparvoli et al., 1994), and the spatial and temporal pattern of expression of these genes in grape has been studied (Boss et al., 1996a, b, c).
Dihydroflavonol reductase (DFR) is the enzyme which catalyses the reduction of dihydroflavonols to leucoanthocyanidins in the anthocyanin pathway (Dooner et al., 1991; Sparvoli et al., 1994; Holton and Cornish, 1995). The DFR products, the leucoanthocyanidins, are substrates for the next step in the anthocyanin pathway and are also converted into proanthocyanidin in a side pathway (Boss et al., 1996a, c). Proanthocyanidins, which include tannins, are important constituents of wine. Since DFR is a key enzyme in this part of the pathway in grape, the regulation of the expression of the dfr gene was studied. The grape dfr gene was shown to be expressed in all the plant organs: leaves, tendrils, green cane, root, seeds, flowers, berry skin, and berry flesh. The dfr is one of the most strongly expressed anthocyanin genes in all tissues (Boss et al., 1996c) and it is the most highly expressed anthocyanin gene in the berry flesh (Boss et al., 1996b). In the berry skin the dfr gene has two phases of expression, similar to the expression of other anthocyanin genes (Boss et al., 1996b).
Previously published studies on light signal transduction pathways, which effect the expression of anthocyanin genes, have addressed only chs. The regulation of chs, the first gene in the anthocyanin pathway, by light has been studied extensively in various plant species (Mol et al., 1996; Schafer et al., 1997). In this manuscript the study of the control of expression of the dfr gene in grape is reported. The influence of light, calcium and sucrose on the expression of the dfr gene in grape was studied.
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Genomic library construction and screening
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FixII (Stratagene, La Jolla, CA) library was constructed from Vitis vinifera cv. Red Globe genomic DNA, partially digested by Sau3A, according to the manufacturer's instruction manual. The amplified library was screened at high stringency, as described below, with a 32P-labelled probe synthesized from the 5'-end of the dfr coding sequence. An isolated 2659 bp BamHI genomic fragment was cloned into pUC19 for further analysis.
Isolation and analysis of nucleic acids
Grape genomic DNA was isolated (according to the method of Steenkamp et al., 1994). Total RNA was isolated (Chang et al., 1993), and 20–25 µg RNA were fractionated on 1% formaldehyde agarose gels before blotting. DNA and RNA were blotted on nitrocellulose and were hybridized to 32P-labelled gene specific-probes for dfr and uidA. Gene-specific fragments were amplified by PCR. A 5'-dfr specific fragment was amplified by PCR using V. vinifera cv. Red Globe genomic DNA and primers synthesized according to the gene bank sequence of the dfr cDNA. The 3'-end of this fragment was 5 bp downstream to the BamHI site in the cDNA sequence (Fig. 1). The fragments were gel-purified before labelling. Hybridizations were performed at 37 °C in 0.12 M sodium phosphate buffer pH 6.7, 1% SDS, 0.5 M Na+, and 50% (v/v) formamide. Filters were washed at high stringency and autoradiographed on Kodak X-Omat film.
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dfr promoter constructs
The 2659 bp BamHI genomic fragment, which was cloned into pUC19, was sequenced and its restriction map was analysed. The dfr promoter fragment and 5'-deletions of the promoter fragment were further cloned into the Agrobacterium binary vector pBI101, in which the original uidA gene was replaced with uidA gene containing the IV2 plant intron of the ST-LS1 gene from the p35SGUSint vector (Vancanneyt et al., 1990). The introduction of the uidA-intron gene excluded expression of the fused promoter–uidA gene in Agrobacterium, thus eliminating possible background. This vector was designated as pBI101–GUSInt.
Plant materials and cell growth
Red cell suspensions of V. vinifera cv. Gamay Red were derived from red fruits. Young berries at a size of 1 cm were disinfected three times, sliced to pieces of 1–2 mm and were put on solid B5 medium (Gamborg et al., 1968) supplemented with 0.2 mg l-1 kinetin and 0.1 mg l-1 
-naphthaleneacetic acid (NAA), 2% sucrose, 250 mg l-1 casein hydrolysate, 100 mg l-1 myo-inositol, pH 5.9. The callus developed is producing anthocyanin and the colour of this callus is red. Cell suspensions derived from the callus obtained from fruits of V. vinifera cv. Gamay Red were grown in B5 medium supplemented with the compounds indicated above. Cultures were maintained in white light at a fluence rate of 20 µmol m-2 s-1 on a gyratory shaker at 85 rpm and 25 °C. Transformant clones were grown in the presence of 40 mg l-1 paromomycin sulphate and 300 mg l-1 cefotaxime sodium. Cell suspension cultures were transferred weekly to a fresh medium. Embryogenic suspensions were derived from anthers of V. vinifera cv. Superior Seedless. Culture conditions were as previously described (Perl et al., 1996; Lipsky et al., 1997). Transformed Superior seedless plantlets and plants were maintained under white light at fluence rate of 50 µmol m-2 s-1.
Agrobacterium-mediated transformation
Transformations were performed to Gamay Red cell suspensions derived from fruits and to embryogenic Superior Seedless cultures. A single colony of Agrobacterium tumefaciens strain EHA105 harbouring a plasmid was grown for 24 h in YEB medium (pH 7) (Vervliet et al., 1975) at 28 °C and 250 rpm to a density of 1 OD600 in the presence of 50 µg ml-1 kanamycin and 25 µg ml-1 rifampicin. Bacterial cells were precipitated by centrifugation at 4130 g for 5 min, resuspended and subcultured in YEB medium (pH 5.6) in the presence of 100 µM acetosyringone, and incubated for 2 h. For inoculation of plant material, bacteria were precipitated and resuspended in NN medium (Nitsch and Nitsch, 1969) pH 5.8, supplemented with 2% sucrose. Co-cultivation was performed on solid NN medium for 3 d in the dark. Antioxidants were included in the co-cultivation and in the selection media as described previously (Perl et al., 1996). A selection of transformed Superior Seedless calli was started in hormone-free MG liquid medium (Lipsky et al., 1997) supplemented with 300 mg l-1 cefotaxime sodium and 5 mg l-1 paromonycin sulphate. Putative transformed calli were subcultured every 3–5 d in fresh MG medium supplemented with increasing concentrations of paromomycin sulphate up to 40 mg l-1. Plants were regenerated on McCown woody plant medium (Lloyd and McCown, 1980) supplemented with 2% sucrose, 100 mg l-1 myo-inositol, 0.2 mg l-1 indole 3-butyric acid (IBA), 0.25% phytagelTM (Sigma), 0.2% activated charcoal, pH 5.8. Gamay Red cells were selected in solid B5 medium, as described, supplemented with 300 mg l-1 cefotaxime sodium and 20 mg l-1 paromomycin sulphate. Transformed cells was further transferred to fresh B5 medium supplemented with 300 mg l-1 cefotaxime sodium and 30 mg l-1 paromomycin sulphate. Stable transformed lines were finally cultured throughout the study on B5 medium supplemented with 300 mg l-1 cefotaxime sodium and 40 mg l-1 paromomycin sulphate.
ß-glucoronidase activity analysis
ß-glucoronidase (GUS) activity was analysed according to Jefferson et al. (Jefferson et al., 1987) with modifications, using 5-bromo-4-chloro-3 indolyl-glucoronide (X-Gluc, Duchefa Biochemicals, Netherlands) as a substrate for the histochemical assay and 4-methyl umbelliferyl glucoronide (MUG, Sigma, USA) as a substrate for the fluorimetric assay. V. vinifera cv. Gamay Red cells were stained overnight in the presence of 0.1 M sodium phosphate buffer, pH 7, 50 µM potassium ferricyanide, 50 µM potassium ferrocyanide, 1 mg ml-1 of X-Gluc and 0.1% Triton X-100 at 25 °C. Plants and plantlets were stained overnight at 37 °C in solution containing 0.1 M sodium phosphate buffer, pH 7, 0.25 mM potassium ferricyanide, 0.25 mM potassium ferrocyanide, 10 mM Na2EDTA, 1 mg ml-1 X-Gluc, and 0.1% Triton X-100, following a 30 min vacuum treatment in the staining solution. For the fluorimetric GUS assay, Gamay Red cells were lysed in extraction buffer (Jefferson et al., 1987). Extracts were kept at -70 °C and analysed for activity using 3.5 µg protein in 200 µl extraction buffer in the presence of 1 mM MUG at 37 °C for 1 h. Fifty microlitre samples were removed for analysis at different time points, and the reaction was terminated with 950 µl of 0.2 M Na2CO3. Fluorescence was measured with excitation at 365 nm, emission at 455 nm on a Shimadzu spectrofluorophotometer RF-540. Calibrations were done against MU standards.
Light conditions and analysis of expression
Cultures of the V. vinifera cv. Gamay Red clones were grown in B5 medium as described above, under 20 µmol m-2 s-1 white light (F36W/54/DL; GE). All the expression experiments (Figs 4, 5, 6, 7) were performed under these light conditions. For the light induction experiment (Fig. 5) cultures grown for 5 d in light were aliquoted and part of the culture continued to grow under light, while the rest of the culture was dark adapted for 3 d after which, part of the dark-adapted cultures were exposed again to light. For the calcium and sucrose experiments, cultures grown under white light in B5 medium were diluted in B5 medium containing various concentrations of calcium (1–100 mM) or various concentrations of sucrose (20–60 g l-1). The basal concentration of calcium and sucrose in B5 medium is 1 mM and 20 g l-1, respectively. The cultures were grown for 7 d under white light and analysed. All manipulations were performed under green safe light. Samples were collected on Whatmann filters by suction filtration and immediately frozen in liquid nitrogen. RNA was isolated from the various cultures, and analysed by hybridization to uidA and dfr specific probes.
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Autoradiograms, and their related ethidium bromide-stained gels, were scanned and the signal intensities were analysed with TINA software. To normalize the intensity values of the hybridization signals (expressed in OD units), they were divided by the intensity values of the total rRNA (also expressed in OD) in the respective lane in the ethidium bromide-stained gel. The resulting ratios allowed comparison between the intensities of different hybridization signals within the same autoradiogram.
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Results |
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Isolation and cloning of the dfr promoter
The copy number of the dfr gene was determined by Southern blot analysis. Genomic DNA from V. vinifera cv. Red Globe, from which the
FixII genomic library was prepared, was digested by different restriction enzymes and hybridized to the dfr specific probe. The results indicated one copy gene for dfr and the hybridization of a 2.7 kb BamHI fragment. Screening of the Red Globe genomic library with the dfr probe at high stringency resulted in the isolation of a clone containing the 2.7 kb BamHI fragment. This BamHI fragment was cloned into pUC19, mapped for restriction endonuclease sites and sequenced (Fig. 1
). The sequence of the 3'-end of this fragment was identical to the 5'-end of the previously published sequence of the dfr cDNA, but included a 100 bp intron (Fig. 1). Putative TATA box and transcription initiation site (Joshi, 1987) were identified and are indicated in Fig. 1. Analysis of the dfr promoter sequence by the TFsearch ver. 1.3 program (Yutaka Akiyama, 1995, Bioinformatics Center, Institute for Chemical Research, Kyoto University, available from the website at http://www.genom.ad.jp) with threshold of 80.0 points and also by the Bestfit computer program, revealed the existence of putative unit I, which includes the G-box factor (GBF) homologous binding site and the downstream MYB homologous binding site (Grotewold et al., 1994). Unit I was initially identified in the chs promoter and shown to be involved in light induction of the gene (Schulze-Lefert et al., 1989; Schafer et al., 1997; Hartman et al., 1998). Unit I was shown to be necessary and sufficient to confer photoregulation through the phytochromes and the UV-A/blue and UV-B photoreceptors (Weisshaar et al., 1991; Rocholl et al., 1994; Kaiser et al., 1995). Unit I can bind transcription factors of the bZip (GBF) and MYB families (Sablowski et al., 1994; Harter et al., 1994; Giuliano et al., 1988). Another putative MYB homologous binding site is located upstream to the G-box (Fig. 1). The TACCAT sequence located in the promoter was identified as a cis-acting element involved in the control of organ-specific expression (van der Meer et al., 1992). Two putative BoxII cis-element and four homologous SBF-1 transcription factor binding sites (Lawton et al., 1991) also exist in the dfr promoter. Two of the SBF-1 sites are highly homologous to the two BoxII sequences, respectively. In addition, putative sucrose box 2 and sucrose box 3 (Tsukaya et al., 1991) exist in the promoter. Interestingly, an AGAMOUS homologous binding site (Huang et al., 1993) was identified in the dfr promoter (Fig. 1).
For further analysis of the promoter, three promoter fragments were isolated: a 2259 bp BamHI–NciI fragment, a 757 bp HindIII–NciI fragment and a 265 bp XbaI–NciI fragment (Fig. 1). The fragments were blunt-ended at their 3'-end by Klenow fragment enzyme and fused to the uidA gene by cloning into the Agrobacterium binary vector pBI101–GUSint. The isolated fragments were ligated in the vector to the SmaI site at their 3'-end and to the BamHI, HindIII and XbaI sites, respectively, at their 5'-end. All three fragments end at their 3'-end at nucleotide +32 from the putative transcription initiation site.
Expression of the dfr promoter–uidA fusions in plantlets and plants
The full-length dfr promoter–uidA gene fusion (-2227/+32) and the deletion fusions (-725/+32 and -233/+32) were introduced by Agrobacterium transformation into embryogenic suspension derived from anthers of V. vinifera cv. Superior Seedless. Figure 2 shows the expression of uidA, analysed by GUS activity, in primary transformants of all three dfr promoter fusions. The full-length dfr promoter fragment (-2227/+32)–uidA fusion was expressed in all regions of plantlets transformants, the -725/+32 deletion fusion was expressed in leaves and hypocotyl, but its expression was very limited in the plantlet roots; it was expressed only in the root tip and in the root vascular tissue (Fig. 2A). The -233/+32 short deletion was expressed in leaves and hypocotyl, but not in the plantlet roots (Fig. 2A). This shows that sequences between positions -2227 and -233 are involved in regulation of dfr expression in the plantlet roots. Expression of all fusions was seen in Superior Seedless plants (Fig. 2B). The uidA gene was expressed in leaves, stem and roots in all three fusions. In plants of the full-length promoter fusion (-2227/+32) and of the -725/+32 deletion fusion, the gene was expressed in leaves, stems and roots of transformant plants, but a slight reduction in staining intensity was observed in the -725/+32 deletion fusion. In the population of plants of the -233/+32 deletion, staining was apparent in leaves, stems and roots, but individual plants were not completely stained. Staining was apparent in young leaves mostly, roots and stems were less stained and older leaves in particular were usually not stained. This pattern of expression of the -233/+32 deletion fusion is different from the expression pattern of the full-length promoter fusion (–2227/+32) and the -725/+32 deletion fusion. The results show that a sequence involved in developmental expression of the dfr gene was removed by deleting sequences between positions -725 and -233.
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Expression of the dfr promoter–uidA fusions in the red cell suspension originated from fruits
The three dfr promoter fusions were also transformed to the red cell suspension derived from the Gamay Red cultivar fruits. GUS staining of transformants of all three fusions is shown in Fig. 3A. High intensity GUS staining was observed in cell suspensions of the full-length (-2227/+32) promoter fusion and less in the -725/+32 deletion fusion. In the -233/+32 deletion fusion, there was no staining or very low staining was seen rarely only in few cells in the cell population (Fig. 3A, indicated by arrows; this phenomenon should be studied in the future). To verify that the -233/+32 deletion clones are positive clones and that the lack of expression is a result of the deletion, genomic DNA isolated from different deletion clones was digested by SnaBI, which is unique restriction site in the uidA gene, run on a gel, blotted and hybridized to the uidA-specific probe. The results (Fig. 3B) indicated that the -233/+32 fusion clones are positive clones and expression is absent in this deletion fusion.
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Expression level of the fusion constructs
To analyse the expression, Gamay Red cell suspensions of clones of the various fusions were grown in B5 medium as described in the Materials and methods, RNA was isolated, blotted and hybridized to the uidA probe and the dfr probe (Fig. 4A). Expression of the uidA gene was seen in clones of the -2227/+32 fusion and the -725/+32 fusion, but there was no expression of the uidA gene in the -233/+32 fusion. In contrast, the dfr endogenous gene was expressed in all fusions. To quantify the expression, GUS activity in all fusions was analysed by GUS fluorescence (Fig. 4B). The GUS activity in the -2227/+32 fusion clones was 1.7 times higher than in the -725/+32 fusion clones and 17 times higher than in -233/+32 fusion clones. The results of the expression in the transcription level and the GUS activity measured by fluorescence are in agreement with the staining pattern seen in the histochemical staining of the Gamay Red cell suspensions of the various fusion clones.
Differential expression of the -233/+32 deletion
Deletion to -233, was expressed in plantlets and plants (Fig. 2), whereas, in contrast to these results, this deletion inactivated all promoter activity in the red cell suspension originated from fruits (Fig. 3A). Although unit I exists in the -233/+32 deletion fusion there was no expression of this deletion fusion in the red cell suspension originated from fruits. The inactivation of the promoter in the red cell suspension suggests that a sequence that is important for the dfr gene expression in the red cell suspension is missing in the -233/+32 deletion fusion, but the deletion of this sequence does not block the expression of the -233/+32 deletion in plantlets and plants.
Light-induced expression of the dfr gene
Since light is involved in anthocyanin biosynthesis in various plant species, the effect of light on the expression of the dfr gene and the dfr promoter–uidA fusion was studied. Experiments were performed with the red cell suspension cultures, to simplify the experiments. Transformant clones of the full-length (-2227/+32) fusion and the -725/+32 and the -233/+32 deletion fusions of the Gamay Red cultivar were grown for 5 d under white light, then dark adapted for 3 d and exposed to light after dark adaptation. Analysis of RNA isolated from the different cultures by Northern hybridization is shown in Fig. 5. The results show that the dfr gene was expressed in light. In dark-adapted cultures its expression was reduced and the gene was induced by light after dark adaptation and exposure to light for 6 h. The results were similar for both the fused dfr promoter–uidA gene and the endogenous dfr gene. Both the -2227/+32 and the -725/+32 promoter fusions were induced by light (Fig. 5), while the -233/+32 deletion fusion was not expressed in the Gamay Red cultivar. By contrast, the endogenous dfr gene was expressed in all analysed clones of the three fusion constructs and was light induced.
The dfr gene expression is induced by increased external Ca2+
Since there is divergence between the PHY A model, based on studies in tomato and soybean (Bowler et al., 1994a, b), and the blue/UV receptors model, based on studies in Arabidopsis and parsley (Christie and Jenkins, 1996; Frohnmeyer et al., 1997), concerning the effect of Ca2+ on the expression of anthocyanin genes, the effect of calcium on the expression of the dfr gene was studied in grape. The effect of external Ca2+ on the expression of the dfr promoter–uidA gene fusion and the dfr gene in cultures of the -2227/+32 dfr promoter fusion clones was analysed. The results show induced expression of the uidA and the dfr genes in the presence of high external Ca2+ (Fig. 6). Both the dfr gene and the uidA gene are induced by Ca2+ and maximal induction is observed in 50 mM Ca2+. Normalization of the radioactive signal intensity to the total rRNA in each lane show the pattern of induction of the dfr gene by Ca2+. The expression of the dfr gene is low in low concentrations of Ca2+ and it is maximally induced (24-fold) in 50 mM Ca2+. There is an increase in induction of the gene by Ca2+ up to 50 mM and reduction in expression at 100 mM, but the gene is still induced in 100 mM Ca2+ (9-fold) compared to the lowest concentration (1 mM) of Ca2+. To confirm that the induction by Ca2+ is specific, the effect of both Ca2+ and Mg2+ on the expression of the dfr gene was analysed. The dfr gene was induced by Ca2+ but not by Mg2+ (data not shown), indicating that the effect of Ca2+ on the expression is specific.
The dfr gene expression is induced by sucrose
Anthocyanin biosynthesis has been shown to be induced in Vitis in the presence of sucrose (Yamakawa et al., 1983). To determine the effect of sucrose on the expression of the dfr gene, cv. Gamay Red cultures of the -2227/+32 dfr promoter fusion were grown in the presence of different sucrose concentrations. Analysis of RNA isolated from these cultures showed that the dfr promoter–uidA gene fusion and the dfr gene were both induced by sucrose. Maximum expression was obtained in the presence of 30 g l-1 sucrose (Fig. 7). Normalization of the radioactive signal to the total rRNA in each lane shows that there is about 2.5-fold induction of the dfr gene by 30 g l-1 sucrose compared to 20 g l-1 sucrose and with higher sucrose concentrations expression was reduced. To determine whether this effect is unique to sucrose, -2227/+32 fusion clones were grown in the presence of sucrose for 6 d, after which the cultures were incubated in a fresh medium containing various sugars: sucrose, glucose, fructose, maltose, and mannitol and analysed for GUS activity. The analysis of GUS fluorescence activity showed that expression of the fused uidA gene was also induced by glucose and fructose, but not by maltose and mannitol (data not shown). The results of the induction of the dfr gene by sucrose are supported by the presence of sucrose box 2 and box 3 in the dfr promoter (Fig. 1).
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Discussion |
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Anthocyanin biosynthesis pathway and its regulation is one of the best characterized secondary metabolite pathways in plants, but promoter analysis was concentrated in the regulation of the early part of the pathway and to the chs gene. It is also clear that in dicot plants the early and the late part of the pathway (of which dfr is a representative) are regulated separately (Mol et al., 1996
), and it is thus well justified to analyse the structure of the dfr promoter reported here.
A central problem in research on plant gene regulation is the mechanism of tissue- or cell-type-specific gene expression. Using the dfr promoter and promoter deletion fusions the expression of the dfr gene was analysed in grape plantlets and plants, and in grape red cell suspension originated from fruits. The full-length promoter fusion was expressed in all regions of plantlets and plants and in the red cell suspension originated from fruits. These results are in agreement with the dfr gene expression in Shiraze grapevine tissues (Boss et al., 1996b). Deletion to -725, resulted in a slight reduction of GUS staining in plants and a reduction of staining in the Gamay Red cell suspension originated from fruits. These findings were consistent with the 40% decrease in GUS fluorescence activity and the reduction in the mRNA transcript in the red cell suspension. Deletion to -233 resulted in limited expression in plants, but did not change the specificity of expression. In contrast to these results, the specificity of the dfr gene expression in the plantlet roots was affected by the deletion of sequences upstream to -233. Deletion to -725 resulted in limited expression in the plantlet roots and deletion to -233, resulted in complete loss of expression in the plantlet roots. This shows that sequences between positions -2227 and -233 are involved in dfr expression in plantlet roots. Possible involvement of the SBF-1 sites and the BoxII sequences can be suggested. The stepwise deletion of the SBF-1 sites and the BoxII sequences may explain these results. Deletion to -725 removed two of the SBF-1 and one of the BoxII sequences. Deletion to -233 removed all four SBF-1 sites and both BoxII sequences.
Deletion to -233 inactivated all promoter activity in the red cell suspension that is originated from fruits in contrast to the results in plantlets and plants. Although unit I and MYB homologous binding site are present in the -233/+32 deletion fusion, there was no expression of this deletion fusion in the red cell suspension originated from fruits, in contrast to its expression in plantlets and plants. This results show that sequences between position -725 and -233 are important for the expression of the dfr gene in the red cell suspension. The inactivation of the -233/+32 promoter fusion in the red cell suspension suggests the possibility that a specific sequence, which is specifically important for the expression in the red cell suspension, is absent from this deletion fusion. Since the origin of these cells is red fruits and since this cell suspension kept at least some of the characteristics of fruit cells, such as their red colour which reflects their anthocyanin production and their anthocyanin genes expression, it is very possible that this sequence is important for the expression in this type of cells and not in plantlets and plants. It would be possible to analyse this phenomenon only in fruits of mature transgenic plants. One possible sequence which might be involved in the dfr gene expression in fruits is the AGAMOUS homologous binding site (Huang et al., 1993), which is located between position -725 and -233. The Arabidopsis floral homeotic gene AGAMOUS is required for normal flower development (Bowman et al., 1989, 1991; Yanofsky et al., 1990). On the other hand TAG1, a tomato homologue of AGAMOUS has been shown to have a role in fruit development in addition to its role in floral meristem determinacy and floral organ identity (Ishida et al., 1998). The possibility for a role of a MADS box protein, an AGAMOUS-related protein (Richmann and Meyerowitz, 1997; Yanofsky et al., 1990; Ishida et al., 1998), in the expression of the dfr gene in grape fruits should be studied in future in fruits of mature transgenic plants expressing the -233/+32 deletion and as a control in fruits of mature transgenic plants expressing the full length fusion. In addition, the possible involvement of the homologous AGAMOUS binding site should be analysed by a specific deletion or mutation of this AGAMOUS binding site.
Studies on the regulation of the dfr gene by light are rare. This study is the first study, as far as is known, involving the effect of light and calcium on the dfr gene in perennial fruit crops. Both the fused promoter and the endogenous dfr gene were analysed to verify that the fused promoter and the dfr gene are expressed similarly. Both the uidA and the dfr gene were induced by low fluence white light after dark adaptation of the red cell suspension cultures of the full length promoter fusion clones. The genes were induced also by calcium in cultures growing under low fluence white light. Low fluence white light was sufficient to induce the dfr promoter–uidA gene fusion and the endogenous dfr gene in the red cell suspension. The induction of the fusion and the dfr gene in the red cell suspension by white light is consistent with the reported induction of anthocyanin genes under white light in grape seedlings (Sparvoli et al., 1994).
Involvement of different light qualities in the chs gene expression has been shown in different plant species. Under white light and red light chs gene expression is positively controlled by cGMP and negatively by calcium and calmodulin through the PHY A signal transduction pathway (Bowler et al., 1994a, b; Frohnmeyer et al., 1998). On the other hand, under UV-A/blue light and UV-B light, calcium and calmodulin are positive regulators of chs gene expression and cGMP is not involved (Christie and Jenkins, 1996; Frohnmeyer et al., 1997, 1998). Both light and calcium induced the grape dfr gene in this study. Since the white light used included UV-A and blue light, the possibility that the grape dfr gene was induced via the UV-A/blue receptor signal transduction pathway should be considered. These results are also consistent with the findings that anthocyanin accumulation in grape seedlings under blue light was similar to the accumulation under white light and that anthocyanin content was 10 times higher under blue light and white light than under red and far-red light (Sparvoli et al., 1994).
Since calcium has been shown to be involved in chs expression through the UV-A/blue and UV-B signal transduction pathways (Christie and Jenkins, 1996; Frohnmeyer et al., 1997, 1998), the effect of calcium on the expression of the dfr gene was analysed, and it was found that high external calcium induced the grape dfr gene. High concentration (100 mM) of external Ca2+ was also reported to induce gene expression of the TCH genes in Arabidopsis root cell suspension (Braam, 1992). High external calcium is known to lead to increase in internal calcium concentration (Hepler and Wayne, 1985; Poovaiah and Reddy, 1987) and, normally, intracellular Ca2+ concentrations of 1–10 µM are needed to activate various processes. The results of this study show for the first time that external calcium influences anthocyanin gene expression under white light. By contrast, the artificial elevation of cytosolic calcium by using calcium ionophores in the presence of calcium was insufficient to increase the chs transcript level under low white light (Christie and Jenkins, 1996). The positive regulation of anthocyanin gene expression by calcium under UV-light was shown by pharmacological studies of the chs gene expression in Arabidopsis and soybean cell suspension culture and in parsley protoplasts (Christie and Jenkins, 1996; Frohnmeyer et al., 1997, 1998). The induction of the grape dfr gene by white light that included UV-A/blue light and by calcium under white light, suggests the possible involvement of UV receptors signal transduction pathway in the expression of the dfr gene. Further studies of the dfr gene expression by pharmacological experiments might clarify this possibility.
Anthocyanin synthesis is induced by light and high sucrose in Vitis (Yamakawa et al., 1983). The effect of sucrose on the dfr expression in red cell suspensions of the full-length promoter fusion clones was examined. Sucrose induced the expression of the dfr gene and of the dfr promoter–uidA fusion. The grape dfr gene is expressed in various plant organs, including the berry skin and the berry flesh (Boss et al., 1996c). Correlation between anthocyanin accumulation, increase in the sugar content of the berries and expression of the anthocyanin genes, including the dfr gene, in grape has been reported (Boss et al., 1996b; Davis and Robinson, 1996). Glucose and fructose constitute most of the soluble solids in the berries at this stage of development (Davies and Robinson, 1996). Sucrose which is exported to the berries from the leaves, where photosynthesis is active (Ho, 1988; Sonnewald et al., 1994), is converted in the berries to glucose and fructose (Davies and Robinson, 1996). Sucrose, which is a secondary product of photosynthesis, induces the grape dfr gene expression (Fig. 7). The results showed induction of dfr by sucrose, glucose and fructose, but not by maltose and mannitol. The chs gene has been shown to be induced mostly by sucrose, but also by glucose and fructose in leaves in Arabidopsis. High expression of chs in flowers was correlated with the high content of sucrose, glucose and fructose in flowers (Tsukaya et al., 1991). The induction of the grape dfr gene by light and sucrose under light suggest a close interaction between the sucrose and the light signalling pathways. Close interaction between sucrose and light signalling pathways was also reported in the study of the Arabidopsis plastocyanin gene (Dijkwell et al., 1997). Light and sucrose when applied in combination induced the dfr gene in juvenile-phase lamina tissue of ivy, but sucrose did not induce the gene in the dark (Murray et al., 1994).
The existence of unit I and BoxII and sites homologous to the SBF-1 transcription factor binding site in the dfr promoter is consistent with the findings for the light- and calcium-induced expression of the dfr gene. Unit I is the target element for induction by cGMP through the PHY A signalling pathway (Wu et al., 1996). Unit I is sufficient to confer UV-A/blue and UV-B light responsiveness in the chs gene expression (Hartman et al., 1998). Calcium and calmodulin are involved in this response. BoxII is the target for calcium induction in phytochrome A signalling (Wu et al., 1996) and the GT-1 protein has been shown to bind to BoxII and to confer light responsiveness (Lam and Chua, 1990). The SBF-1 protein from bean cells has been identified as a transcription factor, whose binding site is homologous to the GT-1 factor binding site (Lawton et al., 1991).
Light induced the expression of the uidA gene in the full-length dfr promoter fusion and in the -725/+32 deletion fusion construct in the red cell suspension. The -725/+32 fragment is sufficient to confer light induction, but expression is lower in the -725/+32 deletion construct. Loss of 40% GUS activity was seen after deletion of the full-length promoter from -2227 to -725. The -233/+32 deletion fusion is not induced by light, although unit I is present in the -233/+32 deletion fusion. This deletion construct is inactive in the red cell suspension. A possible explanation is that a specific sequence that is important for the expression in the red cell suspension is missing in the -233 deletion fusion.
Induced expression of dfr by sucrose is consistent with the existence of the sucrose boxes 2 and 3 in the dfr promoter. Sucrose boxes 2 and 3 exist in the promoter of the chs-A gene from petunia, which has been shown to be induced by sucrose, glucose and fructose in transgenic Arabidopsis (Tsukaya et al., 1991). These boxes were originally identified (Hattori and Nakamura, 1988) in the sporamin gene family of the sweet potato, which are expressed at high levels when high concentrations of sucrose are applied to stems (Hattori et al., 1990). Tsukaya et al. concluded after sequence alignment that there is a common molecular mechanism of sugar-related regulation, since the sucrose boxes 2 and 3 were found to be present in the chs gene from different plant species, and in other sugar-responsive genes, such as patatin and the gene for proteinase inhibitor II in potato (Tsukaya et al., 1991).
Reduced expression of the -725/+32 deletion fusion in clones of the red cell suspension was seen in cultures grown in B5 medium containing sucrose (Fig. 4A, B). The possibility that part of the reduction in expression was due to the deletion of the sucrose boxes should be studied by more specific deletion. Detailed analysis of specific deletions might show which of the specific promoter elements are involved in sucrose induction.
The expression of the dfr gene is regulated by at least three mechanisms: a light-dependent mechanism, a developmental mechanism and a sugar-related mechanism. Further analysis, such as the deletion or mutation of specific 5' cis-elements, should reveal further details of these regulatory networks.
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Acknowledgments |
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We thank S Farhi for maintenance of the transgenic plants, Dr Y Eshdat for his involvement and helpful comments in this research and Dr V Gaba for his helpful comments and for providing the green light lamp used in this study. This work was supported by grant no. DISNAT 00181 from the German–Israeli Foundation.
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Notes |
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1 The nucleotide sequence data reported will appear in the GenBank Nucleotide Sequence Database under the accession number AF280768.
2 To whom correspondence should be addressed. Fax: +972 3 9669583. E-mail: rachel_gollop{at}yahoo.com
3 Present address: Center for Viticulture Science, Florida A&M University, Tallahassee, FL 32307, USA.
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