Plant Cell Rep DOI 10.1007/s00299-015-1742-8

ORIGINAL PAPER

Responses of rose RhACS1 and RhACS2 promoters to abiotic stresses in transgenic Arabidopsis thaliana Muhammad Ali Khan • Yonglu Meng • Daofeng Liu • Hongshu Tang Suhui Lu¨ • Muhammad Imtiaz • Guimei Jiang • Peitao Lu¨ • Yuqi Ji • Junping Gao • Nan Ma

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Received: 27 September 2014 / Revised: 23 December 2014 / Accepted: 6 January 2015 Ó Springer-Verlag Berlin Heidelberg 2015

Abstract Key message Promoter activities of RhACS1 and RhACS2, two rose genes involved in ethylene biosynthesis, are highly sensitive to various abiotic stresses in an organ-specific manner. Abstract Our previous studies indicated that two rose (Rosa hybrida) 1-aminocyclopropane-1-carboxylic acid synthase genes, RhACS1 and RhACS2, play a role in dehydration-induced ethylene production and inhibition of cell expansion in rose petals. Here, both RhACS1 and RhACS2 promoters were analyzed using histochemical staining and glucuronidase synthase (GUS) gene reporter activity assays following their introduction into transgenic Arabidopsis thaliana plants. It was found that the promoter activities of both genes were strong throughout the course of development from young seedlings to mature flowering plants in various organs, including hypocotyls, cotyledons, leaves, roots and lateral roots. RhACS1 promoter activity was induced by drought in both rosette leaves and roots of transgenic A. thaliana lines, but was reduced following a re-hydration treatment. In contrast, RhACS2 promoter activity was decreased by drought in rosette leaves, while its response pattern was similar to that of RhACS1 in roots. A mannitol treatment induced the activity of both the

RhACS1 and RhACS2 promoters, indicating that both genes are also regulated by osmotic stress. In addition, RhACS2 appeared to be abscisic acid (ABA)-inducible, while RhACS1 was less sensitive to ABA. Finally, four truncated sequences of the RhACS1 promoter were generated and GUS activity assays demonstrated that deleting a 327 bp region between bp 862 and -535 resulted in a substantial decrease of the promoter activity. Taken together, our results suggest that the RhACS1 and RhACS2 promoters respond to abiotic stresses in a developmentally regulated and spatially specific manner. Keywords Rose  RhACS1  RhACS2  Abiotic stress  Arabidopsis thaliana Abbreviations ABA Abscisic acid ACC 1-aminocyclopropane-1-carboxylic acid ACS ACC synthase CHX Cycloheximide GUS b-glucuronidase IAA Indole acetic acid MS Murashige and Skoog SAM S-adenosylmethionine

Communicated by R. Schmidt. M. A. Khan  Y. Meng  D. Liu  H. Tang  S. Lu¨  M. Imtiaz  G. Jiang  P. Lu¨  Y. Ji  J. Gao  N. Ma (&) Department of Ornamental Horticulture, China Agricultural University, Beijing 100193, China e-mail: [email protected] M. A. Khan  M. Imtiaz The University of Agriculture Peshawar, Peshawar 25000, Pakistan

Introduction The developmental processes of flower opening and senescence in many plants, including rose (Rosa hybrida), petunia (Petunia hybrida), carnation (Dianthus caryophyllus) and Phalaenopsis (Phalaenopsis hybrid), are regulated by ethylene (Woltering and Van Doorn 1988). This

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gaseous hormone also modulates many other aspects of growth and development (Abele et al. 1992; Johnson and Ecker 1998) and is involved in biotic and abiotic stress responses (Yang and Hoffman 1984; Ge et al. 2000). The mechanisms of ethylene biosynthesis have been well documented in land plants, and the rate-limiting step is known to be the conversion of S-adenosylmethionine (SAM) to 1-aminocyclopropane-1-carboxylic acid (ACC) by ACC synthase (ACS). ACC is then subsequently oxidized to ethylene by ACC oxidase (Yang and Hoffman 1984). ACS has been established as the key enzyme that regulates ethylene production during normal plant growth and in response to various stresses (Boller et al. 1979; Yu et al. 1979; Wang et al. 2002). ACS enzymes are encoded by multigene families in a taxonomically broad range of plants (Kende 1993; Fluhr and Mattoo 1996; Ge et al. 2000; Wang et al. 2002): the experimental model plant Arabidopsis thaliana has eleven ACS genes (Yamagami et al. 2003), while tomato (Solanum lycopersicum) has eight (Oetiker et al. 1997; Shiu et al. 1998), mung bean (Phaseolus vulgaris) has six (Yoon et al. 1997), potato (Solanum tuberosum) has five (Schlagnhaufer et al. 1997) and rice (Oryza sativa) has five (van der Straeten et al. 2001; Yu et al. 2002). The biosynthesis of ACC and the activity of ACS are known to be influenced by a number of internal and external factors, and the expression of ACS genes is regulated both transcriptionally and post-transcriptionally (Argueso et al. 2007). For example, it has been reported that an elevated level of ACC accumulation can be stimulated by enhanced chilling stress (Wang and Adams 1982), while cadmium and copper treatments elicit ethylene production that is preceded or paralleled by enhanced ACC production (Yu and Yang 1980). Moreover, physical wounding of winter squash mesocarp (Cucurbita maxima) induces an increase in ACS activity, followed by a surge in ACC accumulation and a subsequent elevated rate of ethylene production (Hyodo et al. 1985). Ethylene biosynthesis can also be promoted by the plant hormone indole-3-acetic acid (IAA) and this has been found, in many plant species, to be accompanied by increased ACS activity and ACC content (Abeles 1966; Yu and Yang 1980). The expression of ACS genes can be induced by physiological disorders of various organs, or in response to a range of biotic and/or abiotic environmental factors, including IAA, abscisic acid (ABA), salt, benzyladenine, lithium and copper ions, ozone, aminooxyacetic acid, cycloheximide, protein kinase inhibitors, anaerobiosis (flooding), chilling, wounding, radiation and pathogens (Ge et al. 2000; Wang et al. 2002). Moreover, the expression of different members of ACS multigene families can be differentially regulated in response to these factors. The promoter sequences of various ACS genes have been isolated and characterized to elucidate the associated transcriptional regulatory mechanisms, and histochemical assays of several A. thaliana

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ACS (AtACS) genes in transgenic A. thaliana lines have shown various levels of regulation in different organs, including leaves, roots and flowers (Rodrigues-Pousada et al. 1993; Wang et al. 2005). ACS genes have also be studied in the ornamental plant rose (R. hybrida), the flowers of which are generally categorized as ethylene-sensitive; however, several reports have described variable effects of exogenous ethylene on rose flower cultivars (Reid et al. 1989; Yamamoto et al. 1994). For example, during postharvest handling of roses, ethylene and water deficiency can result in abnormal flower opening, and in cut roses, ethylene treatment can induce a rapid and substantial increase in ethylene production in the gynoecia, which has been attributed to the increased expression of two ACS genes, RhACS2 and RhACS3 (Xue et al. 2008). The RhACS1 gene has also been shown to be associated with petal senescence and wounding, while RhACS2 is mainly related to senescence and RhACS3 has been linked to flower opening (Ma et al. 2005). We previously investigated the mechanism by which ethylene adversely affects the opening of detached rose flowers, and showed that the expression of RhACS1 and RhACS2 is associated with the induction of ethylene biosynthesis during dehydration and rehydration treatments, and that this ethylene is primarily produced by sepals and gynoecia (Liu et al. 2013). Additional evidence of a link between dehydration and RhACS1 and RhACS2 was provided by the identification of a number of promoter cis-elements, including ABA, dehydration-responsive and tissue-specific elements, in a 2,080 bp 50 upstream sequence of RhACS1 (NCBI Accession No. HF562220), and a 1,584 bp 50 -upstream sequence of RhACS2 (NCBI Accession No. HF564634) (Liu et al. 2013). In the present study, we used transgenic A. thaliana plants harboring each of these promoters (Pro-RhACS1 and Pro-RhACS2, respectively) coupled to the glucuronidase synthase (GUS) reporter gene to investigate the regulatory mechanisms of RhACS1 and RhACS2 expression. We assessed the influence of abiotic stresses on the RhACS1 and RhACS2 promoter activity and showed that the two genes operate in a development-dependent manner, and are particularly active in rapidly expanding organs. The activity of the RhACS1 and the RhACS2 promoters was variably affected by different environmental stresses, and deletion experiments showed that the region from -862 to -535 bp of the RhACS1 promoter is essential for its activity.

Materials and methods Construction of the RhACS1 promoter deletion vectors The deletion fragments of the RhACS1 promoter were amplified using a high-fidelity DNA polymerase and gene-

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specific primers. The five sense primers used were: D1: 50 -C ATGCATGCTAGAAATAGTCAATCTCATGCT-30 (SphI site italicized); D1–1: 50 -CCCAAGCTTAAGGGTGTGATG CGAAAT-30 (HindIII site italicized); D1–2: 50 -CCCAAGC TTGTTTCACTCAAATATTGTCATC-30 (HindIII site italicized); D1–3: 50 -CCCAAGCTTGAAATTGTCTCAAA CTTCATCAC-30 (HindIII site italicized); and D1–4: 50 -CC CAAGCTTGACCAATAACCCAAGCAC-30 (Hind III site italicized). The antisense primer used was: DRP (deletion reverse primer): 50 -CGCGGATCCATGTCGTGGTAAGT GTATAATC-30 (BamHI site italicized). The truncated fragments were designed based on a previous cis-element analysis (Liu et al. 2013). Each 50 -end deletion primer was used in combination with the DRP primer to amplify the corresponding deletion product. The amplified fragments were digested with SphI and BamHI (for D1), or with HindIII and BamHI (for D1–1 to D1–4), and inserted into the binary vector pBI121 to replace the cauliflower mosaic virus 35S promoter upstream of the b-glucuronidase (GUS) gene. The five constructs were named: D1 (-1,950, HF562220), D1–1 (-1,431, KP133132), D1–2 (-862, KP133133), D1–3 (-535, KP133134) and D1–4 (-213, KP133135), respectively.

them on a plastic sheet at room temperature for 2 h and then re-watering them for 2 h, during which time the general viability of the plants was assessed visually. These plants were then used for GUS activity analyses. Seedlings grown in MS medium under normal conditions were used as controls. Assays were performed on rosettes and roots from four lines of transgenic A. thaliana per treatment and three plants from each transgenic line were evaluated. GUS histochemical assays For GUS histochemical analyses, plants were incubated in GUS staining solution [75.5 mM sodium phosphate (pH 7.0), 0.1 % Triton X-100, 0.05 mM K3/K4 FeCN, 10 mM Na2-ethylenediaminetetraacetic acid (EDTA), 20 % methanol (v/v) and 50 lg ml-1 5-bromo-4-chloro-3-indolyl glucuronic acid (X-gluc)] at 37 °C overnight. The plants were then cleared using 70 % ethanol and subsequently imaged by a NIKON stereomicroscope. The pBI121 vector containing CaMV35S::GUS was used as a positive control and the pBI121 vector from which the 35S promoter had been excised was used as a negative control. GUS activity assays

A. thaliana transformation Arabidopsis thaliana Columbia (Col-0) plants were transformed using the floral dip method (Clough and Bent 1998) with the Agrobacterium tumefaciens strain GV3101 carrying ProRhACS1 or Pro-RhACS2 fused to the GUS reporter gene (Pro-RhACS1:GUS or Pro-RhACS2:GUS, respectively) or a truncated RhACS1 promoter (D1–1 to D1–4). The independent transformants were screened on MS medium (Murashige and Skoog 1962) containing 50 mg L-1 kanamycin. At least 10 lines carrying each of the constructs described above were selected and two representative T2 generation lines with a single copy insertion were used for further experiments.

Fluorimetric analysis of GUS activity was performed as described by Jefferson (1987). Plants were harvested and immediately homogenized by grinding in 0.2 ml GUS extraction buffer [50 mM sodium phosphate (pH 7.0), 10 mM EDTA, 0.1 % sodium lauryl sarcosine, 0.1 % Triton X-100 and 10 mM b-mercaptoethanol]. Cell debris was removed by centrifugation at 12,000g at 4 °C for 10 min, and the supernatant was collected. The substrate 4-methylumbelliferyl-b-D-glucuronide hydrate (MUG) was added to the supernatant (at a final concentration of 2 mM) and the mixture was incubated at 37 °C for 15 min. GUS activity was analyzed using a Microfluor fluorometer (F4500, Hitachi, Tokyo, Japan) with the emission wavelength set at 455 nm and the excitation wavelength at 365 nm.

Abiotic stress treatments Determination of soluble protein content For hormone and abiotic stress response assays, T2 seeds carrying the Pro-RhACS1:GUS construct were germinated on MS medium containing 3 % (w/v) sucrose and 0.6 % (w/v) agar at 21 °C under fluorescent white light, with a photoperiod of 16/8 h (light/dark). For mannitol treatments, T2 seedlings were transferred 15 days after germination onto MS media containing mannitol (0, 150, 250 or 350 lM) for 4 days. For ABA treatments, 15-day-old T2 seedlings were transferred to an ABA solution (0, 10 or 50 lM) for 4 h as previously described (Kim et al. 2011). For drought treatments, 15-day-old T2 seedlings were subjected to a 2-h dehydration/drought period by placing

The soluble protein content of each sample was determined using the Bradford (1976) method with bovine serum albumin (BSA) as a standard.

Results Analysis of RhACS1 promoter activity To assess the activity of the RhACS1 and RhACS2 promoters during plant development, we stably introduced

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(Fig. 1j). Thus, activity of the RhACS1 promoter was detected in most plant organs. The activity of the RhACS2 promoter was similarly evaluated. As shown in Fig. 2, GUS activity was strong in vernalized seeds and all tested organs of the transgenic plants. In 5-day-old etiolated transgenic seedlings, RhACS2-driven GUS expression was clearly detected in both shoots and roots, and the blue color was more intense in older leaves and roots (Fig. 2d). The majority of the GUS activity was detected in the cotyledons (Fig. 2h) and the leaves (Fig. 2i) of young plants, with a clear netted venation pattern. Intense staining was also observed in the vascular tissues of the roots, lateral roots, and the transition zones between the hypocotyls and roots and root tips (Fig. 2e, g, f), while GUS staining of the florets revealed expression in all parts of the flower. A close examination of the fully opened flowers showed that strong GUS activity was present in the entire inflorescence including sepals, petals, the veins of flower petals and sepals, and stamens, as well as the filaments and carpels (Fig. 2j, k, l). Staining was high throughout the inflorescence suggesting a higher expression of RhACS2 during the reproductive stages of plant growth.

RhACS1 promoter::GUS or RhACS2 promoter::GUS cassettes into the A. thaliana genome by Agrobacteriummediated transformation and measured GUS activity throughout the whole life cycle of the transgenic plants using histochemical staining. As shown in Fig. 1, GUS activity driven by the RhACS1 promoter was detected in vernalized seeds and all other evaluated organs at the earlier stages (3–15 days) of vegetative growth. In 3-dayold plants, GUS activity was stronger in the vascular tissues of the lower portion of the plants, including the hypocotyls and roots, than in the cotyledons (Fig. 1d). The leaves of young plants generally showed a lower staining intensity. Seven- to nine-day-old plants were the same size as wild-type plants and lateral roots associated with the primary roots showed greater GUS activity. GUS activity was higher in 13–15-day-old plants, where a more uniform activity was observed in the vascular tissues of leaves, hypocotyls, lateral roots, root tip and the transition zones between the primordia and lateral roots (Fig. 1d). GUS activity was stronger in the vascular tissues of the transition zones between the hypocotyls and the roots than in the root tips (Fig. 1e, f). Similarly, GUS staining was greater in the transition zones between the primordia and lateral roots than in the tip region of the lateral roots (Fig. 1f, g). Strong staining was also detected in the cotyledons (Fig. 1h) and the leaves (Fig. 1i) of young plants, especially in the vascular tissues. During reproductive growth, GUS activity was weaker in the early stages of inflorescence formation than in vegetative organs such as the leaves and roots (Fig. 1d, g, j, k). Low activity was detected in specific parts of the flower, including sepals and petals (Fig. 1l), and there was also some staining in the tip of the flower petals and sepals. No GUS activity was detected in the stamens or the lower portion of the petals

It has previously been reported that the expression of the RhACS1 gene exhibits a spatial and temporal specificity during dehydration and rehydration in detached rose flowers (Liu et al. 2013). Here, we first analyzed the response of the RhACS1 promoter to drought and rewatering conditions. Fifteen-day-old plants were subjected to a 2-h drought treatment at room temperature and the treated plants were then re-watered for 2 h. The whole

Fig. 1 Histochemical localization of GUS expression driven by the RhACS1 promoter in transgenic A. thaliana. a Positive control (35SGUS) 4-day-old plants, b negative control (pBI121 empty vector), 4-day-old plants, c mature seeds and vernalized seeds, d vegetative

growth: 3, 7, 9, 13 and 15-day-old plants, e transition zones between hypocotyls and roots in 7-day-old plants, f root tips in 7-day-old plants, g lateral roots, h cotyledons, i leaves, j the whole inflorescence, k early inflorescence

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Fig. 2 Histochemical localization of GUS expression driven by the RhACS2 promoter in transgenic A. thaliana. a Positive control (35SGUS) 4-day-old plants, b negative control (pBI121 empty vector), 4-day-old plants, c mature seeds and vernalized seeds, d vegetative

growth: 5, 7, 9, 13 and 15-day-old plants, e transition zones between hypocotyls and roots in 7-day-old plants, f root tips in 7-day-old plants, g lateral roots, h cotyledons, i leaves, j the earlier stage of inflorescence formation, k pistil after removing petals, l flower

plants were then divided into two parts: rosettes and roots. As shown in Fig. 3, the drought treatment induced GUS activity in both the rosettes and roots of all transgenic lines, while the subsequent re-watering treatment reduced the drought-induced RhACS1 promoter activity. ABA is known to be an important signaling hormone associated with plant drought response of plants (Zhu 2002; Seki et al. 2007) and so we evaluated the responses of 15-day-old RhACS1 promoter::GUS transgenic plants to

various concentrations of ABA (0, 10 and 50 lM). After ABA treatments, the plants were divided into rosettes and roots and GUS activity in each was measured. We observed that the effect of ABA was different in rosettes and roots, since GUS activity decreased slightly in the rosettes in response to ABA, while no change was observed in the roots (Fig. 4). We also investigated the response of the RhACS1 promoter to osmotic stress by treating the transgenic RhACS1 promoter::GUS plants with various concentrations of mannitol (0, 150, 250 and 350 lM) on day 15 after germination. As shown in Fig. 5, GUS activity increased as

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Fig. 3 RhACS1 promoter activity in response to drought in transgenic A. thaliana plants. Line #1 and #2 represent different transgenic lines. Assays were performed on rosettes and roots. Plants were grown on medium for 15 days and subsequently drought treated. Assays were performed on rosettes and roots from four transgenic lines per treatment and each transgenic line had three biological replicates per treatment. Bars represent standard error and different letters indicate significant differences (ANOVA, Duncan’s test, p \ 0.05)

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Fig. 4 RhACS1 promoter activity in response to ABA treatment. The plants were treated with ABA (0, 10 or 50 lM). Line #1 and #2 represent different transgenic lines. Assays were performed on rosettes and roots. Each transgenic line had three biological replicates per treatment. Bars represent standard error and different letters indicate significant differences (ANOVA, Duncan’s test, p \ 0.05)

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To test the effect of abiotic stresses on the activity of the RhACS2 promoter, 15-day-old transgenic plants harboring the RhACS2 promoter::GUS cassette were subjected to the same drought, ABA and mannitol treatments as described above. Unlike the RhACS1 plants, the activity of the RhACS2 promoter was substantially lower in rosettes in response to drought, while the re-watering treatment had no effect (Fig. 6). Interestingly, the activity of the RhACS2 promoter in roots was induced by drought and was greatly reduced by re-watering, suggesting an organ-specific response to drought (Fig. 7). GUS activity was also increased by the ABA treatment in both rosette and roots, with a stronger effect in rosettes. The activity of the RhACS2 promoter was strongly induced by transferring the transgenic plants to MS plates with even low concentrations of mannitol (150 lM) in the rosettes, while it decreased as the mannitol concentration increased. A similar pattern was seen in roots (Fig. 8). RhACS1 promoter deletion analysis Since the RhACS1 promoter showed a higher activity for all the treatments than RhACS2, this was selected for further structural analysis. Specifically, to determine the contribution of different regions to the activity of the RhACS1 promoter, a series of promoter deletion constructs were made: D1 (-1,950), D1–1 (-1,431), D1–2 (-862), D1–3 (-535), D1–4 (-213), based on previous

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Fig. 5 RhACS1 promoter activity in response to mannitol treatment. The plants were treated with mannitol (0, 150, 250 or 350 lM) Line #1 and #2 represent different transgenic lines. Assays were performed on rosettes and roots. Each transgenic line had three biological replicates per treatment. Bars represent standard error and different letters indicate significant differences (ANOVA, Duncan’s test, p \ 0.05)

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Fig. 6 RhACS2 promoter activity in response to drought. Line #3 and #5 represent different transgenic lines. Assays were performed on rosettes and roots. Plants were grown on medium for 15 days and subsequently drought treated. Each transgenic line had three biological replicates per treatment. Bars represent standard error and different letters indicate significant differences (ANOVA, Duncan’s test, p \ 0.05) 20

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Fig. 7 RhACS2 promoter activity in response to ABA treatment. Plants were treated with ABA (0, 10 or 50 lM). Line #3 and #5 represent different transgenic lines. Assays were performed on rosettes and roots. Each transgenic line had three biological replicates per treatment. Bars represent standard error and different letters indicate significant differences (ANOVA, Duncan’s test, p \ 0.05)

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the mannitol concentration increased in rosettes, while it declined in the roots.

cis-element analysis of the RhACS1 promoter (Fig. 9; Liu et al. 2013). The fragments were individually fused to the GUS gene coding sequence and transformed into A. thaliana. A quantitative measurement of the GUS activity was then performed using 6-day-old transgenic plants harboring the different transgenes. Deletion of the 519 bp fragment between -1,950 and -1,431 bp caused only a minimal reduction in GUS activity (Fig. 9), as did an additional deletion of the fragment from -1,431 to 862 bp. In contrast, the further deletion of 327 bp from position -862 to -535 resulted in a substantial reduction of GUS activity, with approximately sixfold decrease compared to plants carrying the D1–2::GUS cassette

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Fig. 9 Assays for GUS expression driven by the RhACS1 promoter deletions (D1–1, D1–2, D1–3, D1–4) using a tobacco transient expression system. A schematic diagram of the RhACS1 promoter deletions is shown on the left. The numbers on the left of the construct

diagrams indicate the 50 end of each deletion. The results of the transient expression experiments with the different constructs are shown on the right. The mean standard error values of GUS activity were calculated using seven biological replicates

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Previously, we isolated the rose RhACS1 and RhACS2 gene promoters and identified various important cis-elements, including ABA, dehydration-responsive, and tissue-specific elements (Liu et al. 2013). The RhACS1 promoter contains eight putative MYC (CANNTG) and two MYB

(WAACCA) motifs, while the RhACS2 promoter has six putative MYC motifs and one MYB motif, as well as putative GATA boxes. In this current study, we found that the strongest activity of RhACS1 and RhACS1, based on expression of the GUS reporter, was in the vascular bundles and rapidly expanding organs, such as young cotyledons, leaves, roots and most parts of the flowers. Additionally, the activity of both the RhACS1 and RhACS2 genes was detected at the very early stages of plant development, including vernalized seeds. We previously showed that the expression of RhACS1, a wounding-inducible gene, and RhACS2, a senescence-inducible gene (Ma et al. 2005), exhibits spatial and temporal specificity during dehydration and rehydration in detached rose flowers. Sepals and gynoecia were the main sources of ethylene production in cut rose flowers, in accordance with the organ-specific expression induction patterns RhACS1 and RhACS2 (Liu et al. 2013). Here, we found that the RhACS2 promoter drove a stronger GUS activity in all floral organs of transgenic A. thaliana, including sepals, petals, pistils and stamens, while RhACS1 promoter activity was detected only in the pistils, including the stigma and ovary, and either no activity or low activity was observed in the petals and sepals. These results further indicate that the promoter activities of the RhACS genes are organ specific; an observation that is congruent with the observed organspecific ethylene production in cut rose flowers. Tissue-specific ethylene induction in numerous plant species has been ascribed to the spatial and temporal regulation of ACS genes (Lin et al. 2009). In A. thaliana, a higher promoter activity was observed for AtACS5 and AtACS7 in all vegetative and reproductive organs and tissues, including leaves, roots, root tips, petals, sepals, carpels, stamens, cauline leaves, inflorescence stems and siliques, while AtACS4 expression was undetectable in the petals of open flowers (Wang et al. 2005). Similarly, transgenic A. thaliana

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Fig. 8 RhACS2 promoter activity in response to mannitol treatment. The plants were treated with mannitol (0, 150, 250 or 350 lM). Line #3 and #5 represent different transgenic lines. Assays were performed on rosettes and roots. Each transgenic line had three biological replicates per treatment. Bars represent standard error and different letters indicate significant differences (ANOVA, Duncan’s test, p \ 0.05)

(Fig. 9). We therefore concluded that positive regulatory cis-elements are present in the region between -1,950 and -1,450 bp, and that negative regulatory cis-elements are mainly present in the region between -862 and 535 bp. The 327 bp region between -862 and -535 bp was found to be particularly important for RhACS1 promoter activity.

Discussion

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plants harboring a ProAtACS1:GUS cassette were observed to have a strong GUS activity in the course of development from young seedlings to mature flowering plant, both in the rosette and the root system (Rodrigues-Pousada et al. 1993). In petunia (P. hybrida), the activity of the Ph-ACS2 promoter, as determined by GUS reporter expression, was only observed in mature pollen grains (Lindstrom et al. 1999). As a further example of organ specificity, in carnation (D. caryophyllus), exogenous ethylene treatment was reported to differentially induce the expression of ethylene biosynthetic genes in various floral organs. Specifically, elevated ethylene production was detected in styles due to higher expression levels of DCACS2 and DCACS3, while increased expression of DCACS1 was observed in petals (Jones and Woodson 1999). At the transcriptional level, numerous members of the ACS gene family from various plant species have been shown to be differentially induced in response to various hormones, or biotic and abiotic stress treatments, including IAA, ABA, LiCl, NaCl, CuCl2, cycloheximide (CHX), aminooxyacetic acid (AOA), ethylene, physical wounding, cold temperatures and sunlight (Rodrigues-Pousada et al. 1993; Arteca and Arteca 1999; Ge et al. 2000; Peleg and Blumwald 2011). In the present study, the activity of the RhACS1 promoter was induced by drought in both the rosettes and roots of transgenic A. thaliana lines, but was suppressed by subsequent re-watering. The RhACS2 promoter displayed different activation patterns in different organs, decreasing due to both drought and re-watering in rosettes, while increasing after drought treatment and decreasing after re-watering in roots. The highest GUS activity of drought-affected RhACS1::GUS plants was detected in roots, with approximately twofold higher activity than in rosettes, whereas RhACS2 promoter activity was not significantly different between the rosettes and roots. The drought-induced promoter activity of both RhACS1 and RhACS2 is in accordance with our previous study, where we found that the RhACS1 and RhACS2 genes showed substantially increased levels of expression throughout the dehydration period in an organ-specific manner (Liu et al. 2013). Similarly, ABA affected the RhACS1 promoter activity differently in rosettes and roots, with a slightly reduced RhACS1 activity in rosettes after treatment and no change of activity in roots. The RhACS2 promoter was more sensitive to the ABA treatment and an increasing concentration of ABA lead to an increase in GUS activity in both rosettes and roots. Furthermore, a gradual increase in GUS activity driven by the RhACS1 promoter was detected in rosettes following an increase in mannitol concentration, whereas an early induction in roots by low mannitol concentrations was followed by a decline at higher concentrations. Similarly, for the RhACS2 promoter, the GUS activity first increased when mannitol was

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applied, but then gradually decreased at higher concentrations in both rosettes and roots. We conclude that RhACS2 is sensitive to ABA, mannitol and drought treatments and that there is a difference in induction patterns between rosettes and roots. ABA is known to be a key hormone in regulating the responses of plants to drought stress. Here, RhACS2 promoter activity was induced by drought in roots and suppressed after re-watering, supporting the observations made with ABA. We also evaluated the publicly available A. thaliana expression data from the e-FP browser for the homologs of RhACS1 (AtACS2, with 65 % amino acid sequence identity with RhACS1) and RhACS2 (AtACS6, with 72 % amino acid sequence identity with RhACS2) to compare the expression patterns. AtACS2 expression is listed as being induced by both drought and ABA, but down-regulated by mannitol in both rosettes and roots. The expression and regulation of RhACS1 and AtACS2 therefore appear to be different, other than that they are both induced by drought. In contrast, while AtACS6 expression is listed as being rapidly and strongly induced by drought in roots, while the RhACS2 promoter did not show evidence of a response to this treatment in rosettes. Expression of AtACS6 is induced by mannitol and ABA in both rosettes and roots and RhACS2 and AtACS6 therefore share expression patterns in response to these abiotic stresses in addition to the spatial-specific responses to drought. These results suggest that the regulation and expression of RhACS2 and AtACS6 are more conserved between rose and A. thaliana than RhACS1 and AtACS2. Previous reports have also shown induction or differential regulation of different AtACS promoters using other hormones in light-grown A. thaliana seedlings, including IAA- and ABA-induced expression of AtACS5, ACC-, ABA- and jasmonic acid (JA)-induced expression of AtACS4, and gibberellins-, ACC-, salicylic acid- and brassinosteroid-induced expression of AtACS7 (Wang et al. 2005; Tang et al. 2008). Similarly, AtACS6 expression is also enhanced in response to various hormonal and environmental stresses, such as IAA, LiCl, NaCl, CuCl2, CHX, physical wounding and touch stimuli (Arteca and Arteca 1999). Finally, the expression of two Nicotiana tabacum ACS genes, NtACS2 (sharing 64 and 71 % amino acid identity with RhACS1 and RhACS2, respectively) and NtACS4 (sharing 49 and 50 % amino acid identity with RhACS1 and RhACS2, respectively) was also reported to be substantially affected by ABA, JA, cold temperature, ethylene, wounding and treatment with electrical current (Ge et al. 2000). Several members of the ACS gene family have also been shown to be regulated by auxin fluctuations, as reviewed by Peleg and Blumwald (2011). In the present study, we also performed a promoter deletion study of RhACS1. A series of RhACS1 deletions were fused to the GUS reporter gene and transformed into

Plant Cell Rep

A. thaliana. GUS activity was barely reduced when promoter fragments between (D1) and (D1–2) were deleted, but deleting 327 bp from position -862 (D1–2) to -535 (D1–3) resulted in a substantial reduction in GUS activity, suggesting that this 327 bp promoter fragment is critical for RhACS1 promoter activation. We detected several important cis-elements in a previous study in this 327 bp promoter fragment, including GATA boxes, CAAT, MYBIAT, MYBCORE, Inr and others (Liu et al. 2013). These elements are known to be involved in mediating tissue-specific expression (Lam and Chua 1989; Vieweg et al. 2004); however, additional studies are needed to verify the involvement of these elements in controlling the organ specificity of the RhACS1 promoter. In summary, we characterized the RhACS1 and RhACS2 promoters in transgenic A. thaliana lines and found that strong activities of both the promoters were associated with rapidly growing or expanding organs, and that the promoter activities were differentially influenced by hormone and abiotic stress treatments. Moreover, the 327 bp region between -862 and -535 of the RhACS1 promoter was found to be an important region for promoter activity. Considering that the RhACS1 and RhACS2 genes play crucial roles in ethylene biosynthesis, the present work will prove useful for further studies of the mechanism by which ethylene biosynthesis is regulated at the precursor level. Author contribution statement M. A. K. wrote the most of the manuscript and conducted experiments; Y. M. conducted most of the experiments; D. L., H. T., M. I., G. J., P. L., and Y. J. contributed by conducting various experiments presented in the manuscript; and J. G., N. M. supervised and helped in writing the manuscript. Acknowledgments This work was supported by the National Natural Science Foundation of China, Grant 31372095 and Grant 31130048, and the 948 project (2011-G17) of the Ministry of Agriculture. We thank PlantScribe (www.plantscribe.com) for carefully editing this manuscript. Conflict of interest conflict of interest.

All the authors declared that they have no

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