Phytochemistry 103 (2014) 22–31

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PhDAHP1 is required for floral volatile benzenoid/phenylpropanoid biosynthesis in Petunia  hybrida cv ‘Mitchell Diploid’ Kelly M. Langer, Correy R. Jones, Elizabeth A. Jaworski, Gabrielle V. Rushing, Joo Young Kim, David G. Clark, Thomas A. Colquhoun ⇑ Plant Innovation Program, Department of Environmental Horticulture, University of Florida, Gainesville, FL 32611, USA

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Article history: Received 8 February 2014 Received in revised form 28 March 2014 Available online 6 May 2014 Keywords: Petunia Petunia  hybrida Solanaceae Gene characterization Shikimate pathway DAHP synthase Phenylpropanoid Floral volatiles

a b s t r a c t Floral volatile benzenoid/phenylpropanoid (FVBP) biosynthesis consists of numerous enzymatic and regulatory processes. The initial enzymatic step bridging primary metabolism to secondary metabolism is the condensation of phosphoenolpyruvate (PEP) and erythrose-4-phosphate (E4P) carried out via 3-DEOXY-D-ARABINO-HEPTULOSONATE-7-PHOSPHATE (DAHP) synthase. Here, identified, cloned, localized, and functionally characterized were two DAHP synthases from the model plant species Petunia  hybrida cv ‘Mitchell Diploid’ (MD). Full-length transcript sequences for PhDAHP1 and PhDAHP2 were identified and cloned using cDNA SMART libraries constructed from pooled MD corolla and leaf total RNA. Predicted amino acid sequence of PhDAHP1 and PhDAHP2 proteins were 76% and 80% identical to AtDAHP1 and AtDAHP2 from Arabidopsis, respectively. PhDAHP1 transcript accumulated to relatively highest levels in petal limb and tube tissues, while PhDAHP2 accumulated to highest levels in leaf and stem tissues. Through floral development, PhDAHP1 transcript accumulated to highest levels during open flower stages, and PhDAHP2 transcript remained constitutive throughout. Radiolabeled PhDAHP1 and PhDAHP2 proteins localized to plastids, however, PhDAHP2 localization appeared less efficient. PhDAHP1 RNAi knockdown petunia lines were reduced in total FVBP emission compared to MD, while PhDAHP2 RNAi lines emitted ‘wildtype’ FVBP levels. These results demonstrate that PhDAHP1 is the principal DAHP synthase protein responsible for the coupling of metabolites from primary metabolism to secondary metabolism, and the ultimate biosynthesis of FVBPs in the MD flower. Ó 2014 Elsevier Ltd. All rights reserved.

1. Introduction Overall, plant derived volatile organic compounds possess numerous biological and ecological functions like defense, signaling, and reproduction (Dudareva et al., 2013). Many angiosperm species are capable of producing a wide range of volatile molecules from floral tissues that can potentially function to facilitate sexual reproduction and/or defend against florivores (Kessler et al., 2013). The model plant, Petunia  hybrida cv ‘Mitchell Diploid’ (MD), is a doubled haploid petunia that synthesizes and emits relatively large quantities of a select number of floral volatile benzenoid and phenylpropanoid (FVBP) compounds from cells of the corolla limb tissue (Colquhoun et al., 2010b; Underwood et al., 2005; Van Moerkercke et al., 2012). Detected FVBPs from head space analyses include: phenylacetaldehyde (13), 2-phenethyl acetate, 2-phenylethanol (14), phenethyl benzoate (15), benzaldehyde (18), benzyl ⇑ Corresponding author. Address: 1523 Fifield Hall, Gainesville, FL 32611, USA. Tel.: +1 352 273 4584; fax: +1 352 392 3870. E-mail address: ucntcme1@ufl.edu (T.A. Colquhoun). http://dx.doi.org/10.1016/j.phytochem.2014.04.004 0031-9422/Ó 2014 Elsevier Ltd. All rights reserved.

acetate, benzyl alcohol (17), benzyl benzoate (16), methyl benzoate (20), methyl salicylate (22), isoeugenol (29), and eugenol (28) (Fig. 1) (Boatright et al., 2004; Colquhoun et al., 2010b; Koeduka et al., 2006; Kolosova et al., 2001a; Underwood et al., 2005; Verdonk et al., 2003). The last common metabolite in petunia FVBP production is the amino acid, phenylalanine (12) (Schuurink et al., 2006). In plants, the aromatic amino acids tyrosine, tryptophan, and phenylalanine (12) all share the last common metabolite, chorismic acid (9), which is the end product of the plastid localized shikimate pathway. The shikimate pathway is composed of seven enzymatic steps and is conserved between plants and microorganisms (Light and Anderson, 2013; Maeda and Dudareva, 2012). The initial enzymatic reaction of this pathway is an aldol condensation of the glycolytic intermediate, phosphoenolpyruvate (PEP) (2), and the pentose phosphate pathway intermediate, erythrose-4-phosphate (1) (DeLeo and Sprinson, 1975; Herrmann and Weaver, 1999; Huisman and Kosuge, 1974; Rubin and Jensen, 1985). This reaction is catalyzed by the metalloenzyme 3-DEOXY-D-ARABINOHEPTULOSONATE-7-PHOSPHATE (DAHP) synthase to produce

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Fig. 1. A schematic representation of FVBP biosynthesis in a MD flower limb cell. 3-DEOXY-D-ARABINO-HEPTULOSONATE-7-PHOSPHATE (DAHP) synthase is the first enzyme of the shikimate pathway located in the plastid organelle. DAHP synthase catalyzes an aldol condensation of the pentose phosphate pathway intermediate, erythrose-4phosphate (E-4-P) (1), and the glycolytic intermediate, phosphoenolpyruvate (PEP) (2), thereby bridging primary and secondary metabolism. The shikimate pathway culminates in the formation of chorismate (9). Chorismate (9) is converted into prephenate (10) by CHORISMATE MUTASE1 (CM1), which is the first committed step in phenylalanine (12) production. Phenylalanine (11) is transported out of the plastid by an unknown mechanism. Once in the cytosol, phenylalanine (12) can enter the phenylpropanoid pathway with the conversion to (E)-cinnamic acid (23) by PHENYLALANINE-AMMONIA LYASE (PAL), or the first branch of the FVBP pathway via PHENYLACETALDEHYDE SYNTHASE (PAAS). (E)-cinnamic acid (23) is the last shared metabolite between FVBP branch 2 and 3, and can either continue in the phenylpropanoid pathway, or can be imported in peroxisome organelles by an unknown mechanism. Once in the peroxisome, CINNAMATE:COA LIGASE/ACYL-ACTIVATING ENZYME (CNL/AAE) converts (E)-cinnamic acid (23) to cinnamoyl-CoA, which is the first reaction a b-oxidation pathway leading to volatile benzenoid production. Branch 3 of the FVBP pathway ultimately produces the volatile constituents of common clove oil, eugenol and isoeugenol (25).

3-deoxy-d-arabino-heptulosonate-7-phosphate (3) and inorganic phosphate (Friedrich and Schlegel, 1975; Garner and Herrmann, 1984; Stephens and Bauerle, 1991). Numerous DAHP synthases have been identified and characterized at some level in microorganisms, e.g., genetic analysis of DAHP synthase mutants in Salmonella typhimurium (DeLeo and Sprinson, 1975), allosteric regulation of DAHP synthase isozymes in Pseudomonas genera (Whitaker et al., 1981), and the quaternary structure of DAHP synthase as a tetramer in Escherichia coli (Shumilin et al., 1999). Although still being discovered in plants, the predicted amino acid sequence of plant DAHP synthases have been recognized to share approximately 20% similarity to their bacterial homologs (Entus et al., 2002; Herrmann, 1995). DAHP synthases are separated into two types (types I and II) based on phylogenetic comparisons. Most plants and a few microorganisms retain type II DAHP synthases, which have specific and unique allosteric binding sites (Webby et al., 2005). Plant DAHP synthases appear to be plastid localized as suggested by the conservation of an N-terminal chloroplast transit peptide, which can be cleaved upon entrance into the plastid (Zhao et al., 2002; Zybailov et al., 2008). In both Solanum tuberosum

and Arabidopsis thaliana subcellular fractionalization and plastid import assays have demonstrated plastid localization of both DAHP isoforms, respectively (Dyer et al., 1990; Entus et al., 2002). Previous work from Solanum lycopersicum identified multiple genes for putative DAHP synthases that were differentially regulated through multiple tissues (Gorlach et al., 1993), which suggested differential functions. The shikimate pathway is an essential precursor to the phenylpropanoid pathway and ultimately, thousands of phenylpropanoid derivatives. The first step in aromatic amino acid synthesis, DAHP synthase can potentially regulate the overall carbon flux through the shikimate pathway (Jones et al., 1995). The purpose of this study was to identify any DAHP synthase homologs in petunia, and characterize their role in the production of phenylpropanoid compounds in MD flowers. It is hypothesized here that the genes encoding DAHP synthase may have distinct roles in P.  hybrida as suggested in earlier studies in A. thaliana and S. lycopersicum where DAHP synthases have been identified with distinct transcript accumulation profiles under various conditions, indicating distinct biological roles (Gorlach et al., 1993; Keith et al., 1991).

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2. Results and discussion 2.1. Identification of PhDAHP1 and PhDAHP2 After perusing the publically available nucleotide databases for petunia sequences similar to AtDAHP1 and AtDAHP2 from Arabidopsis (NCBI, SGN, and the 454 database), two distinct contigs were identified. The petunia contigs were incomplete compared to AtDAHP1 and AtDAHP2. A SMART RACE library constructed from total RNA of P.  hybrida cv ‘Mitchell Diploid’ (MD) petal limb and leaf tissue was utilized to clone a full-length transcript for each contig, individually named 3-DEOXY-D-ARABINO-HEPTULOSONATE7-PHOSPHATE1 (PhDAHP1) synthase and 3-DEOXY-D-ARABINOHEPTULOSONATE-7-PHOSPHATE2 (PhDAHP2) synthase. PhDAHP1 full-length transcript was 1942 nucleotides (nt) in length and contained a 1599 nt coding region (Fig. S1). PhDAHP2 full-length transcript that was 1927 nt in length, and contained a 1536 nt coding region (Fig. S2). PhDAHP1 and PhDAHP2 sequences were submitted to GenBank under accession numbers, JQ955569 and JQ955570, respectively. The predicted amino acid sequence for PhDAHP1 and PhDAHP2 were 533 and 512 amino acids in length, and had a calculated molecular mass of 58,894 and 57,061 kDa, respectively. Both PhDAHP1 and PhDAHP2 were predicted to be localized to the plastid (Predator v. 1.03) and contained an N-terminal chloroplast transit peptide (cTP) of approximately 80 to 61 amino acids, respectively (ChloroP 1.1, Fig. 2). The PhDAHP cTPs shared 16.3% identity. Additionally, both putative petunia DAHP synthase proteins (minus cTP) contained a Class-II DAHP synthase family domain (cl03230), a phospho-2-dehydro-3-deoxyheptonate aldolase domain (PLN02291), and shared 84.8% identity. An alignment of DAHP synthase protein sequences from P.  hybrida (PhDAHP1 and PhDAHP2), Nicotiana tabacum (NtDAHP), S. tuberosum (StDAHP), Solanum torvum (SlvDAHP), Morinda citrifolia (McDAHP), Populus trichocarpa (PtDAHP), A. thaliana (AtDAHP1 and AtDAHP2), S. lycopersicum (SlDAHP), Lotus japonicas (LjDAHP), Oryza sativa (OsDAHP), Ricinus communis (RcDAHP) and also a bacterial homolog from Helicobacter pylori (HpB8DAHP) demonstrated highly conserved features (Fig. 2). Overall, the N-terminal amino acids (cTP, except for bacterial homolog HpDAHP) were quite variable, whereas the core sequence was strongly conserved. Table 1 communicates all identity comparisons, where PhDAHP1 shared the greatest identity with DAHP synthase homologs from Solanaceous species (approximately 90%), while PhDAHP2 shared approximately 74–81% identity with all DAHP homologs used in the alignment (except HpB8DAHP). In short, the molecular and bioinformatic approaches conducted suggest MD retains at least two individual DAHP synthase isoforms.

2.2. PhDAHP1 and PhDAHP2 transcript accumulation analyses Spatial and floral developmental transcript accumulation analyses were executed using quantitative reverse transcriptase polymerase chain reaction (qRT-PCR) with a comparative CT (DDCT) method (Fig. 3). Total RNA from MD root, stem, stigma, anther, leaf, petal tube, petal limb, and sepal tissue was isolated, purified, and quantified. PhDAHP1 transcript accumulated to an approximate 19 times higher level in petal limb and an eight times higher level in the petal tube tissue compared to root tissue (Fig 3A). PhDAHP2 transcript accumulated to an approximate five times higher level in leaf and three times higher level in stem tissues compared to root. The floral developmental series used total RNA isolated from whole MD flower tissue, which was harvested at six of 11 consecutive stages (Colquhoun et al., 2010b). The six stages included

flower tissue of a non-elongated bud (stage 1), a 3 cm elongated bud (stage 3), fully elongated bud directly prior to flower opening (stage 5), directly after flower opening (stage 7), flower open day 3 (stage 9), and a floral senescence (stage 11) (Fig. 3B). PhDAHP2 transcript accumulation remained at relatively low levels, and was fairly constitutive throughout the flower developmental stages. PhDAHP1 transcript demonstrated a relatively low level of accumulation during closed floral bud stages, however, PhDAHP1 transcript accumulated to levels approximately 19 times greater in open flower, stage 9, compared to stage 3, closed floral bud stage (Fig. 3B). Transcriptional profiling of the PhDAHPs indicates PhDAHP1 as the primary player in FVBP biosynthesis. Much like other FVBP gene network members that have a direct biochemical role (Colquhoun et al., 2010b), PhDAHP1 transcript accumulates to relatively high levels in flower limb tissue during open flower stages of development (Fig. 3). Whereas, PhDAHP2 transcript accumulates to highest levels in stem and leaf tissues (Fig. 3A). The regulation of PhDAHP1 transcription appears coordinated with the FVBP gene network and coincides with FVBP biosynthesis and emission.

2.3. Chloroplast import assay To test the in silico predicted localization of PhDAHP1 and PhDAHP2 proteins, the full-length coding sequence for PhDAHP1 and PhDAHP2 were cloned into pGEMÒ-T Easy vectors in the SP6 direction, in vitro transcribed, and translated. The radiolabeled translation product was incubated with isolated and purified chloroplasts (Pisum sativum) in a protein import assay (Fig. 4). The PhDAHP1 and PhDAHP2 radiolabeled protein products were similar in mass to the predicted 59 kDa and 57 kDa, respectively. PhDAHP1 associated with the isolated chloroplast and was protected by the thermolysin treatment, which indicated that PhDAHP1 was successfully imported into the plastid (Fig. 4). Imported PhDAHP1 was detected at a smaller mass compared to the original radiolabelled product suggesting proper chloroplast import processing, i.e., cleavage of the N-terminal transit peptide (Martin et al., 2009). Similarly, PhDAHP2 was associated with the isolated chloroplast and was protected from thermolysin treatment; however, the importation of PhDAHP2 appeared less efficient compared to that of PhDAHP1. Additionally, PhDAHP2 demonstrated an intermediate processing form in the chloroplast much like that of the positive import control, PsSCY2 (Fig. 4). Both PhDAHP1 and PhDAHP2 were able to enter the chloroplast and were detected in all fractionated samples. Like other plant species (Dyer et al., 1990; Gorlach et al., 1993; Keith et al., 1991; Wang et al., 1991), two distinct DAHP synthase like transcripts were identified in MD petunia (Figs. S1 and S2). The most variable region of PhDAHPs is the N-terminal amino acids (Fig. 2), which contain the predicted cTP. The N-terminus of PhDAHP1, much like the relationship of AtDAHP1 to AtDAHP2, has two smaller regions of additional amino acids when compared to PhDAHP2. The discrepancy in N-terminal amino acid content may be responsible for the difference in plastid import efficiency between PhDAHP1 and PhDAHP2 (Fig. 4), which could result in a fraction of PhDAHP2 in the cytosolic space at any given condition or time. Previous work in Nicotiana observed DAHP activity in the cytosolic space, but did not identify the responsible gene (Ganson et al., 1986). Perhaps corollary, both Arabidopsis and petunia, a cytosolic CHORISMATE MUTASE (CM) have been identified and characterized, but any conclusions as to a function for the cytosolic localization were speculative at best (Colquhoun et al., 2010a; Eberhard et al., 1996). It may be functionally useful for the plant cell to retain certain enzymatic redundancy in the cytosol during periods of development, stress, or recycling of plastids.

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Fig. 2. Predicted amino acid sequence alignment for DAHP synthase proteins from various species. The represented sequences are from Petunia  hybrida cv ‘Mitchell Diploid’ (PhDAHP1 and PhDAHP2), Nicotiana tabacum (NtDAHP), Solanum tuberosum (StDAHP), Solanum torvum (SlvDAHP), Morinda citrifolia (McDAHP), Populus trichocarpa (PtDAHP), Arabidopsis thaliana (AtDAHP1 and AtDAHP2), Solanum lycopersicum (SlDAHP), Lotus japonicas (LjDAHP), Oryza sativa (OsDAHP), Ricinus communis (RcDAHP) and also one bacteria homolog (HpB8DAHP1). As described (Larkin et al., 2007), the multiple alignment mode of the ClustalX 2.0 program software was used to align the sequences. The different colors in the alignment signify groups of amino acids: red – K and R; orange – G; magenta – D, C, and E; yellow – P; green – S, N, Q, and T; cyan – A, L, V, I, F, M, and W; blue – Y and H. Displayed above the alignment, ‘‘*’’ signifies a fully conserved residue, ‘‘:’’ signifies a ‘strong’ conservation, and ‘‘.’’ signifies a ‘weak’ conservation. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

Table 1 Predicted protein sequences of various DAHP synthase homologs aligned using ClustalX 2.0 program software. The sequences represented in this table include Petunia  hybrida cv ‘Mitchell Diploid’ (PhDAHP1 and PhDAHP2), Nicotiana tabacum (NtDAHP), Solanum tuberosum (StDAHP), Solanum torvum (SlvDAHP), Morinda citrifolia (McDAHP), Populus trichocarpa (PtDAHP), Arabidopsis thaliana (AtDAHP1 and AtDAHP2), Solanum lycopersicum (SlDAHP), Lotus japonicas (LjDAHP), Oryza sativa (OsDAHP), Ricinus communis (RcDAHP) and also one bacteria homolog (HpB8DAHP1). The values signify the percentage of identity between the homologs.

AtDAHP1 HpDAHP OsDAHP AtDAHP2 LjDAHP PhDAHP2 NtDAHP SlDAHP StDAHP StvDAHP PhDAHP1 McDAHP

HpDAHP

OsDAHP

AtDAHP2

LjDAHP

PhDAHP2

NtDAHP

SlDAHP

StDAHP

StvDAHP

PhDAHP1

McDAHP

PtDAHP

54

77 53

75 53 76

75 54 78 80

74 54 74 80 81

75 52 78 78 78 77

74 52 78 79 79 77 91

74 52 78 79 78 77 92 97

73 51 77 78 77 76 91 96 99

76 51 78 80 78 77 91 89 89 88

76 52 79 79 80 78 83 83 84 83 83

76 52 81 79 80 78 84 83 83 82 83 84

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Fig. 4. Plastid import assay. Radiolabeled translation products for PhDAHP1, PhDAHP2, and PsSCY2 were incubated with isolated pea (Pisum sativum) chloroplasts. After import, the isolated chloroplasts were treated with thermolysin. Proteolysis was terminated and the intact chloroplasts were then repurified, washed, lysed, and fractionated. The inner envelope protein, PsSCY2 (Skalitzky et al., 2011), was used as a positive control. The translation products (tp), chloroplast (Cp), thermolysin treated chloroplasts (Cp*), stromal extracts (SE), and total membrane fraction (M) were analyzed with SDS–PAGE and fluorography. The molecular weight marker shown on left.

plants looked and grew similar to control MD plants and set viable seed when self pollinated. The T0 lines were screened for a reduction in endogenous PhDAHP synthase transcript using sqRT-PCR with transcript specific primers. Total RNA was isolated and purified from whole corollas at stage 8 from all transgenic and control plants. Numerous T0 ir-PhDAHP1 plants exhibited reduced PhDAHP1 transcript accumulation, while PhDAHP2 transcript appeared consistent with controls (Fig. S4). In line, numerous T0 ir-PhDAHP2 plants exhibited reduced PhDAHP2 transcript accumulation, while PhDAHP1 transcript appeared consistent with controls. To directly test the hypothesis that a specific petunia DAHP synthase functions in the production of floral fragrance, floral volatiles

Fig. 3. PhDAHP1 and PhDAHP2 transcript accumulation analysis (one-step qRTPCR). Spatial analysis was performed using MD root, stem, stigma, anther, leaf, petal (P.) tube, petal (P.) limb, and sepal tissues that were harvested at 16:00 h (A). Floral developmental analysis used MD flowers from six intermittent stages of an 11 sequential stage series that was collected in one day at 16:00 h (B). Total RNA (50 ng) was used per reaction in all cases. Histograms are representative of multiple experiments and multiple biological replicates, and analyzed by the DDCt method with PhFBP1 and Ph18S as internal references (mean ± SE; n = 3). Significant differences were supported by a one-way ANOVA (t-distribution) P < 0.05.

2.4. RNAi mediated suppression of PhDHS1 and PhDHS2 Transcript accumulation of endogenous PhDAHP1 and PhDAHP2 was transgenically downregulated in the MD genetic background utilizing a standard RNAi-mediated gene silencing approach with the hypothesis that a reduction of PhDAHP1 protein would result in a reduction of overall floral volatile benzenoid/phenylpropanoid emission. RNAi constructs were generated using 287 nt and 226 nt fragments at the 50 end of the coding regions for PhDAHP1 and PhDAHP2 as RNAi inducers. The two fragments used were approximately 50% identical to each other (Fig. S3) suggesting that the fragments may be individually specific and would not influence the counter PhDAHP transcript. Leaf disc transformation procedures (Jorgensen et al., 1996) generated approximately 50 independent PhDAHP1 RNAi (ir-PhDAHP1) plants and 50 independent PhDAHP2 RNAi (ir-PhDAHP2) plants. The vast majority of transgenic

Fig. 5. PhDAHP1 and PhDAHP2 transcript accumulation analysis between MD and a representative selection of a segregating T1 ir-PhDAHP1 line. One-step qRT-PCR used with 50 ng of total RNA isolated from stage 8 flowers at 16:00 h, and optimized transcript specific primers. PhFBP1 and Ph18S served internal controls. Histograms are representative of multiple experiments and multiple biological replicates, and analyzed by the DDCt method with (mean ± SE; n = 3).

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Fig. 6. Volatile emission analysis of MD and a representative selection of T0 plants. Each volatile collection was conducted at 18:00 h and used developmental stage 8 flowers (mean ± SE; n = 3). Twelve major petunia volatile compounds were identified and quantified (ng/g fresh weight/hour). Samples denote multiple experiments and biological replications.

were collected and analyzed from the transgenic plants along with control MD plants. Using the floral volatile benzenoid, methyl benzoate (20) as a screening proxy, numerous ir-PhDAHP1 plants with reduced endogenous PhDAHP1 transcript accumulation (Fig. S4)

exhibited a reduced emission of the main constituent of the floral volatile bouquet, methyl benzoate (20), when compared to control MD (Fig. S5). In contrast, numerous ir-PhDAHP2 plants with reduced endogenous PhDAHP2 transcript accumulation (Fig. S4),

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did not exhibit a reduced level of methyl benzoate (20) emission when compared to controls (Fig. S5). All of the T0 ir-PhDAHP1 plants exhibiting both transcriptional and metabolic phenotypes were self pollinated and produced viable T1 progeny. Multiple T1 ir-PhDAHP1 lines segregated in an expected manner for the transgene and phenotype. As a representative, ir-PhDAHP1 line 48 T1 plants were used for a quantitative transcript accumulation assay of endogenous PhDAHP1 and PhDAHP2 transcript levels compared to MD (Fig. 5). Multiple plants demonstrated an approximate 80% reduction in DAHP1 transcript (ir-48-4 and ir-48-7), while DAHP2 transcript levels remained similar to controls. When the same plants are analyzed for total FVBP emission profiles, a similar profile is shown (Fig. 6). The majority of the volatile compounds emitted by ir-48-4 and ir-48-7 flowers were reduced compared to control flowers. These include phenylacetaldehyde (13), 2-phenylethanol (14), benzyl alcohol (13), and benzaldehyde (18), which were almost completely reduced. While the volatile compounds isoeugenol (29) and methyl salicylate (22) were the least effected (if at all) compared to MD, these data support the involvement of PhDAHP1 in FVBP biosynthesis. Specific downregulation of the endogenous PhDAHP1 transcript through RNAi-mediated gene silencing reduced FVBP emission in a similar fashion as other pre-phenylalanine enzymes (PhCM1 and PhADT1) when downregulated in petunia (Colquhoun et al., 2010a; Maeda et al., 2010). The most reduced volatiles are the volatiles derived directly from phenylalanine (12), and the least reduced volatiles are the furthest removed from phenylalanine (12) (Fig. 6). Phenylacetaldehyde (13) and 2-phenylethanol (14) were nearly undetectable in the ir-PhDAHP1 background. Differing levels of reduction are observed for the benzenoids, but seemingly PhBSMT drives the majority of carbon to the production of methyl benzoate (20). Similar to what has been shown under carbon limiting situations in other transgenic petunias (Colquhoun et al., 2012, 2010a; Maeda et al., 2010), isoeugenol (29) levels were the least reduced FVBP in the ir-PhDAHP1 lines (Fig. 6). A plausible interpretation of these volatile profile results under limiting substrate availability is that the greatest demand for phenylalanine is the PAL/C4H endomembrane bound complex (Achnine et al., 2004). Assuming, during peak accumulation of phenylalanine (12) for eventual FVBP biosynthesis in floral limb tissue of an open MD flower there are three relatively large draws from the massive phenylalanine (12) pool: PhPAAS, soluble PhPAL, and PhPAL/PhC4H complexes (Fig. 1). As was discussed in Maeda et al., 2010, the apparent Km for PhPAAS using phenylalanine as a substrate is much higher than the Km for soluble PAL (consensus number from various other species). However, the kinetic values and specific associations for the PAL/C4H complex are unknown and most plant species contain multiple PAL and C4H isoforms. Noteworthy here, two recombinant C4H isoforms from P. trichocarpa demonstrate much greater enzymatic efficiency when in a hetrodimeric complex (Chen et al., 2011). At best, a PAL/C4H complex should exist in petunia based on work done in Nicotiana and potato demonstrating the activities and interaction of both PAL and C4H in microsomal fractions (Achnine et al., 2004; Deshpande et al., 1993; Rasmussen and Dixon, 1999). Another key finding from those studies is the quantity of PAL isoform in complex with C4H may be determined by the amount of C4H protein available. This could suggest that all C4H protein is in a complex with PAL isoforms, leaving specific amounts of soluble PAL available for (E)-cinnamic acid (23) production to supply volatile benzenoid formation. In petunia, it is clear that (E)-cinnamic acid (23) is sequestered, accumulates in peroxisomes, and is the initial substrate for the boxidation pathway, which leads to volatile benzenoids (Colquhoun et al., 2012; Klempien et al., 2012). Therefore, under reduced phenylalanine (12) conditions, the PAL/C4H complex could produce p-coumaric (24) acid at a more or less wildtype rate,

while soluble PAL isoforms out compete PhPAAS to generate (E)-cinnamic acid (23) that is then sequestered into peroxisomes. 3. Conclusions Over a decade ago, the genes, transcripts, and proteins responsible for the production of specific petunia FVBPs were first identified and characterized (Kolosova et al., 2001a; Negre et al., 2003). Four years ago, enzymes (post-chorismic acid) connecting the shikimate to the phenylpropanoid pathways were characterized (Colquhoun et al., 2010a; Maeda et al., 2010). Here, a specific DAHP synthase was characterized in petunia that appears responsible for the coupling of primary metabolism to secondary metabolism with the overarching function to supply carbon for FVBP biosynthesis in floral limb tissue of an open flower. Two individual MD transcripts were identified with strong homology to other plant and bacterial DAHPs. PhDAHP1 transcription is regulated similarly to other FVBP genes, recombinant PhDAHP1 is able to enter plastids, and a reduction inFVBP emission is observed when PhDAHP1 transcript is down-regulated via RNAi. The work presented here, supports for the specific involvement of PhDAHP1 in FVBP biosynthesis. These results empirically demonstrate a specific function in a specific tissue and developmental state for a DAHP synthase isozyme. 4. Experimental 4.1. Plant material Inbred P.  hybrida cv ‘Mitchell Diploid’ (MD) plants were utilized as a ‘wildtype’ or as a control in all experiments (Mitchell et al., 1980). All petunia plants were grown in glass greenhouses to reproductive maturity from seed as previously described (Dexter et al., 2007a). 4.2. PhDAHP1 and PhDAHP2 cloning Petunia nucleic acid sequences with high similarity to A. thaliana AtDAHP1 (At4g39980) and AtDAHP2 (At4g33510) were collected using publicly available databases: the National Center for Biotechnology Information (www.ncbi.nlm.nih.gov), Sol Genomics Network (http://solgenomics.net), and the 454 petunia database (http://140.164.45.140/454petuniadb). The resulting sequences were cataloged and constructed into contigs representing partial nucleotide sequences for PhDAHP1 and PhDAHP2 using Vector NTIÒ Advance 11 software (Invitrogen, Grand Island, NY). SMARTER™ RACE technology was then utilized to clone the fulllength PhDAHP1 and PhDAHP2 transcripts, via the SMARTER™ RACE cDNA amplification kit (Clontech Laboratories Inc., http:// www.clontech.com) according to the manufacturer’s protocol. DNA Primers (PhDAHP1 forward primer 50 -cagcaacaccctccatctc-30 ; PhDAHP1 reverse primer 50 -agcatttgaacctttccacgta-30 and PhDAHP2 forward primer 50 -atccggggtctgtgtacattctt-30 ; PhDAHP2 reverse primer 50 -aagtaggggaagggtagcaaga-30 ) were designed approximately 80–100 nucleotides 50 and 30 of the deduced 1599 nucleotide coding region for PhDAHP1 and the deduced 1536 nucleotide coding region for PhDAHP2. Replicates of the expected, approximate 1692 and 1633 nucleotide products were amplified using AdvantageÒ 2 Polymerase Mix (Clontech Laboratories Inc., http:// www.clontech.com) and purified using QIAquick™ Spin Columns (http://qiagen.com). Amplicons for PhDAHP1 and PhDAHP2 were then ligated into a pGEMÒ-T-easy vector (Promega, Madison, WI), transformed into One ShotÒ Mach1™ T1R chemically competent E. coli (Invitrogen Corp., http://invitrogen.com), and multiple clones from multiple amplicons were isolated and sequenced

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(Big Dye V1–2; University of Florida Interdisciplinary Center for Biotechnology Research) to at least a 6X coverage to check for errors. 4.3. RNA isolation and tissue collection For all cases, total RNA was extracted as previously described (Verdonk et al., 2003) and subjected to TURBO™ DNA-free™ treatment (Ambion Inc., Austin, TX) followed by purification with the RNeasyÒ Mini protocol for RNA cleanup (Qiagen, Valencia, CA). Total RNA was then quantified via a NanoDrop™ 2000c spectrophotometer (Thermo Scientific, Wilmington, DE) and 50 ng lL1 dilutions were generated and kept at 20 °C. All petunia tissue collections were done as previously described (Colquhoun et al., 2010b); the following gives a brief description. The spatial transcript accumulation analysis consisted of total RNA isolated from petunia root, stem, stigma, anther, leaf, petal tube (stage 8), petal limb (stage 8), and sepal tissues from multiple MD plants harvested at 16:00 h. The floral, developmental transcript accumulation analysis utilized total RNA from MD petunia flowers collected at 11 consecutive stages beginning at a small bud to floral senescence from multiple greenhouse-grown plants at 16:00 h. For all tissue collections, individual samples consisted of three flowers. For the comparative transcript accumulation analysis between control MD and transgenic ir-PhDAHP1 and irPhDAHP2 plants, developmentally staged (stage 8) flowers were collected at 16:00 h. All samples were frozen in liquid N2 and stored at 80 °C. Total RNA was then isolated for all samples, with multiple biological replicates. 4.4. Transcript accumulation analysis All transcript accumulation analyses were conducted multiple times with multiple biological replicates and equivalent results were observed. sqRT-PCR was performed using the Qiagen OneStep RT-PCR kit (Qiagen Co., Valencia, CA, USA) with total RNA (50 ng). In all sqRT-PCR experiments the gene specific primers were used in multiple reactions at different cycle numbers to qualify initial observations. To visualize RNA loading concentrations, samples were amplified with Ph18S primers (forward primer 50 ttagcaggctgaggtctcgt-30 ; reverse primer 50 -agcggatgttgcttttagga30 ) and analyzed on an agarose gel. The following gene specific primers were designed and utilized for the visualization of mRNA levels corresponding to PhDAHP1 and PhDAHP2, respectively: PhDAHP1 (forward primer 50 -taccgagaactagctaacag-30 ; PhDAHP1 reverse primer 50 -tacgtggaaaggttcaaatgct-30 ) and PhDAHP2 (forward primer 50 -gactacaacagacttctgga-30 ; PhDAHP2 reverse primer 50 -tcttgctacccttcccctactt-30 ). DDCt Quantitative qRT-PCR was performed and analyzed using StepOnePlus™ real-time PCR system (Applied Biosystems, Foster City, CA). Power SYBRÒ Green RNAto-Ct™ 1 and 2-Step kits (Applied Biosystems, Foster City, CA) were used to amplify and detect the products according to protocol by the manufacturer. The following, qRT-PCR primers were constructed in Primer ExpressÒ software v2.0 (Applied Biosystems, Foster City, CA): PhDAHP1 (forward primer 50 -gtcttgctacccttcccctac-30 ; PhDAHP1 reverse primer 50 -gtcttgctacccttcccctac-30 ) and PhDAHP2 (forward primer 50 -cggtgacaaaacaagaatgg-30 ; PhDAHP2 reverse primer 50 -ttcaagatgccttgcttcac-30 ). Optimization of primers was conducted and demonstrated gene specificity during melt curve analysis. 4.5. PhDAHP1 and PhDAHP2 RNAi To test the gene function of PhDAHP1 and PhDAHP2 directly, RNAi-induced gene silencing was utilized. For PhDAHP1, a 287 nt sequence at the 50 end of the coding sequence was developed for

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the RNAi inducing fragment. (PhDAHP1 forward primers 50 -gctctagatcactatagggcaagcagtggta-30 , 50 -cgggatcctcactatagggcaagcagtggta30 , and PhDAHP1 reverse primers 50 -ggaattcacactcccattgtctcagcc-30 , 50 -ggaattcagaatggctggacaatttgc-30 ). For PhDAHP2, a 226 nt sequence at the 50 end of the coding sequence was developed for the RNAi inducing fragment. (PhDAHP2 forward primers 50 -gctctagacggggtctgtgtacattcttca-30 , 50 -cgggatcccggggtctgtgtacattcttca-30 , and PhDAHP2 reverse primers 50 -ggaattccacatctcagccgttcaat-30 , 50 ggaattcagaaggatggcgtgaagcta-30 ). In planta expression of this fragment was driven by a constitutive promoter, pFMV. Fifty independent PhDAHP1 and 50 independent PhDAHP2 RNAi (ir-PhDAHP, inverted repeat) plants were generated in the MD background by leaf disc transformation (Jorgensen et al., 1996). Further details of the technical cloning have been previously described (Dexter et al., 2007a; Underwood et al., 2005).

4.6. Chloroplast import assay Full-length coding sequences for PhDAHP1 and PhDAHP2 were cloned into a pGEMÒ-T Easy vector (Promega, Madison, WI) in the SP6 direction. The chloroplast import assay was performed as described previously (Colquhoun et al., 2010a; Martin et al., 2009). Briefly, in vitro transcription and translation using wheat germ TNTÒ (Promega, Madison, WI) resulted in radiolabeled, recombinant PhDAHP1, PhDAHP2, and PsSCY2. These recombinant proteins were incubated individually with isolated pea (P. sativum) chloroplasts for 15 min. Subsequent to incubation; the isolated chloroplasts were treated with thermolysin (100 lg ml1) for 40 min at 4 °C. Proteolysis was ended by the addition of EDTA to a final concentration of 10 mm. The intact chloroplasts were then re-purified by centrifugation (8 min at 3200g) through a 35% Percoll™ (GE Healthcare, Waukesha, WI) gradient. Chloroplasts were washed, lysed, and fractionated into total membranes and stromal extracts by centrifugation for 18 min at 15,000g. The translation products, chloroplasts, thermolysin-treated chloroplasts, stromal extracts and total membranes were analyzed via SDS–PAGE and fluorography.

4.7. Volatile emission For all floral volatile emission experiments, emitted floral volatiles from excised flowers were collected at 16:00 h and quantified as previously described (Dexter et al., 2007a; Underwood et al., 2005). Briefly, flower samples were loaded into thin walled glass tubes (2.5 cm i.d., 61 cm long, and 300 mL volume) attached to a push–pull dynamic headspace volatile collection system with a column containing approximately 50 mg HaySep Q 80–100 porous polymer adsorbent (Hayes Separations Inc., Bandera, TX) at the tube outlet to capture volatile organic compounds over a period of one hour, all at room temperature. Volatiles were eluted from the column with 150 lL methylene chloride with nonyl acetate (5 lL) as an elution standard. Two microliter of the elution samples were run on an Agilent 7890A series gas chromatograph flame ionization detector (GC-FID). Peaks from the chromatography traces of compounds within a sample elution were inferred using Chemstation software (Agilent Technologies, Santa Clara, CA). Peak areas of detected analytes were used to determine the mass of each compound within a sample by calculations that adjusted for the nonyl acetate elution standard and the original biological sample mass (Schmelz et al., 2001, 2004). GC–MS (mass spectrometry) was used to confirm the presence of compounds and identify retention times in conjunction with chemical standards (Sigma–Aldrich, St. Louis, Missouri) run on the GC-FID.

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