- Research article
- Open Access
Expression of flavonoid 3’-hydroxylase is controlled by P1, the regulator of 3-deoxyflavonoid biosynthesis in maize
© Sharma et al.; licensee BioMed Central Ltd. 2012
- Received: 24 January 2012
- Accepted: 29 October 2012
- Published: 1 November 2012
The maize (Zea mays) red aleurone1 (pr1) encodes a CYP450-dependent flavonoid 3’-hydroxylase (ZmF3’H1) required for the biosynthesis of purple and red anthocyanin pigments. We previously showed that Zmf3’h1 is regulated by C1 (Colorless1) and R1 (Red1) transcription factors. The current study demonstrates that, in addition to its role in anthocyanin biosynthesis, the Zmf3’h1 gene also participates in the biosynthesis of 3-deoxyflavonoids and phlobaphenes that accumulate in maize pericarps, cob glumes, and silks. Biosynthesis of 3-deoxyflavonoids is regulated by P1 (Pericarp color1) and is independent from the action of C1 and R1 transcription factors.
In maize, apiforol and luteoforol are the precursors of condensed phlobaphenes. Maize lines with functional alleles of pr1 and p1 (Pr1;P1) accumulate luteoforol, while null pr1 lines with a functional or non-functional p1 allele (pr1;P1 or pr1;p1) accumulate apiforol. Apiforol lacks a hydroxyl group at the 3’-position of the flavylium B-ring, while luteoforol has this hydroxyl group. Our biochemical analysis of accumulated compounds in different pr1 genotypes showed that the pr1 encoded ZmF3’H1 has a role in the conversion of mono-hydroxylated to bi-hydroxylated compounds in the B-ring. Steady state RNA analyses demonstrated that Zmf3’h1 mRNA accumulation requires a functional p1 allele. Using a combination of EMSA and ChIP experiments, we established that the Zmf3’h1 gene is a direct target of P1. Highlighting the significance of the Zmf3’h1 gene for resistance against biotic stress, we also show here that the p1 controlled 3-deoxyanthocyanidin and C-glycosyl flavone (maysin) defence compounds accumulate at significantly higher levels in Pr1 silks as compared to pr1 silks. By virtue of increased maysin synthesis in Pr1 plants, corn ear worm larvae fed on Pr1; P1 silks showed slower growth as compared to pr1; P1 silks.
Our results show that the Zmf3’h1 gene participates in the biosynthesis of phlobaphenes and agronomically important 3-deoxyflavonoid compounds under the regulatory control of P1.
The maize (Zea mays) flavonoid biosynthesis provides an excellent system to study gene interaction in plants because of its extensive characterization at genetic, biochemical, and molecular levels . Different flavonoid compounds share the same basic skeleton of the flavan-nucleus consisting of two aromatic rings with six carbon atoms (ring A and B) which are interconnected by a hetero-cyclic ring with three carbon atoms (ring C). Maize produces 3-hydroxyflavonoids (anthocyanidins) and 3-deoxyflavonoids which include phlobaphenes, 3-deoxyanthocyanidins, and C-glycosyl flavones. These compounds are synthesized in different tissues and this spatial distribution depends on the genetic constitution of the plant. Anthocyanins can accumulate in most plant parts whereas phlobaphenes are predominantly found in kernel pericarp (outer layer of ovary wall), cob-glumes (palea and lemma), tassel glumes, and husk . The 3-deoxyanthocyanins and C-glycosyl flavones primarily accumulate in silks [3–5]. However, in some high altitude maize lines C-glycosyl flavones can also accumulate in leaves  indicating genetic diversity for developmental accumulation of flavonoid metabolites.
The 3-hydroxy- and 3-deoxy-flavonoids in maize are regulated by independent sets of transcription factors. Accumulation of 3-hydroxyflavonoids (anthocyanins) is controlled by two sets of duplicated genes: colorless1 (c1)/purple leaf1 (pl1) are members of the R2R3-MYB family of transcription factors , and booster1 (b1)/red1 (r1) are members of the basic helix-loop-helix (bHLH) family [8, 9]. Studies have shown that C1 or PL1 proteins interact directly with R1 or B1 to activate transcription of anthocyanin biosynthetic genes in seed and plant body, respectively [10, 11]. In contrast, 3-deoxyflavonoid pathway genes are regulated by pericarp color1 (p1), which encodes an R2R3-MYB transcription factor . The p locus is a complex of duplicated MYB-homologous genes p1 and p2 on chromosome 1 . The p locus is a major QTL for the biosynthesis of C-glycosyl flavones [14, 15] and 3-deoxyanthocyanidins in silks .
Three flavonoid biosynthetic genes; colorless2 (c2), chalcone isomerase1 (chi1), and anthocyaninless1 (a1) encode chalcone synthase (CHS), chalcone isomerase (CHI), and dihydroflavonol 4-reductase (DFR), respectively. These three genes are common to the anthocyanin and phlobaphene pathways, but are independently regulated by the corresponding sets of transcription factors [10, 17, 18]. In vitro and in vivo studies have shown that C1 + R1 or P1 can direct high level of expression from promoters containing the C1/R1 or P1 binding sites identified previously in the a1 and c2 gene promoter [12, 19–21].
Pr1; P1cob-glumes accumulate luteoforol
Genotype and phenotype of different lines developed and used in this study
Two flavan 4-ols, luteoforol and apiforol have been implicated as precursors of phlobaphene pigments that accumulate in maize genotypes carrying functional p1 or p2 genes. Cob glumes were used to perform biochemical characterization of flavan 4-ols. The dark red cob glumes from Pr1/Pr1; P1 wr/P1 wr had maximum absorption (λ max) at 552 nm, while light red cob glumes from pr1/pr1; P1 wr/P1 wr plants had λ max at 535 nm (Figure 2B). These absorption spectra correspond to luteoforol and apiforol, respectively . To further confirm if the Zmf3’h1 plays a role, flavan 4-ols were converted into their corresponding 3-deoxyanthocyanidins by acid treatment of methanolic extracts (Figure 2B). Extracts from Pr1/Pr1; P1 wr/P1 wr converted to luteolinidin (λ max 498 nm) indicating the presence of luteoforol in the methanolic extracts. Similarly, extracts of pr1/pr1; P1 wr/P1 wr cob glumes were converted into apigeninidin (λ max 482 nm) indicating presence of apiforol in the extract. Acid treated methanolic extracts from cob glumes of Pr1/Pr1; P1 rr/P1 rr and pr1/pr1; P1 rr/P1 rr also had maximum absorption wavelengths corresponding to luteolinidin and apigeninidin, respectively. No detectable flavan 4-ols or corresponding 3-deoxyanthocyanidins accumulated in cob glumes of pr1/pr1; p del2/p del2 or Pr1/Pr1; p del2/p del2. This was in accordance with p1 gene’s function as a regulator of phlobaphene biosynthesis . Taken together, these results show that cob glumes from pr1; P1 plants accumulate apiforol whereas luteoforol accumulates in cob glumes of Pr1; P1 plants. This result also indicates that the accumulation of apiforol and luteoforol is influenced by a flavonoid 3’-hydroxylase acting in parallel for the conversion of naringenin to eriodictyol.
P1 regulates the transcription of Zmf3'h1
P1 binds to two sites in the Zmf3’h1promoter
P1 binds the Zmf3 ′ h1 promoter in vivo
pr1; P1plants do not accumulate luteolinidin in silks
Silks of pr1; P1plants have reduced maysin accumulation
Pr1; P1silks is detrimental to corn earworm larvae development
To determine the biological relevance of differential accumulation of maysin and apimaysin in silks of Pr1 and pr1 plants, we performed insect silk feeding bioassays. Corn earworm (Helicoverpa zea Boddie) larvae (neonate stage) were fed on fresh silks (Additional file 1: Figure S1) collected from the same Pr1 and pr1 plants that were used for HPLC analysis. Larvae fed on Pr1 silks had lower body weight and took longer time to pupate as compared to those fed on pr1 silks (Figure 7C). These results are in agreement with the accumulation of higher amounts of maysin in Pr1 silks. Interestingly, larvae fed on pr1/pr1; P1 wr/P1 wr silks showed lower weight and longer time to pupate as compared to those fed on Pr1/Pr1; P1 wr/P1 wr silks, even though the Pr1 silks had higher levels of maysin. This anomaly could possibly be because of the accumulation of exceptionally higher level of apimaysin in pr1/pr1; P1 wr/P1 wr (see Figure 7B, panel II). As shown in previous studies, apimaysin has insecticidal activity against lepidopteron insects, although, apimaysin’s activity levels were lower than maysin . Overall, combined data from HPLC and larvae feeding bioassay indicated that a functional Zmf3’h1 participates in the accumulation of 3’-hydroxylated C-glycosyl flavones that affect the growth of corn earworm larvae.
The pr1 locus has been extensively used as phenotypic marker in maize genetics research because of its role in determining kernel aleurone color by hydroxylation of anthocyanin compounds (3-hydroxyflavonoids) [24, 37]. However, little is known about the function and regulation of the pr1 encoded ZmF3’H1 in 3-deoxyflavonoids pathway. The 3-deoxyflavonoids include phlobaphene pigments [38, 39] and agronomically important C-glycosyl flavones and 3-deoxyanthocyanidins which provide resistance against various biotic stresses [4, 31, 40–42]. Maize p1 gene regulates the 3-deoxyflavonoid biosynthetic pathway [12, 17]. Here, we describe the first direct evidence of the involvement of Zmf3’h1 in the 3-deoxyflavonoid pathway and its regulation by P1 MYB transcription factor. We have demonstrated that the dark red cob glumes of Pr1; P1 plants accumulates luteoforol as compared to apiforol accumulating in light red cob glumes of pr1; P1 plants. Further, gene expression analysis confirmed that transcription of Zmf3’h1 requires p1 gene expression in pericarps, cob glumes, and silks. Interestingly, the detection of Zmf3’h1 transcripts in silks of P1 ww lines suggest that in addition to p1, the paralog p2 is also involved in the regulation of Zmf3’h1 expression in silks. Additionally, the absence of Zmf3’h1 transcripts in p del2 allele which has a deletion of p1 and p2 genes , supported this hypothesis.
P1 is a R2R3-MYB protein and directly regulates the expression of flavonoid biosynthetic genes. Binding of P1 to the cis-regulatory elements of the a1 and c2 gene promoter has been well characterized [12, 21]. Sequence analysis of the Zmf3’h1 promoter shows the presence of similar conserved P1 binding sites. Further, EMSA results demonstrated the in vitro binding ability of P1 to these cis- sites, while ChIP experiments confirmed that Zmf3’h1 is an immediate direct target of P1. In addition to P1 binding sites, Zmf3’h1 promoter also contains anthocyanin regulatory element (ARE), a conserved sequence present in other anthocyanin biosynthetic genes .
Underlining the importance of Zmf3’h1 in maize biotic stress resistance, our work further added that Pr1; P1 plants accumulate significantly higher levels of the antifungal compound luteolinidin in silks. Luteolinidin is known to be toxic towards fungi and it accumulates at higher level in sorghum lines resistant to the anthracnose fungus [43, 44]. In sorghum, attempted penetration of Cochliobolus heterostrophus leads to up regulation of a f3’h gene and sequential accumulation of luteolinidin . The 3-deoxyanthocyanidin pathway in sorghum requires a MYB protein encoded by yellow seed1 (y1), an ortholog of maize p1[33, 45, 46]. Similar to the regulation of Zmf3’h1 by p1, sorghum f3’h is regulated by y1. It remains to be tested if silk extracts containing higher luteolinidin glycosides show resistance to fungal pathogens of maize.
Zmf3’h1 participates in the biosynthesis of the 3’-hydroxylated C-glycosyl flavones with Pr1; P1 silks accumulating higher level of maysin compared to pr1; P1 silks. Structurally, apimaysin and maysin are highly related and differ only by B-ring hydroxylation in position 3’ [47, 48]. Unexpectedly, the accumulation of maysin and apimaysin in Pr1/Pr1; P1 wr/P1 wr and pr1/pr1; P1 wr/P1 wr silks did not exactly follow the inverse correlation. Apimaysin level in pr1/pr1; P1 wr/P1 wr silks increased to a substantially higher level as compared to the maysin level in a Pr1/Pr1; P1 wr/P1 wr line. One possibility is that the apimaysin is acting as a substrate for another enzyme and is converted into a product that we were not able to detect in the analysis. We also measured rhamnosylisoorientin which has been shown to have insecticidal activity [49, 50]. Rhamnosylisoorientin is a C-glycosyl flavone which is present upstream of maysin. Importantly, Pr1 silks have higher level of rhamnosylisoorientin as compared to pr1 in the presence of P1 rr and P1 ww, respectively. Genetic variation at p locus is significantly correlated with maysin accumulation. Genotypes carrying functional p1 or p2 alone accumulate less amount of maysin than maize lines that have both p1 and p2 genes .
Most of the steps in the formation of C-glycosyl flavones are unknown. It is possible that formation of C-glycosyl flavones does not entirely follow a single linear pathway but demonstrates shunting of substrate flow to alternate pathways through which maysin and apimaysin are formed separately [16, 52, 53]. The 3’, 4’-hydroxylated flavonoids, such as maysin and luteoforol could also be formed as a result of hydroxylation of naringenin to eriodictyol by F3’H and then subsequent conversion into intermediates leading to formation of these compounds (Figure 1) [28, 54]. This could also explain the higher level of rhamnosylisoorientin in Pr1 plants. A recent study by Morohashi et al (2012) have shown the isolation and cloning of a FNS/F2H encoding gene capable of converting flavanones to 2-hydroxy flavanones, a previously unknown step in the formation of C-glycosyl flavones . They have also proposed the formation of 4’ and 3’, 4’- hydroxylated compounds through alternate pathways where a F3’H can perform hydroxylation of naringenin to eriodictyol. The accumulation of different levels of flavones and 3-deoxyanthocyanidins in functional p1 alleles could be attributed to polymorphic structural genes at different loci: functional c2, whp1, and a1 genes have a positive effect on maysin accumulation [16, 51].
The significance of flavonoid defence compounds, 3-deoxyanthocyanidin and C-glycosyl flavone has been well established [5, 15, 55]. The current study attempted to unravel the role of regulatory and biosynthetic genes involved in the synthesis of these flavonoids in order to tailor resistant plants. Through transgenic and non-transgenic studies, it established that functional p1 and p2 genes can induce biosynthesis of these compounds [14, 17]. The current study along with a previous report  demonstrates that Zmf3’h1 plays a significant role in generating diversity in anthocyanin, phlobaphene, 3-deoxyanthocyanidin, and C-glycosyl flavone compounds. It will be informative to further analyse the action of Zmf3’h1 at specific steps for the biosynthesis of related phenylpropanoid compounds.
Maize genetic stocks
Standard maize genetic nomenclature is used in the current study . Alleles of the maize (Zea mays) p1 have been identified based on their expression in the floral organs and are named according to their pericarp and cob-glumes pigmentation: P1 wr (white pericarp, red cob), P1 rr (red pericarp, red cob), and P1 ww (white pericarp, white cob) (Figure 3A) [57–60]. The maize inbred lines W23 (genotype P1 wr Pr1 c1 r g), W22 (P1 wr Pr1 C1 R1), and other genetic stocks MGS 14273 (P1 wr pr1 C1 R1) and MGS 14284 (P1 ww pr1 C1 R1) were kindly provided by the Maize Genetics Co-operation Stock Centre (USDA-ARS, University of Illinois, Urbana, IL). The P1 ww [4Co63] inbred line was obtained from the National Seed Storage Laboratory (Fort Collins, CO), while P1 rr 4B2, P1 ww 1112 and p del2 genetic stocks were obtained from Dr. Thomas Peterson, Iowa State University, Ames, IA [61, 62]. The p del2 deletion mutant was derived from P1 vv 9D9A and has a deletion encompassing both p1 and p2[14, 63]. All p1 alleles except p del2 and P1 ww 1112 are in 4Co63 genetic background. Our genetic tests have shown that all these p stocks carry a functional Pr1 allele and their pigmentation phenotypes are presented in Table 1. To develop F2 populations, pr1 MGS14273 plants were crossed with P1 wr, P1 rr 4B2, P1 ww, and p del2 and progenies were grown from selfed F1 plants. These F2 populations showed a 3:1 segregation for purple to red aleurones. To develop homozygous Pr1 and pr1 stocks containing different p alleles, plants from F2 ears showing desirable pericarp, cob-glumes, and kernel aleurone pigmentation phenotypes (see Table 1) were subjected to six subsequent cycles of self-pollination and selection. To confirm the presence of Pr1 or pr1, PCR based genotyping was done using primers in the promoter region .
Analysis of flavan 4-ols
To detect the presence of flavan 4-ols, 500 mg of cob-glumes were macerated with a plastic grinder in an Eppendorf tube containing 1 mL of 30% HCl/70% butanol (v/v) and incubated for 60 min at 37°C . Samples were spun for 10 min at 14,000 rpm and the absorption spectra of the supernatants were determined using a Shimadzu UV-mini 1240 spectrophotometer (Shimadzu Corporation, Columbia, MD) [65, 66]. Apiforol and luteoforol are flavan 4-ols previously described from maize and sorghum that give flavylium ions in acidic butanol with a λ max of 535 and 552 nm, respectively . To confirm the identity of the major flavan 4-ols in cob glumes of Pr1 and pr1 alleles in the genetic background of different p alleles, methanol extract were treated with aqueous HCl. This converts flavan 4-ols such as apiforol and luteoforol to their corresponding 3-deoxyanthocyanidins (i.e. apigeninidin and luteolinidin). The treated Pr1 extracts had λ max of 498 nm that shifted in alcoholic AlCl3 to a shoulder at 546 nm. The addition of HCl restored its absorption to 498 nm. The results of our samples were verified using commercial standards for apigeninidin and luteolinidin (Extrasynthese, Genay Cedex, France). The commercial sample of apigeninidin had a λ max of 475 nm and did not respond to AlCl3, whereas luteolinidin had a λ max of 495 nm that shifted in AlCl3 to 546 nm and reverted to 498 nm upon re-addition of HCl.
RNA gel blot analysis
Silks were collected 2 d after emergence, and pericarps and cob glumes were dissected 20 DAP. To isolate total RNA, tissues were ground in liquid nitrogen and then extracted using Tri-Reagent (Molecular Research Centre Inc., Cincinnati, OH). RNA gel blot hybridizations were performed as described previously . Probe fragments used for RNA gel blot analysis were: plasmid pC2 containing a maize c2 cDNA , pCHI1 containing a maize chi1 cDNA , pA1 with a maize a1 cDNA , pF3’H1 containing Zmf3’h1 cDNA, and pP1 containing full length p1 cDNA . The p1 probe used here can recognize both p1 and p2 transcripts . Filters were stripped by washing thrice in a boiling solution of 0.1% (w/v) SDS before re-hybridization.
Protein expression and purification
NHis6-P1MYB used for EMSA was expressed in Escherichia coli and affinity purified using Ni-NTA beads under natural condition as described previously . Briefly, IPTG induction of 1-liter culture, the cells were harvested by centrifugation and re-suspended in 20 ml of SB buffer (50 mM sodium phosphate, pH 8.0, 100 mM NaCl, and 100 μg/ml phenylmethylsulfonyl fluoride) and passed twice through a French press. The cell lysate was centrifuged and the supernatant was filtered through Mira-cloth (Calbiochem). One ml of 50% slurry Ni-NTA beads (Qiagen) was incubated with the cell lysate supernatant for 2 h with gentle rocking at 4°C. The beads were gently harvested by centrifugation, re-suspended in five ml of SB, and loaded onto a column. The column was washed with SB five times and WB (50 mM sodium phosphate, pH 8.0, 300 mM NaCl, 1% Tween 20, 5 mM 2-mercaptoethanol, 10 mM EDTA, and 10% glycerol) three times. The protein was eluted with five washes of five column volumes of WB containing 50 mM imidazole. The elutions were then dialyzed against A-0 buffer (10 mM Tris pH 7.5, 50 mM NaCl, 1 mM DTT, 1 mM EDTA, and 5% glycerol) and stored at -80°C until further use.
Electrophoretic mobility shift assay (EMSA)
EMSA was performed as previously described . The two Zmf3’h1 promoter fragments used as probes for EMSA were generated by PCR amplification using the following primer pairs: Pr1-1, 5′- GAGTGGGTTGTGGGATTGTT-3′ and 5′- ACCGTAAGGCCAACTCCAAC-3′; Pr1-2, 5′- GCCCGCGAAGAAAAATATAA-3′ and 5′- CCACTTGCGTGCTTCATCTA-3′, in which one of the primer was radioactively labeled with [γ-32P]ATP by using T4-polynucleotide kinase. The radioactively labeled DNA fragments were purified by PAGE and quantified by scintillation counter. Ten ng of purified P1MYB was incubated with an equal molar amount of probes (with radioactivity ~105 CPM) for 30 min and the P1MYB-probe complex was separated by PAGE. After PAGE, the gel was dried onto Whatman paper and then subjected to autoradiography at -70°C.
Chromatin immuno-precipitation assay (ChIP)
ChIP experiments using pericarps were performed as previously described . Real-time PCR was used to detect enriched DNA fragments after ChIP experiments with a minor modification. To adjust for different PCR efficiencies as a consequence of the presence of inhibitory compounds in chromatin obtained from P1 rr pericarps, equal amounts of the pPHP611 plasmid  were spiked in the real-time PCR reaction buffer (Epicentre). Copy number of pPHP611 plasmid  in each reaction was estimated by using the normalization primer sets directed to the β-lactamase gene responsible for Ampr in the plasmid. The primer sets used for ChIP-qPCR were the following: qChIP-ZmCopia-F, 5′-CGATGTGAAGACAGCATTCCT-3′, qChIP-ZmCopia-R, 5′-CTCAAGTGACATCCCATGTGT-3′, qChIP-ZmAct1-5UTR-F, 5′-TTTAAGGCTGCTGTACTGCTGTAGA-3′, qChIP-ZmAct1-5UTR-R, 5′-CACTTTCTGCTCATGGTTTAAGG-3′, qrt Zmf3′h1Prom-A1, 5′-AGATCGCGGGTAGGTAGGAG-3′, and qrt Zmf3’h1Prom-B1, 5′-ACTGGTGGCGAGGGTGTAGT-3′. The following primer sets were used to detect Ampr gene: ChIP-Amp-F, 5′-GTAGTTATCTACACGACGGGGAGT-3′, and ChIP-Amp-R, 5′-ATCAGTGAGGCACCTATCTCAGC-3′.
Analysis of C-glycosyl flavones and 3-deoxyanthocyanidins
Primary ear shoots were covered prior to silk emergence to prevent random pollination. Silks were collected on ice 2 d after emergence from the ear and subsequently freeze dried. Silk samples were then shipped on dry ice to the Richard B. Russell Research Centre (USDA-ARS, Athens, Georgia) for biochemical analysis. Flavones were extracted with 125 mL methanol at -20°C for 14 d. Concentration of flavones were determined by reversed-phase HPLC [16, 74] and expressed as percent dry silk weight. For 3-deoxyanthocyanidins, silks were extracted for 24 h at -20°C with 10 mL of 1% HCl-Methanol (v/v). Their levels were detected at 495 nm by HPLC, with the same column and solvent program used for flavone analysis. Commercial standard of luteolinidin hydrochloride (different than the luteolinidin standard used for spectrophotometric analysis of cob-glumes) was used for quantification (Roth-Atomergic Chemicals Corp., Farmingdale, N.Y.). Chrysin was used as an internal reference standard. Total 3-deoxyanthocyanidin concentration was calculated as the sum of three distinct luteolinidin glycoside peaks a, b, and c .
Corn earworm (Helicoverpa zea Boddie) eggs were obtained from Benzon Research Company, Carlisle, PA. Eggs were incubated at 28°C. They hatched after 48 h to produce neonate larvae. Maize lines with Pr1 or pr1 alleles in genetic backgrounds of four different p1 alleles; P1 wr, P1 rr, P1 ww, and p del2 were grown during the summer of 2007. Silks collected 2 to 3 d after emergence were pooled from 20 field grown plants per genotype. The experiment was conducted as a randomized complete block design with 30 replications and two cups per replicate. Freshly collected silks were filled into 1 oz. plastic diet cups containing 10 mL of 2.5% (w/v) agar to prevent silk drying. Instead of adding silk extracts to the artificial insect diet, fresh silk tissues were used to maximize the resemblance to natural larval feeding conditions. One neonate larvae was introduced into each cup and larvae were allowed to feed in a controlled environment maintained at 28°C, 75% RH, and a photoperiod of 14/10 h (light/dark). Larval weights were recorded after eight days of feeding and larvae were subsequently transferred to artificial diet  until pupation.
We thank Scott Harkcom and the Penn State Agronomy Farm staff for their help with land preparation and tending the maize crop. We are thankful to one anonymous reviewer for critical comments to improve the manuscript. This work was supported in part by research support to S.C. under Hatch projects 4144, 4154, 4452, and 4430 of the College of Agricultural Sciences, Pennsylvania State University, a USDA-NRI-2007-35318-17795 award and by the Agricultural and Food Research Initiative Competitive Grant of the USDA National Institute of Food and Agriculture 2011-67009-30017 to SC and 2010-65115-20408 to EG. This project was also supported by NSF DBI-0701405 to E.G. M.S. was supported by graduate research assistantship from the Department of Crop & Soil Sciences, Pennsylvania State University.
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