Quantitative Trait Loci Controlling Amounts and Types of Epicuticular Waxes in Onion

in Journal of the American Society for Horticultural Science
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  • 1 Department of Horticulture, 1575 Linden Drive, University of Wisconsin, Madison, WI 53706
  • 2 Vegetable Crops Unit, Agricultural Research Service of the U.S. Department of Agriculture; and the Department of Horticulture, 1575 Linden Drive, University of Wisconsin, Madison, WI 53706

Natural variation exists in onion (Allium cepa L.) for amounts and types of epicuticular waxes on leaves. Wild-type waxy onion possesses copious amounts of these waxes, whereas the foliage of semiglossy and glossy phenotypes accumulates significantly less wax. Reduced amounts of epicuticular waxes have been associated with resistance to onion thrips (Thrips tabaci Lindeman), an important insect pest of onion. A segregating family from the cross of waxy and semiglossy onions was used to map single nucleotide polymorphisms (SNPs) and identify chromosome regions affecting amounts and types of epicuticular waxes as measured by gas chromatography–mass spectrometry. The amount of the primary epicuticular wax on onion leaves, hentriacontanone-16, was controlled by one region on chromosome 5. One region on chromosome 2 affected concentrations of several primary fatty alcohols. Results indicate that the region on chromosome 2 may be associated with the acyl reduction pathway, and the region on chromosome 5 may affect the decarbonylation pathway of epicuticular wax biosynthesis. Because lower amounts of epicuticular waxes are recessively inherited, SNPs tagging regions on chromosomes 2 and 5 will be useful for marker-assisted breeding to vary amounts and types of epicuticular waxes on onion foliage with the goal to develop cultivars resistant to onion thrips.

Abstract

Natural variation exists in onion (Allium cepa L.) for amounts and types of epicuticular waxes on leaves. Wild-type waxy onion possesses copious amounts of these waxes, whereas the foliage of semiglossy and glossy phenotypes accumulates significantly less wax. Reduced amounts of epicuticular waxes have been associated with resistance to onion thrips (Thrips tabaci Lindeman), an important insect pest of onion. A segregating family from the cross of waxy and semiglossy onions was used to map single nucleotide polymorphisms (SNPs) and identify chromosome regions affecting amounts and types of epicuticular waxes as measured by gas chromatography–mass spectrometry. The amount of the primary epicuticular wax on onion leaves, hentriacontanone-16, was controlled by one region on chromosome 5. One region on chromosome 2 affected concentrations of several primary fatty alcohols. Results indicate that the region on chromosome 2 may be associated with the acyl reduction pathway, and the region on chromosome 5 may affect the decarbonylation pathway of epicuticular wax biosynthesis. Because lower amounts of epicuticular waxes are recessively inherited, SNPs tagging regions on chromosomes 2 and 5 will be useful for marker-assisted breeding to vary amounts and types of epicuticular waxes on onion foliage with the goal to develop cultivars resistant to onion thrips.

Onion thrips is an important insect pest of onion causing damage to foliage and reducing bulb and seed yields (Diaz-Montano et al., 2010; Elmore, 1949). Thrips also vector serious diseases such as iris yellow spot virus (Gent et al., 2004; Shock et al., 2008). Onion thrips populations can increase rapidly during warmer conditions requiring frequent pesticide applications, and consequently the insect has developed resistance to pyrethroid and organophosphate insecticides (Allen et al., 2005; Herron et al., 2008; Shelton et al., 2006). Because it is desirable to reduce pesticide use and increase integrated pest management strategies in vegetable crops (Eigenbrode and Trumble, 1994), genetic resistance to onion thrips would be beneficial for growers and, combined with fewer pesticide applications, offers a potentially affordable and sustainable management strategy for this important pest.

Reduced amounts of leaf epicuticular waxes (often referred to as glossy or bloomless phenotypes) have been associated with insect resistance in several crops. Bloomless sorghum [Sorghum bicolor (L.) Moench.] showed resistance to greenbug [Schizaphis graminum Rondani (Starks and Weibel, 1981)], glossy wheat (Triticum aestivum L. emend. Thell.) to grain aphid [Sitobion avenae F. (Lowe et al., 1985)], and glossy Brassica oleracea L. to cabbage worm (Artogeia rapae L.) and cabbage aphid (Brevicoryne brassicae L.) (Stoner, 1990). An explanation for these observations is that insects respond to cues from the chemistry of the waxes, and the broad diversity of compounds found in epicuticular waxes (long-chain fatty acids, esters, ketones, alkanes, and alcohols) may help insects to identify their host plants (Blaney and Chapman, 1970; Eigenbrode and Espelie, 1995; Städler, 1986; Thibout et al., 1982).

The biosynthetic pathway for epicuticular waxes has been studied in several plants (Kunst and Samuels, 2003). De novo fatty acid synthesis occurs in the plastid (Ohlrogge and Browse, 1995) and a substantial portion of fatty acids in epidermal cells is allocated toward wax production (Liu and Post-Beittenmiller, 1995). C18 fatty acids destined for wax synthesis are catalyzed into long chain molecules by fatty acid elongases associated with the endoplasmic reticulum (Fehling and Mukherjee, 1991). In leek (Allium ampeloprasum L.), peak activities of these enzymes were detected in basal regions of leaves immediately before the accumulation of epicuticular waxes (Rhee et al., 1998). Two main pathways have been suggested for production of epicuticular waxes (Millar et al., 1999). Long-chain fatty acids are converted into primary alcohols and esters through the acyl reduction pathway or into secondary alcohols and ketones through the decarbonylation pathway (Millar et al., 1999). Aldehydes are an intermediate step in both pathways and alkanes are either a product or intermediate step in the decarbonylation pathway. Enzymes that catalyze many of the later steps of these pathways are not well understood (Kunst and Samuels, 2003).

Natural variation exists in onion for amounts and types of epicuticular waxes (Damon et al., 2014; Molenaar, 1984). The most prevalent epicuticular waxes on the foliage of wild-type (waxy) onion are ketones (hentriacontanone-16), alkanes (2-methyl octacosane, 1-ethenyloxy octadecane, and heptacosane), and fatty alcohols (heptadecanol-1, hexacosanol-1, octacosanol-1, and triacontanol-1) (Damon et al., 2014). Amounts of these waxes are significantly less on leaves of the glossy (lowest amounts of waxes) and semiglossy (intermediate amounts of waxes) phenotypes relative to waxy plants (Damon et al., 2014). Although glossy onions show significant resistance to onion thrips (Alimousavi et al., 2007; Damon et al., 2014; Jones et al., 1934; Molenaar, 1984), this phenotype is not commercially useful because of susceptibility to leaf pathogens, excessive transpiration, and spray injury (Baker, 1982; Mohan and Molenaar 2005; Wirth et al., 1991). Semiglossy onions possess epicuticular wax amounts intermediate between the waxy and glossy phenotypes (Damon et al., 2014) as well as show significant resistance to onion thrips (Damon et al., 2014; Diaz-Montano et al., 2010, 2012). Therefore, this phenotype may be useful for integrated management of this important pest.

Genetic analyses of amounts and types of epicuticular waxes may reveal genes controlling the different branches of the wax–biosynthetic pathway and lead to a better understanding of epicuticular wax accumulation in plants. Quantitative trait loci (QTL) controlling amounts of epicuticular waxes have been mapped in several crops, including canola [Brassica napus L. (Pu et al., 2013)], rice [Oryza sativa L. (Srinivasan et al., 2008)], maize [Zea mays L. (Liu et al., 2012)], and sorghum (Burow et al., 2009). Mapping of QTL controlling amounts and types of waxes in onion would provide markers associated with onion thrips resistance and contribute to our understanding of the inheritance of different foliar wax phenotypes. In this study, we developed a waxy by semiglossy mapping population to identify QTL controlling amounts and types of epicuticular waxes in onion.

Materials and Methods

Genetic materials.

‘DehyA’ is an S-cytoplasmic male-sterile inbred line with white bulbs and wild-type waxy foliage (gift of Olam International, Hanford, CA). ‘B5351C’ is an inbred line of yellow onion selected from the open-pollinated population ‘Colorado #6’, restores male fertility in S cytoplasm, and has semiglossy foliage (Damon et al., 2014). A single bulb from the cross of ‘DehyA’ by B5351C was self-pollinated to produce the segregating family. F2 seeds were planted in Jan. 2012 in a greenhouse on the campus of the University of Wisconsin (UW)–Madison. Initial photoperiod was set at 12 h and was increased on the first of each subsequent month after the spring equinox to the natural daylength in Madison, WI. Temperatures were 25 °C days and 20 °C nights.

DNA isolation.

In Feb. 2012, 120 F2 plants were transplanted into 20-cm pots and individually labeled. In March, fresh tissue was harvested from each plant into polystyrene tubes and frozen in liquid nitrogen. Samples were stored at –80 °C for ≈48 h before lyophilization and then subsequently stored at –20 °C. Samples were individually ground using a mortar and pestle with ≈20 mL of liquid nitrogen and 0.1 g of silicon dioxide (Sigma-Aldrich, St. Louis, MO). Ground samples were placed in 50-mL polypropylene centrifuge tubes and DNA was isolated using the DNeasy Plant Maxi DNA isolation kit (Qiagen, Hilden, Germany). DNA concentrations were spectrophotometrically determined and 1 μg of DNA from each plant was electrophorized through a 0.8% agarose gel to assess intactness and quantity.

Gas chromatography–mass spectrometry analysis and data collection.

In Apr. 2012 at approximately the eight-leaf stage, three samples were collected from a single leaf from each of the 120 F2 plants for a total of 360 samples. In May 2012 (approximately the 14-leaf stage), the same procedure was followed to yield another 360 samples. In June 2012 (≈20-leaf stage), two samples were collected from an older leaf and two from a younger leaf from each of the 120 plants for a total of 480 samples. Visual phenotyping of F2 plants as either waxy or semiglossy was completed in the greenhouse in 2012. Plants were allowed to mature during July and August in the greenhouse, after which bulbs were dipped in a solution of azoxystrobin fungicide (Quadris; Syngenta, Greensboro, NC) and allowed to air-dry on the greenhouse bench for 24 h. Bulbs were stored at 5 °C until Apr. 2013 and then planted in the field at the UW Horticulture Research Station near Arlington, WI. As a result of storage conditions or unfavorable weather, numerous F2 bulbs rotted in the field. In June 2013, two samples were collected from different leaves from each of 86 plants for a total of 172 samples. In July 2013, two samples were collected from different leaves from each of 74 plants for a total of 148 samples. In total, five samplings were taken from the same F2 plants across two environments to yield a total of 1520 samples. All leaf samples were prepared for gas chromatography–mass spectrometry (GC-MS) as described by Damon et al. (2014) to identify and quantify epicuticular waxes. Identification was made by GC-MS solution post-run software (Shimadzu, Kyoto, Japan) comparing mass spectra with a library to reveal the closest match. Quantities of specific waxes were estimated by the area under peaks as previously described by Damon et al. (2014). All peak areas were normalized to an internal standard (docosane) by dividing by the docosane peak area.

Statistical analysis and QTL mapping.

A mixed model analysis of variance (ANOVA) with the repeated measures option was performed in SAS (Version 9.3; SAS Institute, Cary, NC) to generate restricted maximum likelihood estimates of variance components for the calculation of phenotypic and genotypic correlations among waxes (Holland, 2006). Variance components were also used to calculate broad-sense heritability on an entry mean basis for each epicuticular wax. Phenotypic variances for heritability and correlations were calculated by dividing genotype by environment variance by number of environments and dividing error variance by number of environments multiplied by number of samplings. For QTL mapping, mixed-model ANOVAs with repeated measures were performed in SAS to compute peak area least square means for each F2 plant. Individual plants were experimental units and the three samplings in 2012 and two samplings in 2013 were considered as repeated measures. GC-MS data for each wax was considered as unique dependent variables. Genotypes and samplings were considered as fixed and environments and subsamples as random variables. Departure from normality of model residuals was corrected by square root transformation.

DNAs from 119 F2 plants were used to genotype of 324 SNPs using the KASPar platform (LGC Genomics, Beverly, MA) as described by Duangjit et al. (2013). Linkage maps were created using the Joinmap software (Van Ooijen and Voorrips, 2001) with the Kosambi function. For phase-unknown SNPs, both phases were analyzed and the phase that placed the marker on the map with flanking markers was retained. Cosegregating markers were removed and data from 255 SNPs were combined with least squared means of GC-MS data from 119 plants for composite interval mapping using the R software (Broman et al., 2003). Two covariates and a window size of 10 cM were used in the composite interval model for each of eight waxes. The first covariate was a marker just outside the assigned window next to the QTL; the second covariate was a genomic position with a non-significant effect [i.e., not above the 0.05 logarithm-of-odds (LOD) threshold]. One thousand permutations of each analysis were run to determine the LOD threshold at the 0.05 significance level (Churchill and Doerge, 1994). For each QTL detected, percent of variance explained, maximum LOD score, 1.5 LOD confidence intervals, and additive and dominance effects were calculated.

Results and Discussion

Genetic map.

Genetic mapping of 324 SNPs using DNAs from 119 F2 progenies of ‘DehyA’ × B5351C yielded eight linkage groups. Linkage groups were aligned with genetic maps from two other onion mapping populations (OH1 × 5225 and BYG15-23 × AC43) described by Duangjit et al. (2013), allowing for the assignment of groups to the eight chromosomes of onion (Supplemental Table 1). Of the 324 segregating SNPs, 308 fit the expected 1:2:1 segregation ratio and 16 showed significant segregation distortion (P < 0.05); this level of segregation distortion is at the level expected by random chance. SNPs with segregation distortion at P < 0.0001 were discarded. SNPs with distorted segregation ratios between 0.05 and 0.0001 were concentrated in four areas on chromosomes 1, 4, 5, and 8 (Supplemental Table 1). Of the 324 SNPs, 214 also segregated in the OH1 × 5225 population and 142 were in common with the BYG15-23 × AC43 map (Duangjit et al., 2013). Here we report positions of 66 previously unmapped SNPs (Supplemental Table 1), adding to the 713 previously reported by Duangjit et al. (2013). As a result of missing parental genotypes or heterozygosity in the parental lines, 23 SNPs remained as phase-unknown and were added to the linkage map in both phases, keeping the phase for which the marker was assigned to a linkage group. Total length for the genetic map from ‘DehyA’ × B5351C was 7.8 Morgans.

Mapping of epicuticular wax differences.

Damon et al. (2014) detected significantly different amounts of eight waxes (hentriacontanone-16, heptacosane, heptadecanol-1, hexacosanol-1, octacosanol-1, 1-ethenyloxy octadecane, 2-methyl octacosane, and triacontanol-1) on the foliage of semiglossy and waxy onions. These same eight waxes were identified on the foliage of F2 progenies from ‘DehyA’ by B5351C (Table 1). Because onion is biennial, we sampled the same F2 plants during the bulb (2012) and seed (2013) production years. Although the mapping population performed poorly in the 2013 field environment resulting in a reduced data set, coefficients of variation for the two most prevalent waxes were relatively low (less than 20%) and comparable between the 2 years (13.2 in 2012 and 15.4 in 2013 for hentriacontanone-16 and 19.8 in 2012 and 16.4 in 2013 for octacosanol-1). Significant phenotypic and genotypic correlations were detected among waxes (Table 2). Hexacosanol-1, octacosanol-1, and triacontanol-1 were positively correlated with each other and negatively correlated with heptacosane. Hentriacontanone-16 was positively correlated with heptadecanol-1 and both of these had little or no correlation with octacosanol-1 or triacontanol-1. Broad-sense heritabilities on an entry-mean basis ranged from 0.26 to 0.86 for the eight waxes (Table 1). The high heritabilities for amounts of most waxes likely contributed to our ability to measure significant differences in the 2013 field environment despite reduced progeny numbers.

Table 1.

Broad-sense heritabilities (H2), positions in centiMorgans (cM) on chromosomes (Chr) of quantitative trait loci (QTL), percent of variance explained, additive and dominance effects, maximum logarithm-of-odds (LOD) scores, 0.05 LOD significance thresholds, and single nucleotide polymorphisms (SNPs) flanking 1.5-LOD confidence intervals of QTL affecting amounts of epicuticular waxes in onion and visual phenotypes in the F2 family from ‘DehyA’ crossed with B5351C.

Table 1.
Table 2.

Significance of genotypic (above diagonal) and phenotypic (below diagonal) correlations among eight epicuticular waxes on onion foliage from the segregating family from ‘DehyA’ crossed with B5351C.

Table 2.

QTL were detected controlling amounts of the eight epicuticular waxes, explaining from 17% to 65% of the phenotypic variation (Table 1). Maximum LOD scores ranged from 4.7 to 31.3 and 1.5 LOD confidence intervals ranged from 3 to 10 centiMorgans (cM). Hentriacontanone-16 and triacontanol-1 showed the highest heritability, percent of variance explained, and LOD scores (Table 1). Amounts of hentriacontanone-16 and 1-ethenyloxy octadecane mapped to a region on chromosome 5 (22 to 28 cM) with higher amounts from the waxy ‘DehyA’ parent (Fig. 1; Table 1). Amounts of three fatty alcohols (triacontanol-1, octacosanol-1, hexacosanol-1) and 2-methyl octacosane were controlled by a region on chromosome 2 from 68 to 75 cM (Fig. 1; Table 1) with variation from the semiglossy parent reducing amounts of fatty alcohols and increasing the amount of 2-methyl octacosane. No interactions were detected between the regions chromosomes 2 and 5, indicating that QTL independently affect concentrations of hentriacontanone-16 and 1-ethenyloxy octadecane (chromosome 5) vs. triacontanol-1, octacosanol-1, hexacosanol-1, and 2-methyl octacosane (chromosome 2). Amounts of heptacosane and heptadecanol-1 mapped to both regions on chromosomes 2 and 5 (Table 1). Higher amounts of heptacosane at both regions were conditioned by the semiglossy parent, whereas higher amounts of heptadecanol-1 at both regions were from the waxy parent. There was no evidence of epistasis or interaction between QTL on chromosomes 2 and 5 controlling amounts of heptadecanol-1 and heptacosane. Because three genotypic classes were present in the F2 population, additive and dominance effects were estimated (Table 1). QTL affecting amounts of hentriacontanone-16, heptadecanol-1, and 1-ethenyloxy octadecane on chromosome 5 were mainly additive. QTL for amounts of hexacosanol-1, octacosanol-1, triacontanol-1, and heptacosane on chromosome 2 showed dominance toward the lower values of the semiglossy (B5351C) parent (Table 1).

Fig. 1.
Fig. 1.

Quantitative trait loci (QTL) on chromosomes 2 and 5 affecting amounts of epicuticular waxes on onion foliage as determined by gas chromatography–mass spectrometry and visual phenotyping. Logarithm-of-odds (LOD) thresholds at P < 0.05 ranged from 3.98 to 4.15 for the various waxes and the horizontal line indicates the most stringent threshold at 4.15.

Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 139, 5; 10.21273/JASHS.139.5.597

Visual scores of F2 plants as waxy vs. semiglossy revealed the same position as QTL on chromosome 5, but not on chromosome 2 (Table 1). Mapping of the visual phenotype to the same region on chromosome 5 as amounts of hentriacontanone-16 supports the conclusion of Damon et al. (2014) that crystals of hentriacontanone-16 are primarily responsible for the visible waxy phenotype. The additive effect of the QTL on chromosome 5 controlling amounts of hentriacontanone-16 (Table 1), and therefore the visual phenotype, is consistent with greater waxiness of progenies between waxy and semiglossy parents.

Studies of the biosynthetic pathway for epicuticular waxes in plants have revealed that ketones (such as hentriacontanone-16) are derived from the decarbonylation pathway with aldehydes and alkanes as intermediates (Kunst and Samuels, 2003). The MAH1 gene in Arabidopsis thaliana (L.) Heynh. is a midchain alkane hydroxylase involved in the production of secondary alcohols and ketones in the decarbonylation pathway; mah1 mutants are deficient in these waxes and have increased amounts of alkanes (Greer et al., 2007). The QTL on onion chromosome 5 may be associated with the decarbonylation pathway with the semiglossy region reducing production of hentriacontanone-16. The mainly additive effects of the QTL on chromosome 5 support partial blockage of the decarbonylation pathway in heterozygous plants, possibly as a result of production of both functional and non-functional gene products.

Primary alcohols such as hexacosanol-1, octacosanol-1, and triacontanol-1 are derived from the acyl-reduction pathway (Kunst and Samuels, 2003). The CER4 gene in A. thaliana has been characterized as a fatty acyl-coenzyme A reductase involved in the production of primary fatty alcohols and cer4 mutants were deficient in primary alcohols with slightly elevated levels of other waxes (Rowland et al., 2006). Because the QTL on chromosome 2 from the semiglossy parent are associated with reduced amounts of primary alcohols, this region may encode an enzyme involved in the acyl-reduction pathway. However, the large dominance effect of the QTL on chromosome 2 from the semiglossy parent suggests that this region may encode a regulatory factor affecting genes located elsewhere in the genome (Pu et al., 2013).

Unlike observations in A. thaliana, alkane accumulation in onion appears to correspond more closely with obstruction of the acyl-reduction pathway than the decarbonylation pathway. Jenks et al. (1995) showed that a reduction in primary alcohols resulted in an increase in aldehydes and hypothesized the blockage of an aldehyde reductase in the acyl-reduction pathway. However, Vioque and Kolattukudy (1997) suggested that fatty acids are converted immediately to primary alcohols without release of an aldehyde intermediate. Kunst and Samuels (2003) suggested that the increase in aldehydes resulted from an increased flux of fatty acyl precursors into the decarbonylation pathway. In the semiglossy B5351C parent, such a flux of acyl precursors toward decarbonylation may be associated with the additional blockage of the decarbonylation pathway, resulting in accumulation of heptacosane. The odd chain length of heptacosane should result from decarbonylation (Greer et al., 2007); however, the lack of a significant interaction between regions on chromosomes 2 and 5 suggests that the increase in heptacosane is independent of the step(s) in the decarbonylation pathway affected by the QTL on chromosome 5.

The association of heptadecanol-1 with the chromosome 5 region is not easily explained, because primary alcohols are expected to result from the acyl-reduction pathway. However, this alcohol differs from those associated with the QTL on chromosome 2 by being shorter and having an odd (C17) chain length; odd chain length is typical of secondary alcohols (Kunst and Samuels, 2003). These observations suggest that mechanisms other than the acyl-reduction pathway may be involved in heptadecanol-1 production and, as a result of the highly significant association with chromosome 5, the decarbonylation pathway is implicated. Interestingly, amounts of another unusually short wax, 1-ethenyloxy octadecane (C20), was also significantly associated with chromosome 5.

This study provides strong evidence that two chromosome regions in onion independently and significantly affect the two branches of the biosynthetic pathway for epicuticular waxes. The lack of interaction between these two regions suggests that the QTL do not control early steps in the biosynthesis of epicuticular waxes such as fatty acid elongation and are not involved in any broad regulatory control of the pathway. Therefore, characterization of genes associated with these QTL would likely contribute to a greater understanding of subsequent steps in the pathway, where precursor fatty acids are used to produce diverse groups of epicuticular waxes.

Onion cultivars with semiglossy foliage are commonly grown in many arid and semiarid regions of the world; waxy cultivars predominant in more temperate areas. Backcross conversion of waxy onions to the semiglossy phenotype is complicated because crosses of semiglossy and waxy plants produce visually waxy progenies. Because onion is biennial, introgression of QTL affecting amounts and types of epicuticular waxes will be expedited by molecular tags to avoid selfing plants to reveal less waxy genotypes. Three SNPs on chromosome 5 (i26178_547, i29989_644, and i32901_1333) and two on chromosome 2 (i28284_1005 and i33533_568) are located within the 1.5 LOD confidence interval of the highly significant QTL controlling amounts of epicuticular waxes in onion (Table 1). If amounts of hentriacontanone-16 were a major factor affecting parasitism by onion thrips (Damon et al., 2014), one could select for reduced amounts of hentriacontanone-16 (QTL on chromosome 5) and increased amounts of fatty alcohols (QTL on chromosome 2) to potentially produce plants with adequate amounts of epicuticular waxes to perform well in the field and show resistance to onion thrips, which would be useful toward integrated management of this important insect pest.

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  • Pu, Y., Gao, J., Guo, Y., Liu, T., Zhu, L., Xu, P., Yi, B., Wen, J., Tu, J., Ma, C., Fu, T., Zou, J. & Shen, J. 2013 A novel dominant glossy mutation causes suppression of wax biosynthesis pathway and deficiency of cuticular wax in Brassica napus BMC Plant Biol. 13 215

    • Search Google Scholar
    • Export Citation
  • Rhee, Y., Hlousek-Radojcic, A., Ponsamuel, J., Liu, D. & Post-Beittenmiller, D. 1998 Epicuticular wax accumulation and fatty acid elongation activities are induced during leaf development of leeks Plant Physiol. 116 901 911

    • Search Google Scholar
    • Export Citation
  • Rowland, O., Zheng, H., Hepworth, S.R., Lam, P., Jetter, R. & Kunst, L. 2006 CER4 encodes an alcohol-forming fatty acyl-coenzyme A reductase involved in cuticular wax production in Arabidopsis Plant Physiol. 142 866 877

    • Search Google Scholar
    • Export Citation
  • Shelton, A.M., Zhao, J.Z., Nault, B.A., Plate, J., Musser, F.R. & Larentzaki, E. 2006 Patterns of insecticide resistance in onion thrips (Thysanoptera: Thripidae) in onion fields in New York J. Econ. Entomol. 99 1798 1804

    • Search Google Scholar
    • Export Citation
  • Shock, C., Feibert, E., Jensen, L., Krishna Mohan, S. & Saunders, L. 2008 Onion variety response to Iris yellow spot virus HortTechnology 18 1063 1068

  • Srinivasan, S., Gomez, S.M., Kumar, S.S., Ganesh, S.K., Biji, K.R., Senthil, A. & Babu, R.C. 2008 QTLs linked to leaf epicuticular wax, physio-morphological and plant production traits under drought stress in rice (Oryza sativa L.) Plant Growth Regulat. 56 245 256

    • Search Google Scholar
    • Export Citation
  • Städler, E. 1986 Oviposition and feeding stimuli in leaf surface waxes, p. 105–121. In: Juniper, B.E. and T.R.E. Southwood (eds.). Insects and the plant surface. Edward Arnold, London, UK

  • Starks, K.J. & Weibel, D.E. 1981 Resistance in bloomless and sparse-bloom sorghum to greenbugs Environ. Entomol. 10 963 965

  • Stoner, K.A. 1990 Glossy leaf wax and plant resistance to insects in Brassica oleracea under natural infestation Environ. Entomol. 19 730 739

  • Thibout, E., Auger, J. & Lecomte, C. 1982 Host plant chemicals responsible for attraction and oviposition in Acrolepiopsis assectella. Proc. 5th Intl. Symp. Insect–Plant Relationships. p. 107–115

  • Van Ooijen, J.W. & Voorrips, R.E. 2001 JoinMap® 3.0, software for the calculation of genetic linkage maps. Plant Res. Intl., Wageningen, The Netherlands

  • Vioque, J. & Kolattukudy, P.E. 1997 Resolution and purification of an aldehyde-generating and an alcohol-generating fatty acyl-CoA reductase from pea leaves (Pisum sativum L.) Arch. Biochem. Biophys. 340 64 72

    • Search Google Scholar
    • Export Citation
  • Wirth, W., Storp, S. & Jacobsen, W. 1991 Mechanisms controlling leaf retention of agricultural spray solutions Pestic. Sci. 33 411 420

Supplemental Table 1.

Positions, observed segregations, and probabilties for fits to expected 1:2:1 ratio for single nucleotide polymorphisms (SNPs) segregating in the F2 family from DehyA by B5351C.

Supplemental Table 1.

If the inline PDF is not rendering correctly, you can download the PDF file here.

Contributor Notes

We gratefully acknowledge grant number 2008-51180-04875 from the USDA Specialty Crops Research Initiative, a Monsanto Graduate Fellowship to S.J.D., and the gift of ‘DehyA’ by Olam International, Hanford, CA.

Names are necessary to report factually on available data; however, the U.S. Department of Agriculture (USDA) neither guarantees nor warrants the standard of the product, and the use of the name by USDA implies no approval of the product to the exclusion of others that may also be suitable.

Corresponding author. E-mail: mjhavey@wisc.edu.

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    Quantitative trait loci (QTL) on chromosomes 2 and 5 affecting amounts of epicuticular waxes on onion foliage as determined by gas chromatography–mass spectrometry and visual phenotyping. Logarithm-of-odds (LOD) thresholds at P < 0.05 ranged from 3.98 to 4.15 for the various waxes and the horizontal line indicates the most stringent threshold at 4.15.

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  • Ohlrogge, J. & Browse, J. 1995 Lipid biosynthesis Plant Cell 7 957 970

  • Pu, Y., Gao, J., Guo, Y., Liu, T., Zhu, L., Xu, P., Yi, B., Wen, J., Tu, J., Ma, C., Fu, T., Zou, J. & Shen, J. 2013 A novel dominant glossy mutation causes suppression of wax biosynthesis pathway and deficiency of cuticular wax in Brassica napus BMC Plant Biol. 13 215

    • Search Google Scholar
    • Export Citation
  • Rhee, Y., Hlousek-Radojcic, A., Ponsamuel, J., Liu, D. & Post-Beittenmiller, D. 1998 Epicuticular wax accumulation and fatty acid elongation activities are induced during leaf development of leeks Plant Physiol. 116 901 911

    • Search Google Scholar
    • Export Citation
  • Rowland, O., Zheng, H., Hepworth, S.R., Lam, P., Jetter, R. & Kunst, L. 2006 CER4 encodes an alcohol-forming fatty acyl-coenzyme A reductase involved in cuticular wax production in Arabidopsis Plant Physiol. 142 866 877

    • Search Google Scholar
    • Export Citation
  • Shelton, A.M., Zhao, J.Z., Nault, B.A., Plate, J., Musser, F.R. & Larentzaki, E. 2006 Patterns of insecticide resistance in onion thrips (Thysanoptera: Thripidae) in onion fields in New York J. Econ. Entomol. 99 1798 1804

    • Search Google Scholar
    • Export Citation
  • Shock, C., Feibert, E., Jensen, L., Krishna Mohan, S. & Saunders, L. 2008 Onion variety response to Iris yellow spot virus HortTechnology 18 1063 1068

  • Srinivasan, S., Gomez, S.M., Kumar, S.S., Ganesh, S.K., Biji, K.R., Senthil, A. & Babu, R.C. 2008 QTLs linked to leaf epicuticular wax, physio-morphological and plant production traits under drought stress in rice (Oryza sativa L.) Plant Growth Regulat. 56 245 256

    • Search Google Scholar
    • Export Citation
  • Städler, E. 1986 Oviposition and feeding stimuli in leaf surface waxes, p. 105–121. In: Juniper, B.E. and T.R.E. Southwood (eds.). Insects and the plant surface. Edward Arnold, London, UK

  • Starks, K.J. & Weibel, D.E. 1981 Resistance in bloomless and sparse-bloom sorghum to greenbugs Environ. Entomol. 10 963 965

  • Stoner, K.A. 1990 Glossy leaf wax and plant resistance to insects in Brassica oleracea under natural infestation Environ. Entomol. 19 730 739

  • Thibout, E., Auger, J. & Lecomte, C. 1982 Host plant chemicals responsible for attraction and oviposition in Acrolepiopsis assectella. Proc. 5th Intl. Symp. Insect–Plant Relationships. p. 107–115

  • Van Ooijen, J.W. & Voorrips, R.E. 2001 JoinMap® 3.0, software for the calculation of genetic linkage maps. Plant Res. Intl., Wageningen, The Netherlands

  • Vioque, J. & Kolattukudy, P.E. 1997 Resolution and purification of an aldehyde-generating and an alcohol-generating fatty acyl-CoA reductase from pea leaves (Pisum sativum L.) Arch. Biochem. Biophys. 340 64 72

    • Search Google Scholar
    • Export Citation
  • Wirth, W., Storp, S. & Jacobsen, W. 1991 Mechanisms controlling leaf retention of agricultural spray solutions Pestic. Sci. 33 411 420

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