Abstract
Vegetable crops can be significant sources of nutritionally important dietary carotenoids, and Brassica are sources that also exhibit antioxidant and anticarcinogenic activity. The family Brassicaceae contains a diverse group of plant species commercially important in many parts of the world. The six economically important Brassica species are closely related genetically. Three diploid species (B. nigra, B. rapa, B. oleracea) are the natural progenitors of the amphidiploid species (B. juncea, B. napus, B. carinata). The objective of this study was to characterize the accumulation of important dietary carotenoid pigments among the genetically related Brassica species. High-performance liquid chromatographic quantification revealed significant differences in carotenoid and chlorophyll pigment concentrations among the Brassica species. Brassica rapa accumulated the highest concentrations of antheraxanthin [0.79 mg/100 g fresh weight (FW)], lutein (8.89 mg/100 g FW), and zeaxanthin (0.75 mg/100 g FW). The highest concentrations of β-carotene (4.41 mg/100 g FW) and total chlorophyll (125.9 mg/100 g FW) were found in B. juncea. Brassica nigra accumulated the highest concentrations of 5,6-epoxylutein (0.41 mg/100 g FW) and violaxanthin (2.28 mg/100 g FW), whereas B. oleracea accumulated the highest concentrations of neoxanthin (2.10 mg/100 g FW). For many of the pigments analyzed, the amphidiploids B. carinata and B. napus accumulated significantly less carotenoid concentrations than the diploid species and B. juneca. Brassica convey unique health attributes when consumed in the diet. Identification of genetic relationships among the Brassica species would be beneficial information for improvement programs designed to increase carotenoid values.
The family Brassicaceae (Cruciferae) represents a diverse group of plant species commercially important in many parts of the world. The plants produce condiment mustard; leafy, stored, processed, and picked vegetables; seed oils for margarine, salad oils, cooking oils, and industrial uses; animal fodders; and green manure crops (Williams and Hill, 1986). The six economically important Brassica species in world production are genetically related. Three diploid species—B. nigra, B. rapa, B. oleracea—are the natural progenitors of the amphidiploids species: B. juncea, B. napus, and B. carinata. The “cole crops,” such as broccoli, brussels sprouts, cabbage, cauliflower, curly kale, and kohlrabi, make up B. oleracea. Oil seed types, as well as Chinese cabbage, turnip, and pak-choi are found within B. rapa. Brassica nigra, popularly known as black mustard, is used for its seed oil content. The cross between B. nigra and B. oleracea produces B. carinata. It is a tall, leafy plant mainly restricted to Ethiopia. The cross between B. oleracea and B. rapa produces B. napus. Rutabagas are a variety of B. napus, as well as the economically important oilseed rape. The cross between B. rapa and B. nigra produces B. juncea, the group known as mustard. The mustards are consumed in the southern United States as mustard greens, or are used as a condiment or spice (Williams and Hill, 1986).
The three common diploid species in the genus Brassica have haploid numbers of 8 (B. nigra), 9 (B. oleracea), and 10 (B. rapa) (Nwankiti, 1970). It is believed that these numbers represent a phylogenetically ascending series (Manton, 1932). The higher chromosome number species in the genus are believed to be amphidiploids and have haploid numbers of 17 (B. carinata), 18 (B. juncea), and 19 (B. napus). The genetic arrangement of the diploid and amphidiploid Brassica species is credited to the U (1935). The diagram is referred to as the triangle of U (Nwankiti, 1970) (Fig. 1). Study of interspecific hybridization and subsequent pairing within the genus led to the designation of two distinct groups: 1) monogenomic diploids of B. nigra (2n = 16; BB), B. oleracea (2n = 18; CC), and B. rapa (2n = 20; AA); and 2) amphidiploids of B. juncea (2n = 36; AABB), B. carinata (2n = 34; BBCC), and B. napus (2n = 38; AACC)(Prakash and Hinata, 1980).
Cytogenetic relationships between the diploid and amphidiploid Brassica species. Adapted from U (1935).
Citation: HortScience horts 42, 3; 10.21273/HORTSCI.42.3.461
Carotenoids are C40 isoprenoid polyene plant secondary compounds that form lipid-soluble yellow, orange, and red pigments (Zaripheh and Erdman, Jr., 2002). The carotenoids can be divided into two groups: 1) hydrocarbon carotenes (C40H56) and 2) their oxygenated derivative, referred to as xanthophylls. Carotenoids span the thylakoid membranes of chlorophyll (Chl) complexes and function in accessory roles for light harvesting, photoprotection, and structural stabilization (Demmig-Adams et al., 1996; Tracewell et al., 2001). Carotenoid pigments protect photosynthetic structures by quenching excited triplet Chl (3Chl) to dissipate excess energy (Frank and Cogdell, 1996) and by binding singlet oxygen (1O2) to inhibit potential oxidative damage (Demmig-Adams et al., 1996; Tracewell et al., 2001). Carotenoids are produced in the plastids and are derived from isopentenyl diphosphate (IPP). In the first step in biosynthesis, IPP is isomerized to dimethylallyl diphosphate, which becomes the substrate for the C20 geranylgeranyl diphosphate (GGPP) (Bramley, 2002). The first step unique to carotenoid biosynthesis is the condensation of two molecules of GGPP to form the first C40 carotenoid, phytoene, via phytoene synthase (Gross, 1991). The carotenoid pathway branches at the cyclization reactions of lycopene to produce carotenoids with either two β-rings (e.g., β-carotene, zeaxanthin, antheraxanthin, violaxanthin, and neoxanthin) or carotenoids with one β-ring and one ε-ring (e.g., α-carotene and lutein)(Cunningham, 2002) (Fig. 2).
A simplified version of the carotenoid biosynthetic pathway in plants. DMAPP, dimethylallyl pyrophosphate; GGPP, geranylgeranyl pyrophosphate; IPP, isopentenyl pyrophosphate. Ethyl beta-apo-8′-carotenoate used as internal standard.
Citation: HortScience horts 42, 3; 10.21273/HORTSCI.42.3.461
There has been increased interest in the nutritional and medicinal importance of dietary carotenoids (Faulks and Southon, 2005; Frazer and Bramley, 2004; Yeum and Russell, 2002). Dietary intake of carotenoids has been associated with reduced risk of lung cancer and chronic eye diseases, including cataract and age-related macular degeneration (Johnson et al., 2000; Le Marchand et al., 1993). Studies have indicated that consumption of a variety of vegetables providing a mixture of carotenoids was more strongly associated with disease reduction than individual carotenoid supplements (Johnson et al., 2000; Le Marchand et al., 1993). Plants can be significant sources of carotenoids in the diet, and Brassica vegetables are relatively abundant sources that exhibit antioxidant and anticarcinogenic activity (Kurilich et al., 1999). Therefore, the objective of this study was to characterize the variation in accumulation of important dietary carotenoids among the six genetically related Brassica species.
Materials and Methods
Seeds of rapid-cycling populations of B. nigra, B. rapa, B. oleracea, B. juncea, B. napus, and B. carinata (Crucifer Genetics Cooperative, Department of Plant Pathology, The University of Wisconsin, Madison, Wisc.) were sown into 2.5 × 2.5-cm growing cubes (Grodan A/S, Dk-2640, Hedehusene, Denmark) and germinated in a growth chamber room (Environmental Growth Chambers, Chagrin Falls, Ohio) at 23 ºC/16 ºC day/night under a 16-h photoperiod on 2 Dec. 2004 and again on 11 Jan. 2005. Photosynthetically active radiation at the canopy height was 1000 μmol·m−2·s−1 (LI-188B; LI-COR, Inc., Lincoln, Neb.). The first true leaves appeared 4 to 7 d after planting (DAP). Nutrients were applied as needed with 200 mg·L−1 Peter's 20N–6.9P–16.6K water-soluble fertilizer (The Scotts Company, Marysville, Ohio). At 14 DAP, the plantlets were transferred to 11-L containers (Rubbermaid Inc., Wooster, Ohio) filled with 9 L of a nutrient solution (Hoagland and Arnon, 1950). Elemental concentrations of the nutrient solutions were N (105 mg·L−1), P (15.3 mg·L−1), K (117.3 mg·L−1), Ca (80.2 mg·L−1), Mg (24.6 mg·L−1), S (32.0 mg·L−1), Fe (0.5 mg·L−1), B (0.25 mg·L−1), Mo (0.005 mg·L−1), Cu (0.01 mg·L−1), Mn (0.25 mg·L−1), and Zn (0.025 mg·L−1). Solutions were aerated with an air blower (model 25E133W222; Spencer, Winsor, Conn.) connected to air stones. Water was added daily to maintain initial solution volumes in each container.
Each experimental replication consisted of three randomized blocks, with plots consisting of individual containers holding six plants of one Brassica species, each placed into 2-cm round holds set at 10.6 × 9.5-cm spacing on the lid. Plants were harvested just before anthesis 27 DAP (B. rapa, B. juncea. B. nigra) and 34 DAP (B. carinate, B. napus, B. oleracea). Leaf tissue was removed from six plants for each Brassica species and was immediately stored at –80 ºC before lyophilization. The frozen plant samples were lyophilized for a minimum of 72 h (model 6L FreeZone; LabConCo, Kansas City, Mo.).
Carotenoid/chlorophyll analysis.
Plant pigments were extracted from freeze-dried tissues according to Kopsell et al. (2004) and analyzed according to Emenhiser et al. (1996). A 0.10-g subsample was rehydrated with 0.8 mL ultra pure H2O at 40 ºC for 20 min. After incubation, 0.8 mL of the internal standard ethyl-β-8′-apo-carotenoate (Sigma Chemical Co., St. Louis) was added to determine extraction efficiency. A total of 2.5 mL tetrahydrofuran (THF) stabilized with 25 ppm 2,6-Di-tert-butyl-4-methoxyphenol was also added. The sample was homogenized in a Potter–Elvehjem (Kontes, Vineland, N.J.) tissue grinding tube using ≈25 insertions with a pestle attached to a drill press (Sears, Roebuck and Co., Hoffman Estates, Ill.) set at 540 rpm. During homogenation, the tube was immersed in ice to dissipate heat. The tube was then placed into a clinical centrifuge for 3 min at 500 g n. The supernatant was removed and the sample pellet was resuspended in 2 mL THF and homogenized again with the same extraction technique. The procedure was repeated for a total of four extractions to obtain a colorless supernatant. The combined supernatants were reduced to 1.5 mL under a stream of nitrogen gas (N-EVAP 111; Organomation, Berlin, Mass.) in a water bath set at 40 ºC, and was brought up to a final volume of 5 mL with methanol (MeOH). A 2-mL aliquot was filtered through a 0.2-μm polytetrafluoroethylene (PTFE) filter (model Econofilter PTFE 25/20; Agilent Technologies, Wilmington, Del.) using a 5-mL syringe (Becton, Dickinson and Company, Franklin Lakes, N.J.) before analysis using high-performance liquid chromatography (HPLC).
An Agilent 1100 series HPLC unit with a photodiode array detector (Agilent Technologies, Palo Alto, Calif.) was used for pigment separation. Chromatographic separations were achieved using an analytical scale (4.6 mm i.d. × 250 mm) of 5 μm and a 200-Å polymeric C30 reverse-phase column (ProntoSIL, MAC-MOD Analytical Inc., Chadds Ford, Pa.), which allowed for effective separation of chemically similar carotenoid compounds (Fig. 3). The column was equipped with a guard cartridge (4.0 mm i.d. × 10 mm) and holder (ProntoSIL), and was maintained at 30 ºC using a thermostat column compartment. All separations were achieved isocratically using a binary mobile phase of 11% methyl tert-butyl ethanol, 88.9% MeOH, and 0.1% triethylamine (v/v). The flow rate was 1.0 mL·min−1, with a run time of 53 min, followed by a 10-min equilibration before the next injection. Eluted compounds from a 20-μL injection loop were detected at 453 nm (carotenoids and internal standard), 652 nm (Chl a), and 665 nm (Chl b), and data were collected, recorded, and integrated using ChemStation Software (Agilent Technologies). Internal standard percent recovery ranged from 62% to 100%, with a mean for all samples at 80%. Peak assignment for individual pigments was performed by comparing retention times and line spectra obtained from photodiode array detection using external standards (antheraxanthin, neoxanthin, 5,6-epoxylutein, lutein, violaxanthin, zeaxanthin from Carotenature, Lupsingen, Switzerland; β-carotene, Chl a, Chl b from Sigma Chemical Co.). The concentration of the external pigment standards were determined spectrophotometrically using the following E 1cm 1% values: antheraxanthin, 2349 in ethanol (ETOH), λmax = 446 nm; neoxanthin, 2270 in ETOH, λmax = 438 nm; 5,6-epoxylutein, 2463 in ETOH, λmax = 440 nm; lutein, 2468 in ETOH, λmax = 440 nm; violaxanthin, 2499 in ETOH, λmax = 441 nm; zeaxanthin, 2540 in ETOH, λmax = 452 nm; β-carotene, 2592 in hexane, λmax = 452 nm; Chl a, 819 in ETOH, λmax = 665 nm; and Chl b, 441 in ETOH, λmax = 649 nm (Davies and Köst, 1988). Spinach standard reference material (Slurried Spinach 2385, National Institute of Science and Technology, Gaithersburg, Md.) was used for method validation.
High-performance liquid chromatogram of Brassica juncea at 453 nm using a C30 column. Retention times for the pigments were violaxanthin, 5.520 min; neoxanthin, 5.813 min; antheraxanthin, 7.586 min; 5,6-epoxylutein, 7.894 min; chlorophyll b, 8.511 min; lutein, 9.334 min; zeaxanthin, 11.309 min; chlorophyll a, 13.900 min; ethyl-β-8′-apo-carotenoate (internal standard), 19.326 min; and β-carotene, 51.463 min. High-performance liquid chromatographic conditions are described in the text.
Citation: HortScience horts 42, 3; 10.21273/HORTSCI.42.3.461
Data sets were analyzed by the GLM procedures of SAS (Cary, N.C.) with species means separated by Duncan's multiple range test (α = 0.05). Data were recorded on both a fresh weight (FW; in milligrams per 100 g) and dry weight (DW; in milligrams per gram) basis.
Results and Discussion
Carotenoid pigment accumulation in leaf tissues differed among the Brassica species (Table 1). Lutein accumulation ranged from 8.89 mg/100 g FW (B. rapa) to 5.65 mg/100 g FW (B. carinata). Leaf tissue lutein averaged 8.64 mg/100 g FW among the diploid Brassica species (B. rapa, B. nigra, B. oleracea) and 7.35 mg/100 g FW among the amphidiploid species (B. juncea, B. carinata, B. napus). Carotenoid 5,6-epoxides are widespread in nature. Their conversion is mediated by a mixed-function oxygenase, which requires nicotinamide adenine dinucleotide phosphate (NADPH) and O2 as cofactors (Goodwin, 1980). The accumulation of 5,6-epoxylutein ranged from 0.41 mg/100 g FW (B. nigra) to 0.25 mg/100 g FW (B. carinata). The accumulation of 5,6-epoxylutein averaged 0.40 mg/100 g FW for the diploid Brassica species and 0.33 mg/100 g FW for the amphidiploid species.
Mean values for carotenoid pigments (measured in milligrams per 100 g fresh weight) for the six genetically related Brassica species grown in a controlled environment.z
The accumulation of β-carotene was similar for B. rapa, B. juncea, B. nigra, and B. oleracea, and was significantly less for B. carinata and B. napus (Table 1). The accumulation of β-carotene averaged 3.93 mg/100 g FW for the diploid Brassica species and 3.14 mg/100 g FW for the amphidiploid species. Insertion of the oxygen function in the carotenoid biosynthetic pathway occurs after the cyclization of lycopene (Cunningham and Gantt, 1998). Zeaxanthin forms from the hydroxylation of β-carotene. Only one Brassica species (B. rapa) accumulated significantly higher zeaxanthin concentrations than all the other species. The accumulation of zeaxanthin averaged 0.46 mg/100 g FW for the diploid Brassica species and 0.28 mg/100 g FW for the amphidiploid species. Epoxidation of zeaxanthin, via antheraxanthin, results in the formation of violaxanthin. The rearrangement of the epoxide group is the proposed mechanism for the conversion of violaxanthin to neoxanthin (Gross, 1991). Antheraxanthin accumulation ranged from 0.79 mg/100 g FW (B. rapa) to 0.41 mg/100 g FW (B. napus). The accumulation of antheraxanthin averaged 0.62 mg/100 g FW for the diploid Brassica species and 0.55 mg/100 g FW for the amphidiploid species.
The built-in ability for photoprotective energy dissipation within the photosynthetic apparatus of higher plants occurs through reversible changes in the violaxanthin content in the leaves. The xanthophyll, or violaxanthin, cycle is involved in the deepoxidation of violaxanthin in high light conditions, and the reepoxidation of zeaxanthin in dark conditions via antheraxanthin (Demmig-Adams and Adams, 1996; Gross, 1991). The accumulation of violaxanthin in B. juncea and B. nigra was significantly higher than the other Brassica species. The accumulation of violaxanthin averaged 1.71 mg/100 g FW for the diploid Brassica species and 1.47 mg/100 g FW for the amphidiploid species. The accumulation of neoxanthin also differed among the Brassica species. The accumulation of neoxanthin averaged 2.00 mg/100 g FW for the diploid Brassica species and 1.68 mg/100 g FW for the amphidiploid species.
The Brassica species accumulated much higher concentrations of Chl pigments. Chlorophyll pigments differed among the Brassica species (Table 2). Chlorophyll a accumulation ranged from 100.1 mg/100 g FW (B. juncea) to 52.4 mg/100 g FW (B. carinata). Leaf tissue Chl a averaged 93.4 mg/100 g FW among the diploid Brassica species and 73.6 mg/100 g FW among the amphidiploid species. Chlorophyll b accumulation ranged from 28.8 mg/100 g FW (B. oleracea) to 18.3 mg/100 g FW (B. carinata). The accumulation of Chl b averaged 26.2 mg/100 g FW for the diploid Brassica species and 22.5 mg/100 g FW for the amphidiploid species. Total Chl concentrations ranged from 125.9 mg/100 g FW (B. juncea) to 71.7 mg/100 g FW (B. carinata). Total leaf tissue Chl averaged 119.5 mg/100 g FW among the diploid Brassica species and 96.1 mg/100 g FW among the amphidiploid species.
Mean values for chlorophyll pigments (measured in milligrams per 100 g fresh weight) for the six genetically related Brassica species grown in a controlled environment.z
Vegetable tissues can be dried and encapsulated for use as lutein and β-carotene dietary antioxidant supplements. Pool-Zobel et al. (1997) and Müller et al. (1999) conducted experiments to determine whether carotenoids could protect against DNA damage and oxidative stress. The researchers used dried spinach powder (obtained from Völpel GmbH, Königmoos, Germany) to assess antioxidant potential. The lutein concentration for the dried spinach was 11.3 mg/10 g, whereas the β-carotene concentration was 3.1 mg/10 g. The dried spinach increased serum lutein and β-carotene twofold over the 2-week feeding intervention, and significantly reduced the number of DNA strand breaks in human subjects. This research suggests nutritional impacts from consuming carotenoids from dried plant materials (Müller et al., 1999). Therefore, carotenoid accumulations for the Brassica species in the current study were expressed on a DW basis (Table 3). Lutein accumulation ranged from 0.87 mg·g−1 DW (B. rapa) to 0.52 mg·g−1 DW (B. carinata). The accumulation of 5,6-epoxylutein ranged from 0.04 mg·g−1 DW (B. nigra, B. juncea, B. rapa) to 0.02 mg·g−1 DW (B. carinata). The accumulation of β-carotene ranged from 0.43 mg·g−1 DW (B. juncea) to 0.22 mg·g−1 DW (B. carinata). Zeaxanthin accumulation ranged from 0.04 mg·g−1 DW (B. rapa) to 0.02 mg·g−1 DW (B. napus, B. carinata). Antheraxanthin accumulation ranged from 0.08 mg·g−1 DW (B. rapa) to 0.04 mg·g−1 DW (B. napus). Violaxanthin accumulation ranged from 0.22 mg·g−1 DW (B. nigra) to 0.10 mg·g−1 DW (B. napus, B. carinata). Neoxanthin accumulation ranged from 0.20 mg·g−1 DW (B. juncea, B. oleracea) to 0.12 mg·g−1 DW (B. carinata). For each of the carotenoid pigments analyzed on a DW basis, the diploid Brassica species accumulated higher concentrations, on average, than the amphidiploid species (Table 3). Dry weight carotenoid values measured in the Brassica species would make them comparable sources to previously used supplements (Müller et al., 1999; Pool-Zobel et al., 1997).
Mean values for carotenoid pigments (measured in milligrams per gram dry weight) for the six genetically related Brassica species grown in a controlled environment.z
Genomes are subject to change through mutation of the genetic systems that control meiosis and chromosome morphology. Interspecific hybridization between the diploid Brassica species is easily accomplished and can be used to make comparisons of meiotic behaviors among diploid, amphidiploid, and amphidiploid hybrids. Attia and Röbbelen (1986) studied cytogenetic relationships within cultivated species of Brassica and discovered a high degree of pairing and chiasmata formation for the amphihaploid B. napus. Brassica carinata had very low meiotic pairing and chiasmata compared with B. napus, with slightly higher pairing observed for B. juncea. Morphological (Prakash and Hinata, 1980) and chromosomal (Armstrong and Keller, 1982) differentiation identified B. napus to be the most distantly related of the genomic species. Cunha et al. (2004) used chloroplast DNA (cpDNA) to identify polymorphisms between the six genetically distinct Brassica species. The study concluded that cpDNA polymorphisms could discriminate among the three diploid Brassica species, but were unable to identify polymorphisms among the three amphidiploid species. Research shows that B. rapa and B. oleracea are very closely related species (Harberd and McArthur, 1980). In most cases for the current study, carotenoid pigment accumulation among these two Brassica species did not differ (Table 1).
Earlier studies revealed structural divergence among the monogenic Brassica species was based upon reductions in chromosome pairing in the haploids and production of hybrids (Prakash and Chopra, 1993). Based upon a phylogenetic tree, the amphidiploid species B. carinata is more closely related to its maternal diploid donor B. nigra than to the paternal donor B. oleracea, B. juncea is more closely related to the maternal donor B. rapa than to the paternal donor B. nigra, and B. napus is almost equally related to the maternal and paternal donors of B. rapa and B. oleracea respectively. The results suggest that the maternally donated cytoplasmic genomes have some effect on nuclear genome evolution in the amphidiploid species (Cunha et al., 2004; Song et al., 1996). Large genomic diversity was found among and within the three cultivated amphidiploid species of Brassica (Song et al., 1996).
In the current study, genetic differences were identified for carotenoid and Chl pigment concentrations within and among the diploid and amphidiploid Brassica species. Brassica species also produce glucosinolate secondary compounds that have important anticarcinogenic properties in humans. Quantitative and qualitative differences in glucosinolate production have been linked to genome differences among rapid-cycling Brassica species (Hill et al., 1988). For many of the pigments analyzed in the current study, the amphidiploids B. carinata and B. napus accumulated significantly less carotenoid concentrations than the diploid species and B. juneca. The six genetically related Brassica species share common evolutionary ancestry, and breeding techniques are currently used for interspecific hybridizations and species improvement. Brassica species convey unique health attributes when consumed in the diet, and are shown to accumulate high levels of carotenoids (USDA, 2005). Therefore, identification of the genetic relationships that exist among the Brassica species for carotenoid concentrations would be beneficial information for improvement programs designed to increase nutritional values.
Literature Cited
Armstrong, K.C. & Keller, W.A. 1982 Chromosome pairing in haploids of Brassica oleracea Can. J. Genet. Cytol. 24 735 739
Attia, T. & Röbbelen, G. 1986 Cytogenetic relationship within cultivated Brassica analyzed in amphihaploids from the three diploid ancestors Can. J. Genet. Cytol. 28 323 329
Bramley, P.M. 2002 Regulation of carotenoid formation during tomato fruit ripening and development J. Expt. Bot. 53 2107 2113
Cunha, C. , Tonguc, M. & Griffiths, P.D. 2004 Discrimination of diploid crucifer species using PCR-RFLP of chloroplast DNA HortScience 39 1575 1577
Cunningham F.X. Jr 2002 Regulation of carotenoid synthesis and accumulation in plants Pure Appl. Chem. 74 1409 1417
Cunningham F.X. Jr & Gantt, E. 1998 Genes and enzymes of carotenoid biosynthesis in plants. Annu. Rev. Plant Physiol Plant Mol. Biol. 49 557 583
Davies, B.H. & Köst, H.P. 1988 Chromatographic methods for the separation of carotenoids 1 185 Köst H.P. , Zweig G. & Sherma J. CRC handbook of chromatography, plant pigments. Vol. 1 Fat soluble pigments. CRC Press, Inc Boca Raton, Fla
Demmig-Adams, B. & Adams W.W. III 1996 The role of xanthophyll cycle carotenoids in the protection of photosynthesis Trends Plant Sci. 1 21 26
Demmig-Adams, B. , Gilmore, A.M. & Adams W.W. III 1996 In vivo functions of carotenoids in higher plants FASEB J. 10 403 412
Emenhiser, C. , Simunovic, N. , Sander, L.C. & Schwartz, S.J. 1996 Separation of geometric carotenoid isomers in biological extracts using a polymeric C30 column in reverse-phase liquid chromatography J. Agr. Food Chem. 44 3887 3893
Faulks, R.M. & Southon, S. 2005 Challenges to understanding and measuring carotenoid bioavailability Biochem. Biophys. Acta 1740 95 100
Frank, H.A. & Cogdell, R.J. 1996 Carotenoids in photosynthesis Photochemistry 63 257 264
Frazer, P.D. & Bramley, P.M. 2004 The biosynthesis and nutritional uses of carotenoids Prog. Lipid Res. 43 228 265
Goodwin, T.W. 1980 The biochemistry of the carotenoids 2nd ed Chapman and Hall Publishers London
Gross, J. 1991 Carotenoids 75 278 Pigments in vegetables: Chlorophylls and carotenoid AVI/Van Nostrand Reinhold New York
Harberd, D.J. & McArthur, E.D. 1980 Meiotic analysis of some species and genus hybrids in the Brassiceae 65 87 Tsunoda S. , Hinata K. & Gómez-Campo C. Brassica crops and wild allies, biology and breeding Jpn. Sci. Soc. Press Tokyo
Hill, C.B. , Carlson, D.G. & Williams, P.H. 1988 Glucosinolates in rapid-cycling Brassicas Cruciferae Nwsl. 13 50 51
Hoagland, D.R. & Arnon, D.I. 1950 The water culture method for growing plants without soil Calif. Agricultural Expt. Sta. circ. no. 347. Univ. of California Berkeley, Calif
Johnson, E.J. , Hammond, B.R. , Yeum, K.J. , Qin, J. , Wang, X.D. , Castaneda, C. , Snodderly, D.M. & Russell, R.M. 2000 Relation among serum and tissue concentrations of lutein and zeaxanthin and macular pigment density Amer. J. Clin. Nutr. 71 1555 1562
Kopsell, D.A. , Kopsell, D.E. , Lefsrud, M.G. , Curran-Celentano, J. & Dukach, L.E. 2004 Variation in lutein, β-carotene, and chlorophyll concentrations among Brassica oleracea cultigens and seasons HortScience 39 361 364
Kurilich, A.C. , Tsau, G.J. , Brown, A. , Howard, L. , Klein, B.P. , Jeffery, E.H. , Kushad, M. , Walig, M.A. & Juvik, J.A. 1999 Carotene, tocopherol, and ascorbate in subspecies of Brassica oleracea J. Agr. Food Chem. 47 1576 1581
Le Marchand, L. , Hankin, J.H. , Kolonel, L.N. , Beecher, G.R. , Wilkens, L.R. & Zhao, L.P. 1993 Intake of specific carotenoids and lung cancer risk. Cancer Epidemiol Biomarkers Prev. 2 183 187
Manton, I. 1932 Introduction to the general cytology of the Cruciferae Ann. Bot. (Lond.) 46 509 556
Müller, H. , Bub, A. , Watzl, B. & Rechkemmer, G. 1999 Plasma concentrations of carotenoids in healthy volunteers after intervention with carotenoid-rich foods Eur. J. Nutr. 38 35 44
Nwankiti, O. 1970 Cytogenetic experiments with Brassica napocampestris Hereditas 66 109 126
Pool-Zobel, B.L. , Bub, A. , Müller, H. , Wollowski, I. & Rechkemmer, G. 1997 Consumption of vegetable reduces genetic damage in humans: First results of a human intervention trial with carotenoid-rich foods Carcinogenesis 18 1847 1850
Prakash, S. & Chopera, V.L. 1993 Genome manipulations 108 133 Labana K.S. , Banga S.S. & Banga S.K. Breeding oilseed Brassicas Monogr. Theoretical Appl. Genet. 19 Springer-Verlag, New York
Prakash, S. & Hinata, K. 1980 Taxonomy, cytogenetics and origin of crop Brassicas Opera Bot. 55 1 57
Song, K. , Tang, K. , Osborn, T.C. & Ling, P. 1996 Genome variation and evolution of Brassica amphidiploids. Proc. Intl. Symp. on Brassicas. Ninth Crucifer Genetics Workshop Acta Hort. 407 35 44
Tracewell, C.A. , Vrettos, J.S. , Bautista, J.A. , Frank, H.A. & Brudvig, G.W. 2001 Carotenoid photooxidation in photosystem II Arch. Biochem. Biophys. 385 61 69
U, N 1935 Genome-analysis in Brassica with special reference to the experimental formation of Brassica napus and particular mode of fertilization Jpn. J. Bot. 7 389 452
USDA 2005 U.S. Department of Agriculture, Agricultural Research Service, National Nutrient Database for Standard Reference, release 18 18 Nov. 2005<www.nal.usda.gov/fnic/foodcomp/Data/SR18/sr18.html>
Williams, P.H. & Hill, C.B. 1986 Rapid cycling populations of Brassica Science 232 1385 1389
Yeum, K.-J. & Russell, R.M. 2002 Carotenoid bioavailability and bioconversion Annu. Rev. Nutr. 22 483 504
Zaripheh, S. & Erdman J.W. Jr 2002 Factors that influence the bioavailability of xanthophylls J. Nutr. 132 531S 534S