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Brassica Juncea Descriptive Essay

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E.S. Oplinger1, E.A. Oelke2, D.H. Putnam2, K.A. Kelling1, A.R. Kaminsid1, T.M. Teynor3, J.D. Doll1, and B.R. Durgan2


Department of Agronomy, College of Agricultural and Life Sciences and Cooperative Extension Service, University of Wisconsin-Madison, WI 53706.
2Department of Agronomy and Plant Genetics, University of Minnesota, St. Paul, MN 55108.
3Center for Alternative Plant and Animal Products, University of Minnesota, St. Paul, MN 55108. July 1991.

I. History:

Mustard (Brassica spp.), a native to temperate regions of Europe, was one of the first domesticated crops. This crop's economic value resulted in its wide dispersal and it has been grown as a herb in Asia, North Africa, and Europe for thousands of years. Ancient Greeks and Romans enjoyed mustard (sinapis) seed as a paste and powder. In about 1300, the name "mustard" was given to the condiment made by mixing mustum, which is the Latin word for unfermented grape juice, with ground mustard seeds.

Mustard has been a major specialty crop in North America since supplies from western Europe were interrupted by World War II. California and Montana were the major production areas until the early 1950s. Production of mustard in the Upper Midwest began in the early 1960s. Mustard is currently grown on approximately 250,000 acres annually in the United States. North Dakota has the largest share of domestic production. Canadian mustard production increased for twenty years until it peaked in the mid-1980s. Alberta, Manitoba, and Saskatchewan currently grow a large share of the world's mustard crop. The French people are the largest consumers of mustard (1.5 lbs/person/year), and buy approximately 70% of the annual Canadian production.

Three types of mustard, yellow, brown, and oriental, are grown in North America. Yellow mustard (Brassica hirta) comprises about 90% of the crop in the Upper Midwest. In Europe, yellow mustard is also known as white mustard (Sinapis alba - an older botanical name).

Brown and oriental mustards (Brassica juncea) are grown on limited acres. This crop is commonly produced in a rotation with small grains.

II. Uses:

More than 700 million lbs of mustard are consumed worldwide each year. Yellow mustard is usually used for prepared or table mustard, a condiment, and as dry mustard. Dry mustard is frequently used as a seasoning in mayonnaise, salad dressings, and sauces. Flour made from yellow mustard is an excellent emulsifying agent and stabilizer, and consequently, it is used in sausage preparation. Brown and oriental mustards are also used as oilseed crops. However, the strong flavor of this high-protein oilseed has made it unpopular in the livestock feed and vegetable oil markets of North America. As a result, mustard produced in North America is used primarily as a spice or condiment.

III. Growth Habit:

Mustard is an annual herb with seedlings that emerge rapidly, but then usually grow slowly. Plants cover the ground in 4 to 5 weeks with favorable moisture and temperature conditions. The tap roots will grow 5 ft into the soil under dry conditions, which allows for efficient use of stored soil moisture. Plant height at maturity varies from 30 to 45 in. depending on type, variety, and environmental conditions.

Flower buds are visible about five weeks after emergence. Yellow flowers begin to appear 7 to 10 days later and continue blooming for a longer period with an adequate water supply. A longer flowering period increases the yield potential. About half of the flowers produce dark, reddish-brown seeds that are retained in pods of 0.5 to 0.75 in. in length. Flowers pollinated during the first 15 days of the flowering period produce most of the seed.

IV. Environment Requirements:

A. Climate:

Mustard is a cool season crop that can be grown in a short growing season. Varieties of yellow mustard usually mature in 80 to 85 days whereas brown and oriental types require 90 to 95 days. Seedlings are usually somewhat tolerant to mild frosts after emergence, but severe frosts can destroy the crop. Mustard, especially the brown and oriental types, has a partial drought tolerance between that of wheat and rapeseed. Moisture stress caused by hot, dry conditions during the flowering period frequently causes lower yields.

B. Soil:

Mustard can be raised on variable soil types with good drainage, but is best adapted to fertile, well-drained, loamy soils. Soils prone to crusting prior to seedling emergence can cause problems. This crop will not tolerate waterlogged soils since growth will be stunted. Dry sand and dry, sandy loam soils should also be avoided.

C. Seed Germination:

Seed will germinate at a soil temperature as low as 40°F.

V. Cultural Practices:

A small grain crop following mustard in the rotation will usually yield more than when following continuous small grain. Mustard has several of the same diseases and insect pests as flax, oilseed rape (canola), sweet clover, soybeans, field peas, lentils, and sunflowers. Therefore, crops from this group should be avoided in the same rotation as mustard. Cereal grains are not very susceptible to the pest and disease problems of mustard.

A. Seedbed Preparation:

The seedbed should be firm, fairly level, and free of weeds and previous crop residue. Soil is firm enough for seeding when only a shallow depression of a heel is made when someone stands on the soil. Shallow tillage, just deep enough to kill weeds, should keep soil moisture

close to the surface and leave a firm seedbed. If necessary, the seedbed should be packed before planting to obtain a firm seedbed. Firm seedbeds with adequate moisture allow shallow planting and encourage rapid, uniform seed germination and emergence of seedlings. A number of growers in North Dakota have also successfully planted mustard in standing small-grain stubble and minimum-tilled stubble.

B. Seeding Date:

Planting should occur as early in the season as the environmental conditions allow. The soil temperature should be at least 40 to 45°F at a depth of 1 in. If seedlings are damaged by frost after emergence, 4 or 5 days may pass before the full extent of the damage is known. Plants should recover if the growing points are not destroyed. An earlier seeding date allows plants to benefit from the spring moisture in establishing a good canopy before weeds emerge, and to avoid heat stress during summer that causes flower or pod abortion. Early seeding also reduces the risk of damage from fall frosts that can reduce crop yields and quality. The recommended seeding date in northern Minnesota and Wisconsin is May 1 to 25. Seeding later than May 15 frequently results in lower yields.

C. Method and Rate of Seeding:

Yellow mustard, which has approximately 100,000 seeds/lb, is solid seeded with a grain drill at a rate of 8 to 14 lbs/acre. The higher rate should be used on heavy, fertile soils or on those where emergence is difficult. Brown and oriental mustards have 200,000 seeds/lb and should be solid seeded at a rate of 5 to 7 lbs/acre. Seed is small and must be planted shallow at a 1/2 to 1 in. depth. If very dry soil conditions exist seeding depth should be increased to 1 1/2 in. If mustard stands are poor, quick decisions for reworking and reseeding should be made.

D. Fertility Requirements:

Mustard generally responds to nutrient additions in a similar way as does rape or canola. Soil tests should be used to determine nutrient need. Optimal (medium) soil test levels are about 15 to 20 ppm Bray P, and 80 to 100 ppm K. At these levels fertilizer should be applied at a rate of about 45 lbs/acre P2O5 and 80 lb/acre K2O. When fertilizer is banded, the bands should be placed below and to the side of the seed furrow. Mustard responds well to nitrogen additions with optimum yields occurring at about 100 to 120 lb/acre N. Where mustard follows legumes or manure additions, appropriate credits should be taken.

Work in the western U.S. shows that mustard responds well to sulfur(s) on low S-supplying soils. Sulfur fertil-sandy soils in northwestern Wisconsin and northern Minnesota which have not been manured in the past two years.

On soils deficient in boron (testing at less than 0.5 ppm B), apply 0.5 to 1 lb/acre in a uniform broadcast application. Never band B near the seed.

Soils with a pH near neutral (7.0) are desired for this crop. Nevertheless, an alkaline pH and slightly saline soils are tolerated. Mustard has a tolerance to soil salinity that is similar to barley.

E. Variety Selection:

Varieties of yellow mustard are usually earlier maturing, lower yielding, and shorter in height than brown or oriental varieties. Yield differences among the types of mustard are reflected usually in the prices offered by contracting firms. Contracting firms usually supply growers with the appropriate varieties. Some mustard varieties include:


—Yellow mustard. Similar to Ochre in field performance. Originated in Germany. Distributed by Northern Sales Co. Ltd., Winnipeg. Licensed in 1974.


—Yellow mustard. Released by Colman Foods, Norwich, England in 1970. Distributed by Minn-Dak Growers Association, Grand Forks, ND.


—Yellow mustard. Released by Agriculture Canada, Saskatoon. Licensed in 1981.


—Yellow mustard. Similar to Kirby in field performance but has a high mucilage content desired by processors. Released by Colman Foods Norwich, England in 1978. Distributed by Minn-Dak Growers Association, Grand Forks, ND.

Carrow 85

—Oriental mustard. Undesirable small seed. Released by Colman Foods, Norwich, England in 1980.


—Oriental mustard. Released by Agriculture Canada, Saskatoon. Licensed in 1977.

Lethbridge 22A

—Oriental mustard. Released by Agriculture Canada in 1967, Lethbridge. Licensed in 1974.


—Brown mustard. Released by Agriculture Canada, Saskatoon. Licensed in 1976.

Varietal trials for mustard in your area should be consulted for yield and other agronomic characters as in Minnesota (Table 1).

F. Weed Control:

Weeds can greatly reduce mustard yields. Weed seeds, which are difficult to remove, can cause high losses during seed cleaning and lower market grades. Good weed control is based on preparation of a clean field and shallow seeding to encourage quick, uniform emergence. Young mustard seedlings do not compete well with weeds and the early establishment of a uniform, vigorous crop helps control annual weeds. The crop cannot be cultivated after emergence.

Mustard, especially the oriental and brown types, should be grown on land with as little wild mustard as possible to avoid costs of removal and loss of tame mustard seeds. Wild mustard seed can be mechanically separated from the larger-seeded yellow type, but separation is not possible with the smaller-seeded brown and oriental types. Wild mustard seed often reduces the crop quality to the sample grade. Production of rapeseed and mustard on the same fields is also not recommended since seed mixtures can occur easily and degrade both crops.

Control of perennial weeds such as Canada thistle, field bindweed, and quackgrass, should be started in the fall or prior to planting in the spring. Control Canada thistle by applying Roundup (glyphosate) before the last killing frost in the fall when it is still actively growing. Spring treatment of Canada thistle prior to planting is not usually adequate for complete control. Apply Roundup to quackgrass when it is at least 8 in. tall and growing actively. Allow 3 days between application and tillage. Control field bindweed with Roundup when it is actively growing on moist sod and is at or past full bloom, preferably in late summer or fall the year before planting mustard. Spring treatment to control perennial weeds before planting is usually not practical.

Trifluralin (Treflan EC) is labeled for use on mustard in Minnesota and North Dakota for control of a wide variety of grasses and broadleaf weeds; however, it will not control wild mustard. The rate of herbicide applied will vary with the soil type, organic matter content, and species of weeds that need to be controlled. Check the label for the correct rate to use on your fields. Trifluralin must be applied before seeding and incorporated thoroughly in the soil for maximum effectiveness. Mustard is sensitive to the broadleaf herbicides used on cereal crops, such as 2,4-D and MCPA, and spray drift from adjacent fields must be avoided. Crops that can be sprayed with 2 4-D or MCPA should follow mustard in the rotation so volunteer plants can be controlled.

G. Diseases and Control:

This crop is vulnerable to several diseases, among which the most serious are Sclerotinia stalk rot (white mold), downy mildew, white rust, leaf spots, and mosaic virus. Good cultural practices are the most effective control measures for diseases. These practices should include keeping records of disease occurrence, compliance with the proper crop rotation, control of host plants for diseases in fallow fields and non-crop areas, and use of seed treatments.

Sunflower, rapeseed, canola, safflower, soybeans, crambe, and drybeans should not be grown in rotation with mustard since they have similar disease problems. If these crops are produced in rotation with mustard, damage from these diseases can increase to economic levels. Numerous broadleaf weeds can also serve as hosts or sources of infection for these diseases. Wild mustard, pigweed, field pennycress, and shepherd's purse are examples of predominant hosts. One of the best methods to avoid serious disease problems in mustard (and leaf diseases of small grains) is to produce this crop in a small grain rotation. Mustard should be spaced four or more years apart in the crop rotation to avoid problems with soil-borne diseases. After potato or flax crops, one year should pass before mustard is raised on the same field due to the presence of root rot or damping-off pathogens.

H. Insects and Other Pests:

Growers should monitor fields closely to detect insect problems that can result in significant yield losses. Flea beetles and caterpillars of the diamondback moth have been the most serious pests. Young seedlings can be seriously damaged by flea beetles soon after emergence. Feeding activity of adult beetles causes a shot-holed appearance to cotyledons and the first true leaves. Damaged plants may die or suffer a reduction in vigor. Hot, sunny weather encourages feeding while cool, damp conditions slows insect feeding and promotes crop growth. Injured plants may wilt and die during hot, dry weather, which results in mild to severe yield losses. Serious crop damage does not usually occur once the crop develops beyond the seedling stage since vigorous plants can outgrow beetle defoliation.

Losses caused by flea beetles can be minimized by practicing the proper cultural methods. A well-tilled, firm seedbed with adequate fertility should permit young seedlings to outgrow beetle damage during the vulnerable stages early in the season. The presence of a few flea beetles or scattered shot-holing is not cause for serious concern. However, if beetles are numerous and feeding damage is present on most cotyledons, prompt control may be necessary. Malathion EC at 1 1/4 lbs/acre, Carbaryl (Sevin) at 1 lb/acre, Ethyl parathion 8E at 1/2 lb/acre, and Thiodan EC at 3/4 lb/acre can be used for control of flea beetles. Read labels for waiting period and correct timing.

Larvae of diamondback moths eat leaves, flowers, and green seed pods. Malathion used at 2 1/2 lbs/acre will control these caterpillars. Damage from sugar beet nematodes can be avoided when sugar beets are not grown on the same field two years before or after mustard, since mustard is a host plant for this nematode. Consult local Extension bulletins for further information on the control of insects and other pests.

I. Harvesting:

The normal maturation of the crop, wind, and rain do not cause shattering before cutting. However, the actual harvesting operations can cause great shattering losses when the plants are overripe. Yellow mustard does not shatter readily and can be straight combined if the crop has matured uniformly (10% moisture) and is free of green weeds. If the crop is weedy or uneven in maturity it should be swathed. Swathing, if deemed necessary or preferred, should be done when 60 to 70% of the seed has turned yellow-green. Plants should be cut just beneath the height of the lowest seed pods. The swath will then settle into the stubble and reduce the chance of being blown by high winds. Yellow mustard does not cure quickly. Straight combining is therefore recommended at 12 to 13% moisture, followed by artificial drying, to obtain uniform quality and highest yield.

Brown and oriental varieties will shatter more readily when ripe and should be swathed. The swathing should begin after the general leaf drop and when the overall field color has changed from green to yellow or brown. Pods sampled from the middle of racemes from several plants, in arm representing the average maturity, should be examined for physiological maturity. About 75% of the seeds may have reached the mature color of yellow or brown. The remaining green seeds will mature in the swath before combining.

Swathing should be done under conditions of high humidity or when morning dew is on ripe pods to decrease shattering losses. Windrows tend to be bulky and subject to scattering by the wind. A roller or steel drum should be used to press the swath into the stubble. The combine should be adjusted so seeds are threshed completely by using the lowest cylinder speed, which is set at approximately 600 RPM, and the appropriate cylinder opening. The reel may cause shattering when straight combining, but it can be removed or lifted above the plants it the stand is good. If the reel is needed, remove half of the bats and reduce its speed. Cylinder speed may need to be adjusted during the day as crop moisture content may vary.

J. Drying and Storage:

When the mustard seed reaches a moisture content of 10% or less it can be stored safely. The harvested seed should be handled carefully since it will crack easily when moved in and out of storage. The damaged seed becomes dockage and is a loss to the grower. Air temperatures for seed drying should not exceed 150°F and seed temperature should stay below 120°F. Use of drying equipment designed for corn or wheat may require some modification when drying mustard. A fine screen will be needed to prevent loss of the smaller seed. Storage bins must be free of cracks or holes.

VI. Yield Potential and Performance Results:

Mustard yields in the Upper Midwest have been variable due to differences among varieties, cultural practices, and environmental conditions. Yields for research trials in Minnesota have ranged from 868 to 1,861 lbs/acre (Table 1.). The MINN-DAK Growers Association reported that yields in the past few years were low due to weather conditions. Growers were producing 800 to 900 lbs/acre of yellow mustard, while brown and oriental mustards were yielding 1,000 to 1,100 lbs/acre. A fair estimate for the yield potential of production fields in the Upper Midwest would be 800 to 1,000 lbs/acre. However, yields of 1,400 lbs or more per acre are possible in areas with favorable growing conditions.

Table 1. Average yield and other agronomic characteristics of yellow, brown, and oriental varieties of mustard in Minnesota field trials*.


Yield at 3 Locations


Test weight

Seeds Number
(1,000 lb)

Days from
planting to








Yellow (Brassica hirta)


































Oriental (Brassica juncea)












Brown (Brassica juncea)












*Data from Varietal Trials of Farm Crops, 1990 edition, Minnesota Agricultural Experimental Station, Univ. of Minnesota, Report 24 (AD-MR-1953).
1One year of data; 2Two years of data; 3Three years of data; 4Oven-dry basis, average of four years/location; 51=erect, 9=horizontal.

VII. Economics of Production and Markets:

The cash production costs are less due to lower seed and pesticide costs than for hard red spring wheat. In 1991 the cash costs were estimated at $67.00 for hard red spring wheat and $56.00 for mustard for northwestern Minnesota.

Mustard is produced as a specialty grain and should be grown under contract to guarantee a selling price and market for the producer. A contract is made by the grower with the shipper to supply seed of a specified quality for delivery at a future date. Contract prices for mustard seed in the Upper Midwest for 1991 were 10.5 cents/lb (up to a certain poundage/acre, such as 600 lbs) for yellow mustard and 9 cents/lb for brown and oriental mustards (up to 800 lbs/acre). Contract prices for Canadian mustard in 1991 were approximately 11 cents/lb for yellow mustard for the first 500 to 1,000 lbs/acre, 9 cents/lb for brown mustard for the first 700 to 1,200 lbs/acre, and 8 cents/lb for the first 500 to 1,200 lbs/acre of oriental mustard.

Consumption of mustard has been steady and growth of the mustard market is directly related to population growth. This stability of demand is due to the lack of any real substitutes for mustard. Consumers will not substitute for mustard as this would not save much money. There is a limited number of alternative markets when a surplus is produced.

VIII. Information Sources:

  • Mustard Production in Manitoba. 1980. J.R. Rogolsky, Agriculture Manitoba, Agdex #140-10.
  • Flax, Mustard, Spring Rape: Alternative Crops for
  • Idaho's Cooler Region? 1982. D.L Auld, GA. Murray, G.F. Carnahan, LA. Benson, and B.W. Studer, University of Idaho Cooperative Extension Service, Current Information Series No. 524.
  • Tame Mustard Production. 1987. LL. Helm, and A.A. Schneiter, Circular A-935, Noah Dakota State University Extension Service, Fargo. ND.
  • The Mustard Growers Manual. Rob Tisdale (ed.)., compiled by Seana C. Forhan, The Mustard Association, distributed by Manitoba Agriculture.
  • Fertilizer Recommendations for Agronomic Crops In Minnesota. George Rehm and Michael Schmitt, Minnesota Extension Service, AG-MI-3901.
  • Varietal Trials of Farm Crops. 1990. Minnesota Agricultural Experiment Station, University of Minnesota. Report 24 (AD-MR-1953).
  • Production of Mustard in Northern Idaho. 1991.
  • Stephen Guy, Current Information Series 889. University of Idaho Cooperative Extension Service, Moscow, Idaho.

The information given in this publication is for educational purposes only. Reference to commercial products or trade names is made with the understanding that no endorsement for one product over other similar products is implied by the Minnesota and Wisconsin Extension Services.



Extensive mapping efforts are currently underway for the establishment of comparative genomics between the model plant, Arabidopsis thaliana and various Brassica species. Most of these studies have deployed RFLP markers, the use of which is a laborious and time-consuming process. We therefore tested the efficacy of PCR-based Intron Polymorphism (IP) markers to analyze genome-wide synteny between the oilseed crop, Brassica juncea (AABB genome) and A. thaliana and analyzed the arrangement of 24 (previously described) genomic block segments in the A, B and C Brassica genomes to study the evolutionary events contributing to karyotype variations in the three diploid Brassica genomes.


IP markers were highly efficient and generated easily discernable polymorphisms on agarose gels. Comparative analysis of the segmental organization of the A and B genomes of B. juncea (present study) with the A and B genomes of B. napus and B. nigra respectively (described earlier), revealed a high degree of colinearity suggesting minimal macro-level changes after polyploidization. The ancestral block arrangements that remained unaltered during evolution and the karyotype rearrangements that originated in the Oleracea lineage after its divergence from Rapa lineage were identified. Genomic rearrangements leading to the gain or loss of one chromosome each between the A-B and A-C lineages were deciphered. Complete homoeology in terms of block organization was found between three linkage groups (LG) each for the A-B and A-C genomes. Based on the homoeology shared between the A, B and C genomes, a new nomenclature for the B genome LGs was assigned to establish uniformity in the international Brassica LG nomenclature code.


IP markers were highly effective in generating comparative relationships between Arabidopsis and various Brassica species. Comparative genomics between the three Brassica lineages established the major rearrangements, translocations and fusions pivotal to karyotype diversification between the A, B and C genomes of Brassica species. The inter-relationships established between the Brassica lineages vis-à-vis Arabidopsis would facilitate the identification and isolation of candidate genes contributing to traits of agronomic value in crop Brassicas and the development of unified tools for Brassica genomics.


Extensive genome sequencing and genetic mapping studies have been performed on members of the Brassicaceae family which contains the most widely studied model species, Arabidopsis thaliana (At) and many economically important vegetable and oilseed crops belonging to the genus Brassica. An avowed goal of structural and functional genomics of At is to develop improved strategies for precision breeding of crop plants related to the model species. Since the genome size of Brassica species (529–696 Mb for the diploids and 1068–1284 Mb for the polyploids) [1] is much larger than that of At (125 Mb), there is a high probability that novel gene interactions have evolved in the Brassicas through the processes of sub-functionalization and/or neo-functionalization of paralogs [2-4]. Comparative mapping between At and Brassica species, coupled with the base knowledge of mutation-based functional analysis in At and QTL mapping in crop Brassicas, could greatly contribute towards a better understanding of the genetic architecture for the conserved as well as the evolved traits of agronomic value in the Brassicaceae.

The three diploid Brassica species, B. rapa (n = 10, AA), B. nigra (n = 8, BB) and B. oleracea (n = 9, CC) and the two allopolyploids, B. napus (AACC) and B. juncea (AABB), have been subjected to extensive genetic mapping using molecular markers to identify loci associated with various qualitative and quantitative traits of agronomic interest [5-12]. Some of the mapped quantitative traits like pod size, pod number, pod density, seed size, seed number per pod and oil content are of great importance in improving the yield of the oilseed Brassica species [11,13,14].

Recent attempts to develop a unified comparative genomics system in the Brassicaceae has recognized the existence of 24 conserved genomic blocks [15] which is an extension of 21 syntenic blocks identified in B. napus in an earlier study [16]. Comparative mapping studies between members of family Brassicaceae [16-20], At and Arabidopsis lyrata [21], At and Capsella rubella [22] and the identification of an ancestral karyotype (AK) [23] have also stimulated interest in the evolutionary processes involved in the diversification of different lineages in the Brassicaceae and variations in chromosome number of different species vis-à-vis their ploidy status. Most of the earlier studies on comparative mapping in Brassica species [16,18-20,24] have relied on the use of RFLP markers. However, the deployment of RFLP markers in large segregating populations is a rather cumbersome and laborious process. In recent years, several studies have highlighted the immense potential of polymorphisms in intron sequences for the development of markers for genetic mapping [25-27].

In the present study, we have successfully used PCR-based Intron Polymorphism (IP) markers for the development of a comparative map between B. juncea, related Brassica species (B. napus and B. nigra) and At. We also analyzed the segmental structure of the A and B genomes of B. juncea in terms of the 24 genomic blocks (A-X) proposed earlier [15]. We compared colinearity of the B. juncea and At genomes and also analyzed synteny between the A, B and C genomes of the Brassica species. Additionally, homoeologous linkage groups of the three genomes were identified. On the basis of homoeology among the three genomes, we propose a re-designation of the B genome linkage groups assigned earlier for B. nigra [18] and B. juncea [11]. The comparative map between B. juncea and At developed in this study, in conjunction with the extensive information available from functional genomics studies of the At genome, will greatly facilitate the identification of candidate genes and novel gene interactions responsible for the domestication and evolution of the yield influencing traits in B. juncea and other crop Brassicas.


Comparative map of B. juncea and Arabidopsis thaliana (At) using Intron Polymorphism (IP) markers

Single copy genes from Arabidopsis (At) [28], physically located at an approximate distance of 100–200 kb, were used to design PCR primers spanning intronic sequences. In cases where large genomic regions were devoid of single copy genes, primers were designed from multiple copy genes. Primers for PCR amplification were designed from exon sequences which showed strong nucleotide conservation between At and the corresponding EST or GSS sequences described for any Brassica species [29].

Of the 1180 primer pairs thus designed, 383 (32%) showed polymorphism between the B. juncea lines, Heera and Varuna, parents of the DH mapping population used in this study and in the earlier mapping studies on B. juncea [11,30]. Genotyping using the 383 polymorphic primer pairs (for primer sequences see Additional file 1) generated 486 loci in B. juncea of which 67% were scored as co-dominant markers and the remaining 33% were scored as dominant markers. These 486 loci were incorporated into the framework map developed by Pradhan et al. [30]. Additionally 34 RFLP markers placed earlier on the B. juncea map [11] were assigned corresponding At loci by subjecting the available sequence data to NCBI BLASTN search and identifying the most significant BLASTN hit as the source At gene. A linkage map of B. juncea consisting of 533 At loci (486 IP, 34 RFLP and 13 gene markers) and covering a total genetic length of 1992.2 cM is shown in Figure ​1, ​2, ​3, ​4, ​5. The ten A genome linkage groups (LGs) of the B. juncea map were designated A1–A10 and correspond to the N1–N10 linkage groups of B. napus [16]. The remaining eight B genome LGs were designated B1–B8 based on homoeology between the A, B and C genomes as determined in the present study. The corresponding linkage group nomenclature proposed earlier for the B. nigra genome (G1–G8) [18] and the B genome of B. juncea (J11–J18) [11] is also given in parentheses in Figure ​4, ​5 and Table ​1.

Figure 1

Genetic map of B. juncea showing three linkage groups of the A genome (A1, A2 and A3). The corresponding nomenclature followed earlier in B. juncea (J1–J3) [11] and B. napus (N1–N3) [16] is given in parentheses. Each genetic locus bears...

Figure 2

Genetic map of B. juncea showing five linkage groups of the A genome (A4, A5, A6, A7 and A8). The corresponding nomenclature followed earlier in B. juncea (J4–J8) [11] and B. napus (N4–N8) [16] is given in parentheses. Each genetic locus...

Figure 3

Genetic map of B. juncea showing two linkage groups of the A genome (A9 and A10). The corresponding nomenclature followed earlier in B. juncea (J9 and J10) [11] and B. napus (N9 and N10) [16] is given in parentheses. Each genetic locus bears the name...

Figure 4

Genetic map of B. juncea showing five linkage groups of the B genome (B1, B2, B3, B4 and B5). The corresponding nomenclature followed earlier for the B genome of B. juncea (with a prefix J) [11] and B. nigra (with a prefix G) [18] is given in parentheses....

Figure 5

Genetic map of B. juncea showing three linkage groups of the B genome (B6, B7 and B8). The corresponding nomenclature followed earlier for the B genome of B. juncea (with a prefix J) [11] and B. nigra (with a prefix G) [18] is given in parentheses. This...

Table 1

Characteristics of B. juncea map based on the distribution of 533 At loci derived from the five chromosomes of Arabidopsis thaliana

The 533 At loci mapped with variable frequencies to all the 18 linkage groups of B. juncea (Table ​1). Linkage group A3 contained the highest number of markers (66) while B5 had the least with only 9 markers. Overall, 63% of the markers mapped to the A genome (A1–A10) at an average marker density of 0.34 and average marker interval of 2.9 cM (Table ​1). The B genome (B1–B8) contained the remaining 37% of the markers with a marker density of 0.24 at an average marker interval of 4.7 cM, indicating that there is less polymorphism in the B genome as compared to the A genome in the intronic regions. As a result a number of unmapped islands were found in the B genome (an island is defined as a region with a gap of ≥ 20 cM between adjacent markers) (Table ​1).

An uneven distribution of At loci originating from each Arabidopsis chromosome was observed in the genome of B. juncea. Among the 10 LGs of the A genome (A1–A10), all the linkage groups except A2, A6, A7, A8 and A10 contained At loci from each of the five Arabidopsis chromosomes (At C1–At C5). A2 and A6 were devoid of loci from At C3 and At C4 respectively. A7 did not contain any locus from At C4 and At C5. The linkage group A8 was composed of markers from At C1 and At C4 while A10 was composed of markers from At C1 and At C5 (Table ​1). Linkage group A9 was found to be the most chimeric linkage group consisting of 12 genomic blocks followed by A3 with 10 blocks. Linkage group A10 was least chimeric with only 3 blocks followed by A5 with 4 blocks and A1 and A4 with 5 blocks each. The simplest genomic organization was observed in linkage groups A1 (primarily consisting of At C3 and At C4 loci), A4 (composed primarily of At C2 and At C3 loci), A8 (composed of At C1 and At C4 loci) and A10 (composed of At C1 and At C5 loci). Uneven distribution of the At loci was also observed in the B genome (B1–B8) of B. juncea of which linkage groups B3, B4 and B8 contained loci derived from all the five chromosomes of Arabidopsis (Table ​1).

The organization of the B. juncea linkage map with respect to the At genome was also studied on the basis of the distribution of 24 genomic blocks (A-X) described for a hypothetical ancestor of the At and Brassica lineages by Schranz et al. [15]. This approach facilitated the identification of conserved blocks between At and B. juncea. A conserved block was defined as a region that contained at least two At loci from the same block region. In some instances, a block was recognized even with a single mapped marker if one or more markers of the same block were found mapping at the corresponding region in earlier maps for the A and B genomes [16,18]. Using this criteria, a total of 67 genomic blocks were identified in the A genome of B. juncea with an average of 2.8 paralogous blocks for each block recognized in the hypothetical ancestral species (Table ​2). As compared to the A genome, we identified a lesser number of blocks (42) in the B genome (Table ​2). Among the 42 blocks identified in the B genome of B. juncea, the larger blocks viz. E, F, J, R and U were observed to be represented by three paralogous blocks within the B genome (Table ​2).

Table 2

Distribution of the 24 genomic blocks (A-X) in the A and B genomes of B. juncea

Comparative block arrangement in the A genomes of B. juncea and B. napus

The A genome of B. juncea (present study) was compared with the A genome of B. napus [16] based on the arrangement of the 24 genomic blocks. For comparison, all the RFLP markers mapped on B. napus were converted to corresponding At loci based on the information available in the supplementary Table S1 of Parkin et al. [16] and assigned to different blocks (A-X) (see Additional file 2). The comparative block arrangement in the A genomes of B. juncea and B. napus has been shown in Figure ​6a.

Figure 6

Comparative block arrangement in (a) the A genome of B. juncea (A1–A10; present study) and B. napus (N1–N10) [16] and (b) the B genome of B. juncea (B1–B8; present study) and B. nigra (G1–G8) [18]. Blocks identified in...

In terms of the arrangement of the blocks, the A genomes of both B. juncea and B. napus were essentially collinear. We identified five new blocks (M-N in A1, G-H in A3 and H block in A9) in the A genome of B. juncea which were not detected in the corresponding A genome LGs of B. napus (Figure ​6a). All these new blocks were identified in regions with a reasonably high density of IP markers. Additionally, we also established the presence of some blocks (O-P in A2/N2, N in A3/N3, X and H in A9/N9) (Figure ​6a) in the A genome of both B. juncea and B. napus which were not designated earlier in B. napus by Schranz et al. [15]. Block status to these regions were assigned as we observed the presence of At loci from these blocks not only in the homoeologous LGs of the A and C genomes of B. napus [16] but also in the corresponding LGs of the A genome of B. juncea in the present study (blocks marked with asterisk in Figure ​6a). Blocks L, W and X observed in the linkage groups N2, N3 and N6 of B. napus (Figure ​6a) could not be detected in the homoeologous A genome LGs of B. juncea in the present study. This, in all probability, is due to the absence of IP markers representing these blocks in the map. These regions show up as large gaps in the map shown in Figure ​1, ​2, ​3.

Potential centromeric regions were predicted in the linkage groups A3, A4, A5 and A7 of B. juncea based on the non-rearranged blocks that flank the centromere in AK and At as proposed by Schranz et al. [15]. The I-J non-rearranged blocks in A4 represent the conserved centromeric regions between the Brassica species and AK, while the G-H non-rearranged blocks in A7 represent the conserved centromeric regions between the Brassica species, AK and At. Similar centromeric regions were predicted for the corresponding N4 and N7 LGs of B. napus [16]. In A3 and A5 of B. juncea, alternate centromere sites could also be predicted (Figure ​6a) due to the presence of mapped markers corresponding to the pericentromeric regions in At. In N3 of B. napus, a conserved centromere was predicted between the J-I blocks [15]. However, our mapping data on B. juncea suggests that the centromeric region is present between the G-H blocks. All the five markers representing the G-H blocks in A3 were derived from the pericentromeric region of At C2 (Figure ​1). Moreover, A3 shared the block arrangement of F-G-H with the ancestral karyotype (AK chromosome 3) [15] unlike in At where the F and G-H blocks are located on different chromosomes. The possibility of a centromere between the J and I blocks in A3 is further weakened by the fact that the orientation of the J block is inverted with respect to the I block in A3 of B. juncea which is otherwise non-rearranged in AK [15]. In N5 of B. napus, a centromere is predicted between the B-C blocks [15]. Based on the presence of several markers from the pericentromeric region between the J and I blocks in A5 of B. juncea, an alternate probable site of the centromere has been predicted in this linkage group.

Comparative block arrangement in the B genomes of B. juncea and B. nigra vis-à-vis At

Comparative organization of the A genomes of B. juncea and B. napus (described above) based on the block boundaries defined by Schranz et al. [15] confirmed that the set of 24 genomic blocks (A-X) can be used to delineate the genomic organization of Brassica genomes. The same block definition was therefore used to study the segmental organization of the B genome of B. juncea. The segmental structure of the B genome (B1–B8) of B. juncea in relation to the At genome is represented schematically in Figure ​4, ​5. Since our study allows for a detailed gene-to-gene alignment between the B genome of Brassica and At which has not been reported earlier, the description of the genomic organization of the eight linkage groups (B1–B8) with respect to At is described in detail below.

B1 (G7/J17): B1 predominantly consists of two long stretches of collinear genes from At C3 (block F), one each at the two ends of the linkage group. The two F blocks constitute about 62% of the total mapped area. The colinearity of gene order in comparison with At is suggestive of at least one inversion each in both the F blocks signifying that inversions occurred prior to the diversification of the two blocks. The orientations of the F blocks are inverted with respect to each other. The presence of duplicated F blocks on a single linkage group appears to be a unique feature of the B. nigra genome and has not been observed in the A and C genomes. The middle segment of B1 (comprising around 10% of the LG) has markers from block Q (At C5) and block D (At C1).

B2 (G5/J15): This LG consists of four blocks, E-M-R-U, with the E block (At C1) constituting 20% of the top segment and the U block (At C4) constituting 54% of the lower segment of this linkage group. Except for minor rearrangements, the gene order in the U block is highly collinear with its corresponding At C4 region. These rearrangements also explain the presence of markers from the adjacent R and M blocks in this region.

B3 (G8/J18): B3 is constituted of blocks N-M-O-P-J-W-R. Blocks N and M, constituting the top segment of B3, share colinearity of gene order with their counterparts in At. Blocks O and P harbour at least one inversion each which explains the break in colinearity of the gene order as compared with At. The middle segment of B3 is represented by block J (At C2). A minimum of two inversions within the J block could have resulted in the reshuffling of gene markers as compared with At. Blocks R and W constitute the terminal region of B3. A mixture of markers from At C2, At C3 and At C4 is seen between the J and W blocks. Three of these markers viz. At3g47370, At4g07666b and At4g09820 belong to blocks M and P. Their placement between the J and W blocks could be a consequence of the inversions described above.

B4 (G6/J16): B4 consists of small collinear stretches contributed by all the five chromosomes of At. The block order is represented by E-T-N-A-J, with an X block insertion within the J block. The top of the LG is made up by blocks E, T and N. The arrangement of the genes in the N block is suggestive of rearrangements within the block.

B5 (G1/J11): This linkage group consists of two major blocks: F, constituting 30% and J contributing 12% of the total mapped area. The gene order in these two blocks are collinear with the F and J blocks of At.

B6 (G4/J14): About 70% of the total mapped area in the lower part of the B6 linkage group is made up of blocks from At C1 in the order C-B-A-D. The upper part of the linkage group shows the presence of the K block from At C2 while the remaining portions above the K block could not be assigned to any syntenous block as no At loci could be mapped to this region.

B7 (G3/J13): This LG consists predominantly of gene markers from At C1, representing ~53% of the mapped area. The block order is A-B-U-A-E-H. The distribution of the At C1 specific markers is suggestive of the presence of three large collinear regions separated by large gaps and constituting the top, middle and terminal segments of B7. The top portion of this LG comprises of an A block harboring a minimum of one inversion breaking its colinearity with its counterpart in At. The middle segment is formed by a collinear stretch of genes belonging to the B and U blocks. The terminal segment of B7 comprises another A block followed by a long stretch of genes belonging to block E. The gene order in this block indicates two inversions since its divergence from At.

B8 (G2/J12): This is a highly chimeric linkage group comprising nine blocks [A-G-H-I-W-R-U (with an N-Q segment inserted within the U block)] with loci derived from all the five chromosomes of At. The terminal segment of the linkage group is consists of a U block from At C4 with insertions from the N and Q blocks. The gene order of At loci in this block revealed the presence of two sub-blocks ordered in opposite orientations. This break in colinearity can be explained by at least one major inversion which would also explain the appearance of two regions harboring markers from the Q block. The middle portion of this LG is formed of a large collinear region from At C5 (block R-W). Above the R-W block, there is a contiguous stretch of markers derived from blocks G, H and I. This contiguous block arrangement, G-H-I, is observed in At C2 whereas it is shared between linkage groups 3 (G-H) and 4 (I) in AK with a predicted centromere between the G and H blocks in both At and AK [15]. On the basis of the foregoing, a conserved region constituting the centromere could be predicted between the G and H blocks in B8.

Lagercrantz [18] reported a comparative map in B. nigra by mapping 284 RFLP loci generated from 160 At DNA fragments. To compare the block arrangement in the B genome of B. juncea with its diploid progenitor B. nigra, the RFLP loci of the B. nigra map, wherever possible, were converted to orthologous At loci and assigned to different blocks (A-X) (see Additional file 3). For the conversion, the RFLP probes whose GenBank sequences could be retrieved, were subjected to NCBI BLASTN search to assign the best corresponding homologous At loci. For the other RFLP probes (which mapped on different At physical maps available at TAIR [31], the corresponding map position in the AGI physical map was determined and the closest At locus name was assigned to the RFLP probe. The comparative block arrangement observed in this study and that reported by Lagercrantz [18] is shown in Figure ​6b. Although a general colinearity is observed between the two maps, we identified some new blocks (T in B4 and G, H, I in B8) in the B genome of B. juncea which were not detected in the earlier study. Conversely, we also identified some blocks in the B. nigra map [18] (C in G1, V in G2, U in G3, Q and V in G4, W in G5 and S in G6) which could not be identified in the B genome of B. juncea in this study (Figure ​6b and Additional file 3). These discrepancies in block arrangements between the two maps could be due to a limited coverage of At loci in the B genome of B. juncea (Table ​1) and our inability to convert some of the At RFLP loci described earlier [18] in the B. nigra map. Further saturation of the B genome map would facilitate greater accuracy in assigning block status to these regions found to be different between the two genomes. Among the different blocks identified in the B genome, F, J, R and U were the major blocks with the maximum coverage. These were represented as three homoeologous blocks in both the maps. A similar triplication for three of these blocks (F, J and R) was also reported in the previous study by Lagercrantz [18].

Identification of homoeology between the A, B and C genomes of Brassica species

Considerable homoeology has been reported between the LGs of the A and C genomes [16]. With the mapping information for the B genome derived from the present study, we were able to compare the block-based architecture of the LGs of the three diploid Brassicas to understand the extent of homoeology shared between the three genomes. This analysis provided insights into the prominent rearrangements that shaped the three genomes and also identified those ancestral blocks which remained unaltered in all the three diploid Brassica genomes. For this comparative study, the RFLP loci of the C genome of B. napus [16] were converted to their corresponding At loci (Additional file 4). Figure ​7 provides an overall view of the homoeology between the three genomes.

Figure 7

The block arrangements in the A and B genomes are based on the consensus block arrangement of the A genomes of B. juncea (A1–A10; present study) and B. napus (N1–N10) [16] and the B genomes of B. juncea (B1–B8; present study) and...

The comparative map of the A (A1–A10), B (B1–B8) and C (C1–C9) genomes of Brassica (Figure ​7) is based on the consensus block arrangements identified in the comparative map of the A genome of B. juncea (A1–A10 of the present study) and B. napus (N1–N10) reported earlier [16], B genome of B. juncea (B1–B8; present study) and B. nigra (G1–G8) reported earlier [18] and C genome of B. napus (N11–N18) [16]. This comparative analysis shows that all the linkage groups of the C genome except N16 and N17 [16] correspond to LGs of the A genome based on the extent of the homoeology between them. Hence, we propose that N16 and N17 should be designated as C7 and C6 respectively (Figure ​7). Based on the homoeology observed among the A, B and C genomes in our study, a similar nomenclature is proposed for the B genome (B1–B8). This homoeology-based designation will enable greater accuracy and uniformity in the international nomenclature of the three diploid progenitor genomes.

All the linkage groups belonging to the three diploid Brassica species could be divided into ten categories or groups (Group1–10) based on the extent of homoeology between them (Figure ​7). Group 1 consists of A1/B1–B2/C1. A1 was entirely homoeologous to C1, both being constituted by the block arrangement F-T-U. The F-T-U arrangement was specific to the Rapa/Oleracea lineage. Interestingly, this F-T-U arrangement was found repeated in both the A (A3) and the C (C3) genomes (Group 3, Figure ​7) but was absent from the B genome. The linkage group B1 shared the F block with A1/C1 in Group1. Due to the presence of two F blocks in B1, this linkage group also had homoeology with Group 3. B2, which shared the U block with A1 and C1, could also be placed in this group. In Group 2 (A2/B2/C2), A2 and C2 were completely homoeologous, while B2 showed homoeology with A2 and C2 for the block motifs R-W-E-O-P. One inversion in B2 could explain the separation of the E block from the R-W block combination (Figure ​8). Group 3 (A3/B1–B3/C3) members shared the common block arrangement R-W-J-I-P-O. B3 was almost entirely composed of this block arrangement, while an additional F-T-U block was present in A3 and C3. Members of Group 4 (A4/B4/C4) shared the block motif J-I-S-N-T. A4 and B4 appeared to be homoeologous along their entire length, while C4 had acquired an additional J block. The presence of two J blocks in C4 was a C genome-specific rearrangement. Due to the presence of an additional J block, C4 has also been placed in group 5. In Group 5 (A5/B5/C4–C5), A5 and B5 were homoeologous along their entire length sharing the block motif J-C-F. C5 shared partial homoeology (blocks F-C) with A5 and B5. The terminal J block present in A5 and B5 was however absent in C5. In Group 6 (A6/B6/C5–C6), A6 and B6 were homoeologous along their entire length sharing the block arrangement C-B-A-V-K-L-Q. One inversion in either of the two genomes could explain the reverse orientation of the blocks constituting the lower segment of A6–B6 (blocks V-K-L-Q). Two C genome LGs (C5 and C6) are also components of this group. C5 shared the A-B-C block arrangement with A6/B6 while C6 was homoeologous for the block arrangement V-K-L-Q with A6/B6. Members of Group 7 (A7/B7/C7) shared homoeology for a large E block while Group 8 (A8/B7/C8) members were homoeologous for blocks A-B-U. Group 9 (A9/C8–C9) had A9 sharing homoeology with C9 for the blocks O-Q-X-H-D-V which constituted the top half of both the linkage groups. The lower segment of A9 shared the block arrangement N-I-H-A with C8. No LG from the B genome seemed to possess any major block homoeologous with A9. Members of group 10 (A10/B8/C9) shared the blocks R-W which constituted a major portion of the linkage group in all the three genomes.

Figure 8

Major rearrangements between the rapa/oleracea and nigra genomes (a, b) and the rapa and oleracea genomes (c, d). Boxes bearing the same color represent homoeologous blocks while the hatched boxes represent large gaps (≥10 cM regions devoid of...

Karyotype changes that led to the divergence of A, B and C genomes of Brassica species

The identification of homoeologous chromosomes among the three genomes of Brassica sp. (A, B and C) in the present study allows us to predict the possible, macro-level karyotype changes that led to the divergence of the A, B and C genomes. It has been predicted that the Nigra (B) and the Rapa/Oleracea (A/C) lineages separated from each other about 7.9 Mya [32] followed by the splitting of the Rapa and Oleracea lineages. Three linkage groups of the B genome (B4, B5 and B6; Figure ​7) have retained similar block organization as their corresponding A genome LGs (A4, A5 and A6; Figure ​7). For the remaining five B genome chromosomes (B1, B2, B3, B7 and B8), we propose two types of changes: (1) Rearrangements without any change in the chromosome number: blocks constituting four LGs of the A/C genome (A1–C1/A2–C2/A3–C3/A10) could be reshuffled to explain the block arrangement in four LGs in the B genome (B1/B2/B3/B8) (Figure ​8a). No reduction in chromosome number seems to have taken place here. In the process however, both the F-T-U block motifs of the A/C genome lost their identity in the B genome while a unique LG with two F blocks emerged in the B genome. (2) Rearrangements with variations in chromosome number: the genomic block arrangement of B7 could be derived by fusing two linkage groups from the A/C (A7–C7/A8–C8) genome (Figure ​8b). This also explains the difference of one LG between the A/C and B genomes.

A high level of similarity between the A and C genomes was established in earlier studies [16]. However, it was not clear whether the changes that were observed between the two occurred in one of the lineages after their divergence or occurred independently in both the lineages. The similar organization of A1–C1, A2–C2, A3–C3 and A7–C7 LGs of the A and C genomes (Figure ​7) indicated the absence of any structural changes in these chromosomes after the divergence of the A and C genomes. We therefore predict the following types of changes in the evolution of the A and C genomes: (1) Rearrangements related to the C4, C5 and C6 chromosomes are specific to the C genome (Figure ​8c) and occurred after the diversification of the A and C genomes since the corresponding LGs of the A and B genomes are identical (Figure ​7). These changes, mainly translocations, break the homoeology between the A and C genomes (Figure ​8c). The remaining two LGs (C8 and C9) appear to be made up of rearranged blocks constituting three LGs of the rapa genome (A8, A9 and A10; Figure ​8d). The block arrangement of C9 can be achieved by fusing half of A9 with almost the entire A10 LG. Similarly, the other half of A9 and the entire A8 represent the block arrangement of the C8 chromosome (Figure ​8d). These rearrangements would have contributed to the difference of one chromosome between the A and C genomes.


In this study, we generated a detailed comparative map between B. juncea and Arabidopsis thaliana containing 533 At loci including 486 IP markers. Although exons of At and Brassica species share around 75–90% homology [33], tapping polymorphism available in the intronic sequences is an efficient method to generate PCR-based markers from otherwise highly conserved genes. Thirty two percent of the primers designed in this study were found to be polymorphic between the B. juncea lines, Varuna and Heera, used as parents of the mapping population. Screening the amplified fragments for SNPs could lead to a further increase in the number of polymorphic loci. Previous reports on comparative analysis between At and Brassica species have been based mostly on RFLP probes [16,18-20,24]. Screening large segregating populations with RFLP markers is rather cumbersome. In contrast, the polymorphisms obtained using IP markers were easily discernable on simple 1.5–2.0% agarose gels and would therefore enable rapid screening of large segregating populations. Additionally, being genic in origin, IP marker-based genetic maps would directly reflect the syntenic relationship between the two species, B. juncea and At. Since the IP markers used in this study were designed from conserved sequences between At exons and available Brassica EST/GSS sequences, they also possess enormous potential for wider applicability across various Brassica species. As the At genome is partially duplicated [34], the use of multicopy At loci to establish syntenic relationships in other species would raise issues of paralogy and orthology. In contrast, analyzing the distribution of At singletons in Brassica would give an unambiguous representation of various rearrangements that have occurred between the two lineages since their divergence. The major emphasis in this study was therefore on generating IP PCR primers predominantly from single copy loci in Arabidopsis. Our genomic block definitions were based mostly on At singletons in contrast to earlier studies [16]. Irrespective of the approach used to define synteny, the number of times each block is represented in the A genome of B. juncea was found to be highly similar to that observed for B. napus. This observation supports the earlier hypothesis which advocated the occurrence of duplications in the Arabidopsis genome prior to its divergence from the Brassica lineage [16].

In the present study, we analyzed the segmental organization of the B. juncea genome based on the 24 conserved genomic blocks described by Schranz et al. [15]. These blocks represent the conserved regions common to the AK, At and B. rapa [15]. Block order obtained for B. juncea in this study was compared with the available maps for the A and C constituent genomes of B. napus [16] and the B genome of B. nigra [18]. A high level of conserved macro-level colinearity was observed between B. juncea and its diploid progenitors. The rapa (A) genomes of both B. juncea and B. napus were found to be highly comparable (Figure ​6a). Similarly, the B genome of B. juncea appeared to maintain similar genomic block architecture as its diploid counterpart in B. nigra (Figure ​6b). This signified the absence of large scale perturbations during the formation of the allopolyploid Brassicas contrary to earlier reports by many groups working on synthetic Brassica polyploids [35-37]. Additionally, the conserved identity of both the constituent diploid genomes in the Brassica polyploids suggests the involvement of a strong and active gene action which inhibits pairing between the homoeologous chromosomes. Thus, survival of the allopolypoids B. napus, B. juncea and B. carinata, in all probability, was based on hybrid vigour and suppression of pairing of the homoeologous chromosomes.

Defining the constitution of the A, B and C Brassica genomes based on these genomic blocks facilitated the reconstruction of the ancestral karyotype to decipher the evolution and diversification of the Brassica lineage. Common block arrangements shared between the three lineages suggest an ancestral origin. Several such conserved block motifs were identified in this study. These include blocks R-W-E-O-P (Group 2); blocks R-W-J-I-O-P (Group 3) and blocks J-I-S-T-N (Group 4) (Figure ​7). Since the two lineages within the tribe Brassiceae (Nigra lineage and Rapa/Oleracea lineage) diverged approximately ~7.9 Mya [32], the block motifs shared by the A and B genomes would also be ancestral. Thus, the block motifs shared between the members of Group 5 (blocks J-C-F) and Group 6 (blocks C-B-A-V-K-L-Q) (Figure ​7) are speculated to be ancestral in origin although their identity in the Oleracea lineage is lost. Based on the comparative genome analysis of the three diploid Brassica species carried out in this study the constitution of the ancestral Brassica karyotype (ABK) in terms of the block arrangement could be predicted for at least five LGs (ABK2–ABK6; see Additional File 5). Earlier studies on chloroplast DNA analysis [38,39] clearly established that the B. rapa/B. oleracea are very closely placed in one lineage and B. nigra belongs to a different lineage. Our results suggest significant similarity between the A and B genome for five linkage groups. We therefore propose that besides the five LG putative ancestor, there were divergent parents (female) involved in the evolution of the two major Brassica lineages.

Comparative analysis of block organization revealed certain major rearrangements (translocations and fusions) that were pivotal to karyotype diversification in the different Brassica lineages. Complete homoeology in terms of block organization was seen for three linkage groups each between the rapa (A) and oleracea (C) genomes (A1–C1, A2–C2 and A3–C3; Figure ​7) and the rapa (A) and nigra (B) genomes (A4–B4, A5–B5 and A6–B6; Figure ​7). This implies that although the Rapa and Oleracea lineages are more closely related, certain block arrangements have been retained by the Rapa and Nigra lineages while being lost from the C genome. If the rearrangements in the block organization of the remaining LGs are compared, rearrangements leading to lineage-specific karyotype diversification become evident. Our study also highlights certain rearrangements (Figure ​8c) that appear to have originated in the Oleracea lineage after its divergence from Rapa. Since the chromosome number of the common Brassica ancestor remains uncertain, one cannot discern whether the splitting of chromosomes led to the successive increase in chromosome number in the three Brassica lineages or fusions led to a reduction in the chromosome number. However, in the present study, the rearrangements leading to the difference of one LG each between the A-B and A-C lineages could be deciphered (Figure 8b, 8d).

Previous studies suggested that two reciprocal translocations, three chromosome fusions and at least three inversions were instrumental in the derivation of At from the ancestral karyotype-AK [40,41]. In the process, several new block boundaries were created. We compared the genomic block organization of AK, At and the three Brassica genomes to identify new/common block junctions to gain further insight into the evolution of the crucifer genomes. In the three Brassica genomes analyzed, block R was found to be almost always associated with block W (Figure ​7) and block Q was associated with block X. These associations are characteristic of the Brassica genome and are not found in AK or At [15]. The rapa/oleracea A3/C3 LGs shared the ancestral pattern of blocks F-G-H (AK chromosome 3) [15] while in At, F and G-H occupy different chromosomes. Similarly, with the extent of saturation available for the three genetic maps of Brassica, the C-D, K-G and Q-S block fusions characteristic of At are lacking in Brassica. This suggests that different rearrangements led to the formation of the Arabidopsis and the Brassica ancestor. A more detailed view of karyotype evolution would be available once all the centromeres for the Brassica genomes are identified.

The identification of homoeology based on the conserved 24 genomic blocks would help build a unified comparative genomics system in the Brassicaceae as has been done for the Gramineae family by the generation of the 'Crop-Circle' [42-44]. This will facilitate the transfer of information from one species to another for which IP markers would be most useful. With the elucidation of genes involved in many biochemical pathways and the availability of gene expression data, information generated in the model species Arabidopsis holds enormous potential for application in breeding of Brassica crops. Establishment of syntenic relationships between At and Brassica would be highly beneficial for the identification of candidate genes contributing to traits of agronomic value from corresponding regions in At and also serve as an exhaustive resource to generate more markers for fine mapping in syntenic regions of Brassicas.


The present study conclusively establishes the efficacy of IP markers for the development of a comparative map between a model plant and a polyploid crop species. The comparative genome analysis performed in this study has contributed significantly to our understanding of the homoeology shared between the A, B and C Brassica genomes. The study also identifies, at the macro-level, the evolutionary rearrangements leading to karyotype diversification among the three genomes. Additionally, some of the putative ancestral Brassica-specific block motifs were identified. Syntenic relationships thus established between B. juncea, B. napus, B. nigra and At will facilitate precision breeding and identification and positional cloning of candidate genes contributing to traits of agronomic value in Brassica species.


A total of 1180 primer pairs (247 from At C1, 193 from At C2, 214 from At C3, 292 from At C4 and 234 from At C5) were designed. Genes having larger introns were preferred as the incidence of indels is expected to be higher in such cases. The expected amplicon size for the designed primer pairs was approx. 500 bp-1 kb in A. thaliana. To reduce non-specific amplifications in B. juncea, primer pairs designed from At singletons were first tested on At DNA to optimize conditions for PCR which result in a single amplicon. For most of the primer pairs screened on At DNA, the optimum PCR condition was: initial denaturation at 94°C for 5 min, followed by 30 cycles of denaturation at 94°C for 30 s, annealing at 55°C for 30 s and elongation at 72°C for 1 min followed by a final extension at 72°C for 5 min. Amplified PCR products were analyzed on 1.2–2% w/v agarose gel to score for polymorphism either on the basis of size/length difference or presence/absence (dominant) of PCR products. A mapping population consisting of 123 doubled haploid lines derived from a cross between Varuna (an Indian cultivar) and Heera (an east European line) was used for the construction of the linkage map [30]. IP markers were added to a framework map of B. juncea described earlier by Pradhan et al. [30] and Ramchairy et al. [11] using the program Joinmap version 2.0 [45,46].


At: Arabidopsis thaliana; At C1–C5: Chromosomes of At (1–5); AK: Ancestral karyotype; cM: centiMorgan; IP: Intron polymorphism; LG: Linkage group.

Authors' contributions

PP, AJ, NCB, LP and SS designed intron spanning primers from one each of the five At chromosomes and mapped the IP markers on the F1DH population. PP wrote the paper and put together the figures. The comparative analysis of the A, B and C genomes of Brassica were done by PP. VG contributed to some of the mapping experiments and organized the laboratory work. AKP looked at all the mapping data and helped in organizing the mapping information and the manuscript. DP conceived the project and contributed to the writing of the manuscript. All authors read and approved the final manuscript.

Supplementary Material

Additional file 1:

Sequence data of the intron spanning primers used in the present study. This file contains the sequence information of all the intron spanning primers which generated polymorphic IP markers deployed to develop the B. juncea genetic map in the present study.

Click here for file(92K, xls)

Additional file 2:

Comparative genome organization of the A genome of B. juncea (A1–A10; present study) and B. napus (N1–N10) [16]. This file contains the map of the A genome of B. napus [16] with the RFLP loci converted to their corresponding At (A. thaliana) loci and a detailed comparison (in terms of the At loci arrangement) of this map with the A genome of B. juncea (present study).

Click here for file(652K, ppt)

Additional file 3:

Comparative genome organization of the B genome of B. juncea (B1–B8; present study) and B. nigra (G1–G8) [18]. This file contains the map of the B genome of B. nigra [18] with the RFLP loci converted to their corresponding At (A. thaliana) loci and a detailed comparison (in terms of the At loci arrangement) of this map with the B genome of B. juncea (present study).

Click here for file(481K, ppt)

Additional file 4:

The C genome map of B. napus [16] with the RFLP probes converted to their corresponding At loci.

Click here for file(350K, ppt)

Additional file 5:

Putative ancestral Brassica karyotype (ABK2–ABK6). The file contains the figurative representation of the putative ancestral Brassica karyotype (ABK2–ABK6) predicted in our study. This is based on the conserved group structure of groups 2, 3, 4, 5 and 6 (Figure ​7). Only partial organization of the putative linkage groups ABK2 and ABK3 is shown representing the common blocks shared between the three diploid Brassica genomes.

Click here for file(29K, ppt)


The work was financially supported by Dhara Vegetable Oil and Food Company Ltd (DOFCO), a subsidiary of the National Dairy Development Board (NDDB), India. Partial support came from the UGC-SAP programme. SS was funded by a fellowship from the Department of Science and Technology (DST) under the SERC Fast Track Scheme for Young Scientists. LP was supported by a research fellowship from the Council of Scientific and Industrial Research (CSIR). Arundhati Mukhopadhyay, N Arumugam and Y S Sodhi contributed to the development of the mapping population used in the study and Jagdish Verma maintained it in the field.


  • Johnston JS, Pepper AE, Hall AE, Chen ZJ, Hodnett G, Drabek J, Lopez R, Price HJ. Evolution of genome size in Brassicaceae. Ann Bot. 2005;95:229–235. doi: 10.1093/aob/mci016.[PMC free article][PubMed]

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