The Biology of Brassica napus L. (Canola/Rapeseed)

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Biology Document BIO2017-03: A companion document to Directive 94-08 (Dir94-08), Assessment Criteria for Determining Environmental Safety of Plant with Novel Traits

Plant and Biotechnology Risk Assessment Unit
Plant Health Science Division, Canadian Food Inspection Agency
Ottawa, Ontario

Table of contents

1 General Administrative Information

1.1 Background

The Canadian Food Inspection Agency's Plant and Biotechnology Risk Assessment (PBRA) Unit is responsible for assessing the potential risk to the environment from the release of plants with novel traits (PNTs) into the Canadian environmentFootnote 1. The PBRA Unit is also responsible for assessing the pest potential of plant imports and plant species new to Canada.

Risk assessments conducted by the PBRA Unit require biological information about the plant species being assessed. Therefore, these assessments can be done in conjunction with species-specific biology documents that provide the necessary biological information. When a PNT is assessed, these biology documents serve as companion documents to Dir94-08: Assessment Criteria for Determining Environmental Safety of Plants with Novel Traits.

1.2 Scope

This document is intended to provide background information on the biology of Brassica napus, its identity, geographical distribution, reproductive biology, related species, the potential for gene introgression from B. napus into relatives and details of the life forms with which it interacts.

Such information will be used during risk assessments conducted by the PBRA Unit. Specifically, it may be used to characterize the potential risk from the release of new varieties/genotypes of B. napus that qualify as PNTs into the Canadian environment with regard to weediness/invasiveness, gene flow, plant pest properties, impacts on other organisms and impact on biodiversity.

2 Identity

2.1 Name(s)

Brassica napus L. (USDA-ARS 2017)

2.2 Family

Brassicaceae (alt. Cruciferae) (mustard family) (USDA-ARS 2017)

2.3 Synonym(s)

Brassica napus includes a number of recognized forms, subspecies and varieties: Brassica napus L. subsp. napus, Brassica napus L. subsp. napus forma annua (Schübl. & G. Martens) Thell., Brassica napus L. subsp. napus forma napus, Brassica napus subsp. rapifera Metzg. and Brassica napus L. var. pabularia (DC.) Alef. (USDA-ARS 2017).

Synonyms for Brassica napus and its forms, subspecies and varieties include: Brassica campestris subsp. napobrassica (L.) Schübl. & G. Martens, Brassica campestris subsp. napus (L.) Hook. f. & T. Anderson, Brassica campestris [unranked] biennis Schübl. & G. Martens, Brassica campestris[unranked] pabularia DC., Brassica napobrassica (L.) Mill., Brassica napus f. biennis (Schübl. & G. Martens) Thell., Brassica napus subsp. napobrassica (L.) Jafri, Brassica napus subsp. oleifera (Delile) Sinskaya, Brassica napus subsp. rapifera Metzg. ex Sinskaya, Brassica napus var. annua W. D. J. Koch, Brassica napus var. biennis (Schübl. & G. Martens) Rchb., Brassica napus var. napobrassica (L.) Döll, Brassica napus var. oleifera Delile, Brassica napus var. sahariensis A. Chev. and Brassica oleracea var. napobrassica L. (USDA-ARS 2017).

2.4 Common name(s)

Brassica napus and its forms, subspecies and varieties are commonly known in English as annual rape, Argentine canola, canola, colza, Hanover-salad, oilseed rape, rape, rapeseed, rape kale, rutabaga, Siberian kale, summer rape, swede, Swede rape, Swedish turnip and winter rape (Gulden et al. 2008; USDA-ARS 2017).

Unlike many other crops, the products called "rapeseed" and "canola" do not come from a single species and they do not necessarily refer to the same variety. The term "rapeseed" refers to oilseeds from the species B. napus and B. rapa, while the term "canola" refers to specific varieties of rapeseed bred to produce edible material for human and animal consumption. These edible varieties must contain less than 2% erucic acid and 30 µmol glucosinolates per gram of air-dried oil-free meal. Recent canola varieties also include the related species B. juncea (L.) Czern. (Canola Council of Canada 2014c; Canola Council of Canada 2014d).

2.5 Taxonomy and genetics

Brassica napus is a member of the Brassicaceae family, which consists of approximately 25 tribes, 338 genera and 3709 species (OECD 2012). It is included within the Brassiceae tribe, which consists of 9 subtribes, 48 genera and 240 species (OECD 2012).

The Brassicaceae family includes many well-known plants, such as the model plant Arabidopsis thaliana (mouse-ear cress), the weedy relative Sinapis arvensis (wild mustard) and vegetable crops such as B. napus (rutabaga, Siberian kale), B. rapa (Chinese cabbage, pai-tsai, mizuna, Chinese mustard, broccoli raab and turnip), B. oleraceae (cabbage, broccoli, cauliflower, Brussels sprouts, Kohlrabi, collards, kale) and Raphanus sativus (radish). Examples of condiment crops of the Brassicaceae family include B. nigra (black mustard), B. carinata (Ethiopian mustard), B. juncea (brown or Indian mustard), Armoracea rusticana (horseradish) and a number of other minor pot herbs and salad vegetables (Downey and Rimmer 1993; OECD 2012; Rakow 2004).

B. napus is an amphidiploid that resulted from the interspecific hybridization between B. oleracea L. and B. rapa (Downey and Rimmer 1993; OECD 2012). This cytogenetic relationship was first proposed in 1935 as the U triangle (Nagaharu 1935). The triangle depicts the three monogenomic diploids B. nigra (B genome, n=8), B. oleraceae (C genome, n=9) and B. rapa (A genome, n=10) and the three digenomic species B. carinata (BC genome, n=17), B. juncea (AB genome, n=18) and B. napus (AC genome, n=19). This cytogenetic relationship is believed to have evolved naturally, without cultivation (Downey and Rimmer 1993; Gulden et al. 2008; Kays and Dias 1995; OECD 2012; Vaughan 1977). Because tetraploid Brassica species share a genome with their diploid parents, gene flow can continue in both directions. For more information on this topic, refer to Section 5: Related Species of Brassica napus of this document.

Taxonomic position (USDA-NRCS 2014):

Kingdom:
Plantae (plants)
Subkingdom:
racheobionta (vascular plants)
Superdivision:
Spermatophyta (seed plants)
Division:
Magnoliophyta (flowering plants)
Class:
Magnoliopsida (dicotyledons)
Subclass:
Dilleniidae
Order:
Capparales
Family:
Brassicaceae (mustard family)
Tribe:
Brassiceae
Genus:
Brassica L. (mustard)
Species:
Brassica napus L. (canola/rapeseed)

2.6 General description

Brassica napus is an annual or biennial species (Gulden et al. 2008). The stems are erect, simple to freely branched, glabrous or sparsely hairy and can grow up to 1.5 m tall. Leaves are waxy with a glabrous underside and often have an enlarged base that partially clasps the stem. Inflorescences are racemes which form on the main and axillary branches. Flowering begins at the base of the raceme and the buds form above the open flowers. The pale yellow flowers have 4 sepals and 4 diagonally opposite obovate petals arranged in the form of a cross when viewed from above. The stamens are tetradynamous with four long and two short stamens in each flower. The ovary is superior (Callihan et al. 2000; Gulden et al. 2008; OECD 2012).

The fruit is a linear cylindrical silique with slight constrictions at regular intervals and dehiscent valves in the lower 4–10 cm segment of the fruit. The seeds are arranged in a single row in the fruit. The upper 3.5–5.0 mm thick segment of the silique is narrow and is usually seedless (Callihan et al. 2000; Gulden et al. 2008; OECD 2012).

There may be 15 or more seeds per silique. Seed formation begins in the lower one-third branches of the main stem. Seeds are translucent and turn green when full sized. At maturity, seeds are spherical, reddish brown, brown or black, 1.8–2.7 mm in diameter with a finely net-veined surface (Gulden et al. 2008).

The cotyledons are conduplicate (i.e. folded longitudinally around the radicle in the seed). During germination, seedlings emerge with the growing point situated between the cotyledons. At the seedling stage, the cotyledons are 6–12 mm wide, kidney-shaped and have a deep, wide rounded notch at their tip (Gulden et al. 2008).

The seedlings grow to form a rosette of 5–6 leaves with the older leaves at the base and the smaller younger leaves in the centre of the rosette. The basal rosette leaves are short-petiolate, toothed, glaucous, ovate to elongate and entire to lobed with 1 to 5 pairs of small lateral lobes and a large terminal lobe. Bolting, stem extension, branching and upper leaf formation is the next stage of plant growth (Gulden et al. 2008).

Differentiation between Brassica napus and close relatives can be difficult as they may share similar morphological features (OECD 2012). Despite this, there are differences between B. napus and B. rapa. The leaves of B. napus are hairless, smooth, fleshy and bluish-green in colour while those of B. rapa are yellowish-green. The base of B. napus leaves partially clasp the stem while those of B. rapa fully clasp. In B. napus, the buds are borne above the open florets while in B. rapa the buds are borne below the open florets in the raceme. The flowers of B. rapa are smaller and darker yellow compared to the flowers of B. napus (Gulden et al. 2008).

3 Geographical Distribution

3.1 Origin and history of introduction

The centre of origin for Brassica napus is uncertain; some scholars believe it originated in Mediterranean Europe while others support multiple centres of origin (OECD 2012; Rakow 2004).

Rapeseed (including B. napus) has been cultivated in Asia, Europe and Northwestern Africa since ancient times as a source of oil for food, lamps, soap and later for industrial purposes. In Canada, rapeseed has been commercially grown in the western provinces since the middle of the 20th century, serving as a supply of lubricant for steam engines. Cultivation of rapeseed in Canada increased dramatically during World War II, to secure a steady supply of industrial lubricant. Following the end of the war, rapeseed production declined as there was reduced demand for industrial lubricants (Canola Council of Canada 2014c; Rakow 2000).

The war also disrupted the Canadian supply of edible oil and as a result Canadian plant breeders began to focus on developing edible oil alternatives. By the mid-1950s, rapeseed oil was fully recognized in the Western hemisphere as edible oil and breeding programs began to focus on the development of low-erucic acid and low-glucosinolate varieties for human and animal consumption. In 1974, the first agronomically viable low-erucic acid/low-glucosinolate variety was released in Canada and in 1978, the term "canola," derived from "Canadian oil," was adopted to identify these varieties (Canaola Council of Canada 2011; Daun 1993b). Originally a trademark, "canola" is now a generic term for edible varieties of rapeseed (Canola Council of Canada 2011). In Canada, the term canola is officially defined as: "An oil that must contain less than 2% erucic acid and less than 30 µmol of glucosinolates per gram of air-dried oil-free meal" (Canola Council of Canada 2014c; GOC 2014; Rakow 2000).

Canola is now ranked amongst the top oilseed crops in the world. Canola oil is also the most widely used edible oil in Canada's domestic market. Canola meal, a by-product of the seed oil extraction crushing process, has also found widespread acceptance as a livestock, poultry, swine and fish feed (Daun 1993b; Eskin 2013). Rapeseed is still cultivated to a lesser degree in North America for industrial purposes (e.g. high-quality lubricants, hydraulic fluid, plastics, etc.) and for use in food processing (e.g. candy bars, or as an emulsifier in peanut butter, etc.) (Canadian Grain Commission 2014; Canola Council of Canada 2014a; Canola Council of Canada 2014c; Canola Council of Canada 2014d).

3.2 Native range

The native range of Brassica napus is believed by some to be the coastal Mediterranean and European Atlantic region (Tsunoda 1980); however, an alternate view considers that the species is native to a wide range of countries due to multiple centres of origin (Downey and Rimmer 1993; OECD 2012; Rakow 2004).

3.3 Introduced range

Asia:
Brassica napus is cultivated in China, Japan, Kazakhstan, Afghanistan, Iran, India and Pakistan. It has become naturalized in China, Japan and Afghanistan (USDA-ARS 2017; USDA-ERS 2014).
Africa:
B. napus is cultivated in Ethiopia, Kenya, Tanzania, Mali and Zimbabwe (USDA-ARS 2017).
Australia:
B. napus is cultivated and naturalized widely in medium and high rainfall areas of southern Australia (Australian Government 2008; USDA-ARS 2017; USDA-ERS 2014).
Canada:
B. napus was introduced into Canada from Eurasia and is considered naturalized. It is present in all provinces of Canada, and the Northwest Territories. In Canada the largest acreage for canola production is Saskatchewan followed by Alberta, Manitoba, British Columbia, Ontario, New Brunswick, Prince Edward Island and Nova Scotia (Brouillet et al. 2010; Canola Council of Canada 2014b; Gulden et al. 2008; USDA-ARS 2017; USDA-ERS 2014; VASCAN 2014).
Europe:
B. napus has is cultivated and considered naturalized throughout Europe. It is present in agricultural, disturbed and urban habitats in Austria, Denmark, Estonia and Lithuania. In Britain, B. napus was first recorded in the wild in 1660 and is considered established in disturbed areas, roadsides, waste and cultivated ground. B. napus is an established species in Denmark, Netherlands and Norway and has also been reported in Germany. However, the species is not considered to be invasive in most of these European countries (NOBANIS 2014; USDA-ARS 2017; USDA-ERS 2014).
Mexico:
B. napus is cultivated and has become naturalized in Mexico (USDA-ARS 2017; USDA-ERS 2014).
New Zealand:
B. napus is cultivated and has become naturalized in New Zealand (USDA-ARS 2017; USDA-ERS 2014).
Russia:
B. napus is cultivated in eastern and western Siberia and in the far eastern part of the Russian Federation (USDA-ARS 2017; USDA-ERS 2014).
United States:
B. napus is cultivated and has become naturalized in the United States. It is present in several of the contiguous States of the country; Idaho, Minnesota, Montana, North Dakota, Oklahoma, Oregon, Washington, Colorado and Kansas (USDA-ARS 2017; USDA-ERS 2014; USDA-NRCS 2014).
Central & South America:
The species has become naturalized in Central America, Ecuador - Galapagos Islands, Argentina and Chile (USDA-ARS 2017).

3.4 Potential range in North America

Brassica napus is estimated to be hardy to USDA Plant Hardiness Zone 1 so long as there are sufficient warm days during the growing season (Daily et al. 2012; Margarey et al. 2008; Canola Council of Canada, 2017). This potential range captures every Canadian province and territory.

3.5 Habitat

In Canada, Brassica napus is grown in ecoregions characterized by mean summer temperatures that range from 13.0°C to 16.0°C, mean winter temperatures that range from -14.5°C to -8.0°C and mean annual precipitation of up to 700 mm (e.g. Manitoba, Saskatchewan and Alberta). When calculated at a base temperature of 0°C, the species requires an average of 1560 growing degree-days to mature (Gulden et al. 2008).

The cultivation of B. napus is not suitable in soils with pH less than 5.5 or more than 8.3, in waterlogged soils, or in soils with electrical conductivity values greater than 6 dS per meter. The optimal temperature for B. napus germination is 20°C. Unfavourable environmental conditions (e.g. temperature fluctuations, low soil moisture, prolonged darkness) may induce secondary dormancy in seeds (Gulden 2003; Gulden et al. 2003b, 2008; Johnson et al. 2004).

4 Biology

4.1 Reproductive biology

Brassica napus reproduces by seed and does not exhibit vegetative reproduction under field conditions (Andersson et al. 2010). It is reported to be predominantly self-fertile, but also shows a considerable degree of receptiveness to pollen from other varieties and a tendency to outcross (Hüsken and Dietz-Pfeilstetter 2007; Légère 2005). Under field conditions, cross-pollination occurs mainly through physical contact with neighbouring plants, but pollen is also transferred over longer distances by wind and insects (Eastham and Sweet 2002; Hüsken and Dietz-Pfeilstetter 2007).

In terms of reproductive structures, flowers are produced in racemes, with one raceme developing at the very top of the plant, followed by axillary racemes (Lamb 1989). Flowering in Canada occurs between 45 and 70 days after seeding and lasts 14–21 days (Canola Council of Canada 2014d). Based on typical seeding in Canada, flowering normally occurs during the months of June and July (Canola Council of Canada 2014d).

Petals drop after fertilization and a cylinder-shaped silique is formed (Downey 1983). Upon maturation, this pod contains between 15 and 40 seeds. Seed pods take 35–45 days to fill (Canola Council of Canada 2014d). Of Brassica oilseed crops, B. napus is most susceptible to silique shattering (Downey 1983). The seed is predominantly embryo tissue and contains less endosperm than cereal grains. The seed coat can be black or yellow and consists of an outer epidermis layer. Leaf senescence begins as the pods mature, starting with the leaves at the base of the plant (Canola Council of Canada 2014d).

4.2 Breeding and seed production

Agronomic Performance Traits

Canadian breeding programs for improvement of Brassica napus started during the 1940s, with the first varieties registered in 1954 (McInnis 2004). Agronomic traits of this first variety included uniform early maturity, high seed yield and high oil content.

In 1991, a high-yielding B. napus variety was developed through a conventional cross between yellow-seeded and black-seeded B. napus lines (Rakow et al. 1999). Although yellow-seeded varieties were typically lower-yielding, they were also associated with improved meal quality and therefore a high-yielding yellow-seeded canola variety became a breeding priority (Rakow et al. 1999).

In general, B. napus is less cold-tolerant than B. rapa, one of the original crossing parents of B. napus (Hayward 2012).

Seed Quality Traits

Throughout the 1960s, low erucic acid and low glucosinolate traits became high priorities for the Canadian breeding program, to produce edible oil suitable for human and animal consumption. The low erucic acid and low glucosinolate traits were introgressed from a European Brassica napus forage rape variety and a Polish B. napus rapeseed variety, respectively (McInnis 2004).

Other names for B. napus oilseeds with a comparable nutritional profile include Double Zero (00) rapeseed and LEAR (Low Erucic Acid Rapeseed) oil (Health Canada 2003).

Herbicide Tolerance Traits

The first genetically engineered canola variety was introduced in 1995 and possessed tolerance to glufosinate-ammonium (phosphinothricin) based herbicides. Since then, herbicide tolerance (HT) traits have been developed for four active ingredients (i.e. glyphosate, glufosinate-ammonium, bromoxynil, imidazoline).

Tolerance to glyphosate, glufosinate-ammonium and bromoxynil was obtained by introducing genes from other organisms into the B. napus genome, through recombinant DNA technologies, whereas tolerance to imidazoline was obtained by mutagenesis (Johnson et al. 2004; Simard et al. 2002). In 2010, 47% of canola acreage in Canada was glyphosatetolerant, 46% was glufosinate-ammonium tolerant and 6% was imidazolinonetolerant (Canola Council of Canada 2014b).

Disease Resistance Traits

Varieties with disease resistance have been available in Canada since the late 1990s, with most early work focussed on resistance to blackleg (Leptosphaeria maculans (Desmaz.) Ces. et De Not.) (Kutcher et al. 2013a). Clubroot (Plasmodiophora brassicae Woronin) has been documented throughout Western Canada since 2003 (Strelkov and Hwang 2014). B. napus varieties with resistance to clubroot are widely used in commercial production in Canada, although the clubroot pathogen has developed resistance against some varieties exhibiting a single resistance trait locus/gene (Strelkov et al. 2016).

Seed Production and Varietal Registration

The Canadian Seed Growers Association has developed varietal purity standards for pedigree seed production of Foundation, Registered and Certified seed (Canadian Seed Growers Association 2005). Canola is an internationally regulated standard for oilseed rape, which was trademarked in 1978 by the Western Canadian Oilseed Crushers' Association (McInnes 2004). The trademark was transferred to the Canola Council of Canada in 1980.

4.3 Cultivation and use as a crop

Seeding

Brassica napus may be fall or spring-seeded. In Western Canada, the short growing seasons and harsh winters only allow for the cultivation of spring genotypes (Gulden et al. 2003b; Gulden et al. 2004). B. napus should be seeded at 12–25 mm depth into a moist, firm seedbed (Canola Council of Canada 2014e). Broadcast seeding is generally not recommended, but is still commonly used where soil conditions prevent the use of seed drills (Canola Council of Canada 2014e). Row spacing at 23–30 cm was found to optimize yield (Kutcher et al. 2013b). Stand density should be 70–100 plants per meter square (Canola Council of Canada 2014e).

It should be noted that while B. napus seed may be marketed as greater than 90% germination rates, observed rates of germination in soil may be closer to 50% (Harker et al. 2003).

Like all Brassica crops, B. napus plants emerge, with a pair of cotyledons, 4–7 days after seeding (Lamb 1989).

Fertilization

To avoid significant seed damage and seedling mortality, fertilizer should be applied at seeding in a side band or mid-row band (Canola Council of Canada 2014e).

Safe rates of seed-placed nitrogen for canola are 11 kg per hectare in medium-textured soils. Applied nitrogen rates should be increased when openers spread seed and fertilizeover more of the seedbed and when seeding into heavy-textured (clay) soils (Canola Council of Canada 2014e). Polymer-coated urea and urease inhibitors, which slow the release of ammonia and ammonium from urea fertilizer, can increase seed safety and allow for higher rates of nitrogen to be placed with the seed.

Weeds

Common weeds in canola production in Western Canada include volunteer canola, cleavers (Galium spp.), Canada thistle (Cirsium arvense (L.) Scop.), sow thistle (Sonchus oleraceus L.), green foxtail (Setaria viridis (L.) P. Beauv.), quackgrass (Elymus repens (L.) Gould), wild oat (Avena fatua L.), volunteer wheat (Triticum aestivum L.) and volunteer barley (Hordeum vulgare L.) (Canola Council of Canada 2014e). In addition, weeds include Capsella bursa-pastoris (L.) Medik. (shepherd's-purse), Sinapsis arvensis (L.) DC. (wild mustard), Diplotaxis muralis L. (annual wall-rocket) and Sisymbrium officianale (L.) Scop. (hedge mustard) (Buczacki and Ockendon 1979).

Harvest

Brassica napus can be direct combined or swathed at harvest. Swathing involves cutting the crop at 50-60% seed colour change on the main stem and is more common than direct combining in Canadian canola production (Canola Council of Canada 2014f).

Uses of Brassica napus

Brassica napus is used to generate high quality vegetable oil for human consumption and as a high-protein meal to feed cattle, poultry, swine and fish (Canola Council of Canada 2014d). B. napus varieties are being developed as biofuel feedstock as well.

Canola oil is renowned for its nutritional and culinary qualities and is used in 80% of the salad oil market, 56% of the shortening market and 42% of the margarine market (Canola Council of Canada 2014d). Canola oil is also used in deep frying, baking, sandwich spreads, coffee whiteners and creamers. It is also used in cosmetics, printing inks, suntan oils, oiled fabrics, plasticizers, plastic wraps, pesticides and industrial lubricants.

Insect pests and diseases affecting cultivated B. napus are discussed in Section 6.

4.4 Gene flow during commercial seed and biomass production

Rates of outcrossing in Brassica napus, from neighbouring plants in the field or from pollen dispersed by wind or insects, have been estimated to occur between 12 to 55% with a mean outcrossing rate of 30% (Beckie et al. 2003; Damgaar and Kjellsson 2005; Eastham and Sweet 2002; Halfhill et al. 2004; Hüsken and Dietz-Pfeilstetter 2007; Légère 2005; Yoshimura et al. 2006). The level of outcrossing varies according to variety, local topography, environmental conditions and the availability of insect pollinators. Differences in experimental design may also result in variability of outcrossing rates reported among studies (Salisbury 2002b). Cleistogamy in B. napus can decrease intra-specific hybridization (Gruber et al. 2012).

Pollen dispersal from genetically modified B. napus has been measured up to 2000 m from source (Cai et al. 2008). Most pollen is dispersed within 4.5 m of the parent plant, with a maximum dispersal rate at 1.4 m. The dispersal rate declines sharply with distance from source between 0–33 m and then remains relatively constant from 33–2000 m, at approximately 0.015% (Rieger et al. 2002). Based on predictions from pollen dispersal modeling, wind is thought to be the main vector of long-distance gene flow in B. napus (Hoyle et al. 2007).

In Canada, gene flow between B. napus plants at 30 meter separation distances has been reported at rates of 0.03% (Staniland et al. 2000). In Australia, similar results have been observed (Rieger et al. 2002).

Herbicide tolerance trait stacking in volunteer B. napus can occur from intra-specific hybridization (Hall et al. 2000; Knispel et al. 2008). In Alberta, multiple-herbicide tolerance was observed in canola volunteers located in fields that had contained glufosinate-ammonium and imidazolinone tolerant varieties of canola and adjacent to where glyphosate tolerant varieties of canola were cultivated (Hall et al. 2000). In southern Manitoba, 88%, 81% and 31% of B. napus populations growing along the edges of fields and roadways showed glyphosate, glufosinate-ammonium and imidazolinone tolerance, respectively (Knispel et al. 2008).

Glyphosate drift outside of agricultural fields can occur from 5 to 400 m from spray boundaries, as measured by the USDA in Corvallis, Oregon, United States. The glyphosate rates resulting from the observed drift distances were significantly diminished (0.1–0.001x recommended field application concentration) when compared to recommended field rates (Londo et al. 2010). When low levels of glyphosate drift off of agricultural land on to surrounding fields, it may facilitate herbicide tolerant genotypes in volunteer populations (Londo et al. 2010).

4.5 Cultivated Brassica napus as a volunteer weed

Brassica napus is not listed as a noxious weed in the Weed Seeds Order, 2016. However, volunteer canola plants are considered a weed in managed ecosystems in Canada (Harker et al. 2006, 2015; Légère et al. 2001). Volunteers compete with crops for water, nutrients and sunlight, negatively impacting yields (Thomas 2003). A recent increase in the occurrence of volunteer canola in Western Canada may be due to increases in the area planted to the crop, as well as high levels of seed lost at harvest and the widespread adoption of reduced tillage practices in many regions (Gray et al. 1996; Lawson et al. 2006).

Feral populations of genetically engineered, herbicide-resistant B. napus have been reported in Canada (Beckie et al. 2003, Beckie and Owen, 2007; Beckie and Warwick 2010; Hall et al. 2005; Yoshimura et al. 2006), Great Britain (Crawley and Brown 1995), France (Pessel et al. 2001), Australia (Rieger et al. 2002), Japan (Aono et al. 2006; Kawata et al. 2009; Katsuta et al. 2015) and the United States (Schafer et al. 2011; Munier et al. 2012). In Switzerland, high densities of genetically modified canola plants were found growing near ports and along railway lines and roads (Schulze et al. 2015).

B. napus seed can persist in the soil, thereby contributing to the seed bank and volunteer populations in subsequent years. Volunteers compete with crops for water, nutrients and sunlight, negatively impacting yields and are considered a weed in the agricultural system (Thomas 2003).

Volunteer canola density and persistence are influenced by environmental factors and agronomic practices (Gulden et al. 2003b; Lutman et al. 2003; Pekrun et al. 1998; Légère et al. 2001; Simard et al. 2002; Harker et al. 2006). For example, sandy soils retain a larger proportion of dormant canola seeds than clay soil, because they contain less water causing the seed to degrade less readily (Pekrun et al. 1998). Furthermore, weather can have an impact, as canola seed pods shatter very easily, especially if the crop is harvested late in the season, is subject to heavy rainfall, strong winds, hail or lodging (Lutman et al. 2003; OGTR 2011a). Studies have reported viable canola seeds persisting for 4 to 5 years (Légère et al. 2001; Simard et al. 2002; Harker et al. 2006). In the process of harvesting spring canola, seed losses average 5.5% or about 2,000 to 3,600 seeds per meter square. This contributes to a considerable seedbank (Gulden et al. 2003a; Légère et al. 2001; Warwick et al. 2003b). Even if losses are reportedly low in terms of yield percentage, the small seed size of canola results in significant seedbank additions (Lawson et al. 2006).

The persistence and quantity of canola volunteers in subsequent crops is also influenced by seed dormancy (OECD 2012). Seed dormancy prevents an intact viable seed from germinating under favorable conditions (Bewley 1997). There are two types of seed dormancy termed primary and secondary (Hillhorst and Toorop 1997; OECD 2012). Primary dormancy is associated with the development of the seed (Hillhorst and Toorop 1997). It has been defined as a state that prevents seed germination during the seed maturation process and for a period of time after the seed has been removed from the parent plant (Bewley 1997; Hillhorst and Toorop 1997; OECD 2012). A post-ripening period is required in order to break the primary dormancy and enable the seed to germinate (Baskin and Baskin 2004; Bewley 1997).

Secondary dormancy is defined as a reduction in germinability that occurs after the seed is separated from the parent plant (Hillhorst and Toorop 1997; OECD 2012). B. napus seed is highly germinable when shed on the ground. As with most other crops, B. napus seed shows no signs of primary dormancy at the time of seed shed or at maturity (Gulden et al. 2004; Pekrun et al. 1997). Volunteer B. napus seeds are able to persist and stay viable in the soil seedbank for several years by developing secondary dormancy. Induction of secondary dormancy in B. napus is possible in response to sub-optimal germination conditions such as large temperature fluctuations, long exposure to darkness, osmotic stress and limited oxygen (Gulden et al. 2004; Pekrun et al. 1997, 1998). It has been suggested that persistence and the potential for secondary seed dormancy of B. napus in Western Canada can vary between genotypes of Canadian cultivars (Gulden et al. 2004; Harker et al. 2006; OGTR 2011a). Among a group of 16 Canadian genotypes, the contribution of genotype to secondary seed dormancy of B. napus ranged from 44 to 82% (Gulden et al. 2004). Secondary seed dormancy can be reversed by several factors, such as exposure to light, cold stratification, alternating temperatures and exogenous application of gibberellic acid (Baskin and Baskin 2004; Bewley 1997; Hillhorst and Toorop 1997). There are low dormancy genotypes available on the market as well as recommendations for minimizing volunteers on agricultural lands (Gruber et al. 2004b, 2010; Gulden 2003; Gulden et al. 2003b; Harker et al. 2006; Pekrun et al. 1997).

Conventional tillage incorporates seeds into the soil, potentially triggering secondary dormancy and increasing existing seedbanks (Gulden et al. 2003b; Harker et al. 2006; Pekrun et al. 1998). In contrast, seeds that remain on or near the soil surface in a zero tillage system have been reported to not exhibit secondary dormancy (Pekrun et al. 1997, 1998), although it has been suggested that in some cases secondary dormancy may occur in zero tillage systems as a result of large quantities of crop residues creating shade conditions (Légère et al. 2001; Simard et al. 2002). Still, other studies have suggested that tillage system does not influence the persistence of volunteer B. napus (Légère et al. 2001; Simard et al. 2002).

B. napus volunteers can emerge from seed deposited from improperly cleaned farm machinery, seed immigration from neighbouring fields, existing seedbanks and spills from transport vehicles (Pivard et al. 2008; Knispel and McLachlan 2010; Katsuta et al. 2015). On a year-to-year basis, the density of volunteers is highest in the first year after B. napus cultivation, with seedbanks declining up to 99% before the second year after cultivation (Gulden et al. 2008; Gulden 2003; Pekrun et al. 1998; Simard et al. 2002; Harker et al. 2006). In some cases in Western Canada, volunteer B. napus is the most prevalent weed species for 1–3 years following a B. napus crop (Harker et al. 2015).

With the commercialization and widespread adoption of herbicide-tolerant B. napus varieties over the past two decades, studies have evaluated conventional and herbicide- tolerant canola cultivars for their respective seed persistence in soil. There are no indications that the modifications conferring herbicide tolerance influence the number of seeds shed at harvest, seed dormancy, or seed persistence when comparing herbicide-tolerant and conventional counterparts (Crawley et al. 2001; Gruber et al. 2004b; Hails et al. 1997; Lutman et al. 2005) and secondary dormancy was similar in herbicide-tolerant and conventional varieties (Gruber et al. 2004b).

4.5.1 Cultural/mechanical control

Management of volunteer B. napus begins at the time of the preceding season's harvest with producers ensuring minimal harvest losses. This can be aided by properly calibrating combines and sealing leaks.

The seedbank persistence of volunteer B. napus is influenced by the vertical distribution of seed in the soil. Avoiding deep burial of seed will limit the persistence of B. napus seedbanks.

Cultural and mechanical control of volunteer B. napus has been described in great detail (Beckie et al. 2004). Recommendations on how to manage multiple-herbicide tolerant canola volunteers are as follows:

(1) Leave seeds on or near the soil surface as long as possible after harvest to reduce secondary dormancy.

(2) Use tillage or an effective herbicide burndown treatment immediately before seeding.

(3) Use silage and green manure to prevent seed set in volunteers.

(4) Isolate fields of B. napus with different herbicide tolerance traits to reduce trait stacking through intraspecific hybridization.

(5) Follow canola with a cereal crop and rotate canola in a 4-year diverse crop rotation.

(6) Scout fields for volunteers not controlled by weed management treatments and prevent seed set.

(7) Use pedigreed seed to reduce probability of the presence of off-types with different herbicide tolerance traits.

(8) Reduce seed loss during harvest by swathing at the correct crop development stage as well as properly adjusting combine settings.

Adoption of proper management practices and removal of escaped volunteer B. napus plants prior to flowering will help to prevent seedbank replenishment and long-term persistence of this weed.

4.5.2 Chemical control

Brassica napus varieties with tolerance to specific herbicides were first introduced in Canada in 1995. Four different herbicide tolerance traits have been marketed so far, including glyphosate (Roundup, Group 9), glufosinate-ammonium (Liberty, Group 10), imidazolinone (Clearfield, Group 2) and bromoxynil (Group 6) (Johnson et al. 2004; Simard et al. 2002). It should be noted that bromoxynil-tolerant canola is no longer commercially available (Johnson et al. 2004).

Herbicide-tolerant volunteer B. napus is becoming more common (Beckie 2015; Harker et al. 2015a, 2015b; Kumar and Jha 2015) with canola volunteers increasing from 16th to 4th most prevalent weed in Western Canada between 2003 and 2014 (Beckie 2015). Approaches in controlling volunteer canola typically involve deploying herbicides that the volunteers are susceptible to (Beckie et al. 2004).

Available data suggests that, in terms of chemical control measures, canola volunteers are best controlled at the two- to three-leaf stage and using group 4 herbicides, such as 2,4-D or MCPA, applied alone or in a tank mix in the first year following a canola crop (Beckie et al. 2004; Légère et al. 2006; Harker et al. 2006). To control canola volunteers that have single-HR, multiple-HR or stacked traits, available evidence suggests that group 4 and group 14 (carfentrazone, saflufenacil) herbicides are effective control strategies as pre-seeding (Beckie and Owen 2007). Chemical volunteer in-crop control strategies are relatively limited for canola (Beckie et al. 2004), though volunteers can also be controlled using herbicides in group 2 (florasulam, flucarbazone, tribenuron, thifensulfuron and pyroxsulam) and group 6 (bromoxynil and bentazon).

Recommendations for canola production include crop rotation cereals in a 3- to 4- year rotation (Rimmer et al. 2003; Cathcart et al. 2006). Short rotations (0- to 1-year break between canola crops) have become increasingly common in Western Canada because of the high commodity price and economic return of canola relative to cereals and pulses as well as the ability to use alternative herbicide modes of action for controlling Group 1- or 2-resistant weeds (Kutcher et al. 2013; Strelkov et al. 2011).

4.5.3 Integrated weed management

Integrated weed management (IWM) employs a combination of cultural, mechanical and chemical weed control approaches to manage weed populations, maximize crop yields and reduce reliance on single weed control techniques (Swanton and Weise 1991).

Steps for growers to control canola volunteers have previously been described at length, as follows (Thomas 2003):

(1) Rotate canola with cereal, pea and forage crops. Lengthening the rotation depletes the volunteers from the weed seed bank. Diversification permits the use of a wider selection of herbicides. Stacked volunteers that have up to three herbicide tolerance traits are susceptible to other common herbicides, including 2,4-D and MCPA and various new products. Group-6 Basagran can control all canola volunteers in peas, flax, beans and other broadleaf crops.

(2) Scout rotation fields for volunteer canola not controlled by herbicide application. Early detection allows time for control before seed set.

(3) Apply herbicides early to control canola pre-emergence. If volunteers germinate in a field planned for canola, carfentrazone (group 14), glyphosate (group 9) and amitrole (Group 11) are registered for pre-seed application before canola, providing alternatives to glyphosate alone. Spraying volunteer canola at the 2–4 leaf stage is more effective than at the 5–6 leaf stage.

(4) Keep detailed notes on herbicides and herbicide-tolerant systems used on each field.

(5) Leave canola seeds on the soil surface after harvest. Delay fall tillage until a few weeks after harvest. Tillage delay allows seeds to germinate through the fall and resulting plants to die over the winter, or to be eaten by birds and insects. Also, seed dormancy is reduced near the soil surface and exposure to winter elements. If B. napus seed is incorporated into the soil it may develop secondary dormancy and persist for up to four years.

(6) Reduce harvest losses. The fewer seeds lost, the lower the volunteer seedbank. The minimum harvest loss is typically about 12 kg per acre, which is 5 times the typical seeding rate. On many fields, harvest losses are 20 times the seeding rate, or more.

(7) Avoid tillage in the spring, to allow for maximum volunteer emergence. However, if using a conventional tillage system, growers can maximize weed control benefits by shortening the interval between tillage and seeding operations.

4.5.4 Biological control

Biological control methods for canola volunteers have not been developed.

4.6 Means of movement and dispersal

Seed dispersal of Brassica napus can occur through pod shatter, spillage of seed during transportation, activities of seed-eating bird and mammalian herbivores and by wind and/or water (Bailleul et al. 2012; Devos et al. 2012; Haile et al. 2014; Kawata et al. 2009; OECD 2012; Panter and Dolman 2012; Pivard et al. 2008; Twigg et al. 2009; von der Lippe and Kowarik 2007; Wang et al. 2007).

Spillage during seed transportation contributes up to 15% of canola seed dispersal. Several studies have shown that feral populations of B. napus result from spillage along transport routes, along railway lines, riverbanks, field margins and near grain silos. Feral populations may be as little as a few individuals to more than a thousand plants per location (Devos et al. 2012; Gulden et al. 2008; Pivard et al. 2008).

5 Related species of Brassica napus

Members of the Brassica genus that are present in Canada and can hybridize with B. napus include B. carinata, B. juncea, B. nigra, B. oleracea and B. rapa (OECD 2012; Warwick et al. 2014).

B. carinata is an annual that is cultivated in Alberta, Saskatchewan and Manitoba (Getinet 1986; Sask Mustard 2013).

B. juncea is an annual species that is introduced and naturalized throughout Canada with the exception of Yukon and Nunavut. It is regularly identified in cultivated wheat, oat, potato, rape fields, orchards, and as escape weeds in irrigation ditches and spring runoff areas, near grain elevators, and on road margins (Brouillet et al. 2010+; Canola Council of Canada 2014a; CFIA 2014; Gan et al. 2008; May et al. 2010; OECD 2012; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2009; Warwick et al. 2014).

B. nigra is an annual species that is introduced and naturalized in British Columbia, Alberta, Saskatchewan, Manitoba, Ontario, Quebec, New Brunswick, Nova Scotia and Newfoundland. As a weed, it can be found in fields, orchards, gardens, riverbanks, roadsides, waste spaces and ballast (Brouillet et al. 2010+; Commission 2014; OECD 2012; USDA-ARS 2017; Wanasundara 2008; Warwick et al. 2014).

B. oleracea is a biennial species that is introduced in British Columbia, Alberta, Ontario and Quebec. It is rare for it to escape from cultivation, and is mainly found in agricultural plots, near driftwood in British Columbia, roadsides and waste spaces (AAFC 2012; Brouillet et al. 2010+; OECD 2012; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

B. rapa is an annual species that is introduced and naturalized in all provinces and territories except Nunavut. It can be found in open woods, meadows, ballast, on riverbanks, slopes, and beaches, alongside roadways and in waste spaces (Brouillet et al. 2010+; Canola Council of Canada 2014a; CFIA 2014; Gulden et al. 2008; OECD 2012; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

Outside the genus Brassica, species that may hybridize naturally with B. napus in Canada include: Barbarea vulgaris, Capsella bursa-pastoris, Descurainia sophia, Diplotaxis erucoides, Diplotaxis muralis, Diplotaxis tenuifolia, Eruca vesicaria subsp. sativa, Erucastrum gallicum, Rorippa indica, Raphanus raphanistrum, Raphanus sativus, Sinapis alba and Sinapsis arvensis (OECD 2012; Warwick et al. 2009).

B. vulgaris is a biennial species that is introduced and naturalized in all provinces of Canada, but none of the territories. The species grows in gardens, meadows, pastures, open woods and bogs, along stream sides, shores and roadsides and in waste places. B. vulgaris can establish in soils that range from sand, gravel and clay, to rich loam (Brouillet et al. 2010+; MacDonald and Cavers 1991; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

C. bursa-pastoris is an annual species that is introduced and naturalized in all provinces and territories of Canada. The species grows in gardens, orchards, vineyards, woods, pastures, meadows, flats and beaches, around hot springs, docks and fish houses, along roadsides and railways and in waste places (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

D. sophia is an annual or biennial species that is introduced and naturalized in most provinces and territories of Canada, except Nunavut and Labrador. It can be found growing in sagebrush, on ranges, flats, beaches, sloughs and ballast, around barnyards, silos and wharves, along roadsides and railways and in waste places (Best 1977; Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

D. erucoides is an annual or biennial species that is ephemeral, not established or recurring in Quebec. It is found growing on ballast, around fish houses and along roadsides (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

D. muralis is an annual or biennial species that is introduced and naturalized in Alberta, Saskatchewan, Manitoba, Ontario, Quebec, New Brunswick, and Nova Scotia. It is present in British Columbia. It grows on sand, gravel, clay and loam, in disturbed prairie fields, parklands, gardens, shores, harbours, ditches, around fish houses, along shores, roadsides and railways and in waste places (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

D. tenuifolia is a perennial species that is introduced in British Columbia, Ontario and Quebec and ephemeral in New Brunswick and Nova Scotia. The species grows on sand, gravel and clay, in fields, river and lake shores, in gravel pits, around ports, on ballast and cinders, along roadsides and railways and in waste places (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

E. vesicaria subsp. sativa is an annual species that is introduced and naturalized in British Columbia, Alberta, Saskatchewan, Manitoba and Ontario. It is ephemeral to Quebec. It is found in cultivated alfalfa fields as a rare escape and seed contaminant, and occasionally along roadsides and waste spaces (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

E. gallicum is an introduced, naturalized annual, winter annual in British Columbia, Alberta, Saskatchewan, Manitoba, Ontario, Quebec, New Brunswick, Prince Edward Island, Nova Scotia, Newfoundland, and the Northwest Territories. It is considered weedy and can be found in gardens, orchards, grain, mustard and sunflower fields, pastures, woods, thickets, shores, and flats. It grows in ballast, along grain elevators, roadsides and in waste spaces (Brouillet et al. 2010+; GOC 2005; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014; Warwick and Wall 1998).

R. indica is an annual, biennial or perennial species native to British Columbia but has not been reported growing in other regions of Canada (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

R. raphanistrum is an introduced, naturalized annual, or biennial which can be found in British Columbia, Alberta, Saskatchewan, Manitoba, Ontario, Quebec, New Brunswick, Prince Edward Island, Nova Scotia, Newfoundland and Laborador. It is a weed in grain, rape, potato, cabbage, hay, clover, pea, bulb and hop fields, in gardens, orchards, woods, cliffs, outcrops, beaches, and dunes. It grows in sand, grass, gravel, clay, sandy loam, and can be found along wharves, roadsides railways and waste spaces (Brouillet et al. 2010+; GOC 2005; Plants of Canada Database 2014; USDA-ARS 2017; Warwick and Ardath 2005; Warwick et al. 2014).

R. sativus is an introduced annual in British Columbia and Saskatchewan, and ephemeral in Mantioba, Ontario, Quebec, New Brunswick, Nova Scotia and Newfoundland. It is found in gardens, grain, rape and corn fields, orchards, riverbanks, flats, by wharves and roadsides. It grows on loamy, sandy soil (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

S. alba is an introduced annual found in all areas of Canada except Northwest Territories, Nunavut, Nova Scotia and Newfoundland and Laborador. It is considered a weed and can be found in fields, farmyards, disturbed prairies, irrigated land, ballast, talus wharf, roadsides, railways and waste spaces (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

S. arvensis is an introduced found in all areas of Canada except for Nunavut. It is considered weedy and is found in grain, hay, rape, potato, and fruit fields, gardens orchards, clearings, river valleys and shores. It also grows on ballast, gravel, sand and can be found near grain elevators, roadsides, railways and waste spaces (Brouillet et al. 2010+; GOC 2005; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2000; Warwick et al. 2014).

5.1 Inter-species/genus hybridization

The extent of inter-species and inter-genus hybridization of B. napus with related species is influenced by factors such as flowering synchronicity, physical proximity, size of pollen source, pollen viability, pollination vectors, sexual compatibility, breeding systems, environmental factors, etc. In vitro techniques have also been used to generate B. napus hybrids through ovary, ovule and embryo culture and through protoplast fusion. Tables 1 and 2 summarize the results of inter-species and inter-genus hybridization of B. napus with related species.

Table 1: Reports of experimental inter-species and inter-genera hybridization (sexual crosses) between Brassica napus and related species
Cross Female Cross Male Description Reference
B. carinata B. napus Successful. (Fernandez-Escobar J. et al. 1988)
B. carinata B. napus Successful. 26 seeds harvested. (Getinet A. et al. 1997)
B. carinata B. napus 1 F1 seed produced from unreported number of crosses. (La Mura et al. 2010)
B. carinata B. napus 1.56 seeds per pollination. (Niemann et al. 2014)
B. napus B. carinata Successful. (Fernandez-Escobar J. et al. 1988)
B. napus B. carinata Successful. (Chen and Heneen 1992)
B. napus B. carinata Successful. On average, cross yielded 8.2 F1 seeds per pollination. (Rashid. A. et al. 1994)
B. napus B. carinata 90 siliques from 110 pollinated flowers. (Gupta 1997)
B. napus B. carinata 0-0.6 seeds per pollination. No pollen in F1. (Chang et al. 2007)
B. napus B. carinata 4 F1 seed produced from unreported number of crosses. (La Mura et al. 2010)
B. napus B. carinata 3.44 seeds per pollination. (Niemann et al. 2014)
B. napus B. carinata Successful only in 1 or 2 B. carinata genotypes. 15-20% of F1 demonstrated male sterility. (Sheikh et al. 2014)
B. gravinae B. napus Unsuccessful. (Nanda Kumar et al. 1988)
B. napus B. gravinae Unsuccessful. (Nanda Kumar et al. 1988)
B. juncea B. napus Successful. Inheritance study for resistance to blackleg. (Roy 1978)
B. juncea B. napus Successful. (Anand et al. 1985)
B. juncea B. napus Successful. (Dhillon et al. 1985)
B. juncea B. napus Successful. Inheritance study for resistance to white rust. (Subudhi and Raut 1994)
B. juncea B. napus Successful. Obtained an average of 4.6 F1 seeds per pollination. (Frello et al. 1995)
B. napus B. juncea Successful. Inheritance study on male sterility. (Fan et al. 1986)
B. napus B. juncea Successful. F1 hybrids plants were vigorous with pollen fertility of 29%. Siliques contained 1 to 2 seeds. (Prakash and Chopra 1990)
B. napus B. juncea Successful. An average of 2.1 seeds was obtained from each pollinated bud. (Rashid. A. et al. 1994)
B. maurorum B. napus Successful. Number of seeds formed was 7; number of hybrid seedlings established was 2; hybrids were pollen sterile. (Chrungu. B et al. 1999)
B. nigra B. napus Successful. (Struss et al. 1991)
B. nigra B. napus Unsuccessful. (Diederichsen and Sacristan 1988)
B. nigra B. napus Unsuccessful. (Kerlan et al. 1992b)
B. napus B. nigra Successful. Stability of Bt content study. (Zhu and Struss 1991)
B. napus B. nigra Unsuccessful. (Diederichsen and Sacristan 1988)
B. napus B. nigra Unsuccessful. (Kerlan et al. 1992b)
B. oleraceae B. napus Successful. 4-19 seeds produced. (Chiang et al. 1977)
B. oleraceae B. napus Unsuccessful. (Kerlan et al. 1992b)
B. napus B. oleraceae Successful. (Chiang et al. 1977)
B. napus B. oleraceae Unsuccessful. (Ayotte et al. 1987)
B. rapa B. napus Successful. Yielded 278 seeds including 122 hybrids. (Palmer 1962)
B. rapa B. napus Successful. Fertile F1 plants produced. (Lammerink 1970)
B. rapa B. napus Successful. Produced 35 seeds of which 8 seeds germinated and grew into plants. (Kamala 1976)
B. rapa B. napus Successful. Produced 52 viable seeds. (Beversdorf et al. 1980)
B. rapa B. napus Successful. (Anand and Downey 1981)
B. rapa B. napus Successful. Inheritance study of clubroot resistant. (Gowers 1982)
B. rapa B. napus Successful. Varying numbers of F1 hybrid plants obtained from crosses between B. rapa and 147 B. napus cultivars and lines. (Pellan-Delourme and Renard 1987)
B. rapa B. napus Successful. Numerous hybrid seeds produced. (Brown and Brown 1996)
B. rapa B. napus Successful. Numerous hybrid seeds produced. (Hauser et al. 1997)
B. rapa B. napus Successful. Numerous hybrid seeds produced. (Hauser et al. 1998b)
B. rapa B. napus Successful. Hand pollination and flower culture yielded 3 hybrids out of 230 pollinated flowers (1.5%). (Metz et al. 1997)
B. rapa B. napus Successful. The frequency of F1 hybrids ranged from 19% to 100%. (Halfhill et al. 2001)
B. rapa B. napus Successful. Hybridization frequency ranged from 16.9% to 0.7%. (Halfhill et al. 2002)
B. rapa B. napus Successful. (Hansen et al. 2001)
B. rapa B. napus Successful. Average number of seeds produced was 7.6 per silique. (Jenkins et al. 2005)
B. rapa B. napus Successful. (Zhu et al. 2004)
B. napus B. rapa Successful. Produced 226 seeds, 165 plants including 160 hybrids. (Palmer 1962)
B. napus B. rapa Successful. (Nwankiti 1970)
B. napus B. rapa Successful. Numerous seeds produced. (McNaughton 1973)
B. napus B. rapa Successful. Study on resistance to clubroot race 3. (Johnston 1974)
B. napus B. rapa Successful. Produced 87 seeds of which 10 seeds germinated & grew into plants. (Kamala 1976)
B. napus B. rapa Successful. Numerous seeds obtained. (Mackay 1977)
B. napus B. rapa Successful. (Beversdorf et al. 1980)
B. napus B. rapa Successful. (Anand and Downey 1981)
B. napus B. rapa Successful. (Chen et al. 1990)
B. napus B. rapa Successful. (Goring et al. 1992)
B. napus B. rapa Successful. Produced on average 4 seeds per silique. (Cheng et al. 1994)
B. napus B. rapa Successful. Numerous seeds produced. (Brown and Brown 1996)
B. napus B. rapa Successful. Average seed set per pollination was 9.8. (Mikkelsen et al. 1996b)
B. napus B. rapa Successful. (Hauser et al. 1997)
B. napus B. rapa Successful. Numerous hybrid seeds produced. (Hauser et al. 1998b)
B. napus B. rapa Successful. (Hu et al. 1997)
B. napus B. rapa Successful. (Verma et al. 2000)
B. napus B. rapa Successful. Long pod character from a B. napus introgressed into B. rapa. (Lewis et al. 2001)
B. napus B. rapa Successful. (Liu et al. 2002)
D. erucoides B. napus Successful. 78 pods formed, 74 seeds harvested. Of seeds that germinated, 3 were hybrids. (Ringdahl et al. 1987)
D. erucoides B. napus Unsuccessful. (Vyas et al. 1995)
D. muralis B. napus Successful. (Fan et al. 1985)
D. muralis B. napus Successful. 157 pods formed, 607 seeds harvested. Of seeds which germinated, 31 were hybrids. (Ringdahl et al. 1987)
B. napus D. muralis Successful. (Bijral and Sharma 1996)
D. tenuifolia B. napus Unsuccessful. (Ringdahl et al. 1987)
B. napus E. gallicum Successful. Several viable seeds obtained. 1 hybrid plant obtained per 100 pollinated flowers. (Lefol et al. 1997)
E. gallicum B. napus Unsuccessful. (Lefol et al. 1997)
B. napus R. raphanistrum Successful. The number of viable seeds obtained per 100 harvested seeds was 2. (Lefol et al. 1997)
Successful. Produced on average 0.12 seeds per 100 flowers and 0.78 seeds per plant. (Chèvre et al. 1998)
R. raphanistrum B. napus Unsuccessful. (Kerlan et al. 1992b)
R. raphanistrum B. napus Unsuccessful. (Lefol et al. 1997)
R. raphanistrum B. napus Unsuccessful. (Warwick et al. 2003a)
B. napus R. raphanistrum Unsuccessful. (Kerlan et al. 1992b)
R. sativus B. napus Successful. Several hybrid plants were obtained. (Paulmann and Röbbelen 1988)
R. sativus B. napus Unsuccessful. (Lelivelt et al. 1993a)
R. sativus B. napus Unsuccessful. (Metz et al. 1995)
B. napus R. sativus Successful. Many intergeneric hybrids were produced. (Huang et al. 2002)
B. napus R. sativus Unsuccessful. (Lelivelt et al. 1993a)
S. alba B. napus Unsuccessful. (Lelivelt et al. 1993b)
S. alba B. napus Unsuccessful. (Lelivelt et al. 1993b)
S. alba B. napus Unsuccessful. (Sridevi and Sarla 1996)
B. napus S. alba Unsuccessful. (Lelivelt et al. 1993b)
B. napus S. alba Unsuccessful. (Sridevi and Sarla 1996)
S. arvensis B. napus Successful. 1 hybrid obtained. (Moyes et al. 2002)
S. arvensis B. napus Unsuccessful. (Kerlan et al. 1992b)
S. arvensis B. napus Unsuccessful. (Bing et al. 1995)
S. arvensis B. napus Unsuccessful. (Moyes et al. 1999)
B. napus S. arvensis Successful. (Inomata 1988)
B. napus S. arvensis Successful. (Moyes et al. 1999)
B. napus S. arvensis Successful. (Moyes et al. 2002)
B. napus S. arvensis Unsuccessful. (Kerlan et al. 1992b)
B. napus S. arvensis Unsuccessful. (Bing et al. 1995)
B. napus S. pubescens Successful. (Inomata 1994)
Table 2: Reports of experimental inter-species and inter-genera hybridization (artificially through culturing of the ovary, ovules and embryo or through protoplast fusion) with Brassica napus and related species
Hybridization Description References
B. napus +
Arabidopsis thaliana
Polyethylene glycol mediated protoplast fusion resulted in calli that produced 29 shoots (each from a different callus) of which 25 were hybrids. 14 out of 19 tested hybrids were female fertile and set seed after backcrossing to B. napus. (Forsberg et al. 1994)
B. napus +
Arabidopsis thaliana
Polyethylene glycol-mediated protoplast fusion resulted in 155 shoots from 1,520 calli of which 109 rooted in vitro and were transplanted to soil. Of the 68 plants that established in soil, 56 plants produced seeds after self-pollination. The remaining plants either did not flower or failed to set seeds. (Yamagishi et al. 2002)
B. napus +
Barbarea vulgaris
Polyethylene glycol-mediated protoplast fusion yielded 1414 calli from 5 experiments of which 102 developed shoots, 9 developed into plantlets of which 6 were hybrids. Mature plants could not be established outside in vitro conditions. (Fahleson et al. 1994a)
B. napus x B. carinata Ovary, ovule culture; 2 F1 seeds from 44 pollinated flowers, F1 hybrids were male sterile. (Sabharwal and Doležel 1993)
B. napus x B. carinata Polyethylene glycol mediated protoplast fusion; 13 plants were regenerated. (Klíma et al. 2009)
B. carinata x B. napus Embryo culture; successful, 9 bivalents observed in 50 cells. (Harberd and McArthur 1980)
B. carinata x B. napus Ovule culture; 17.0-64.1% hybrid yield after varying days of pollination, pollen viability of sample of F1 plants ranged from 0-30%, with most hybrids between 10-20%. (Sacristan and Gerdemann 1986)
B. napus x
B. juncea
Embryos resulting from controlled pollination were cultured in basal media of Murashige & Skoog (MS) and White, percent ovaries showing seed formation ranged from 37.8% to 80%. Hybrids were raised under field conditions to maturity. (Bajaj et al. 1986)
B. napus x
B. juncea
Developing ovules resulting from controlled crosses between B. napus as female and B. juncea as male parent were cultured in MS agar medium. Total of 2346 ovules were cultured and 249 hybrids were obtained. (Sacristan and Gerdemann 1986)
B. napus x
B. juncea
Developing ovules from controlled crosses were cultured in MS medium, young seedlings were transferred to perlite and then to soil and grown to maturity in greenhouse. Out of 95 crosses, the 44 pods that developed yielded 14 F1 seeds. (Sabharwal and Dolezel 1993)
B. napus x
B. juncea
Hybrid embryos resulting from controlled crosses were cultured in MS and B5 media. Cultured embryo-derived plants were transferred to pots with soil. Percent calli obtained ranged from around 36 to 50% (MS media) and 13 to 42% (B5 media). Percent plants obtained were 18.62% (MS media) and 6.66% (B5 media). (Zhang et al. 2003)
B. napus x
B. juncea
Polyethylene glycol-mediated protoplast fusion yielded 800 calli that were isolated by micropipettes of which 13 calli (1.6%) differentiated into shoots and 5 were successfully transferred to the greenhouse. Of the 1500 calli that were flow sorted, 38 (2.5%) differentiated into shoots from which 20 hybrid plants were obtained. (Sundberg and Glimelius 1991)
B. napus x
B. nigra
Controlled crosses followed by ovary culture in E12 medium; transfer of seedlings to MS medium and transfer of plantlets to pots and grown in greenhouse. Percent of embryos obtained for B. napus as a female versus as a male parent was 4.9% & 0.4% and percent plantlets obtained was 3.4% & 0%, respectively. (Kerlan et al. 1992a)
B. napus x
B. nigra
Polyethylene glycol-mediated protoplast fusion yielded 30 hybrids with chromosomes of both species, 20 plants had the sum of the parental chromosome numbers. (Sjodin and Glimelius 1989)
B. napus x
B. nigra
X-ray irradiated protoplasts followed by polyethylene glycol-mediated protoplast fusion yielded a total of 332 hybrid calli from which 30 produced shoots (1-20 per callus). Shoots were transferred into soil and grown in growth chambers. (Gerdemann-Knorck et al. 1995; Gerdemann-Knorck et al. 1994)
B. napus x
B. oleracea
Embryo culture on 77 developing hybrid ovules yielded 9 backcrossed 2 (BC2) plants. (Ayotte et al. 1987)
B. napus x
B. oleracea
Controlled crosses followed by ovary culture in E12 medium; transfer of seedlings to MS medium and transfer of plantlets to pots and grown in greenhouse. Percent embryos obtained for the 2 varieties of B. oleraceae with B. napus as a female parent was 21.5% and 13.2%, with B. napus as a male parent it was 1.8% and 1.2%. Percent plantlets obtained with B. napus as a female parent was 13.7% and 5.7%, with B. napus as a male parent it was 0.7% and 0.2%. (Kerlan et al. 1992a)
B. napus x
B. oleracea
Developing ovules, resulting from controlled crosses were cultured in Nitsch and Nitsch medium. Efficiency of conversion of embryos to plants was 0 to 22%. (Ripley and Beversdorf 2003)
B. napus x
B. oleracea
Hybrid embryos were rescued by ovule culture in a Nitsch and Nitsch medium supplemented with 300 mg l-1 casein hydrolysate, 200 mg l-1 glutamine and 13% sucrose. (Bennett et al. 2008)
B. napus x
B. oleracea
Polyethylene glycol-mediated protoplast fusion yielded 1128 calli. The frequency of calli regenerated shoots was 8.9% and of hybrid plants was 93%. (Sundberg and Glimelius 1991)
B. napus x
B. oleracea
Embryos resulting from controlled crosses were rescued from developing ovules and cultured in Nitsch and Nitsch medium. The resulting hybrids were backcrossed to B. campestris and the progeny again backcrossed to B. campestris. A total of about 350 plants were generated. ( Quiros et al. 1987)
B. napus x
B. rapa
Amphidiploid B. napus lines were resynthesized through reciprocal interspecific crosses between B. oleracea and B. rapa using ovule culture and colchicine treatment. (Zhang et al. 2002)
B. napus x
Capsella bursa- pastoris
Embryos resulting from controlled crosses were cultured in MS medium to obtain F1 plants. From 9248 pollinations, 169 F1 plants were obtained. (Chen et al. 2007)
B. napus +
Descurainia sophia
Polyethylene glycol-mediated protoplast fusion yielded 19 somatic hybrid plants of which 8 did not flower. (Rongzhan et al. 2007a)
B. napus x
Diplotaxis harra
Ovaries resulting from controlled crosses were cultured in Nitsch and Nitsch (NN) basal medium. Three seeds were harvested from 75 cultured ovaries. All seeds germinated, but two died at the seedling stage. One F1 hybrid matured. (Inomata 2005)
B. napus x
Diplotaxis tenuifolia
In-vitro pollination of ovules followed by culturing of 40 ovaries in Murashige and Skoog's media yielded 14 ovaries with enlarged ovules containing embryos at different stages of development. (Zenkteler 1990)
B. napus +
Eruca sativa
Hybrid cells resulting from polyethylene glycol-mediated protoplast fusion were cultured in a modified 8p medium, transferred to K3 medium. The resultant calli were transferred to MS medium for rooting and to obtain hybrid plants that produced seeds. On average 5.4% of the calli obtained after selection differentiated into shoots and 23 hybrids were successfully transferred to the greenhouse. (Fahleson et al. 1988)
B. napus +
Thlaspi perfoliatum
Polyethylene glycol-mediated protoplast fusion yielded 27 hybrid or partially hybrid calli from which 19 plants were grown to maturity. (Fahleson et al. 1994b)
B. napus x
Raphanus raphanistrum
Ovaries obtained from controlled crosses were cultured in E12 medium, seedlings that emerged were transferred to Murashige & Skoog medium and plantlets were transferred to pots and grown in greenhouse. From the 243 embryos produced, 109 interspecific hybrid plants were obtained. (Kerlan et al. 1992a)
B. napus x
Raphanus sativus
Ovules obtained from controlled crosses were cultured in basic White's agar medium and germinated embryos were transferred to Murashige & Skoog medium. Of 14 cross combinations, 5 combinations gave hybrids by ovule culture. (Takeshita et al. 1980)
B. napus x
Raphanus sativus
Siliquae obtained from controlled reciprocal crosses were cultured in Murashige & Skoog medium. Embryos from siliquae were then cultured in Murashige & Skoog medium and resulting plantlets were transferred to soil and grown in greenhouse. No hybrid plants were obtained. (Lelivelt et al. 1993a)
B. napus x
Raphanus sativus
Ovaries resulting from controlled pollination and cultured in B5 media yielded 14 hybrid plants from 58 cultured ovaries. ( Luo et al. 2000)
B. napus x
Raphanus sativus
Polyethylene glycol-mediated fusion of R. sativus cytoplast and B. napus protoplast resulted in fertile and sterile cybrids; 4 out of 10 plants contained novel mtDNA, 1 was male sterile and 3 were male fertile. (Sakai and Imamura 1990)
B. napus x
Raphanus sativus
Polyethylene glycol-mediated protoplast fusion yielded 364 calli. The frequency of calli that regenerated shoots was 5% and frequency of hybrid plants obtained was 100%. (Sundberg and Glimelius 1991)
B. napus x
Raphanus sativus
PEG-induced protoplast fusion resulted in one somatic hybrid. (Lelivelt and Krens 1992)
B. napus x
Raphanus sativus
PEG-mediated protoplast fusion yielded 12 somatic hybrids. (Wang et al. 2006)
B. napus +
Rorippa indica
PEG-DMSO-mediated protoplast fusion and culturing with B5 liquid medium yielded somatic hybrids. Thereafter, 3-way intergeneric hybridization and embryo rescue yielded 890 F2 individuals. (Rongzhan et al. 2007b)
B. napus x
Sinapis alba
PEG-mediated protoplast fusion did not yield plants with recombinant mitochondrial or chloroplast DNA. (Lelivelt et al. 1993b)
B. napus x
Sinapis alba
Ovaries from reciprocal crosses were cultured in E12 medium that yielded 2.2 and 1.9% of interspecific hybrids when S. alba was used as the female and male parent, respectively. (Chevre et al. 1994)
B. napus x
Sinapis alba
Electrically-induced protoplast fusion yielded 7 somatic hybrids. Hybrids were grown to full maturity and set seeds after self-pollination or back-crossing with B. napus. (Wang et al. 2005)
B. napus x
Sinapis arvensis
Ovules obtained from controlled crosses, cultured in M91 medium and transferred to B5 medium yielded one hybrid plant. (Bing et al. 1991)
B. napus x
Sinapis arvensis
Ovaries obtained from controlled reciprocal crosses were cultured in E12 medium, followed by culturing embryos in Murashige and Skoog medium and transferring plantlets in pots for growing in greenhouse. Reciprocal crosses without embryo rescue did not yield any seed. (Kerlan et al. 1991)
B. napus x
Sinapis arvensis
Ovules obtained from controlled crosses and cultured in E12 medium yielded 30 embryos and 3 plantlets from the 808 ovaries cultured with B. napus as the female parent versus no embryos and no plantlets from 732 ovaries cultured with B. napus as the male parent. (Kerlan et al. 1992a)
B. napus x
Sinapis arvensis
Ovaries obtained from controlled crosses were cultured in E12 medium and yielded 1.2% B. napus and 0.1% S. arvensis hybrids per number of flower pollinated. (Lefol et al. 1996)
B. napus x
Sinapis arvensis
Polyethlene glycol-mediated protoplast fusion yielded somatic hybrids (54 symmetric and 4 asymmetric hybrids) with a plant regeneration efficiency of 1.4%. (Hu et al. 2002)

5.2 Potential for introgression of genetic information from Brassica napus into relatives

Brassica napus is thought to have moderate gene introgression potential (Stewart et al. 2003). The level of introgression depends on factors such as sympatry and flowering time, genetic background, performance of the F1 and backcross hybrids, the number of backcrosses performed, habitat suitability, life span of the weedy population and the time-scale of sympatric occurrence (Stewart et al. 2003; OECD 2012; OGTR 2011b).

Trait introgression potential from B. napus into B. rapa is considered to be higher relative to other species because B. rapa is a self-incompatible, allogamous species with a common set of chromosomes with B. napus (Nagaharu 1935; OECD 2012; OGTR 2011b; Salisbury 2002a). Preventing natural introgression of traits from B. napus to B. rapa is not thought to be possible (Hails 2000; Hauser et al. 1998a; Hauser et al. 1998b; OECD 2012; Salisbury 2002a).

B. napus and B. rapa interspecific hybridization has been detected under controlled field and greenhouse conditions as well as in the natural environment (Johnson 1974; Beversdorf et al. 1980; Goring et al. 1992; Lewis et al. 2001). In Canada, hybridization between B. napus and B. rapa has been reported in wild populations and in commercial fields (Warwick et al. 2003b). Similar observations have also been made in other countries, like Denmark (Mikkelsen et al. 1996a; Jørgensen and Andersen 1994, Jørgensen et al. 1996, 1998; Hansen et al. 2001), the United Kingdom (Allainguillame et al. 2009; Wilkinson et al. 2003) and the United States (Halfhill et al. 2003; Halfhill et al. 2001; Neal Stewart Jr. et al. 2003; OECD 2012).

Hybridization rates between B. napus and B. rapa vary from 10% to 90% (Jørgensen et al. 1996) and are most effective when B. napus is the pollinator (Hails 2000; Jørgensen et al. 1996, 1998; OECD 2012; OGTR 2011b; Scott and Wilkinson 1998). For example, when pollen was donated from transgenic, Bt insect resistant B. napus, to wild B. rapa, 27% of the resultant hybrid population contained the Bt trait (Vacher et al. 2004). In another study when B. napus x B. rapa hybrids showed reduced fertility and seed set, as well as low seedling survival rates when compared to either parent (Hauser et al. 1998a). It should be noted that considerable agronomic trait variability was observed; some hybrids were reported to be as fit as parental lines (Hauser et al. 1998b).

B. juncea has the second highest rate of hybridization with B. napus, after B. rapa (Scheffler and Dale 1994). B. napus x B. rapa F1 hybrids possess low viability pollen, although improved fertility with spontaneous backcrossing has been observed (Tsuda et al. 2014). Due to improved performance after backcrossing, it is thought that introgression of genes from B. napus to B. juncea can occur in nature (Bing et al. 1991; Bing et al. 1996; OECD 2012; OGTR 2011b; Scheffler and Dale 1994; Tsuda et al. 2014).

B. napus and B. nigra hybrids demonstrate sterility rates from moderate to high (Bing et al. 1996; Kerlan et al. 1992a). Controlled crosses between the two species have produced no hybrids with B. nigra as the female parent, so trait introgression from B. napus to B. nigra is unlikely (Bing et al. 1991; Bing et al. 1996; Kerlan et al. 1992a; OECD 2012; Salisbury 2002a).

B. napus oilseed varieties could potentially hybridize with B. napus and B. rapa vegetable subspecies. However, vegetable crops are harvested prior to flowering unless they are being grown for seed production (Salisbury 2002a). Controlled crosses between B. napus and B. oleracea have been produced at very low frequencies. Spontaneous hybridization has been observed when wild B. oleracea populations were within 25 m of B. napus fields. Given these conditions, natural introgression from B. napus to B. oleracea is considered to be unlikely (Ayotte et al. 1987; Chiang et al. 1977; OECD 2012; Salisbury 2002a).

Hybridization between B. napus and Raphanus raphanistrum was reported to occur at very low frequencies in field surveys in France (Darmency et al. 1998) and Australia (Rieger et al. 2001). Hybridization was not observed during field surveys in Canada (Warwick et al. 2001), Switzerland (Thalmann et al. 2001) or the United Kingdom (Sweet and Shepperson 1996). Using herbicide-tolerant traits as a measure, hybridization rates were low or unsuccessful when R. raphanistrum was used as the female parent (Salisbury 2002a).

Gene flow between B. napus and Sinapis arvensis has not been detected under natural conditions or co-cultivation experiments (Bing et al. 1996). No hybrids were detected when S. arvensis was used as the female parent or when the species was used as a pollinator under open pollination conditions. All hybrids were reportedly sterile (Bing et al. 1996; Downey 1999; Lefol et al. 1996; Moyes et al. 2002; Salisbury 2002).

Hybridization between B. napus and Erucastrum gallicum is considered to be remote/low since embryo rescue is necessary to obtain hybrids (Lefol et al. 1997). The few hybrids that have been obtained were slow-growing with low pollen viability. No seeds have been produced with E. gallicum as the female parent and plants produced from backcrossed hybrids were identical to E. gallicum (Lefol et al. 1997; OECD 2012; Warwick et al. 2003b; Warwick and Wall 1998). Similarly, hybridization between B. napus and Sinapis alba is also considered to be low since field crosses between the two species have not been detected and hand pollination with embryo or ovule culture are necessary to produce hybrids (Brown et al. 1997; Chevre et al. 1994; Lelivelt 1993; OECD 2012).

5.3 Summary of the ecology of relatives of Brassica napus

Species that are related to Brassica napus and are common weeds in Canada include Brassica juncea, Brassica nigra, Barbarea vulgaris, Capsella bursa-pastoris, Descurainia sophia, Raphanus sativus, Sinapis alba. Raphanus raphinastrum Erucastrum gallicum, S. arvensis (secondary noxious weed). Several of these are classified as noxious weeds under the Weed Seeds Order, 2016, including R. raphanistrum and Barbarea spp. (primary noxious weeds) as well as E. gallicum and S. arvensis (secondary noxious weed) (GOC 2005; Holm et al. 1991; OECD 2012; Warwick et al. 2014).

B. juncea grows throughout Canada as a weed in cultivated fields of wheat, oat, potato and rapeseed. B. juncea is also cultivated in the drier regions of Western Canada for the condiment industry. B. juncea has also been developed as a canola-quality oilseed crop in Canada (Brouillet et al. 2010+; Canola Council of Canada 2014a; CFIA 2014; Gan et al. 2008; May et al. 2010; OECD 2012; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2009, 2014).

B. nigra can be found as a weed in orchards and gardens. It is cultivated in limited areas in Western Canada as a condiment crop (Brouillet et al. 2010+; Commission 2014; OECD 2012; Plants of Canada Database 2014; USDA-ARS 2017; Wanasundara 2008; Warwick et al. 2014).

Similarly, B. oleracea can be found growing as a weed in agricultural fields. B. oleracea vegetable crops (i.e. cabbage, broccoli, cauliflower and Brussels sprouts) are grown as annuals in Canada (AAFC 2012; Brouillet et al. 2010+; OECD 2012; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

B. rapa grows as a weed in grain, hay, mustard, vegetable, potato, corn, bulb crops and sugar beet fields. B. rapa is cultivated in Canada as canola (Brouillet et al. 2010+; Canola Council of Canada 2014a; CFIA 2014; Gulden et al. 2008; OECD 2012; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

Other species of interest that hybridize with B. napus include Barbarea vulgaris, Capsella bursa-pastoris, Descurainia sophia, Diplotaxis erucoides, D. muralis, D. tenuifolia, Eruca vesicaria subsp. sativa, Erucastrum gallicum, Rorippa indica, Raphanus raphanistrum, R. sativus, Sinapis alba and S. arvensis (OECD 2012; Warwick et al. 2009).

B. vulgaris is a noxious weed in Ontario while C. bursa-pastoris is a noxious weed in Manitoba (Brouillet et al. 2010+; MacDonald and Cavers 1991; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014). As a weedy species, C. bursa-pastoris can be found growing in cultivated fields of grain crops, hay, canola, potato and strawberry fields (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

D. sophia is a noxious weed in Manitoba. Plants that germinate in fall can survive the freezing winters of Canada. The species is better adapted to the Canadian Prairies compared with the more humid conditions of eastern Canada. D. sophia is a weed in dry disturbed meadows, grasslands, pastures, prairies, gardens, barnyards and grain and hay fields (Best 1977; Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

D. muralis and D. tenuifolia grow as weeds in grain fields, while E. vesicaria subsp. sativa is a weed in cultivated fields of alfalfa (Medicago sativa L.) (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

E. gallicum is a Class 3 (secondary noxious) weed in the Weed Seeds Order, 2016 and a noxious weed in Manitoba. The species grows as a weed in gardens, orchards, field crops and in pastures (Brouillet et al. 2010+; GOC 2005; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014; Warwick and Wall 1998).

R. raphanistrum is a Class 2 (primary noxious) weed in the Weed Seeds Order, 2016 and grows as a weed in fields of grain crops, rapeseed, potatoes, cabbage, hay, clover, peas, bulbs and hops (Brouillet et al. 2010+; GOC 2005; Plants of Canada Database 2014; USDA-ARS 2017; Warwick and Ardath 2005; Warwick et al. 2014). Similarly, R. sativus grows as a weed in gardens, fields of grain, rape, corn crops and in orchards (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014).

S. alba grows as a weed in crop fields (Brouillet et al. 2010+; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2014). S. arvensis is a Class 2 (primary noxious) weed in the Weed Seeds Order, 2016, a noxious weed in Manitoba and a regional noxious weed in British Columbia. The species grows as a weed in fields of grain crops, hay, rapeseed, potatoes and fruits (Brouillet et al. 2010+; GOC 2005; Plants of Canada Database 2014; USDA-ARS 2017; Warwick et al. 2000; Warwick et al. 2014).

6 Potential Interaction of Brassica napus with Other Life Forms

There are a number of diseases that affect B. napus in Canada, such as blackleg, clubroot, Sclerotinia stem rot (Sclerotinia sclerotiorum (Lib.) Massee), Alternaria leaf spot (Alternaria brassicae (Berk.) Sacc.), A. brassicicola (Schwein.) Wiltshire, Fusarium spp., Rhizoctonia solani, Pythium ultimum Trow. and P. debaryanum Auct. non R. Hesse Downy mildew (Peronospora parasitica) (Humpherson-Jones and Phelps 1989; Jensen et al. 1999; Abdelazher 2003; Tanina et al. 2004).

Major insect pests of B. napus in Ontario include caterpillars, such as imported cabbageworm (Pieris rapae L.), cabbage looper (Trichoplusia ni Hübner), diamond-back moth (Plutella xylostella L.) and cabbage maggot (Delia radicum L.). Mamestra configurata, also known as bertha armyworm, is a major oilseed crop defoliator. Flea beetles (order Coleoptera, genus Phyllotreta) are important leaf feeders, especially on seedlings (Lamb 1989). Lygus spp. (Lygus elisus Van Duzee and Lygus lineolaris Palisot de Beauvois) and aphids (Myzus persicae Sulzer and Brevicoryne brassicae L.) cause damage but generally have a minor impact on yield (Lamb 1989). Swede midge (Contarinia nasturtii Keiffer) is a serious pest of Brassica crops in Ontario and increasing threat to B. napus grown in western Canada (Allen et al. 2008; Hallett et al. 2009).

For a complete list of species associated with B. napus, please refer to Table 3.

Table 3. Examples of potential interactions of Brassica napus with other life forms during its life cycle in a natural environment
Fungus and Fungus-like
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Albugo candida (Pers.) Kunze
(White rust and staghead); synonym: Albugo cruciferarum (DC.) S.F. Gray
Pathogen Present, widespread (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OECD 2012; Thomas 2003)
Alternaria alternata (Fr.:Fr.) Keissl.
(leaf spot or pod rot)
Pathogen Present (Gulden et al. 2008; OECD 2012)
Alternaria brassicae (Berk.) Sacc.
(alternaria black spot)
Pathogen Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OECD 2012)
Alternaria brassicicola (Schwein.) Wiltshire
(alternaria black spot)
Pathogen Present (OECD 2012)
Alternaria raphani Groves & Skolko
(alternaria black spot); synonym: Alternaria japonica H. Yoshii
Pathogen Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OECD 2012)
Aphanomyces raphani Kendrick
(black rot)
Pathogen Present (OECD 2012; Thomas 2003; Ginns 1986)
Botrytis cinerea Pers.:Fr.
(graymold); teleomorph: Botryotinia fuckeliana (de Bary) Whetzel
Pathogen Present (OECD 2012; Thomas 2003)
Cladosporium sp.
(pod rot)
Pathogen Present (Clear and Patrick 1995; Gilbert et al. 2010;
Ginns 1986; Inglis and Boland 1990;
Vaartnou et al. 1974)
Erysiphe polygoni DC.
(powdery mildew)
Pathogen Present, widespread (OECD 2012; Thomas 2003)
Fusarium avenaceum Sacc.
(Fusarium Wilt)
Pathogen Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008)
Fusarium oxysporum Schlechtend.:Fr
(yellows)
Pathogen Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OECD 2012; Thomas 2003)
Fusarium oxysporum Schlechtend.:Fr.f.sp. conglutinans (Wollenweb.) W.C. Snyder and H.N. Hans
(fusarium wilt)
Pathogen Present (OECD 2012)
Leptosphaeria biglobosa R. A. Shoemaker & H. Brun Pathogen Present (Gulden et al. 2008)
Leptosphaeria maculans (Desmaz.) Ces. and De Not
(Blackleg); anamorph: Phoma lingam (Tode:Fr.) Desmaz.
Pathogen Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OECD 2012; OMAFRA 2009)
Mycosphaerella brassicicola (Duby) Lindau in Engl. And Prantl.
(ring spot); anamorph: Asteromella brassica (Chev.) Boerema and Van Kesteren
Pathogen Present; mostly in western Canada (Agriculture and Agri-Food Canada 2005; OECD 2012) (Thomas 2003)
Penthaleus major (Dugés)
(blue oat mite)
Pathogen Unknown (Gulden et al. 2008)
Peronospora parasitica (Pers.:Fr.) Fr.
(downey mildew)
Pathogen Present, widespread (Gulden et al. 2008; Thomas 2003)
Phytophthora megasperma Drechs.
(phytophthora root rot)
Pathogen Unknown (OECD 2012; Thomas 2003)
Pseudocercosporella capsellae (Ellis & Everh.) Deighton
(white leaf spot and gray stem); synonym: Cercosporella brassicae (Faitrey and Roum.) Höhn.
Pathogen Present; widespread (Gulden et al. 2008; OECD 2012; Thomas 2003)
Pythium spp.
(pythium root rot)
Pathogen Present (Agriculture and Agri-Food Canada 2005; OECD 2012)
Pythium debaryanum Auct. Non R. Hesse
(root rot)
Pathogen Present (Vanterpool 1956)
Rhizoctonia solani, Fusarium and Pythium spp.
(seedling disease and root rot complex )
Pathogen Present (Agriculture and Agri-Food Canada 2005; Thomas 2003)
Rhizoctonia solani Kühn
(white blight or brown girding root rot or head rot); teleomorph:
Thanatephorus cucumeris (A. B. Frank) Donk
Pathogen Present; widespread (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OECD 2012)
Sclerotinia sclerotiorum (Lib.) de Bary
(Sclerotinia White Stem Rot)
Pathogen Present; widespread (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OECD 2012; OMAFRA 2009; Thomas 2003)
Verticillium dahlia Kleb.
(verticillium wilt)
Pathogen Present (Eynck et al. 2007; Gulden et al. 2008)
Verticillium longisporum (comb. Nov. Karapappa et al.)
(verticillium wilt)
Pathogen Present (Eynck et al. 2007; OECD 2012)
Chromist
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Plasmodiophora brassicae
Woronin
(Clubroot)
Pathogen Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008)
Phytoplasma
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Candidatus Phytoplasma asteris
(Aster yellows)
Pathogen Present (Agriculture and Agri-Food Canada 2005; Saskatchewan Ministry of Agriculture 2012)
Bacteria
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Pseudomonas syringae pv. maculicola (McCulloch 1911) Young, Dye and Wilkie 1978)
(bacterial pod rot)
Pathogen Present (OECD 2012; Thomas 2003)
Xanthomonas campestris pv. campestris (Pammel 1895) Dowson 1939
(bacterial black rot); synonyms:
Xanthomonas campestris pv. aberrans and
Xanthomonas campestris pv. raphani
Pathogen Present (OECD 2012; Thomas 2003)
Virus
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Beet Western yellows virus
(BWYV)
Pathogen Present; British Columbia (Gulden et al. 2008; OECD 2012; Thomas 2003)
Turnip Mosaic Virus
(TuMV)
Pathogen Present (OECD 2012; OMAFRA 2009; Thomas 2003)
Insect
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Aelothrips fasciatus L.
(banded wing thrip)
Consumer Present (Gavloski et al. 2011; Heming 1985)
Agriotes lineatus L.
(lined click beetle)
Consumer Present (OECD 2012)
Agrotis ipsilon Hufn.
(black cutworm)
Consumer Present (OECD 2012)
Agrotis orthogonia Morrison
(pale western cutworm)
Consumer Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; Thomas 2003)
Amplicephalus inimicus Say
(painted leafhopper)
Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Apis Mellifera L.
(honey bee)
Symbiont or beneficial organism Present, widespread (Gulden et al. 2008; Thomas 2003)
Autographa californica Speyer
(alfalfa looper)
Consumer Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008)
Autographa gamma L.
(silver-Y moth)
Consumer Present (OECD 2012)
Balclutha sp. Kirkaldy
(leafhoppers)
Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Bemisia tabaci Genn.
(tobacco whitefly)
Consumer Present (OECD 2012)
Bombus terrestris L.
(bumblebee)
Symbiont or beneficial organism Present (Gulden et al. 2008)
Brevicoryne brassicae L.
(aphid)
Consumer Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008)
Camnula pellucida Scudder
(clearwinged grasshopper)
Consumer Present (Agriculture and Agri-Food Canada 2005)
Ceratagalia humilis Kirkaldy Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Ceutorhynchus litura Fabricius
(cabbage seed pod weevil)
Consumer Present, widespread (Agriculture and Agri-Food Canada 2005)
Ceutorhynchus neglectus Blatchey
(weevil)
Consumer Present (Dosdall et al. 1999; Gavloski et al. 2011)
Ceutorhynchus obstrictus Marsham
(cabbage Seedpod Weevil)
Consumer Present, widespread (Gulden et al. 2008; OMAFRA 2009; Thomas 2003)
Ceutorhynchus rapae Gyll.
(cabbage curculio)
Consumer Present (OECD 2012)
Contarinia nasturtii Kieffer
(swede midge)
Consumer Present (Gulden et al. 2008; OMAFRA 2009)
Delia floralis Fallén
(turnip root maggot)
Consumer Present (Gavloski et al. 2011; Gulden et al. 2008)
Delia florilega Zetterstedt
(bean seed maggot)
Consumer Present (Gavloski et al. 2011)
Delia planipalpus Stein
(maggot)
Consumer Present (Gavloski et al. 2011)
Delia platura Meigen
(seed corn maggot)
Consumer Present (Gavloski et al. 2011)
Delia radicum L.
(cabbage root maggot)
Consumer Present (Gavloski et al. 2011; Gulden et al. 2008)
Diplocolenus configuratus Uhler Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Discestra trifolii Hufnagel
(clover cutworm)
Consumer Present (Ayre and Lamb 1990; Gavloski et al. 2011)
Entomoscelis americana Brown
(red turnip beetle)
Consumer Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; Thomas 2003)
Epicauta ferruginea Say
(rust coloured beetle)
Consumer Present (Bousquet et al. 2013)
Euscelis maculipennis Delong and Davidson Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Euxoa ochrogaster Guenée
(red-backed cutworm)
Consumer Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008)
Feltia jaculifera Guenée
(dingy cutworm)
Consumer Present (Byers et al. 1990; Gavloski et al. 2011)
Frankliniella tritici Fitch
(flower thrip)
Consumer Present (CABI 2017; Childers and Achor 1995)
Gyponana sp. Germar
(leafhopper)
Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Hadula trifolii Hufn.
(clover cutworm)
Consumer Present (OECD 2012)
Lipaphis pseudobrassicae Davis
(turnip aphis)
Consumer Present (Gavloski et al. 2011)
Liriomyza brassicae Riley
(serpentine leaf miner)
Consumer Present (OECD 2012)
Loxostege sticticalis L.
(beet webworm)
Consumer Present (Agriculture and Agri-Food Canada 2005)
Lygus borealis Kelton
(lygus bug)
Consumer Present (Agriculture and Agri-Food Canada 2005)
Lygus elisus van Duzee
(pale legume bug)
Consumer Present (Agriculture and Agri-Food Canada 2005)
Lygus keltoni Schwartz
(lygus bug)
Consumer Present (Agriculture and Agri-Food Canada 2005)
Lygus lineolaris Palisot de Beauvois
(tarnished plant bug)
Consumer Present (Agriculture and Agri-Food Canada 2005; OMAFRA 2009)
Lytta nuttalli Say
(nuttall blister beetle)
Consumer Present (Bousquet et al. 2013)
Macrosteles quadrilineatus Forbes
(aster leafhopper)
Consumer Present, widespread (Gavloski et al. 2011; Gulden et al. 2008)
Mamestra configurata Walker
(bertha armyworm)
Consumer Present, widespread in west (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; Thomas 2003)
Melanoplus bivittatus Say
(two striped grasshopper)
Consumer Present (Agriculture and Agri-Food Canada 2005)
Melanoplus sanguinipes
Fabricius
(migratory grasshopper)
Consumer Present (CABI 201; Vickery and Kevan 1985)
Meligethes viridescens Fabricius
(bronzed or blossom beetle)
Consumer Present (Gulden et al. 2008; OECD 2012)
Mythimna unipuncta Haworth
(armyworm)
Consumer Present (Gavloski et al. 2011)
Myzus persicae Sulzer
(green peach aphid)
Consumer Present, widespread (Gulden et al. 2008)
Neokolla hieroglyphica Say Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Nezara viridula L.
(green stink bug)
Consumer Present (OECD 2012)
Osmia rufa L.
(red mason bee)
Symbiont or beneficial organism (Gulden et al. 2008)
Peridroma saucia Hübner
(variegated cutworm)
Consumer Present (Gavloski et al. 2011)
Phyllotreta cruciferae Goeze
(crucifer flea beetle)
Consumer Present, widespread (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OMAFRA 2009; Thomas 2003)
Phyllotreta striolata Fabricius
(stiped flea beetle)
Consumer Present, widespread (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OMAFRA 2009; Thomas 2003)
Phyllotreta undulata Kutschera
(lesser striped flea beetle)
Consumer Present (OECD 2012)
Pieris rapae L.
(imported cabbageworm or cabbage white butterfly)
Consumer Present (OECD 2012; Lamb 1989)
Plutella xylostella L.
(diamondback moth)
Consumer Present (Agriculture and Agri-Food Canada 2005; Gulden et al. 2008; OMAFRA 2009)
Psammotettix sp. Haupt Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Psylliodes punctulata Melsh.
(hop flea beetle)
Consumer Present (OECD 2012)
Scaphytopius acutus Say
(sharpnosed leafhopper)
Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Sorhoanus ulheri Oman Consumer Present (Gavloski et al. 2011; Olivier et al. 2007)
Spodoptera exigua Hbn.
(beet armyworm)
Consumer Present (OECD 2012)
Thrips tabaci Lindeman
(onion thrips)
Consumer Present (Gulden et al. 2008)
Thrips vulgatissimus Haliday
(white flower thrips)
Consumer Present (Gulden et al. 2008)
Trichoplusia ni Hbn.
(cabbage looper)
Consumer Present (OECD 2012; Lamb 1989)
Vanessa Cardui L.
(painted lady butterfly)
Consumer Present, widespread (Gulden et al. 2008; Thomas 2003)
Lady Beetle adult and larva Symbiont or beneficial organism Present, widespread (Thomas 2003)
Hover Fly Symbiont or beneficial organism Present, widespread (Thomas 2003)
Lacewing Symbiont or beneficial organism Present, widespread (Thomas 2003)
Parasitic wasp Symbiont or beneficial organism Present, widespread (Thomas 2003)
Soil insects Consumer Present (OECD 1997)
Nematode
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Pratylenchus penetrans (Cobb) Filipjev and Schuurmans Stekhoven
(root-lesion nematode)
Consumer Present in Quebec (Gulden et al. 2008)
Animals
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Animal browsers (e.g. deer, hare, rabits, rodents) Consumer Present (OECD 1997)
Birds Consumer Present (OECD 1997)
Carduelis tristis L.
(American goldfinches)
Consumer Present (Gulden et al. 2008)
Earthworms Symbiont or beneficial organism Present (OECD 1997)
Grus Canadensis (L.)
(sandhill crane)
Consumer Present (Gulden et al. 2008)
Odocoileus virginiamus Zimmerman
(white-tailed deer)
Consumer Present (Gulden et al. 2008)
Slugs Consumer Present (OECD 1997)
Spermophilus richardsonii
(Richardson's ground squirrel)
Consumer Present (Gulden et al. 2008)
Plant
Other Life Forms Interaction with B. napus (pathogen; symbiont or beneficial organism; consumer; gene transfer) Presence in Canada Reference(s)
Brassica napus L.
(canola)
Gene transfer Present (OECD 1997)
Brassica juncea (L.) Czern.
(Indian mustard)
Gene transfer Present (OECD 1997)
Brassica nigra (L.) W.D.J. Koch
(black mustard)
Gene transfer Present (OECD 1997)
Brassica rapa L.
(Bird rape)
Gene transfer Present (Gulden et al. 2008)
Erucastrum gallicum (Willd.) O.E. Schulz
(dog mustard)
Gene transfer Present (Gulden et al. 2008)
Raphanus raphanistrum L.
(wild radish)
Gene transfer Present (Gulden et al. 2008)
Sinapis arvensis L.
(wild mustard)
Gene transfer Present (Gulden et al. 2008)

7 References

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