Canola Production Field Guide

The germination process involves water absorption, swelling, splitting of the seed coat and emergence of the root tip. Cotyledons are pushed through the soil surface by an active hypocotyl. Germination typically takes from four to 10 days, depending on soil temperature and moisture, seed soil contact and planting depth. During this stage, canola is susceptible to many soil-borne pathogens.

Once emerged, the cotyledons open and supply the new seedling with energy. At this stage, the seedling still is vulnerable to soil-borne pathogens and very susceptible to flea beetle injury. The growing point of canola is between the two cotyledons. The exposed growing point is susceptible to spring frosts, soil drifting, insects and hail damage. Canola is a very poor competitor with weeds, and good stand establishment is extremely important.

The first true leaves develop four to eight days after emergence. The plant quickly establishes a rosette, with older leaves at the base increasing in size and smaller, younger leaves developing in the center. During this time, the stem length remains basically unchanged, although its thickness increases. The rosette stage is characterized by an increase in leaf area index. Rapid and abundant leaf growth captures more sunlight and produces more food for the plant, thus producing more dry matter per day and increasing yield potential. A rapidly developing canola canopy encourages root growth, reduces soil moisture evaporation and shades weeds.

Bud formation is triggered as the days lengthen and temperatures rise. A cluster of flower buds becomes visible at the center of the rosette and “bolts,” or lengthens. Secondary branches arise from buds, which develop in the axils of the upper leaves. Secondary branches develop one to four leaves and a flower bud cluster. The canola plant reaches its maximum leaf area index in the bud stage. The vast majority of photosynthesis occurs in the leaves, and their removal results in large yield losses. The vegetative stages (seedling to first flower) for B. napus generally range from 40 to 60 days, depending on environmental conditions.

Flowering begins with the opening of the lowest bud on the main stem and continues for 14 to 21 days. Three to five flowers or more open per day, and 40 to 55 percent of the flowers that open will develop pods. High temperatures coupled with dry conditions can reduce yield potential severely during this stage.

Ripening begins when the petals on the last flower on the main stem drop. By the time flowering is finished, most of the leaves have yellowed and fallen off. Seed fill is completed approximately 35 to 45 days after flower initiation. The crop is considered ripe and ready to swath when 30 to 40 percent of the seeds on the main stem have turned brown. If using a contact herbicide for desiccation, the recommended application timing is when 60 to 75 percent of the seed has turned brown, which is past the stage when swathing typically would be recommended.

Spring B. napus usually matures 85 to 110 days after planting, depending on hybrid and environmental conditions.

(Source: Harper and Berkenkamp. 1975. Revised growth-stage key for Brassica campestris and B. napus. Can. J. Plant Sci. 55: 657-678.)

Canola does well following cereal grains, most legumes or fallow in rotation. Research at various Research Extension Centers indicated that no yield difference occurs if canola is seeded after wheat or soybean. A preferred crop rotation would have at least two cropping years between canola plantings. However, if seeding canola after one year out, we recommend growing a hybrid that is resistant to blackleg.

Canola is susceptible to sclerotinia stem rot. White mold (sclerotinia) infection risk increases if canola is planted in tight rotation with other highly susceptible crops, such as sunflowers or dry edible beans. If planting canola within three years of susceptible crops, a fungicide application may be needed.

Less susceptible crops that could be planted successfully in a close rotation with canola are soybeans, flax, semileafless field peas or lentils. In years when ideal environmental conditions favor air-borne spore movement, all canola plantings without fungicide applied, regardless of rotation intervals, may have economic losses due to sclerotinia.

Canola is certain to shatter seeds, and volunteer plants are likely the next season. Cereals should follow canola to allow the use of certain broadleaf phenoxy herbicides for volunteer canola control.

Corn should not follow canola in a rotation because canola does not support arbuscular mycorrhizal fungi, which aid in corn phosphorous uptake. Also, avoid the production of canola and tame mustard on the same farm. A mixture of the two crops reduces the market value of both. In addition, conventional canola should not be planted on fields with heavy infestations of wild mustard. Roundup Ready, Liberty Link, Clearfield or SU canola can be planted to manage wild mustard in canola fields.

Herbicide carryover from previous crops can injure new canola seedlings. These include sulfonylurea (except the SU canola), imidazolinone (except Imi canola), triazine and PPO classes of herbicides. Always refer to the herbicide label information pertaining to crop rotation restrictions following their use.

Nitrogen (N), phosphorus (P) and potassium (K) requirements of canola and mustard are similar to those of small grains. Sulfur (S) requirements for canola are higher than most crops.

Soil cores should be taken from 0 to 24 inches deep and divided into 0- to 6-inch and 6- to 24-inch samples. P and K should be analyzed on the 0- to 6-inch sample, while N should be tested on each depth. The S soil test is nondiagnostic, so it should not be used to make a decision. The savvy grower will choose to not include S in the soil analysis suite.

All nutrient recommendations, including N, are not dependent on yield. A similar N rate is appropriate for greatest profit/yield in a poor as well as a good production season. Ammonium sources of N may be fall-applied on most North Dakota soils, except on sandy loam or coarser textures, or where flooding is expected in the spring. Spring application may be made preplant or at seeding.

Canola is very sensitive to fertilizer salts. No more than 5 pounds per acre (lb/a) of N is recommended with the seed in 12-inch row spacing for medium-textured soils, but the rate can be increased proportionally with narrow row spacing or increase in seed spread (see table labeled Maximum rates of seed-placed N + K₂O for canola).

General climate map of North Dakota with respect to canola production. In any given year, the line separating cooler, moister areas from warmer, drier areas may move east or west considerably.

P and K recommendations are shown in table labeled N, P and K recommendations. Canola is a good scavenger of P, and a row-starter fertilizer rate of 20 to 30 pounds of P₂O₅ per acre is sufficient for most soil test levels. On light soils where no N is recommended, 11-52-0 (MAP) would be a better seed-placed choice of phosphate because its N component is not as likely to injure seed as 18-46-0 (DAP).

Potassium, if needed, may be added to row starter if final N + K₂O is 10 lb/a or lower, using a double disc opener with 12-inch row spacing (see table labeled Maximum rates of seed-placed K₂O for canola). Broadcasting P and K is acceptable. However, I recommend a small amount of P as a row starter in addition to any broadcast application.

Canola has a special requirement for sulfur. The yield consequences of low soil S levels are very serious in canola production. Yield increases due to sulfur application have been demonstrated in North Dakota (see response table).

The S soil test is not diagnostic for canola or any other crop. Recommended sulfur application rates are 20 to 30 pounds of S per acre.

Canola takes up sulfate-S. The form of sulfur fertilizer may be ammonium sulfate (21-0-0-24S) or other sulfate fertilizers, such as ammonium thiosulfate or potassium thiosulfate. Elemental sulfur forms have not performed well in regional trials and are not recommended because these S sources have little value in the year of application.

Application of elemental S the previous year or previous fall has not proven to be of practical value due to low oxidation rates of S and high probability of sulfate leaching. Sulfur fertilizers are best applied in the spring and should not be applied in the fall.

Canola yield has not been higher with the application of any micronutrient in North Dakota.

A uniform stand of a competitive canola hybrid is the best weed control tool. Canola is not very competitive during early growth but becomes more competitive as it approaches the late-rosette and bolting stages. Competitive canola hybrids may allow a grower to reduce costs by spraying only once or, in some cases, not at all. Photographs of various weeds can be found in the back of this field guide.

• Some weeds are more competitive with canola than others. Canada thistle and wild oat are very competitive early in the growing season. High densities may require a split herbicide application. Research has shown a 400 lb/a yield advantage when wild oat was removed early (three-leaf canola) by PPI and/or POST herbicides, compared with late applications (six-leaf canola).
• NDSU studies have shown that a rotation of wheat followed by canola will reduce Canada thistle density. Suppress Canada thistle with pre- or postharvest glyphosate in the fall prior to seeding canola. Glyphosate (in Roundup Ready canola) and Stinger provide excellent Canada thistle control in canola. Consider a split application if Canada thistle densities are high (greater than one per foot2) or emerge before or about the same time as canola.
• Most postemergence herbicides used in canola can be tank-mixed with labeled pyrethroid insecticides. Be sure to consult the labels of all pesticides before mixing them.

The following information provides general details regarding herbicide rate, weeds controlled, when to apply, etc. This information does not supersede the herbicide label. Always read and follow label instructions.

The herbicides listed were registered for use as of the 2018 growing season. Illegal herbicide use may result in condemnation of the crop and possible fines.

A general rating for herbicide effectiveness on weeds and herbicide persistence in soil is provided in the following tables. Under favorable weather conditions, control may be better than indicated. Under unfavorable conditions, some herbicides rated as good may give erratic and unacceptable results. Also, dry and/or cool weather increases herbicide persistence while wet and/or warm weather reduces herbicide persistence.

Weed control ratings in this section are based on the following scale:

E: Excellent = 90 to 99 percent control
G: Good = 80 to 90 percent control
F: Fair = 65 to 80 percent control
P: Poor = 40 to 65 percent control
N: None = No control

Herbicide persistence ratings are for residues present 12 months after application:

O = Often      S = Seldom      N = None

Minimum interval between application and rain for maximum postemergence weed control in canola.

¹Or generic equivalent.
NCS = Next cropping season after herbicide application.
2CS = Second cropping season after herbicide application.
B or b = Bioassay. Do not plant until field bioassay indicates it is safe.
c = Do not use on soil with pH greater than 7.9.
d = Requires soil pH of 7.9 or less and a 34-month minimum rotation interval and 28 inches of cumulative precipitation.
e = Requires soil pH of 7.9 or less, 22 months and 22 inches of precipitation west of North Dakota Highway 1 or 34 months and 34 inches of precipitation east of Highway 1. These restrictions also apply to Ally Extra* at rates greater than 0.2 oz dry flowable (DF)/a.
h = Any rotational crop may be planted 120 days following application of Banvel* at 1.5 pt/a or less, excluding days when ground is frozen. For all crops and rates greater than 1.5 pt/a, allow 45 days per 1 pt/a of Banvel* excluding days when ground is frozen.
j = Requires 15 inches of cumulative precipitation during the growing season following application. An 18-month restriction applies to Accent*, Resolve*, Prequel and Steadfast applied above rates indicated or if drought follows application. Refer to label for crop rotation restrictions if rates greater than those indicated are used.
u = Must add two months if soil is frozen.
v = Do not include time when soil is frozen. For Verdict: All crops can be planted the spring following application.

Clearfield (imidazolinone resistant) canola hybrids may be planted the season after application. Conventional canola hybrids may be planted the following season after application at 1 pt/a in North Dakota counties of Cavalier, Pembina, Ramsey, Rolette, Towner and Walsh, and Minnesota counties of Kittson, Marshall, Pennington, Red Lake and Roseau.

Canola can volunteer for several years following a canola crop. These volunteers will compete with the succeeding crop and may affect yield, depending on volunteer density. Take steps during harvest to minimize canola seed loss.

Following the canola harvest, seeds that remain on or near the soil surface may germinate in the fall and be killed by frost. Avoid deep tillage that will bury canola seeds several inches deep, where they are less likely to germinate and secondary dormancy is more likely to be induced. If practical, allow time in the spring for canola volunteers to germinate, and control volunteers with shallow tillage or an herbicide application, then seed the new crop as soon as possible.

For no-till small grains, consider adding a labeled herbicide to the glyphosate burn-down application to control emerged glyphosate-resistant canola volunteers. Canola volunteers that emerge before or with the crop may be very large by the time a postemergence herbicide application is made.

Canola volunteers become much more difficult to control with herbicides once they reach the six-leaf to bolting stage. Some herbicides provide excellent control of small volunteers but poor control of bolting canola. Canola volunteers are best controlled when herbicide is applied by the five-leaf stage.

The following tables provide a general rating for herbicide effectiveness on volunteer canola. All postemergence herbicides were applied with recommended adjuvants. Volunteer canola control will improve where densities are lower and canola is smaller.

Volunteer canola control ratings in
this section are based on the following scale:

E: Excellent = 90 to 99 percent control
G: Good = 80 to 90 percent control
F: Fair = 65 to 80 percent control
P: Poor = 40 to 65 percent control
VP: Very poor = Less than 40 percent control
N: None = No control

The crucifer flea beetle, Phyllotreta cruciferae, is the main species of flea beetle that feeds on canola in North Dakota. The striped flea beetle, P. striolata, also feeds on canola. Flea beetles have a single generation per year.

Adult flea beetles overwinter in shelterbelts and leaf litter near last year’s canola fields. Adults start to emerge as temperatures warm up to 58 F. Populations will emerge slowly with cool temperatures or more quickly with warm temperatures.

The tiny, black crucifer flea beetles are about 1/10th of an inch long with a metallic bluish sheen, and move into the canola fields just as the seedlings are emerging. Striped flea beetles have two yellow stripes on their black wing covers and are comparable to the crucifer flea beetles in size.

Adults feed on the cotyledons and first true leaves of seedlings, and cause pitting and holes in the leaves. Damage is most serious to seedling plants and can cause seedling death and significant stand loss.

Eggs are laid in the soil and hatch in 12 days into larvae that feed on the roots of the canola plant. The larvae feed for three to four weeks, pupate for one week and then emerge as the new generation of adult flea beetles in mid-July to early August. These beetles feed on maturing crops by chewing green pods, stems and leaves for several weeks.

Fortunately, populations of summer flea beetles are rarely high enough to cause serious damage. However, extremely high populations feeding on green pods can cause pod shattering and seeds to remain green.

After feeding, adult beetles move into shelterbelts and other grassy overwintering sites. Large populations of the summer generation often indicate that flea beetle pressure may be high next spring.

Insecticides are the most effective control measure for flea beetles. The seedling stage is the most susceptible period, and insecticides are usually applied as a seed treatment prior to planting or as a foliar application to protect the crop from flea beetle feeding injury.

If growers use treated canola seed, seedlings should be protected from flea beetle feeding for most of the susceptible seedling stage. Seed treatments usually provide three to four weeks of protection against flea beetles.

Adult flea beetles emerge during a three- to four-week period in the spring. As a result, field monitoring is critical for any untreated or partially protected fields, or areas with a history of high flea beetle populations. Producers should inspect seedling fields daily for flea beetle injury. Check several locations in the field (for example, edges and center).

Warm (more than 58 F), calm, sunny weather increases feeding activity and movement, while cool, windy, damp weather slows feeding and favors crop growth. In some instances, flea beetles can move quickly and infest large fields by flying; in other instances, beetles invade slowly and walk from plant to plant in a field. Yellow sticky traps also can be used as monitoring tools to indicate when and how many flea beetles are moving into fields.

If the seed treatment did not provide adequate protection or was not used, an application of a foliar insecticide may be necessary. Foliar insecticide applications are recommended when 20 to 25 percent defoliation occurs on the cotyledons and true leaves and/or first true leaves, and beetles are actively feeding in field.

When flea beetle populations are high during hot, dry weather, populations can increase rapidly and fields can be re-infested due to flea beetle movement. Thus, insecticide seed treatment may not provide adequate protection against flea beetles, and a foliar application must be made quickly for effective pest management.

One of the problems producers face is being able to cover a large acreage quickly when flea beetle populations are high or re-infesting fields. Canola usually can compensate for flea beetle injury once it reaches the four to six true leaf stage.

The development of insecticide resistance in Phyllotreta flea beetles is a concern in canola because neonicotinoid insecticide seed treatments have been the primary insecticide used to control flea beetles for more than a decade. In Canada, the striped flea beetles are more tolerant (less mortality) of neonicotinoid seed treatments, compared with the crucifer flea beetles.

Research is underway in North Dakota to determine if neonicotinoid seed treatments still are effective against these two Phyllotreta species and if striped flea beetles are becoming more common than crucifer flea beetles in canola.

For more information, consult the NDSU Extension publication “Integrated Pest Management of Flea Beetles in Canola,” E1234.

The migratory diamondback moth usually arrives in late May or early June in North Dakota. The complete life cycle takes about 32 days from egg to adult. The moth has several generations during a single growing season, so all different life stages (eggs, larvae, pupae, adults) can be found in the field at the same time.

The adult is small, about ½ inch long, drab brown and at rest, the forewings of the male moth form three diamonds — hence the name diamondback moth. Females lay up to 160 eggs during the night. Eggs hatch in five to six days into pale yellowish-green caterpillars with a forked posterior end.

The newly emerged larvae burrow into the leaf and mine the leaf for several days to a week. Then the larvae exit the leaf and feed externally for another seven to 14 days. When disturbed, the larvae thrash backward violently and often drop from the plant on a strand of silk. The larvae pupate for five to 15 days in a white netlike cocoon attached to the leaves, stems or pods.

Larvae feed on the leaves, buds, flowers, seed pods, the green outer layer of the stems and, occasionally, the developing seeds. The amount of damage will depend on the crop stage and larval density and size. Extensive feeding on the flowers will delay plant maturity, cause flowers to abort and the crop to develop unevenly, resulting in reduced seed yield.

As leaves wilt and drop in late July to early August, larvae will feed on the stem, pods and developing seeds. Damaged pods will not fill completely and may shatter. Severely damaged pods appear whitish in contrast to the normal yellowing and browning of ripening undamaged pods.

Sex pheromone traps are useful tools for detecting the flights of the adult diamondback moth. The recommended trap design is the wing trap or delta trap with sticky inserts to capture moths. Traps should be suspended near the crop at the field’s edge. Traps provide an early indication of a possible infestation.

The moth usually has two to three generations in North Dakota. The second generation is the most important because it usually is present when the crop is most susceptible to injury (blooming to early pod). The third generation usually is too late to cause injury, except in the late-planted canola fields.

If high numbers of adults (greater than 100 moths per trap per week) are being captured in the traps during bloom to early pod development, monitor fields for diamondback moth larvae by beating plants to dislodge the larvae. After beating plants, count larvae on the ground or dangling from plants on a silk thread. Again, check several locations per field.

For the early flowering stage, insecticide applications may be justified at larval densities of 10 to 15 larvae per square foot (or one to two larvae per plant). The action threshold for canola at the pod stage is an average of 20 larvae per square foot (two to three larvae per plant). The best pest management strategies to avoid yield losses from diamondback moth include early monitoring of adults and larvae, and judicious use of insecticides only when fields are above thresholds.

A number of natural factors can affect diamondback moth populations negatively. For example, heavy rain can drown many first-generation larvae. Humid conditions associated with rainfall also can favor the development of fatal fungal diseases such as Entomophthorales. Several parasitic wasps and predators (flies, lacewings, minute pirate bugs, spiders and birds) prey on diamondback moth larvae.

For more information, consult the NDSU Extension publication “Diamondback Moth in Canola: Biology and Integrated Pest Management,” E1346.

The adult moth is about 1½ inches long and mainly gray-black with a silvery-whitish kidney-shaped spot and with a silvery-whitish fringe on each forewing. Moths emerge from the overwintering pupae in mid-late June and emergence continues through early August. These night fliers are particularly attracted to blooming canola fields for their nectar and egg-laying sites.

Eggs are laid on the lower side of leaves in clusters of 50 to 500 eggs in a honeycomb pattern and hatch in about one week. The emerging larvae (1/10th of an inch) usually are green. Mature larvae are about 1½ inch long and vary from green to brown to velvety black.

Larvae often hide underneath leaf litter and clumps of soil during the day, which makes them difficult to see. Larvae develop for six weeks and then drop to the ground in mid to late August to pupate. The moth has only one generation per year.

As the canola plant drops its leaves, the mature larvae (greater than ½ inch) begin to feed directly on the pods, which causes economic yield losses and premature shattering. Feeding injury by the mature larvae also accounts for 80 to 90 percent of the plant material consumed during a larvae’s life.

Mature larvae even will continue to feed on pods in the swath. Fortunately, populations are kept low during most years due to natural environmental factors such as harsh winters and a number of biological control agents (diseases and parasites).

Monitoring and Economic Thresholds – Knowing When to Control

Sex pheromone traps can be used to detect bertha armyworms in a general area. The recommended trap design is the green unitrap or bucket trap suspended above the crop canopy near the field’s edge. High trap catches generally indicate the level of larval populations to follow.

Fields should be monitored about two weeks after peak trap catch and scouted regularly to minimize crop losses. Check several locations per field, and continue scouting until an economic threshold is reached or the crop is swathed or direct combined.

The economic threshold will vary from one to three larvae per square foot

Higher-priced canola will take fewer larvae to reach the economic threshold. The key to controlling bertha armyworm is:

• Early detection of adult moths for their presence or absence and relative abundance in an area

• Monitoring fields for young larvae about ½ inch long

• Determining if fields are above economic thresholds for larvae

• Spraying fields above the economic threshold level once the hatch is complete and just before larvae move to the pods. Apply a well-timed insecticide in early morning or late evening, when larvae are actively feeding.

• Using high volumes of water for good coverage of the dense canola canopy.

For more information, consult the NDSU Extension publication “Bertha Armyworm in Canola: Biology and Integrated Pest Management,” E1347.

Lygus bugs consist of several species belonging to the genus Lygus. The tarnished plant bug, Lygus lineolaris, is one of the more common species and is known to feed on more than 200 host plants. Adult Lygus bugs are about ¼ inch in length, and pale green, light brown or dark brown with a distinctive triangular marking on the back.

Lygus bugs overwinter as adults in weedy areas and move into canola fields throughout the season. Adults lay eggs in the stems, leaves and flowers of host plants and then die. Immature nymphs hatch from these eggs. Nymphs are small, green and sometimes confused with aphids; however, Lygus nymphs are very active and move rapidly when disturbed, while aphids do not.

Several generations occur each year, with the second generation occurring in late July to early August. Hot, dry weather favors the buildup of Lygus populations and increases the risk of damage to the canola crop. Immature and adult Lygus bugs feed on growing points, buds, flowers and green pods.

Lygus bugs inject a toxic saliva with their piercing-sucking mouthparts during feeding, causing blasting of flowers or buds and shriveled seeds. Blasted flowers turn white within 24 hours and quickly fall to the ground. The small or damaged seeds are lost during harvest. In severe outbreaks in Canada, yield losses from bud blasting and seed damage have been estimated at 20 percent.

Scout for Lygus bugs just prior to bud formation until seeds within the pod have become firm. Lygus populations can increase suddenly. For example, when an alfalfa (preferred host) field is cut, Lygus bugs will migrate quickly into nearby canola fields, often in high numbers. Use a 15-inch sweep net and make 10 180-degree sweeps at several sampling sites.

The economic thresholds are: 15 Lygus bugs per 10 sweeps during bud stage through petal fall, and 20 Lygus bugs per 10 sweeps after petal fall and through pod ripening. However, if populations are high, control during the early pod ripening stage is usually the most economical.

Several species of cutworms feed on agricultural crops in the northern Great Plains, such as dingy cutworm, red-backed cutworm and pale western cutworm. Adult cutworms are moths about 1½ inches long with brown to gray wings, often with dark markings.

Cutworms have one generation per year. They overwinter as eggs or young larvae, depending on the species. Eggs hatch in April or early May, and young larvae (or caterpillars) feed at night on weeds and volunteer plants before the canola crop emerges. Larvae molt six times and grow larger each time. A mature cutworm larva is about 1½ inches long and as wide a pencil.

Cutworms are most noticeable in canola during late May through the first three weeks of June. After cutworms complete their development in late June, they burrow deeper into the soil and make a small pupal chamber. An adult moth emerges from the pupa in August to early September. Adults mate and females lay eggs on or just below the surface of loose, dry soil, weedy stubble or fallow fields, depending on the species.

Cutworm damage first appears on hilltops, south-facing slopes or in areas of light soil, which warm earlier in the spring. Larvae will cut young canola plants in the seedling to rosette stages. Cut plants can be found drying and lying on the soil surface. As damage continues, fields will have areas of bare soil where the canola has disappeared. In a severe infestation, the entire field can be destroyed.

Scout fields by looking for freshly damaged (cut off) plants. Dig 2 or more inches down around the cut-off plant and search for cutworm larvae. When disturbed, cutworms curl up or hide under soil debris. Canola is more susceptible to cutworm damage than small grains because cut plants do not grow back (grains compensate by tillering).

Three to four cutworms per square yard justify an insecticide treatment. Cutworm larvae are feeding actively at night, so an evening insecticide application is best. As a cultural control technique, weed-free fields and crusted summer fallow fields are less attractive to egg laying adults in late summer.

Grasshoppers are generalists and feed on a wide range of agricultural crops, such as small grains, flax and sunflowers. Grasshoppers overwinter as eggs, and nymphs start to emerge in late April to early May, with peak egg hatch in mid-June.

Nymphs (young grasshoppers) will go through five molts before transforming into adults. The length of time from egg to adult is 40 to 60 days. Adults of crop-damaging species become numerous in mid-July, with egg laying usually beginning in late July and continuing into the fall.

Eggs are deposited in a variety of noncrop areas including ditches, shelterbelts and weedy fall fields. Adults and nymphs feed on green plant material with their chewing mouthparts, creating holes on leaves or pods.

Grasshopper outbreaks usually coincide with several years of low rainfall and drought periods. Cool, wet weather increases the diseases that infect and kill grasshoppers. Scout canola for feeding injury from nymphs in the seedling stage, and in the pod development stage for adults.

Grasshopper damage is often concentrated on field edges and only the field edges will need to be sprayed. Grasshopper thresholds are based on the number of grasshoppers per square yard. Four 180-degree sweeps with a 15-inch sweep net equals 1 square yard. The infestation ratings are listed below in the table. A “threatening” rating would indicate a need to treat with an insecticide.

Several blister beetle species feed on canola, including Lytta nuttalli, a large purplish green beetle; Epicauta fabricii or the ash-gray blister beetle; and Epicauta ferruginea, a smaller rusty-colored, pubescent beetle.
Most blister beetle species have one generation per year.

Adults become active in early to mid-summer and lay eggs in the soil. Eggs hatch in about two weeks into larvae called triungulins, which actively prey on grasshopper egg pods (Epicauta spp.) or bee nests (Lytta spp.).

Blister beetles overwinter as larvae. Adult blister beetles are attracted to blooming canola fields, where they are ravenous feeders, devouring leaves, stems, flowers and pods. Blister beetles are mobile and gregarious, and often congregate in certain spots in a field.

In some instances, blister beetles feed for a short period of time and then migrate to other plants or fields. Alfalfa is an alternative host of blister beetles and they often move into canola fields when the alfalfa is cut. 

The presence of large numbers of blister beetles in spots of a canola field often has concerned producers. However, adult feeding generally is not significant enough to warrant an insecticide treatment. The “High Plains Integrated Pest Management Guide” recommends treatment when 10 adult blister beetles per plant are feeding on the flowers or pods. However, no economic threshold has been set in North Dakota. Spot treatment with foliar insecticides registered in canola is recommended.

Several species of aphids (cabbage aphid, turnip aphid, green peach aphid) infest canola and other plants in the mustard family. Individual aphids are small, approximately 2 to 4 millimeters in length, with a pair of tubelike structures called cornicles protruding from the back. Aphids on canola are usually pale green to grayish green and found in large numbers near the top of individual plants. Infested plants often appear shiny from the honeydew they secrete.

Most aphids migrate into North Dakota from southern states, and some may overwinter here. Aphids arrive in canola during late spring. As a result, late-planted canola may be more susceptible to heavy aphid infestations. Females reproduce asexually and give birth to live young within seven days.

As aphid populations build up and become crowded, winged adults are produced and disperse to begin new colonies. Multiple, overlapping generations of aphids occur within a season.

Aphids have piercing-sucking mouthparts that feed on the plant’s sap, inhibit terminal growth, stunt plant size and reduce seed yield. Aphid infestations often are localized within a field, and usually cause little damage if the infestations occur after pod development.

No thresholds have been established for aphids on canola. In most cases, spraying is not economical because aphids are on the top 2 to 3 inches of the plant, where pods are the smallest and contribute little to the overall yield. However, control may be justified when at least 20 percent of the stems are infested with a cluster of aphids in late flowering or early pod stages.

Scout field edges in upwind areas where aphids tend to be abundant. Note the presence of natural enemies as well as aphids. A treatment may be necessary when the following conditions are met: 1) canola was planted late; 2) plants are still in pod development; and 3) low populations of natural enemies, such as lady beetles, syrphid larvae or lacewing larvae, are present.

Aster Leafhopper (also called six-spotted Leafhopper) are greyish leafhoppers. The adults are 1/8 inch long, and have clear wings and six spots arranged in three rows between the compound eyes. Immature leafhoppers or nymphs are small, wingless versions of adults.

Aster leafhoppers migrate into North Dakota from the southern states and usually arrive in mid-May to mid-June. Leafhoppers are active, jumping and flying when temperatures are above 59 F.

The leafhopper uses its piercing-sucking mouthparts to feed on plant sap. The injury caused by aster leafhopper feeding produces localized necrosis or stippling.

Canola and other Brassica species (mustard) are not a preferred host for food and reproduction of aster leafhopper. It prefers to feed on wheat, barley and vegetable plants (lettuce and carrot), grapevines and herbs. Adults may move between host plants and follow what’s green and available.

Disease Vectors — These insects may be economically important in canola, but less because of their feeding injury and more because they are the principal vector of Aster yellows. Aster yellows has a very wide host range and causes plant disease and economic losses in several vegetables and ornamentals. Aster yellows is caused by a phytoplasm; an organism similar to a bacterium but without cell walls.

Aster leafhoppers acquire the phytoplasm by feeding on an aster yellow-infected plant for a minimum of 30 minutes. Acquisition of the phytoplasm increases with longer feeding times. The aster yellow phytoplasm requires another two weeks’ incubation period within the aster leafhopper before the leafhopper can transmit the disease to new plants.

Consequently, immigrating aster leafhoppers, arriving already infectious, are more likely to vector the disease into fields. However, once it acquires the phytoplasm, a leafhopper remains infectious for an extended period of time. Although the acquisition phase may be long, the leafhopper needs only a very short feeding period to transmit the disease to uninfected plants.

Symptoms of aster yellows in canola take about 21 to 35 days to appear, and sometimes plants are infected but show no symptoms. Flowers are distorted and sterile. Flowering portions of the plant may show excess branching and stunting (shortened internodes), resulting in a “witches broom” appearance. Aster yellow-infected canola plants often are taller than the rest of the plants.

Pods are replaced by inflated round or oval blue-green bladderlike structures. Infected plants produce little seed, but the percent of infected plants is usually quite low (less than 5 percent) most years.

Symptoms on canola can be confused with sulfur deficiency, anthocyanin production (purpling due to crop stress) and growth regulator herbicide damage.

The amount of aster yellows in fields would depend on the number of leafhoppers from local fields and immigrants from southern states, the percent of leafhoppers carrying the aster yellows phytoplasma and the time of arrival of the leafhoppers. Aster leafhoppers may feed on young canola plants at the beginning of the season and then move to grasses and more preferred crops as the canola matures.

Hot and dry conditions are not conducive to the spread of aster yellows. In contrast, abundant rainfall makes plants more succulent, which makes them more attractive to leafhoppers.

To manage aster yellows in canola:

1. Plant early to reduce the incidence of aster yellows by making plants less attractive (more mature) to migrating aster leafhoppers.
2. Control perennial weeds in fields because weeds can be a good source of the aster yellows phytoplasma.
3. Due to the mobility of aster leafhoppers, any insecticide spray program would be a “hit or miss” situation. Leafhoppers move around from field to field and crops to crops, and preventing aster yellows from being vectored would be difficult. As in wheat and barley, no economic threshold has been established in canola.
4. We also do not know how often we would need to spray. The residual of most pyrethroid insecticides is seven to 10 days. However, the economics of multiple and frequent foliar insecticide applications is not practical in canola as it is in high-value vegetable crops.
5. In addition, flowering canola is a major source of honey for honey bees in North Dakota, and any insecticides sprayed during flowering would be deadly for honey bees. We recommend not spraying insecticides on flowering canola to protect pollinators, and only spraying insecticides when bertha armyworms or other insect pests are at economic threshold levels in fields. If foliar-applied insecticides are necessary during flowering, spray in the early morning or late evening, when bees are not actively foraging.
6. Research conducted in snap beans and okra suggests that the neonicotinoid seed treatments should be effective in reducing leafhopper infestations in canola for up to 30 days.

Diptera: Cecidomyiidae

Swede midge is an invasive insect pest from Europe and Asia that was introduced into North America in Ontario, Canada, in 2000, and then first was found in the U.S. in 2004 in New York. It quickly spread in Canada into Quebec in 2003, Nova Scotia and Saskatchewan in 2007, and Prince Edward Island and Manitoba in 2008.

In the U.S., it occurs primarily in the northeast, including Connecticut, Massachusetts, New Jersey and Vermont. In 2014, swede midge was captured in pheromone traps near Winkler, Manitoba, which is close to the North Dakota border (just north of Walhalla), and near the major canola production area of northeastern North Dakota.

As result of this positive detection, NDSU Extension has conducted pheromone trap surveys for swede midge in 2015, 2017 and 2018 in the major canola production areas of North Dakota. Results of the trap surveys are all negative for swede midge in North Dakota so far.

The swede midge adult is a small brown fly about 1/16 inch long with sparse venation on wings. Swede midge is difficult to identify without a microscope and taxonomic expertise because of its small size and similarity to other midges.

The antennae and wing venation of swede midge are two characteristics used for identification. Antennae of males are distinctive, consisting of 12 antennal segments, each with two beadlike “nodes” separated by more slender, stemlike connections. Larvae are cream to yellow, and only about 1/16 inch long when mature.

Hosts of the swede midge include a wide range of species within the family Brassicaceae (or Cruciferae), and include the following: canola (Brassica napus, B. rapa), broccoli (B. oleracea var. italica), cauliflower (B. oleracea var. botrytis), cabbage (B. oleracea var. capitata), radish (Raphanus sativus) and cruciferous weeds (for example, wild mustard, field pennycress, wild radish).

Swede midge overwinter as pupae in the soil of canola fields and other hosts. In the northern Plains, three overlapping generations per year are common, with the first emergence of the overwintering generation occurring in late May and peak emergence in June. The second, or summer generation, emerges mid-July to early August, and the third generation emerges during late August into early September. The complete life cycle takes from 20 to 80 days.

Adults emerge in the spring from the puparium in the soil, and only live one to five days. Females lay 15 to 50 eggs in clusters on the youngest parts of the plant, such as flower buds or leaf stalk bases. Larvae emerge from the egg after three to 10 days. Larvae feed on plant tissue for two to three weeks, then jump or drop to the ground and spin cocoons in the top ½ inch of soil for pupation. The next generation of adults, except for the overwintering generation, will emerge two weeks later.

Swede midge can cause significant yield loss to canola plants during wet, warm weather, which allows the swede midge to emerge earlier, and increases its survival and ability to produce multiple generations. Adults only feed on nectar of flowering plants and do not damage the canola plant.

However, larvae secrete salivary fluids that are toxic to the plant causing abnormal growth and brown scarring of plant tissue (for example: deformed, twisted leaves or malformed growing points on the plant), gall formation and fused flowers that produce no pods. Damage easily can be confused with mechanical damage, heat or cold stress, fertility issues or herbicide damage.

Pest Management

For early detection of swede midge in canola grown in North Dakota, growers should monitor fields with pheromone traps and report any suspicious damage to Extension agents and/or specialists. The Jackson trap and pheromone lures are available commercially for swede midge monitoring from insect trap suppliers (for example, Great Lakes IPM).

Traps are placed low, about 12 inches above the ground, and at field edges from June through maturity. Check traps at least weekly and send any suspect midge samples to NDSU Extension Entomology or the NDSU Plant Diagnostic Laboratory for identification.

Although insecticides are registered for canola, control of swede midge is problematic due to its multiple overlapping generations that could infest canola throughout the season. Insecticide applications need to focus on peak adult populations based on pheromone trap monitoring, and preventing egg laying. No trap economic threshold has been established for monitoring swede midge in canola.

A cultural control strategy is early planting to ensure the crop is growing actively and vigorously by the time adult populations peak. Fall tillage of volunteer canola and cruciferous weeds also can help reduce overwintering populations. Because swede midge is a weak flyer, planting new canola fields about ½ mile away from previously infested fields can help lower infestation levels. However, this does not work well in areas with a high density of canola fields in major production areas.

Parasitoids and predators (for example, parasitic wasps, spiders, ground beetles) that attack swede midge have been recorded in Canada and may help reduce pest populations to some extent. Dry, cool weather also can limit population growth and range expansion, whereas moist, warm weather favors its development.

Alternaria brassicae and Alternaria japonica

Spots may develop at any time during the growth of the plant; however, they are more likely to occur during or after flowering and may be produced on leaves, branches and pods. These spots may be gray, gray with a dark border or black. Leaf spots often are surrounded by a yellowish halo.

Black spots may develop on stems and pods. Severely spotted pods may contain shrunken seeds infested with the fungus. Green seeds often are found next to deeply penetrating pod spots, resulting in green seed formation and low-test weight seed.

Black spot usually is less severe in B. napus (Argentine canola) than in B. rapa (Polish canola). Severely infected pods may split prematurely, resulting in shattering of seed.

Plants are infected directly from infested seeds or from spores produced on infected canola residue. The fungi also infect several weeds in the mustard family and survive on them. Frequent rains, fog or heavy dew and temperatures around 70 F during pod production and ripening favor severe outbreaks of black spot.

Buy certified, disease-free, germination-tested seed that has been cleaned properly. Most seed treatment fungicides effective against blackleg also will help control seed-borne black spot.

Use well-balanced fertilizer applications because stressed plants are more susceptible to black spot. When swathing, avoid a heavy swath; instead, choose a light swath that will dry more rapidly and combine as soon as possible.

Candidatus phytoplasma asteris

Distorted and sterile flowers are formed. Flowering portions of the plant may show excessive branching and stunting (shortened internodes), resulting in a “witches broom” appearance. Pods are replaced by inflated round or oval blue-green bladderlike structures.

Infected plants produce little seed, but the percentage of infected plants usually is quite low. An exception was noted in 1999 and 2012, when infection was noticeable in many fields in North Dakota and Minnesota, and economic losses occurred in a few fields.

The aster yellows phytoplasma does not survive the winter in North Dakota or Minnesota. It is transmitted by the aster leafhopper, Macrosteles fascifrons. The incidence of aster yellows depends on the number of leafhoppers that migrate in from overwintering areas to the south, the percentage of leafhoppers carrying the aster yellows phytoplasma and the time of arrival of the leafhoppers.

The disease rarely causes economic loss. Early planting may reduce the incidence of aster yellows.

Blackleg is caused by two fungal pathogens, Leptosphaeria maculans and L. biglobosa. Leptosphaeria maculans is highly aggressive and is the predominant cause of blackleg in North Dakota and Minnesota. In contrast, L. biglobosa is less damaging than L. maculans, and it is found in North Dakota and Minnesota infrequently.

The L. maculans pathogen can produce races that differ in their virulence to specific canola hybrids. These race isolates are classified as PG2, PG3, PG4 and PGT. In the 1990s, L. biglobosas was considered a mild-affecting race of L. maculans and was named PG1. However today, L. biglobosa is recognized as a distinct species but still can be referred to as PG1. Blackleg races predominant in the region can infect cultivars carrying resistance genes Rlm1, Rlm2 and Rlm3; isolates classified earlier as belonging to PG4 share this ability.

Blackleg can affect canola plants throughout their life cycle; however, economic losses occur mainly when initial infections take place before the plant reaches the four- to six-leaf stage. Earliest infection by the blackleg pathogen often is observed on cotyledon leaves as round to irregular-shaped lesions with a tan or buff color.

Leaf spots enlarge, and small black fruiting bodies called “pycnidia” are formed in the center of the lesions. Pycnidia are small but visible to the unaided eye and resemble ground black pepper flakes in color and size. On true leaves, lesions may be surrounded by a yellow halo.

Once pycnidia start forming, hyphae from the fungus grow inside the plant tissues toward the stem and then downward toward the crown. This growth occurs mostly without causing any visible symptoms. When plants approach the flowering stage, stem lesions become evident as sunken gray to dark gray areas that may be surrounded by a dark or black border.

Pycnidia frequently are produced in the center of stem lesions. As lesions enlarge, they may rupture and girdle the stem, producing the characteristic “blackleg” symptom. Infected stems also exhibit internal discoloration: when split open, the internal tissues of infected stems frequently are black, gray or stained with gray streaks.

Early infection of the stem may result in premature senescence and lodging. Late stem infection can cause plants to be less vigorous but otherwise causes few above-ground symptoms.

Roots, pods and seeds also are susceptible to infection. Infected roots develop internal discoloration; when split open, internal tissues will be black, gray or stained with gray streaks. Root infection causes premature senescence and yield loss but is not always associated with other above-ground symptoms. Infected pods may split open, resulting in seed loss, and seed produced in infected pods may be gray and shriveled.

The blackleg pathogen survives for several years as pycnidia or mycelium on infected crop residues. In the spring, during wet weather, pycnidia release a gelatinous ooze containing pink asexual spores called “conidia” that are dispersed by rain. The conidia are responsible for localized spread of the blackleg fungus, resulting in “hot spots” of infection.

In addition, the blackleg pathogen can produce pseudothecia, fruiting structures that produce sexual spores called “ascospores” on these residues. Ascospores can be dispersed for several miles, but most are deposited much closer to the source. Maximum ascospore discharge occurs the year following a canola crop but can occur until residue is degraded completely.

Temperatures in the 70s F and extended periods of canopy wetness favor infection. Disease development is inhibited by temperatures above 85 F or below 50 F. Plant injury from flea beetle activity, especially during the seedling stage, can increase the incidence and severity of blackleg.

The blackleg pathogen can survive for several years in infected seed. When infected seed is planted, seedlings emerge and may develop cotyledon, leaf and stem infections. Infections from seed can result in early and widespread epidemics. Transport of infected seed to other regions has contributed to the spread of blackleg throughout the world.

Resistance – Host resistance is one of the most effective and least expensive management tools for this disease. We recommend that producers plant different resistant hybrids to extend the shelf life of resistant genes currently available. Continuously utilizing the same resistant hybrid may speed up the development of races capable of infecting them.

We encourage growers to submit blackleg-infested canola stubble to qualified laboratories for identification of races present and then use that information to select hybrids with the appropriate resistance.

Crop rotation – Because the blackleg pathogen survives in crop residue for multiple years, we recommend a four-year crop rotation to most effectively manage the disease. In addition to reducing disease pressure, a longer crop rotation will reduce the likelihood that new races develop.

Disease-free seed – Plant certified disease-free seed. This is especially important when seeding canola in a new field.

Seed treatment fungicides – Numerous fungicide seed treatments are registered for blackleg control in North Dakota. Make sure to use a seed treatment that is effective against blackleg. Metalaxyl alone will not control blackleg. Consult the most recent issue of the “North Dakota Field Crop Fungicide Guide” (Extension publication PP622) when selecting a fungicide seed treatment.

Foliar fungicides – Take economics and disease risk into consideration before applying a foliar fungicide. High-risk factors for blackleg include a susceptible or moderately susceptible hybrid, the known occurrence of virulent races of the blackleg pathogen or a tight crop rotation. Fungicide trials demonstrated that a single fungicide application was able to reduce disease and provide some yield protection in susceptible hybrids but that foliar fungicides did not give adequate control.

If utilized, foliar fungicides should be applied at the two- to four-leaf stage. Further, because many foliar fungicides registered to manage blackleg have the same active ingredient as the seed treatments available, consider applying foliar fungicides with different modes of action than those in the seed treatment. Consult the “North Dakota Field Crop Fungicide Guide” (Extension publication PP622) for available products and always follow label directions.

Control weed hosts – Volunteer canola and wild mustard are hosts to the disease and should be controlled in crops rotated with canola.

This information was adapted from the publication “Blackleg of Canola,” (PP1367) and additional information and photographs can be found in that publication. Fungicide information can be found in the publication “Field Crop Fungicide Guide” (PP622).

Plasmodiophora brassicae

Galls and club-shaped growths appear on roots. The galls are firm and white, later becoming soft and gray-brown. Severely infected plants are stunted and may wilt. Areas of poor plant growth may develop and should be investigated thoroughly at the end of the season for clubroot.

Resting spores of the fungus survive many years in the soil. When soils are wet and susceptible roots are near, the resting spores germinate, producing swimming spores (zoospores) that infect the roots. The disease is spread via soil movement, including soil on equipment, and wind and water erosion. Commonly, this includes movement of soil from field to field during cultivation.

No satisfactory management exists. High pH soils are less conducive to clubroot than acidic soils. In acidic soils, liming with beet lime may reduce severity temporarily.

In North Dakota, the primary mechanism of spread has been through infested soil carried on farming equipment. Avoid introducing soil from infested areas. Always work the clubroot-infested fields last and sanitize your farm equipment after working in clubroot-infested fields.

Although on a yearly basis, the inoculum density is reduced by 50 percent, we recommend producers practice longer crop rotations. Resistant hybrids are available, but judicious use of that resistance will help prolong the effectiveness of the resistance; seeding the same cultivar not more than twice in a five-year period is strongly recommended.

The clubroot pathogen also is known to occur in British Columbia; Alberta; parts of Saskatchewan, Manitoba and Wisconsin; and eastern U.S. and Canada. Farm equipment obtained from those regions should be sanitized before use in North Dakota.

The clubroot pathogen also affects radish, broccoli, cauliflower, cabbage, rutabaga, Chinese cabbage, turnip and arugula, so use caution when using these plants in your crop rotation (for instance in a cover crop mixture) or planting them in your garden. Good weed control is important because clubroot also can infect (and survive on) wild mustard, shepherd’s purse, field pennycress and other brassica weeds.

Hyaloperonospora parasitica

A white, mealy growth develops on the lower surfaces of leaves. The upper leaf surface opposite the infected area turns yellow and could die. Symptoms can develop early in the life of plants and can continue after the canopy has closed; however, under North Dakota and Minnesota conditions, it rarely progresses above the third or fourth true leaves.

The downy mildew fungus is favored by cool, moist weather. It may occur in association with white rust in other regions of the world, and the disease complex usually would be more serious than either disease alone; however, in North Dakota, white rust is not considered a problem.

Downy mildew rarely is observed and has not been of economic concern in North Dakota. When disease is observed, it often is found on young seedlings in densely planted fields when conditions are very wet. As seedlings grow or temperatures rise, the disease usually disappears.

Fusarium oxysporum f.sp. conglutinans

The disease can be observed at almost any growth stage of the plants. Affected plants first will exhibit stunting and foliar discoloration and eventually may wilt, leading to premature death. Pods of infected plants may be reduced in size and have few to no seeds.

Fusarium wilt causes discoloration of the water-conducting vascular tissues in the upper roots and, when the disease is severe, in the lower stems; if a cross-section is cut through roots or lower stem tissues of infected plants, reddish-brown discoloration of the vascular tissues will be apparent. Yellow to brown streaking also may be observed along one side of the stem of infected plants.

The pathogen can survive in the soil as chlamydospores for several years in the absence of a host, but it also can survive infecting cruciferous weeds. Little is known about the infection process, but symptoms may be more severe under hot and dry environmental conditions.

This disease has not yet been observed in the U.S. Crop rotation with cereal crops may reduce inoculum levels in the field. Data collected in Canada have shown that differences in cultivar susceptibility exist.

The fungal pathogen Sclerotinia sclerotiorum causes disease in more than 400 different plant species, including canola, dry beans, soybeans, sunflowers, flax and the pulse crops (peas, lentils, chickpeas).

Symptoms on canola plants do not appear until after flowering. Spores of S. sclerotiorum colonize dead flower petals, and the first visible symptom of SSR is a mushy, light brown target pattern on and around cast petals (colored parts of the flower). Cast flower petals accumulate in the junctions where leaf petioles join the stem.

When one or more of these petals become infected with Sclerotinia, the infection spreads into the leaf petioles (slender stalk connecting the leaf to the plant), branches and stems. Infected areas become bleached, taking on the appearance of dry bone, and they frequently become brittle and shredded. If infection occurs in the main stem, plants may die early and become prone to lodging.

Standing dead plants may be visible in fields with severe SSR infection. After infection is well-established, hard, black structures (sclerotia), which resemble rat droppings, may be produced in infected tissues. The sclerotia are the survival structures of the pathogen and, although they are not produced in every infection, they are a definitive sign of SSR.

The pathogen produces sclerotia (resting structures) in infected stems. Sclerotia fall to the ground during harvest, and they can survive on or in the soil for many years. After the sclerotia have overwintered, 1 to 2 inches of rain in a one- to two-week period will moisten soil enough for sclerotia to germinate and produce small mushroom structures (0.1 to 0.25 inch diameter) called apothecia. If rain does not occur prior to flowering, the soil will be too dry for sclerotia to germinate, no apothecia and spores will be produced, and infection will not take place.

Sclerotia must be in the top 1 to 2 inches of the soil profile to form apothecia; otherwise, they will remain dormant. Apothecia can produce millions of spores (ascospores) that are dispersed by wind; however, most ascospores remain trapped in the canopy near the apothecia. The great majority of spores do not travel more than 150 feet, but some spores will be disbursed significantly farther when the right conditions persist.

Spores can infect floral parts directly, but most infections occur when spores land on dead or dying tissue. Canola petals that die after flowering provide an excellent food source for the SSR spores, and most SSR disease lesions are initiated from infected dead flower petals. If spores are produced, the canopy must be wet for 48 hours or more for the spores to germinate and for infection to take place.

However, even when ample moisture is available, if temperatures are too hot during this time (above 86 F), spores may die before they can begin to grow. If infection occurs, disease development will not occur above 86 F, but it will resume once cooler temperatures return. Similarly, if weather becomes dry, disease progression will slow and increase only when the canopy becomes wet.

Sclerotinia Disease Cycle

Crop rotation – Crop rotation is important to limit the buildup of sclerotia. Try to avoid more than one highly susceptible crop (sunflower, canola, crambe, dry bean, pea or soybean) in a rotation. Semileafless peas support less buildup of sclerotinia than the vining types and may be acceptable in crop rotations. In irrigated trials at Carrington, N.D., no sclerotia were produced in flax; therefore, flax may be a relatively safe crop in rotation with canola. Additionally, corn and small grains are not susceptible to SSR. A preferred rotation would have canola planted after two or more nonsusceptible crops. Despite appropriate crop rotation, some sclerotia can survive for long periods of time and can germinate and cause infection many years after they were produced. Additionally, infection can occur from spores blown in from other fields.

Tillage – Management of SSR with tillage is uncertain. Deep tillage may bury sclerotia deep enough that they do not produce apothecia. However, some data indicate that these sclerotia do not break down as fast as those near the surface and may result in infection if they are brought back near the surface later. Additionally, even if deep tillage is done, it probably is of limited value if used on one or a few fields and not areawide.

Resistance – All canola hybrids are susceptible to some degree. However, some cultivars may be less susceptible than others.

Biological control – The fungus Coniothyrium minitans (sold commercially as Contans WG) can attack and kill sclerotia in the soil. Research conducted at Carrington, N.D., showed that fall application of Contans was effective in reducing viability and vigor of sclerotia. Contans may be of greatest use under high disease pressure or when using cropping practices such as irrigation. Another biocontrol product registered for use in North Dakota (sold commercially as Serenade ASO) is Bacillus subtilis strain QST 2808. Serenade ASO is labelled for spray or fungigation application. Additional biocontrol products likely will become available. Consult the most current “North Dakota Field Crop Fungicide Guide” (PP622) for more information.

Fungicides – Numerous fungicides with diverse modes of action are available, and new products likely will be available in the future. To find products available, consult the most current “North Dakota Field Crop Fungicide Guide” (PP622). Fungicide trials are done frequently at NDSU’s Research Extension Centers in Carrington, Langdon and Minot (North Central REC). For the most current information, visit their websites.

Making the decision to apply (or not to apply) fungicides and spraying at the most appropriate growth stage are the two keys to managing SSR. Several resources can be used when determining whether a fungicide application is warranted, including the Sclerotinia Risk Map, Sclerotinia Risk Calculator and Sclerotinia Stem Rot Checklist. In general, spraying is most beneficial when:

• One to 2 inches (or more) of rain occurred in the two weeks prior to flowering
• Rain or high humidity is expected
• Sclerotinia has been a problem in recent years

Sclerotinia Risk Map

Environmental information generated from the North Dakota Agricultural Weather Network (NDAWN) weather stations are used to determine if conditions are favorable for the germination of sclerotia, development of apothecia and spore release. This information is used to generate a map with color-coded indications of risk throughout North Dakota and the canola-growing areas in Minnesota.

The map is an effective tool to help growers assess their risk of SSR infection. However, producers should keep several things in mind:

• The risk map only applies to you when your canola is flowering. Because the pathogen uses petals as a food source, no infection occurs prior to flowering.
• The map is meant as a regional guide to risk. The Risk Map is only as good as the information source, and even though NDAWN has more than 90 weather stations, the environmental conditions in your field may be different. If you know you have had more/less rain than the NDAWN stations nearest you, your risk may be higher or lower.
• The Sclerotinia Risk Map calculates the likelihood of sclerotia germinating and, thus, spores being available to cause infection. Your fields may have greater or less risk, depending on other management techniques (crop rotation, less susceptible hybrid), field history and weather forecasts (for example, rain, high humidity, temperatures).

The map is available or at this site.

Sclerotinia Risk Calculator

The Risk Calculator is an interactive tool that produces field-tailored estimations of risk using information such as the fields’ nearest NDAWN station, crop rotation and history of Sclerotinia diseases.

The Sclerotinia risk map and risk calculator are available at www.northerncanola.com and www.ag.ndsu.edu/sclerotinia/. These tools also are available through the NDSU Canola Doctor app for smartphone devices. The app can be downloaded from the Google Play Store or Apple App Store for installation on android and iOS devices, respectively.

If you decide to make a fungicide application, remember that the best time to spray is when the plants are between 30 and 50 percent bloom. Fungicide applications after canola has reached the 50 percent bloom stage usually do not provide an economic return.

Resources and Citations

This information was adapted from the publication “Sclerotinia of Canola” (PP1410), and additional information and photographs can be found in that publication. Fungicide information can be found in the publication “Field Crop Fungicide Guide” (PP622)

Read each question and circle the point value assigned to the answer you choose. Count up the points for each question and enter the total for each section. Answer all the questions in this section.

Section one:

If you scored 60 or more in this section, you probably had sclerotinia stem rot in your canola crops. Proceed to section two with a 60 or more score.

Section two:

If you had a high score in section one and more than 50 for section two, you should consider applying a fungicide to protect your crop against sclerotinia stem rot.

Section three:

If you scored 50 or higher in section three, along with high scores from the first and second sections (60 and 50 plus respectively), it may be worthwhile to protect your crop against sclerotinia stem rot. If you scored less than 50 in the last section, it is not likely worth applying a foliar fungicide.

Permission by the Canola Council of Canada to reproduce this checklist is gratefully acknowledged.

Albugo candida

This disease commonly occurs on B. rapa (Polish canola), as well as on brown and oriental mustard (B. juncea), and is of no economic importance in North Dakota and Minnesota. The most obvious symptoms of white rust are the swollen flowering stems and pods. These swollen tissues may be spiny, resulting in the name staghead.

The stagheads initially are green but later turn brown. In wet weather, powdery white spore masses develop on the stagheads, stems and lower surfaces of leaves. Severely diseased leaves may turn yellow and drop. Yellowing leaves may develop “green islands” of tissue over each pustule.

The fungus, Albugo candida, survives as thick-walled spores in the stagheads, many of which fall to the ground during harvest. Many stagheads are broken during combining, resulting in the release of the spores. The following spring, these thick-walled spores germinate and infect the cotyledons and leaves of young plants. Secondary spread from cotyledons and leaves to the flower parts occurs from a second type of spore.

All B. napus (Argentine canola) hybrids are resistant to white rust. Most B. rapa (Polish canola) hybrids are susceptible, but a few newer hybrids are moderately susceptible, moderately resistant or resistant. Do not plant canola on canola if susceptible hybrids are being grown. Control volunteer canola and wild mustard early in the season because the fungus can reproduce on the cotyledons and young leaves.

Swathing canola at the optimum stage of ripening reduces green seed problems and shatter losses, and ensures the quality required for top grades and prices. Inspect fields every two to three days when color change starts to occur in the first-formed pods on the bottom of the main stem.

To determine when a field of canola is ready to swath, examine plants from different parts of the field. The stage of maturity in an evenly maturing field will vary from plant to plant and from area to area within the field. When examining the plants, take into account varying soil types, low-lying areas, available soil moisture and exposed early ripening areas.

Examine only pods on the main stem. Seeds in pods on the bottom third of the main stem were formed earlier and will turn color much sooner than seeds in the pods on the top third of the plant.

Swathing without a negative effect on yield or seed quality can start when about 60 to 70 percent of the seeds in pods on the main stem will have changed color or have started to change color. Seeds with only small patches of color should be counted as color changed.

Remember, the color of the seed is more important than the overall color of the field in determining the stage of maturity. Most of the seeds that have changed color will be from the bottom third of the main stem. When seeds in the bottom pods turn color, seeds in the top, last-formed pods are filled or nearly filled and still green.

Seeds in all pods on a plant complete filling (physiological maturity) at about 40 percent moisture and then slowly turn from green to light yellow or reddish brown, brown or black, depending on the hybrid. In hot (90 F), dry weather, canola seed can go from 10 to 50 percent seed color change in just three to five days or less. Once filled, seeds rapidly lose moisture at about 2 to 3 percentage points or more each day, depending on the weather.

Swathing early can be beneficial if a hard fall frost is expected. Frost fixes the chlorophyll or green color in immature seed, making it difficult to remove during processing. Early fall frosts rarely freeze to ground level. A swathed crop will not only lie below the coldest night temperatures, but much of the seed will benefit from the insulating properties of the swath and residual soil heat, preventing or reducing frost-fixed chlorophyll.

Another sign of the canola being very near the swathing stage is the natural yellowing and senescence of leaves and leaf drop. When canola plants consist only of stems, stem branches and pods, the crop probably is very near the optimum time for swathing. Swathing can begin in B. napus (Argentine canola) at 15 percent seed color change. B. rapa (Polish canola) should be left until 20 to 25 percent seed color change.

The swather should be run just low enough to get all the seed pods, leaving the maximum amount of stubble to anchor the windrow and ensure adequate air circulation through the windrow. Most stubble height varies from 10 to 12 inches in canola fields after swathing.

The canola must flow smoothly through the swather without bunching. Bunching leads to uneven drying and combine plugging. Therefore, a swather for canola must have enough depth of table (40 inches) to handle the crop material.

It also should have a large throat opening at least as wide (40 to 54 inches) as the distance between the two swather canvasses on center-delivery swathers. It should have a vertical clearance for the windrow of at least 30 to 40 inches.

The table canvas should be strong enough to carry the heavy load of material cut and should be run just fast enough to keep the table clean. If possible, canvas speed should be varied, depending on the maturity of the crop cut. A fast canvas tends to produce a hollow, twisted windrow; a slower canvas produces a more compact windrow, but it may bunch and sit high on the stubble. Increase the canvas speed until the windrow is pressed into the stubble.

The reel should be set as high and as far forward as possible. The reel speed should be set to correspond with the forward speed of the swather. Finger reels work best in canola to help bring the material back onto the table and gently handle the ripened canola. For a lodged or leaning canola crop, finger reels are highly recommended for ease of swathing.

Ordinary end dividers, which are long and gently sloping, generally are less prone to plugging than short, abrupt types. When the crop is tall, tangled and lodged or laid across the seeded rows, divider plugging is almost inevitable unless special vertical cutter bars or power blades are fitted on the swather. These can cause minor loss of pods and whole seed tops, but they prevent stops and bunching.

In badly lodged crops, swathing in a direction parallel to the direction in which the crop is leaning may be advantageous. In areas where windrows could be lifted and blown by the wind, a light roller pulled behind the swather will help anchor the windrow in the stubble.

The roller should be set so that it just anchors the windrow into the stubble without shelling any ripe pods. Excessive roller pressure will produce a windrow that is too compact to dry quickly and will be difficult to pick up without shelling the canola. NDSU research has shown that swath pack density and seeding rates had little effect on green seed of canola.

Swathing late, when seed moisture content is much lower (about 80 percent mature seed color), will result in fluffy windrows susceptible to blowing and increased shattering. To reduce shattering losses, overripe fields should be swathed when humidity is high, such as after a rain or heavy dew, or at night.

Determining when to swath unevenly maturing fields is difficult. When checking uneven stands, a producer should do an early count on the ratio of early emerged canola, which is bolting or starting to flower, to the late-emerged flush of young, more immature plants. Knowing the ratio of early to late-emerged canola plants allows producers to make a better decision on how soon to swath or to wait until the later crop catches up. If the stand is 20 to 25 percent early and 75 to 80 percent late, then waiting to cut until later may be the best strategy to reduce the amount of green seed.

Canola should be allowed to cure and ripen from 10 to 14 days in the swath before combining. If combined too early, the chance of increased green seed in the harvested crop is much greater.

While starting on the early side is better for swathing, the same doesn’t necessarily hold true for combining. Hot or windy weather at or after swathing can cause canola seed to be at the appropriate moisture content for combining before it has cured and cleared the green chlorophyll. This occurs because the plant dries before sufficient moisture can move into the seed to finish curing it.

Canola requires at least 20 percent moisture in the seed for the maturing process to take place and eliminate the green seed color. Checking moisture content and green seed count before starting to combine is important. Delayed combining can help clear the green color, particularly if the swath sits through several heavy dews or light rain showers.

Growers can make management decisions to reduce green seed problems:

• Choose fields with better surface drainage and fertility.
• Seed as early as possible in the spring to allow for the maximum ripening time.
• Provide a firm seedbed to achieve the correct depth of planting and good seed-to-soil contact for rapid and even emergence. Do not broadcast seed.
• Swath at the recommended color stage for the weather conditions.
• Maintain adequate fertility levels for canola growth and ripening. Canola stressed from sulfur deficiency will not mature evenly.
• Take soil samples for a general indication of N, P and K.
• Sample plant tissue early during the rosette stage to allow time for corrective sulfur applications.
• Fields with high fertility levels can be expected to delay maturity in years with below-normal growing degree day (GDD) accumulations or heat units (cool years).
• Don’t swath canola if the weather forecast is for extremely hot, dry and windy conditions.
• All canola management decisions should be targeted toward uniform crop maturity.

Most combines work fairly well to harvest canola. Combines should be checked thoroughly before starting on canola. Cover any holes or worn spots in the table/platform or within a combine with duct tape or caulking compound. Leakage can occur easily in the stone trap or top feeder housing, or through lower inspection doors.

The travel speed of the combine should be equal to that of the pickup so a gentle lifting of the swath occurs without tearing or pushing. Set the pickup to rub just under the swath. Cylinder speeds will depend on canola crop conditions.

Speeds of one-half to two-thirds of that used for small grains often are used for canola. The speed should be just fast enough to break open the pods. Speed reduction is important to prevent overthreshing of pods and stems and overloading the sieves.

Cracked canola is caused by impact when the cylinder speed is too fast. Examine the threshed seed for cracked canola. Push your arm into the seeds and observe if cracked canola seed pieces stick to your skin or hair on your arm. Reduce cylinder speeds if excessive cracking occurs.

Fan speed should be set low to avoid blowing canola seed out with the chaff. This will allow large amounts of pods in the return. Start with a low fan speed and increase gradually until the separation of chaff and seed occurs with no canola being blown over the chaffer sieve.

The type of canola affects the aeration system design, operating static pressure, airflow and fan selected. The resistance to airflow (static pressure) of B. napus (Argentine canola) is about twice that of wheat, and for B. rapa (Polish canola), it is about three times that of wheat.

Fans must be selected to operate at the appropriate static pressure. For example, to obtain an airflow rate of 0.2 cubic feet per minute per bushel (cfm/bu) in a 36-foot-diameter bin filled 24 feet deep with B. napus (Argentine canola), you would need a 4.5-horsepower fan operating at 4.4 inches of static pressure.

The B. napus (Argentine) is the most common type of canola grown. Obtaining that same airflow rate through B. rapa (Polish) canola requires a 6.8-horsepower fan operating at 6.6 inches of static pressure. If a 5-horsepower low-speed centrifugal fan is placed on the 36-foot-diameter bin filled 24 feet deep with B. napus (Argentine canola), an airflow rate of 0.23 cfm/bu is obtained. However, if the same fan is placed on the bin of B. rapa (Polish canola), an airflow rate of only 0.18 cfm/bu is obtained.

Natural air/low-temperature drying with an airflow rate of 0.75 cfm/bu will dry canola at moisture contents up to 12 percent. An airflow rate of 1 cfm/bu permits drying canola with initial moisture contents up to 13 percent. Higher airflow rates are not economical due to the large resistance to airflow through canola.

Based on average North Dakota climatic conditions, canola would be expected to dry to about 8 percent in August, 9 percent in September and 9.8 percent in October. The canola moisture content is expected to be lower due to the fan warming the air about 5 F. With this warmed air, the expected moisture contents are: August, 6.6 percent; September, 7.8 percent; and October, 8.2 percent.

Supplemental heat may be added if canola does not reach the recommended storage moisture content using natural air drying. Do not warm the air more than 5 F or the canola will be overdried. The estimated drying time in September using an airflow rate of 1 cfm/bu is about 25 days, and with an airflow rate of 0.75 cfm/bu, it is about 35 days.

High-temperature dryers work for drying canola, but check the screen size on column dryers because most are equipped with a standard-size perforation of 3/32 inch through which canola seed may pass or lodge. Check with the dryer manufacturer before making any changes to be sure fan operation will not be affected adversely.

Drying temperatures need to be limited during high-temperature drying. At moisture contents up to 12 percent, a drying temperature of 180 F can be used with dryers that mix the seed as it is dried. At moisture contents exceeding 12 percent, the dryer temperature needs to be limited to 160 F, even with mixing occurring in the dryer. Without mixing, the dryer temperature needs to be limited to 140 F.

Canola that is used for seed should be dried at temperatures less than 110 F. If the moisture content is above 17 percent, drying the kernels in two passes is advantageous. Too much moisture extracted at one pass can lead to shriveling and cracking of the seed, as well as limit the drying performance.

Overdrying causes cracking of the seed coats. Damaged seeds undergo a marked rise in the level of free fatty acids, causing a reduction in oil quality. Canola offers more resistance to airflow than cereal grains, so airflow and the drying rate will be reduced. Chaff may interfere with the flow of canola in a dryer, which may lead to an overdried area and the potential for fires.

Fire risk when drying canola may be reduced by cleaning the seed to remove light or fine material before drying, removing accumulations of debris from the walls and other areas of the dryer, using wind deflectors to prevent drawing airborne material through the burner, and avoiding overdrying the seed. Frequently check the dryer and periodically clean it to reduce the fire hazard.

Operate augers at full capacity and moderate speeds to avoid excessive damage to the seed. Drag conveyors have been used successfully, but experience with canola suggests that the spacing between paddles should be reduced to maintain satisfactory capacity.

Pneumatic conveyors can handle canola adequately but may have difficulty in feeding and discharging the seed. Kernel damage usually is not a problem for most equipment unless the moisture content is below 7 percent.

U.S. sample grade - canola that:

• does not meet the requirements for U.S. Nos. 1, 2 or 3, or
• has a musty, sour or commercially objectionable foreign odor or
• is heating or otherwise of distinctly low quality

¹Conspicuous admixture is all matter other than canola that is readily distinguishable from canola and remains in the sample after the removal of machine-separated dockage. It is not limited to ergot, sclerotia and stones.

²Inconspicuous admixture. Any seed that is difficult to distinguish from canola. This includes, but is not limited to, common wild mustard (Brassica kaber and Brassica juncea), domestic brown mustard (Brassica juncea), yellow mustard (Brassoca hirta) and seed other than the mustard group.

Factors of most importance in the determination of grades are admixtures and soundness. Grading admixtures include factors such as foreign material, common wild mustard seed, tame brown and yellow mustard seed, earth pellets, sclerotia, ergot and stones. Soundness refers to broken seed not assessed to dockage, seeds distinctly green after cracking, heat damage and odor.

Source: Federal Grain and Inspection Service, USDA.

Duane Berglund, Agronomist Emeritus, NDSU Extension
Patrick Beauzay, Entomology Research Specialist, NDSU Entomologist
Venkat Chapara, NDSU Research Plant Pathologist
Kishore Chittem, NDSU Research Plant Pathologist
Dave Franzen, Soil Science Specialist, NDSU Extension
Kenneth Hellevang, Agricultural Engineer, NDSU Extension
Brian Jenks, Weed Scientist, NDSU North Central Research Extension Center
Hans Kandel, Agronomist, NDSU Extension
Clair Keene, Agronomist, NDSU Extension
Janet Knodel, Entomologist, NDSU Extension
Lesley Lubenow, Agronomist, NDSU Extension
Samuel Markell, Plant Pathologist, NDSU Extension
Luis del Rio Mendoza, NDSU Plant Pathologist
John Nowatzki, Agricultural Machine Systems Specialist, NDSU Extension

R. Zollinger, Weed Specialist Emeritus, NDSU Extension
M. Wunsch, NDSU Research Plant Pathologist
G. Endres, E. Eriksmoen, B. Hanson, M. Ostlie, J. Rickertsen, G. Pradhan and B. Schatz, NDSU Research Extension Center and Extension agronomists

Complete list of NDSU Extension contacts
Complete list of NDSU Research Extension Centers

Hybrid Trial Results – Canola
NDSU Extension canola publications
Sclerotinia Risk in Canola Forecast Program
Canola Insects Information

Hybrid Trial Results - Canola

Northern Canola Growers Association
125 Slate Drive, Suite 4
Bismarck, ND 58503
Phone: 701-223-4124
Toll free: 877-585-1671 Fax: 701-223-4130
Email
Web address

Minnesota Canola Council
4630 Churchill St., Suite 1
St. Paul, MN 55126
Phone: 651-638-9883
Email

U.S. Canola Association
600 Pennsylvania Ave. S.E., Suite 320
Washington, D.C. 20003
Phone: 202-969-8113
Email
Web address

Canola Council of Canada
400 - 167 Lombard Ave., Winnipeg, MB R3B 0T6
Phone: 204-982-2109
Toll free: 866-834-4378
Web address

Manitoba Department of Agriculture and Food
Web address

Canola Growers Manual, Canola Council of Canada
Web address