Chapter 9 - Soil Fertility

Soil Fertility and Canola Nutrition

Profitable canola production relies heavily on adequate plant nutrition, which in turn is affected by management of soil fertility. In addition, the nutritional level of the plant will affect the crop response to stress factors such as disease and adverse weather. Balanced, effective fertilizer management not only contributes to profitable canola yield but also helps to maintain the productivity of the soil resource.

Basic Plant Nutrition

The living plant depends on a number of basic factors for normal growth:

• light
• air
• water
• nutrients
• physical support

Soil plays an important role in all these factors except for light. If any of these basic factors are limiting, plant growth will be reduced or the life cycle may not be completed– this is called the principle of limiting factors. In other words, plant growth potential is limited by the factor in shortest supply.

Yield may be reduced when one nutrient reaches excessive levels that cause toxicity, so the proper balance of nutrients is important. Also, other factors such as improper management or pests can lower yield. Therefore, a systems approach is necessary to integrate all the factors in the best combination to achieve the most economic yield.

Essential Plant Nutrients

The plant’s mineral composition does not simply reflect the elements needed for growth. Plants can selectively absorb required elements for their growth. But they also can take up elements not needed for growth.

The terms essential plant nutrient or essential mineral element were formed to describe the minerals needed by plants to grow and complete life cycles. Essential plant nutrients must be directly involved in some aspect of the plant metabolism such as structural material, enzymes or hormones, and they must not be totally replaceable by another mineral element. For higher plants such as canola, there are 14 essential nutrients (besides CO₂, oxygen and water).

Table 1. Essential Plant Nutrients

Macronutrients	N, P, K, S, Mg, Ca
Micronutrients	Fe, Mn, Zn, Cu, B, Mo, Cl and Ni

Table 1 also indicates that plant nutrients are often classified by the relative amounts needed. Macronutrients are needed in large amounts relative to micronutrients. Table 2 shows the relative amounts of nutrients contained in a typical canola crop. Some nutrients can be accumulated in plants much higher than necessary for growth.

Table 2. Approximate Amounts of Nutrients in the Above- Ground Portion of a 1,960 kg/ha (35 bu/ac) Canola Crop

Element	kg/ha	lb/ac
Nitrogen (N)	112-134	100-120
Phosphorus (P)	1-28	15-25*
Potassium (K)	67-134	60-120*
Sulphur (S)	22-28	20-25
Calcium (Ca)	45-67	40-60
Magnesium (Mg)	13-20	12-18
Iron (Fe)	~1	~1
Chlorine (Cl)	~0.8	~0.7
Manganese (Mn)	~0.2	~0.2
Zinc (Zn)	~0.2	~0.2
Boron (B)	~0.2	~0.2
Copper (Cu)	~0.7	~0.06
Nickel (Ni)	~0.004	~0.004
Molybdenum (Mo)	~0.004	~0.004

* P X 2.3=P₂O₅; K X 1.2=K₂O
**Crop uptake of nutrients is greatly affected by conditions in the soil or
weather (dry, wet, cold, compaction, nutrient imbalances, salinity, etc.).

General Nutrient Uptake

The following discussion outlines nutrient movement into and through the canola plant. The level of most nutrients in the plant sap is much higher than in the water surrounding the roots. For example, a typical N content in a canola plant at the rosette stage would be 5 to 6% N, whereas a fertile soil in the spring would contain about 0.0002% N in a plant available form on a dry weight basis. The N level in the soil solution would be in the range of 0.00002%. Therefore, plant nutrient uptake must be highly selective.

Nutrient uptake begins when plant available forms move from the soil water through pores in the root skin (exodermis) into the free space of the roots. This free space comprises about 5 to 10% of the root’s internal volume. This movement is a passive process (doesn’t require energy from the plant) driven either by diffusion (movement due to differences in concentration) or mass flow (simply carried by water flowing into the roots). The movement is selective since pores into the free space act as a size filter. Many nutrient ion diameters are much smaller than the pores. For example, potassium and calcium are only 10 to 20% of the pore size, and have easy access to the free space. Large diameter substances such as metal chelates, viruses and fungi are restricted from entry by the small pore size.

As plant roots grow, the soil volume and surface area explored increases, which increases the capacity for nutrient absorption. In addition, roots possess a cation exchange capacity (CEC) due to negative charges in cell walls. This root CEC attracts positive ions (cations) like ammonium (NH₄+) but repels negative ions (anions) such as nitrate (NO₃-). (For details on CEC, see section “Soil Properties that Affect Plant Nutrition.”)

After entry into the free space, nutrients move into the cell interior by crossing a plasma membrane found on the inside of cell walls. Another similar membrane is found surrounding a large central storage compartment (vacuole) that usually fills more than 80% of the total cell volume. The plasma and vacuole membranes are effective barriers and are the main sites for nutrient uptake selectivity. These membranes contain carrier systems or ion pumps that transport certain nutrients. Such systems are called active since they require energy from the plant to work. This energy demand for ion uptake by roots is considerable, taking up to 1/3 of the energy during rapid growth. The energy for root activity arises from respiration, which requires carbohydrates and oxygen. This explains why nutrient uptake often stops in flooded soils—there is a lack of oxygen.

Some active uptake systems are constant while others have a rate that can be regulated. As the plant level of nutrients and related compounds increases, the root uptake rate can decrease (negative feedback). In contrast, as plants build tissue, the level of nutrient “building blocks” decreases, and the roots are signalled to increase the uptake rate (positive feedback) for nutrients. Passive ion channels through the membranes allow for selective nutrient movement.

The selectivity of the various transport systems across the membranes is not absolute. There is often competition between ions of similar size and charge. For example, chloride (Cl-) competes with nitrate (NO₃-). This competition between Cl- and NO₃ - is important in certain saline soils with Cl- as a major component of the salt. Most prairie soils contain salt with sulphate as the main anion.

Since cation and anion uptake are regulated differently, plants must be able to compensate for differences in electrical charges that arise from disproportionate uptake of cations and anions. Plant cells maintain a pH in the range 7.3 to 7.6 by either releasing or consuming hydrogen cations (H+), which is achieved by formation or removal of organic acids.

The nutrient journey continues in a path from cell to cell through tiny connecting tubes (plasmodesmata), although some nutrients can continue to move between cells through the free space. The next barrier occurs at the waxy layer (Casparian band) that surrounds the central vascular tissue (phloem and xylem). The phloem and xylem are special tissues that act like highways for nutrient transport from roots to leaves (xylem) and from leaves to growing points and roots (phloem). In young root tips, the Casparian strip is not well formed and thus is an incomplete barrier. The mechanism how ions pass through the Casparian strip and into the xylem (“xylem loading”) is not well understood. There is probably a combination of active (ion pumps) and passive channels for ion movement into the xylem. Xylem loading is regulated separately from root uptake, thus creating a control system for nutrient movement. Nutrients are carried by water up through the xylem. Water flows up through xylem tissues due to a suction-like force created when water evaporates from the leaves, and from slight pressure produced by roots. Once inside the xylem sap, nutrients can be unloaded and reloaded before reaching the end growing points.

Once nutrients reach their targets, there often is considerable recycling, especially for the mobile nutrients such as N. For example, a normal feature of plants appears to be simultaneous import and export of nutrients from leaves. This dynamic nutrient cycling is termed remobilization or retranslocation. In young vegetative plants, nutrient recycling occurs from the mature leaves to roots and young leaves through the phloem. The remobilization ability of different nutrients affects where deficiency symptoms occur. Deficiency symptoms of mobile nutrients such as N will first appear in old tissues. In contrast, deficiency symptoms of nutrients with limited mobility such as sulphur and copper will occur in young tissue, and can hinder flower/seed development.

Nutrient remobilization is particularly important when seeds are forming. At this stage, mobile nutrients are being exported from ageing leaves while nutrient imports are decreasing. Also, root activity and nutrient uptake generally decrease by this stage due to drying soils, nutrient depletion in the soil and a relative shift in the energy supply from roots to developing pods and seeds. Plant parts with a strong energy or nutrient demand are called “sinks.” As a result, old leaves are sacrificed to supply pod and seed growth. Mobile nutrients in the seed have mostly been transferred from other plant tissue—in canola the sources are pods, stems and leaves.

Roots are not the only sites where nutrient uptake can occur. Some nutrients can be absorbed by leaves and other above ground plant parts. Nutrients in the gas form—NH3 (ammonia), NO₂ (nitrogen dioxide) and SO₂ (sulphur dioxide)—can enter leaves through leaf pores (stomata) and then be changed into organic forms. These gases are major air pollution components and in some areas contribute considerably to plant nutrition. In areas with intensive livestock operations, NH3 uptake can contribute 10 to 20% of the nitrogen for adjacent crops. SO₂ is readily absorbed by leaves. In a European field experiment, almost half of the total sulphur (S) taken up by vegetative rapeseed came from atmospheric S compounds, probably SO₂. This may partly explain why S deficiencies have increased in western Canada after environmental regulations enforced cleanup of S emissions from gas plants.

Soil Properties that Affect Plant Nutrition

Soil is a complex mixture of non-living substances (minerals, organic matter, gases and liquids) and living organisms (bacteria, fungi, insects, worms, etc.). These factors influence soil fertility either directly or indirectly.

Soil solids consist of mineral particles, organic matter in varying stages of decomposition and living organisms. Solids make up about half the soil volume, while water and gases make up the other half in the pore space.

Soil mineral particles vary widely in size and are classified by size:

• rocks are larger than 2 mm in diameter (0.08")
• sand particles range from 0.05 to 2 mm (0.002 to 0.08") in diameter
• silt particles range from 0.002 to 0.05 mm (0.00008 to 0.002") in diameter
• clay particles are smaller than 0.002 mm in diameter—(0.00008") in diameter

These particles are made from various mineral types with different elemental composition, which affects weathering processes and thus the release of certain nutrients. Two soils with identical texture could be drastically different in fertility due to differences in mineral composition. Potassium is an example of a plant nutrient whose supply arises from mineral weathering in soil.

The soil colloidal fraction refers to microscopic particles of clay and organic matter. The surface of the colloidal fraction is where most soil chemical reactions occur and it is very important in nutrient supply.

The proportion of sand, silt and clay determines the texture of a soil. Soil texture is grouped into five or more classes (see Chapter 6 for details). The texture influences fertility by affecting moisture holding capacity, air exchange and the CEC. Adequate moisture is key to fertilizer response and potential yield for canola in western Canada (Chapter 4 is devoted to this topic).

The CEC is an important property that influences the soil storage of many plant nutrients. Most nutrients are present in the soil water as positively charged cations. A few are negatively charged anions. The CEC indicates a soil’s ability to hold or store cations. Prairie soil particles typically have a negative charge. The process of electrical attraction that holds cations to negative surfaces of soil colloids is called adsorption (not absorption). The cations are not permanently stuck to the colloidal surface and can be exchanged with other cations. With time certain cations may become “fixed” into forms that are not easily removed from the exchange complexes. Adsorbed cations are not removed by water moving through the soil and can be accessed by plant roots. Cations with a higher positive charge (for example Ca⁺²) are held more tightly than those with a lower charge (for example K+).

Soil negative charges arise due to substitutions in the mineral crystals by elements with smaller positive charge, and due to reactions at the edges. Organic particles also contain a significant number of negative charges. The total particle surface area in a soil increases as particle sizes get smaller. Therefore, a soil high in clay has a much greater surface area than a sandy soil.

A high clay soil also has a bearing on the surface area and negative charge. Clay minerals are microscopic layers of aluminium and silicon crystals formed by weathering of other minerals. Thus clays are called secondary minerals. The type of clay depends on the original minerals and the weathering extent. Fairly “young” clays common in western Canada (such as montmorillonite) have a 2:1 arrangement of silica:alumina crystal sheets, while older clays have a 1:1 arrangement. Generally, 2:1 clays have 10 to 100 times more surface area, negative charges and, consequently, a higher CEC than 1:1 clays. Organic colloids have 10 to 100 times more negative charges and higher CECs than the 1:1 clays. Therefore, soil organic matter levels greatly influence the CEC.

The CEC strongly influences soil fertility. A higher CEC means that more cations, including plant nutrients, can be loosely stored in a plant available form, giving the plant a greater pool of nutrients to draw from. Since most cations are not highly soluble, only small quantities can be dissolved in the soil solution at one time. The CEC soil property allows a reservoir of nutrients to be stored then released to plant roots. This continuous replenishment of nutrients in soil water is very important for several nutrients, including potassium. A high CEC also means that fewer cations will be lost through leaching out of the root zone.

Since soils are predominantly negatively charged, anions [such as nitrate (NO₃-) and sulphate (SO₄^-2)] are repelled by soil colloids and tend to stay in the soil water. They will flow with water and are potentially subject to leaching loss.

Soil organic matter (OM) plays an important role in soil fertility as a plant nutrient storehouse. Not only does OM adsorb many cations due to a high CEC, it also stores nutrients as part of its structure. As the OM is decomposed by soil microbes, nutrients are released from the organic structure into plant available forms—this process is called mineralization. Mineralization from OM is the primary natural source of plant available N and S in prairie soils, and also influences P availability. Mineralization of individual nutrients will be described in later sections. Soil OM also plays a secondary role in soil fertility by improving physical properties such as water holding capacity, infiltration, aggregation (tilth) and buffering pH.

Soil Testing for Nutrient Content

Plant nutrient content in soil varies over years, between fields and even within fields that appear uniform. Soil sampling and analysis methods (“soil testing”) were developed to assess the fertility level and to predict crop response to applied fertilizer or manure. Soil testing is not an exact science due to nutrient variability inherent in most fields and the inability to predict growing season weather. Although soil testing is not exact, it can help estimate soil fertility and give reasonable guides for profitable fertilizer application.

Spatial nutrient variability in fields creates problems for soil testing and fertilizer application. The variability makes it difficult to obtain representative soil samples. Using single fertilizer rates across variable fields results in over-fertilized and under-fertilized areas within the field. Although variable rate fertilization is being researched and developed, most fields still are fertilized with a single rate. In addition, fertilizer response calibrations developed from research sites with low variability will under-predict the optimum fertilizer rate for larger farm fields with more variability.

For meaningful soil test results proper soil sampling is necessary. A sampling error in the field is usually much greater than the analytical error in the lab. Ensure soil samples accurately reflect the overall field. However, intensive soil sampling is not convenient, cost effective or practical. Research shows that the accuracy of a composite soil sample increases with the number of sub-samples taken (see Figure 1). Accuracy refers to how similar the soil sample value is to the true field average (is the sample representative). Precision describes how often the same value can be obtained when repeating the procedure (reproducibility). The common recommendation to sample a field 20 times and mix all the samples into one composite, produces an accuracy of about ±17% for NO₃-, assuming 80% precision. This is an acceptable level of accuracy and precision for fertilizer recommendation purposes in most cases.

Figure 1. Effect of Sub-Sample Number on % Accuracy of Composite Sample for Nitrate-N with 80% Precision

In addition to adequate sample numbers, proper soil sampling techniques include:

• where and how to sample each field (sampling plan)
• proper equipment
• proper sampling time
• correct depth
• proper sample handling

Soil Sampling Plan

Sample individual fields separately. The first step is to assess the field variability and identify representative areas. The type and level of variability can influence the choice of sampling plan. Consider the four basic sampling plans:

1. Random soil sampling uses a random pattern across a field, generally avoiding unusual or problem areas such as hilltops and potholes. Bulk together 20 soil cores into one composite sample, air-dry then send to a soil test lab. This common method is adequate for smaller, relatively uniform fields.
2. Topographic sampling involves separating sets of samples based on topography. Identify the dominant topographic features such as hilltop, midslope and bottom slope, and take 20 core samples for each type. Bulk sub-samples from each type into one composite sample for each landscape type. Send several samples to the testing lab. This method can provide more meaningful results for variable fields, but at additional expense and labour. You must be willing and able to apply different fertilizer rates in the separate landscape areas.
3. Benchmark sampling expands on the topographic sampling concept by considering unique areas based on topography, soil texture and type and typical crop growth. Once these unique areas are identified, each year the samples are taken from the same spot within each benchmark area. There may be one or more benchmark sites within a field, depending on the variability. Each benchmark area then becomes a reference area on which fertilizer recommendations are based. The benchmark sampling area is much smaller than the whole field, and together with sampling in the same spot from year to year, this is assumed to reduce variability of the test results. If the benchmarks are carefully selected to represent the majority of the field, then good soil test results can be obtained.
4. Grid or systematic sampling follows an organized grid pattern, perhaps every 0.2 to 2 ha (0.5 to 5 acres). This method can reveal field nutrient variability and allow for variable rate fertilization and precision farming techniques. However, the sampling and analysis costs are not economic with current field crop prices on the prairies.

Proper Soil Sampling Equipment

Soil sampling to a 60 cm (2') depth can be done with a probe or auger. Do not use flight or screw sampling augers if samples need to be separated by depth since mixing will occur. If testing for micronutrients, ensure the sampling tool is chrome plated or stainless steel and rust-free. Do not bulk samples in metal pails. Clean plastic pails labelled for location and depth work well.

Information sheets, sample bags and shipping boxes are available from soil testing labs and most fertilizer dealers.

Proper Sampling Date

The most accurate sampling time is just prior to seeding. However, this isn’t practical because time is needed to purchase and perhaps place the fertilizer before seeding. Sampling is commonly done in the spring or fall. Spring sampling is done after the soil has thawed and is no longer saturated from snow melt. Fall sampling can begin once the soil has cooled to 5 to 7°C. This helps reduce nutrient content changes due to microbial activity.

The old assumption that N availability does not change over the winter has been proven wrong. Research in Alberta found that available N increased by 56 kg/ha (50 lb/ac) in stubble fields from fall to late winter while the soil was frozen. Summerfallow fields increased by 73 kg/ha (65 lb/ac). This overwinter gain in available N is apparently due to death of soil microbes and subsequent release of available N forms from their ruptured cells. However, the available N gained over the winter was temporary as it was mostly lost in the spring, probably through denitrification. The stubble fields lost 45 kg/ha (40 lb N/ac) in the spring while summerfallow fields lost 73 kg/ha (65 lb/ac). Therefore, the net change from late fall to late spring was minimal.

Late fall sampling tends to more accurately reflect spring NO₃- (nitrate) contents than early fall sampling, especially for Black soils. Alberta research in the 1980’s compared soil samples taken in the fall (early October and early November) ' to spring samples for 26 stubble fields. Early fall samples averaged 34 kg/ha (30 lb/ac) less nitrate N thanspring samples, while late fall samples averaged only about17 kg/ha (15 lb/ac) less. The early fall samples were also more variable in relation to spring samples. Overall, late fall samples more accurately predicted spring nitrate contents and grain yields than early fall. Spring samples were slightly better than late fall samples for predicting grain yield and N uptake.

In contrast, recent research in Manitoba measured very little change in soil nitrate levels in cereal stubble from early September to freeze-up. North and South Dakota extension soil scientists recommend early fall sampling in view of time constraints, but reduce the N recommendation by 0.2 kg (0.5 lb) for each sampling day prior to September 15.

Another experiment in central Alberta compared the effects of sample timing on phosphorus (P) soil test values. On average over 27 sites, the extractable P increased from 28 kg/ha (25 lb/ac) in early October to 49 kg/ha (44 lb/ac) by early November, and to 50 kg (45 lb) by spring (late April to early May). The relationship between early fall and spring P values was not close and, therefore, it was not possible to simply correct the early fall values. In contrast, research on a Brown soil in Saskatchewan over 24 years found both overwinter increases and decreases in soil P tests, but relatively few were significant. An experiment on irrigated alfalfa on a Dark Brown soil near Lethbridge, AB found significant overwinter increases in organic P. The conflicting results may be related to the differences in soil organic matter content and biological activity, and, therefore, potential for microbial changes to the plant available P pool.

In conclusion, early fall sampling can create higher than necessary fertilizer N and P recommendations due to an underestimation of spring nitrate N and available P, especially in the Black and Gray soil zones. Fertilizer response curves have been calibrated only against spring nutrient contents.

One disadvantage to late fall sampling after soil has cooled to 5 to 7°C is that fall fertilizer banding opportunities become more limited. By the time the samples are taken, dried, sent to the lab, analyzed and results returned, the soil may have become frozen or covered with snow. On average over the prairies, soil cools by 1°C every five days in the fall.

Correct Sampling Depth

The appropriate sampling depth depends on the nutrients to be tested. For mobile nutrients such as N and sulphur (S), sampling to the 60 cm (2') depth is usually the most accurate according to research conducted in the 1960’s. The fertilizer response database for N was developed with 0 to 60 cm (0 to 24") samples. However, recent research in Saskatchewan and North Dakota indicates that the 0 to 30 cm (0 to 12") depth may be more accurate for N than either 0 to 15 cm (0 to 6") or 60 cm samples. This recent research and the fact that sampling 60 cm is considerably more difficult, supports the 0 to 30 cm depth as a reasonable recommendation for these areas. In contrast, research in Manitoba has documented that the 0 to 15 cm depth is inferior. Separate samples for 0 to 15 cm, 15 to 30 cm and 30 to 60 cm was often recommended in the past, but was rarely done due to additional expense and time.

For immobile nutrients such as P and potassium (K) and most micronutrients, the ideal depth is 0 to 15 cm because the fertilizer response calibrations are based on that depth. If only the 0 to 30 cm depth is sampled, the soil testing labs must use a correction factor to estimate the value for the 0 to 15 cm depth. Since these nutrients are relatively immobile, they tend to remain at the fertilizer application depth. Therefore, the 0 to 30 cm depth may underestimate these nutrients and lead to high fertilizer recommendations.

Proper Sample Handling

Handle samples carefully to prevent accidental mixing and contamination. Mix each sample and spread on separate pieces of clean paper to dry at room temperature. Use a fan if required and avoid additional heat sources. Once dry, fill soil sample cartons or bags with about 0.5 kg (1 lb) of soil, and label with field number and depth.

Consistency of Soil Test Lab Recommendations

Soil sampling, analysis and interpretation is not an exact science. However, reasonable precision and accuracy is needed in order to make costly fertilizer decisions. Over the years, various growers, agencies or companies have sent duplicate samples to different labs to compare their analysis and recommendations. Unfortunately, widely differing results and recommendations have occurred, causing growers to question the credibility of the labs or the soil testing practice.

Two fundamental reasons contribute to differences in results. First, labs may be using different analytical procedures to measure soil nutrient content. For example, there are several methods to extract soil phosphorus. Or the technique may differ slightly when using a particular method. Over time, labs are harmonizing their test methods and in time this problem may disappear.

Variations in soil test recommendations arise mainly due to the differences in each lab’s interpretation. The recommended fertilizer amounts at the same soil test level can vary significantly from lab to lab. This may be due to using:

• different critical (deficient) soil levels being used
• regional fertilizer response (calibration) data but modifying recommendations to fit a particular philosophy of fertilizer use or economic payback
• recommendations from other regions or countries
• a unique system of fertilizer recommendation not based on regional calibration data or economics

Although consistency among labs has been improving since the first comparison studies in the 1980s, further improvements are needed. Calibration data must continue to be collected to account for changes in fertilizer application techniques and changes in other agronomic practices.

Overall, soil testing is a useful agronomic practice. Use labs that base fertilizer recommendations on economics using regional calibration data. Be prepared to question unusual recommendations based on experience and the local knowledge of qualified agronomists. Keep in mind that the accuracy of fertilizer recommendations will always be limited by sampling challenges and the inability to predict the weather of the upcoming growing season.

Plant and Tissue Testing

Crop nutritional status also can be assessed by plant and tissue analyses. These methods can supplement, but not replace, soil testing. Plant and tissue analyses measure the nutrient content of above ground plant parts during growth. The values are compared to established ranges for inadequate, adequate and excess levels.

Plant tissue testing is suitable for diagnosing crop problems that may be nutritionally related and to identify any nutrients that may be limiting yields. Plant analysis can determine if the fertilizer rate and method of application were adequate to meet crop needs. The disadvantage of tissue sampling is timing—after tissue samples are taken and analyzed, it may be too late to correct deficiencies in the current crop. No reliable interpretative criteria exist for nutrient ranges in seedling canola. Also, nutrient contents usually differ greatly between different plant parts and ages. Therefore, the proper part must be sampled at the proper growth stage.

An adequate sample will contain 50 to 80 plants, depending on the nutrients to be tested and plant part/age. Avoid unusual, dead or stressed plants, as well as those covered with soil or recent sprays. Cut samples with a clean, rust-free knife or scissors. Dry the plant samples on clean paper or plastic at room temperature (do not oven dry). After drying, keep the samples in a paper bag. The following table shows sufficiency levels for most plant nutrients in flowering canola.

Table 3. Plant Tissue Analysis Interpretative Criteria for Canola (whole above ground plant at flowering)

Nutrient	Sufficiency Level
Nitrogen (N) %	> 2.4
Phosphorus (P) %	> 0.24
Potassium (K) %	> 1.4
Sulphur (S) %	> 0.24
Calcium (Ca) %	> 0.49
Magnesium (Mg) %	> 0.19
Zinc (Zn) ppm	> 14
Copper (Cu) ppm	> 2.6
Iron (Fe) ppm	> 19
Manganese (Mn) ppm	> 14
Boron (B) ppm	> 29
Molybdenum (Mo) ppm	> 0.02

Comparing the plant analysis results from two areas of a field that differ visibly in growth can be difficult to interpret because nutrient content differences can be confounded by growth differences. If the two areas differ mainly in deficiency symptoms, then comparative sampling can be useful. In this case, collect the samples soon after the symptoms appear and before major differences in growth and maturity occur. Plant and tissue analyses need to be interpreted by experienced individuals.

How to Diagnose Nutrient Deficiency Symptoms

In moderate to severe nutrient deficiencies, visible symptoms can indicate the specific nutrient that is lacking. Nutrients that are only slightly limiting often do not show visible symptoms, a situation that has been termed “hidden hunger.”

A systematic diagnosis of visible symptoms is needed to correctly identify limiting nutrients. Symptoms usually appear on either old or young leaves depending on the mobility of the nutrient in question. Chlorosis (loss of green colour, yellowing) and necrosis (death of plant tissue, often leading to white or brown colour) are important visible symptoms. Diagnosis under field situations can be complicated by high field variability, multiple deficiencies, and other causes such as weather, pests and herbicide injury. For example, a sulphur deficiency can easily be confused with Group 2 herbicide injury due to similar symptoms.

The diagnosis of nutrient deficiency symptoms is outlined in the Figure 2 flowchart.

Nitrogen (N)

Nitrogen is the most common limiting nutrient (other than water) for canola production. Therefore, a good understanding of this nutrient is needed to efficiently manage fertilizer N and maximize economic returns.

Figure 2. Diagnosing Nutrient Deficiency Symptoms

Role of Nitrogen in the Canola Plant

As shown in the previous section (Table 3), canola, like most crops, contains large amounts of N. Nitrogen is a part of many critical plant components: amino acids and proteins (which form enzymes); genetic material (nucleotides and nucleic acids); and other components found in membranes (such as amines), co-enzymes and others. The majority of the N in green plant tissue is present as enzyme protein in chloroplasts where chlorophyll is located. By harvest, the majority of the N in a canola plant is found as seed protein. The relative N proportions in the plant changes over time and growth stage. The N proportioning closely resembles the dry matter partitioning as shown in Figure 3.

The N level in canola plants is highest in the early seedling stage when young leaves are the majority of the plant’s dry matter. As the plant grows into flowering stages, the overall N level declines due to stem material and leaf loss. By maturity, canola straw contains just 0.5 to 1.5% N while the seed contains 3.4 to over 4% N.

Figure 3. Nitrogen Partitioning in Canola

Nitrogen Effects on Canola Growth

The most obvious N effect is an overall increase in plant growth (height and dry matter). This stimulation occurs early in the vegetative stage and continues into the reproductive stage. Research in England illustrates the canola leaf stimulation, leaf area and pods/seed production by fertilizer N (Figures 4, 5, 6).

Figure 4. Canola Leaf Area Response

Other research generally confirms that N fertilizer mainly increases canola leaf area index, leaf duration, plant weight, growth rates, number of flowering branches, plant height, number of flowers, number and weight of pods and seed yield. Therefore, good N fertility is necessary to produce a large, photosynthetically efficient leaf area that will support high numbers of flowers, pods and seed yield.

Canola Nitrogen Deficiency Symptoms

Healthy canola plants with adequate N have dark green leaves. Nitrogen is mobile within the plant and can be moved from older to younger leaves and pods. Therefore, N deficiency symptoms first show up in older leaves as pale green to yellow colouring, and sometimes purpling. These older leaves tend to die early, turn brown and drop off prematurely. Overall plant growth is slow, with short thin stems, small leaves, and few branches. The amount and time of flowering is restricted, and pod numbers are low. Nitrogen-deficient canola pictures are shown in Figure 19. In healthy canola, plant tissue tests of above ground material at flowering will show more than 2.5% N.

Figure 5. N Fertilizer Effect on Pod, Seed Number and Yield

Figure 6. Effect of N on Flowering Branches

Canola Response to Fertilizer Nitrogen

Canola responds well to applied fertilizer N on deficient soils. Most stubble fields have insufficient N for high canola production and thus require fertilizer or manure application. Research data has been collected that describes crop response to fertilizer in relation to initial soil reserves in western Canada and around the world. This data is used by soil testing labs to predict fertilizer requirements. Research on the prairies has found that profitable dryland canola yield response to fertilizer N is unlikely when the soil contains more than 34 to 45 kg nitrate N/ha (75 to 100 lb nitrate N/ac) in the top 60 cm (2').

Higher yielding winter canola types typically respond to more N fertilizer than winter wheat. In contrast, spring canola types have a similar N fertilizer response to high yielding CPS wheat in western Canada.

An example of typical canola response to fertilizer N on a deficient stubble black soil in Alberta is shown in Figure 7. The vertical line represents the economical level of N fertilizer under medium moisture with $265/tonne ($6/bu) canola, $0.88 kg ($0.40 lb) N and a 2:1 risk ratio.

Figure 7. Canola Response to N Fertilizer

The previous crop influences the crop response to N fertilizer. Different preceding crops vary in the amount of available N removed from the soil and the amount released from crop residue. In addition, the disease break with different rotational crops can affect the yield potential and, therefore, the economic response to fertilizer.

Moisture availability greatly affects the yield response to N fertilizer. Conversely, adequate N is needed for crops to respond to moisture. Canola yield is reduced under extremely dry or wet conditions. Research at Outlook, SK illustrates the synergistic response of canola to moisture and N over eight years (Figure 8).

Figure 8. Canola Response to N and Moisture at Outlook, SK

Under dry soil conditions, root growth and activity are reduced, resulting in less N uptake. In addition, soil microbial growth is slower, which reduces N release from soil organic matter, but also reduces temporary tie-up by soil microbes. Normally, more plant available N is left in the soil after a dry growing season than after wet seasons.

Drought and high temperature stress near flowering and podding dramatically reduce both water and N fertilizer efficiency. This is true for both stubble and fallow crops, although moisture is more often limiting for stubble crops. Unfortunately, crops use the most water during the flowering and podding period when soil moisture reserves and rainfall are usually low. Therefore, the amount and timing of rainfall are important. Studies in western Canada generally find that growing season precipitation increases grain yield two to three times more than equivalent amounts of stored soil moisture.

Heavy N fertilization can reduce canola yields when excellent spring moisture conditions are followed by drought. Under this condition, the N stimulates larger leaves, increases transpiration and moisture use. As a result, soil moisture can be depleted, leaving little for flowering, podding and seed fill. Excessive N fertilization can also reduce yields by promoting lodging, delaying maturity that may increase fall frost damage, and increase foliar disease due to the dense canopy and lodging.

Effect of Nitrogen Fertilizer on Canola Quantity

N fertilization generally increases the protein content of canola seed and meal. However, when N fertilizer is added under conditions of S deficiency, there may not be a protein increase but a rise in free amino acids due to hampered protein synthesis.

In contrast, N fertilization may slightly decrease the canola seed oil content, especially at higher rates. Plant breeders report that oil and protein contents are often inversely related—any attempt to change one causes an opposite change in the other component. Although the seed oil content may decrease at high N rates, the total oil yield per acre still increases because yield increases more than oil level decreases. The overall effect of N fertilization on yield and canola quality based on research in Manitoba is illustrated in Figure 9.

Figure 9. Canola Response to N and Moisture at Outlook, SK

Supply of Nitrogen by Soil and Fertilizer to Canola

Canola plants obtain N from several sources:

• soil supply of nitrate and ammonium arising from soil organic matter decomposition and fertilizer residues of previous years
• nitrate and ammonium released through mineralization during the current growing season
• fertilizer or manure N additions to the current crop
• other minor sources such as available forms deposited during rainfall and lightning, nonsymbiotic N fixation in the root zone, and ammonia uptake by foliage

Since canola plants obtain most of their N from the soil environment, understanding the N cycle is very important to managing N fertilizer effectively.

Nitrogen Cycle and Transformations

Nitrogen is subject to many different processes in soil due to microbial activity and physical forces (Figure 10).

Figure 10. Nitrogen Cycle and Transformations

Transformations that Supply Plant Available Nitrogen

Soil organic matter is a major N reservoir containing several thousand pounds of organic N per acre. This large organic N storehouse needs to be decomposed by soil microbes before becoming available for root uptake. The decomposition process is called mineralization.

The decomposition rate is fairly slow and variable, ranging from about 0.4 to 5% per year. As shown in Figure 10, organic N is first slowly changed to ammonium by bacteria. Then different bacteria rapidly change the ammonium to nitrate, in a two-step process called nitrification. The ammonium is first changed to nitrite (NO2 -), then to nitrate. Under normal soil conditions, the bacterial oxidation of ammonium to nitrite is much slower than nitrite to nitrate. Therefore, very little nitrite is normally found in soil. This is fortunate since nitrite is toxic to plants. The result of these rate differences is that most of the plant available N in the soil tends to be nitrate. This is also why most soil testing labs analyze soil for nitrate and don’t include ammonium.

Since N transformations result from soil microbial activity, soil conditions such as temperature, moisture and acidity will strongly affect the rates at which these processes occur. If soil is cold, saturated or very acidic, then mineralization and other microbial activities will proceed slowly. Cultivation stimulates organic matter decomposition and mineralization of N by improving aeration and physical mixing that gives soil microbes access to new organic matter supplies.

There are several sources of plant available N to the soil system, in addition to soil organic matter decomposition and fertilizer additions (Figure 10). The atmosphere contains 78% N₂ and thus is a huge source of N. But it first must be changed to plant available forms through biological or industrial processes called N₂ fixation. Biological fixation of atmospheric N₂ is performed by certain bacteria or blue-green algae species. Biological N fixation in soil falls into three general types:

• symbiosis with legumes
• associative
• free-living N fixing bacteria

Symbiotic N fixation is much larger relative to the other types. Canola is not a legume and cannot form the symbiosis with rhizobia to fix atmospheric N. However, canola can benefit from residual N fixed by previous legume crops.

Associative N fixation occurs when bacteria just inside the root or on the root surface use root exudates for energy to fix atmospheric N. The plant benefits indirectly when the bacteria dies and its N is released through mineralization. The results of many experiments in Russia and throughout the world on cereal inoculation with associative N fixing bacteria have shown varying and unpredictable responses. This suggests that the interactions between plants and the bacteria are complex, unstable and can vary greatly depending on genotypes. Much less research has been done on canola inoculation with associative N fixing bacteria. The limited research does show that some strains will successfully colonize the roots of Brassica species but often do not significantly influence harvest dry weight or N accumulation.

Transformations that Reduce Plant Available Nitrogen

Several different mechanisms contribute to N loss from soil, and, therefore, lost opportunity for increasing yield. Understanding the conditions that promote such losses can be valuable in avoiding such conditions in the field and improving the N fertilizer efficiency.

Denitrification

One major N loss from soil occurs through a microbial process called denitrification. As shown in Figure 10, nitrate can be changed by certain bacteria to gases such as N₂O and N₂, which escape back to the atmosphere. These soil bacteria have the ability to switch their respiration from using oxygen to nitrate. Since respiration is more efficient using oxygen, these denitrifying bacteria will only switch to nitrate if oxygen is absent, such as in waterlogged soil. Therefore, denitrification becomes significant when soils become saturated. Other secondary factors that encourage denitrification include carbon availability (crop residues), warm soil, and neutral to alkaline pH.

Denitrification accounts for 10 to 50% of the available N losses in prairie soil. Research on the prairies has shown that considerable N denitrification losses occur during spring thaw. For example, research near Edmonton, AB found that 16 to 60% of annual denitrification loss occurred immediately following snowmelt. During spring thaw on the prairies, frozen subsoil is often a barrier for water drainage and the overlying thawed soil becomes saturated.

The second major period of denitrification loss occurs in late spring and early summer during rainfall events that cause soil saturation. Remember that denitrification occurs regardless of the nitrate source—from fertilizer, manure or from decomposition. Effective N fertilizer management strives to avoid having large amounts of nitrate present during spring melts. Summerfallow is especially prone to large denitrification losses since large amounts of nitrate and moisture are stored during the fallow year, which increases the denitrification potential in the next spring thaw. Summerfallow also is a major contributor to leaching losses of nitrate.

Immobilization

Immobilization is the second major N transformation that reduces the plant available N supply. Figure 10 indicates that soil bacteria may use either nitrate or ammonium for their own growth, temporarily tying up the N in the soil organic N storehouse. Immobilization essentially is the reverse of mineralization, and occurs when residues with low N content (like cereal straw) are being decomposed. Since these residues don’t contain enough N for the microbes to make their own protein, they need to use the nitrate and ammonium. Soil microbes thus compete with plant roots for the available N, and plant growth suffers when N supplies are inadequate for both microbial and plant growth needs. The poor crop growth in heavy chaff rows is due in part to immobilization of N and other nutrients by the decomposing microbes. One effective fertilization strategy is to place the fertilizer away from residues and thus avoid immobilization losses. The remaining transformations that reduce plant available N are usually relatively minor.

Table 4. Seedbed Utilization (SBU) of Various Openers*

	Spread Width of Fertilizer in Seed Row
	2.5 cm (1") Disc or Knife			5 cm (2") Spoon or Hoe			7.6 cm (3") Sweep			10 cm (4") Sweep
Row spacing cm	15	23	30	15	23	30	15	23	30	15	23	30
Row spacing "	6	9	12	6	9	12	6	9	12	6	9	12
SBU %	17	11	8	33	22	17	50	33	25	67	44	33

*Although some openers also vertically spread seed and fertilizer, this is not considered in the table since seed should be placed at a consistent depth for uniform germination and emergence. The actual spread width varies with air flow, soil type, moisture, residue and speed, and should be checked under prevailing field conditions.

Leaching

Leaching of nitrate can occur since this form is not adsorbed to the soil and moves readily with soil water. Leaching losses can be significant in sandy soils in high rainfall areas or under summerfallow, but overall leaching probably contributes to less than 10% of the available N losses on the prairies. To reduce leaching losses time fertilizer applications to avoid prolonged exposure to wet conditions, and consider band placement to delay the conversion to the vulnerable nitrate form.

Volatilization

Volatilization occurs when ammonia escapes from the soil to the atmosphere. Such losses happen in a variety of ways. One obvious loss occurs when anhydrous ammonia fertilizer is improperly applied (too shallow or into a too dry or wet soil). Broadcasting urea fertilizer on the surface without incorporation can also lead to significant volatilization losses if significant rainfall (more than 6 mm (1/4") does not occur soon after application. All ammonium based fertilizers are subject to volatilization if broadcast on the surface of soils with high pH, surface lime salts, low soil organic matter, warm temperatures and dry conditions. To reduce volatilization loss, ensure proper fertilizer placement into the soil.

Weeds

Weeds can contribute to poor N fertilizer efficiency by competing with crops for uptake. The competitive ability of the crop for fertilizer uptake can be improved by placing the fertilizer near crop roots rather than broadcasting or random banding.

Erosion

Erosion of topsoil carries significant N and other nutrients away from the field. Use soil conservation techniques to minimize such losses.

A final minor loss mechanism occurs when ammonium is fixed into the crystal structure of certain clays. Some soils contain expanding type clays that allow ammonium to enter within the plates of the crystal structure and become “fixed.” Such ammonium trapped within the crystal lattice is held tightly and unavailable for root uptake.

Nitrogen Fertilizer Management for Canola

Due to the various N losses described in the previous section, N fertilizer use efficiency cannot approach 100%. Generally, research in western Canada has found that N fertilizer use efficiency (fertilizer N recovered in seed) rarely exceeds 50% and often is less than 20%. In the latter case, this means that only 20% of the fertilizer N made it into the seed. Although a small amount of fertilizer N remains in plant parts other than seed, a significant portion is lost. Fertilizer management strives to increase efficiency by increasing crop uptake and decreasing the losses. Fertilizer N management uses two main tools: placement and timing.

Seed Row Placement

Although seed row placement of N fertilizer is an efficient method for uptake, canola is sensitive to seed row N and this limits application rates. Canola seedlings are injured by excessive seed row N by the “salt effect” that reduces water uptake by the seed, and by ammonia toxicity. Greenhouse research at the Agriculture and Agri-Food Canada (AAFC) Beaverlodge, AB Research Centre in 1960 showed that rapeseed was sensitive to seed-placed N (see Figure 11). Subsequent field research confirmed that rapeseed was more sensitive to seed-placed N than cereals. During this period, seed row placement was limited to P and very low rates of N. The common drills during this time were double disc and hoe press drills, which give a very narrow seed spread.

The adoption of conservation tillage seeding systems and air-seeders has greatly influenced fertilizer placement. Development of pneumatic delivery implements (“air seeders” and “air drills”) has facilitated both dry fertilizer banding and direct seeding. Conservation seeding systems limit tillage passes in order to retain surface residues and this reduces the options for fertilizer application. However, newer machines designed for direct seeding have resolved the fertilizer placement issue by either placing the fertilizer away from the seed or increasing the seedbed utilization. Seedbed utilization is the spread width of fertilizer and seed relative to the row spacing. For example, a 7.6 cm (3") spread with 15 cm (6") row spacing creates 50% seedbed utilization. Table 4 outlines the seedbed utilization (SBU) obtained with different openers and row spacing.

Numerous trials have examined the safe seed row N amounts with various openers and configurations. Research conducted in Alberta from 1992-96 illustrates the effect of seed row N on canola emergence and yield. The research involved 32 site years with canola at various Alberta locations. The highest N rate and least SBU reduced canola emergence and yield 90 and 45% of the time, respectively. Sites with limited moisture (due to sandy texture, low seedbed moisture or dry conditions) two weeks after seeding experienced the greatest reduction in emergence and yield with 101 kg N/ha (90 lb N/ac) as urea and low SBU (see Figures 12 and 13).

Figure 11. Seed Row N Fertilizer Effect on Canola Emergence

Figure 12. Effect of Moisture and SBU on Canola Emergence [101 kg N/ha (90 lb N/ac)] in Alberta

Figure 14 shows the approximate safe rates of seed row granular N fertilizer based on prairie research to date. In canola, there is no significant difference in seed row safety between urea (46-0-0), ammonium sulphate (21-0-0-24) or ammonium nitrate (34-0-0). Anhydrous ammonia (82-0-0) must be placed separately from the seed. If moisture conditions are dry, reduce the safe amount of seed row N by half. The N rates are in addition to N contained in seed row phosphate fertilizer.

Band Placement

Band placement away from the seed row can be used to avoid toxicity and to improve fertilizer use efficiency. Banded N fertilizer is usually more efficient than broadcast incorporation because concentrating fertilizer in the band reduces the contact with soil and microbes, and reduces losses due to denitrification and immobilization. The banding benefit varies between soils and years mainly due to differences in moisture and susceptibility to loss. Also, if the band is located near the seed row rather than random, fertilizer loss due to weed uptake is reduced.

Pre-plant banding (or “deep banding”) involves placing granular, liquid or gaseous fertilizer N in a ribbon several inches below the soil surface before seeding. Banding is often done in the fall on fields that tend to be wet in the spring. Fall banding can spread the workload without significantly lowering N efficiency, and often allows growers to buy fertilizer at lower cost than in the spring. The banding depth often is 8 to 10 cm (3 to 4"). Under dry or sandy soil conditions, ensure anhydrous ammonia is placed deep enough to prevent visible gaseous loss, and ensure the soil flows well around the openers to permit a good seal behind the shanks. Spring banding can be shallower due to better moisture and tilth. However, in dry springs the banding operation can reduce seedbed moisture and quality. Make spring band spacings narrower than in the fall.

Figure 13. Effect of Moisture and SBU on Canola Yield [101 kg N/ha (90 lb N/ac)] in Alberta

Figure 14: Approximate Maximum Rates for Seed Row Placement in Canola with Good to Excellent Seedbed Moisture

Spring banded anhydrous ammonia can be immediately followed by seeding, providing there are several inches of vertical separation between the injection point and seed depth. Canola emergence directly over the bands may be slightly reduced, but yields are generally not affected.

Figure 15 illustrates research conducted at the Agriculture and Agri-Food Canada (AAFC) Scott, SK Research Centre on the safety of seeding directly after banding anhydrous ammonia 10 to 15 cm (4 to 6") deep. Yields varied between seeding dates following anhydrous banding over the years with no relationship between yields and plant stands. Proper soil packing over the seed row to firm the soil disrupted by the banding operation was deemed more important to avoid stand reduction than was potential injury from the banded ammonia. No difference was found between banding ammonia parallel to and perpendicular to the seed row.

Figure 15. Effect of Waiting Period after Anhydrous Ammonia Banding on Canola Stand and Yield

Side banding involves placing the fertilizer band to the side and often below the seed during the seeding operation. This method has good N use efficiency and avoids seed row toxicity if the separation is maintained under field conditions. However, side banding at seeding does involve more complicated, costly openers, increased draught and wear.

Mid-row banding involves placing fertilizer between every second seed row or between a paired row during the seeding operation. This method also has good N use efficiency and avoids seed row toxicity if the separation can be maintained under field conditions. Both side and midrow banding can improve N efficiency and yield response by favouring crop versus weed access to the fertilizer.

In recent years, openers have been designed that allow anhydrous ammonia to be side-row or mid-row banded during seeding. Ensure the anhydrous ammonia is horizontally separated from the seed by at least 5 cm (2").

Broadcast Incorporation

N fertilizer can be spread onto the soil surface then incorporated into the soil with a tillage implement. Although this method can be time and labour saving, fertilizer efficiency is usually sacrificed. Under dry conditions, broadcast-incorporated fertilizer can be stranded in dry surface soil, and not accessible by plant roots growing down into moisture. In wet conditions, broadcast-incorporated fertilizer is more vulnerable to losses due to denitrification, immobilization and leaching than banded fertilizer. Broadcasting without incorporation is the least efficient method of applying N fertilizer due to increased loss due to runoff, erosion and volatilization. Broadcasting after crop emergence or topdressing generally has low efficiency, but it can serve as a rescue treatment when poor fertility was not corrected prior to seeding.

Foliar Application

Foliar application is possible, but only a limited amount of fertilizer N can enter through leaves without significant leaf burn. The only practical instance of foliar fertilization for canola on the prairies occurs under irrigation where up to 20% of applied N can be supplied in irrigation water early in the growing season.

Time of Nitrogen Fertilizer Application

Nitrogen fertilizer efficiency is greatly affected by the placement method and the application date. However, differences between placement methods or timing varies widely between years or fields due to variability in weather, soil type and drainage. Generally, N fertilizer applied at or near seeding is the most efficient. The major disadvantages of applying all the fertilizer N at seeding are:

• increased time required to seed and fertilize during the short seeding season
• higher fertilizer prices during the spring season compared to fall
• increased risk of reduced seedbed quality due to extra tillage/soil disturbance at or near seeding
• seed row N limits
• fertilizer applicators being unavailable during the busy spring season

Considerable fertilizer placement/timing research has been conducted on the prairies. Table 5 gives approximate relative efficiencies of the various placement and timing methods.

Table 5. N Fertilizer Effect on Pod, Seed Number and Yield

Time and Placement	Relative Efficiency (%)
	Dry*	Medium	Wet
Spring broadcast-incorporated	100	100	100
Spring branded	120**	110	105
Fall broadcast-incorporated	80	75	65
Fall banded	120	110	85

* These are soil-climate categories based on typical conditions expected in the spring. **Extra tillage associated with spring banding can dry out the seedbed, reduce emergence and yield in some cases.

An extensive survey conducted by Alberta Agriculture Food and Rural Development in 1982 on the practices of above average growers confirmed the banding benefit (top yields were associated with farmers who fall-banded fertilizer N).

Date of Fall Banding

Nitrogen fertilizer banded too early in the fall increases potential for losses on wet, poorly drained soils. Delay banding until soil temperatures have cooled to less than 10°C on well drained sites, and less than 5°C on poorly drained sites.

This will significantly reduce losses by reducing the rate of change from ammonium to the more vulnerable nitrate form. Unfortunately, the opportunity to fall band fertilizer into cold soil is quite short since snow or frozen soil can occur without much warning. On average, fall soil temperatures on the prairies drop 1°C every 5 days. In the Black and Gray soil zones, fall soil temperatures usually decline to 10°C by the last week of September. In these zones, since the banding benefit is significant due to typically wet spring conditions and moderately high N rates, begin fall banding in late September. This is earlier than the normal practice.

Nitrification Inhibitors

Since only the nitrate form is susceptible to losses through denitrification and leaching, methods to delay the conversion from ammonium to nitrate (nitrification) would be beneficial. Banding achieves this delay to some extent. However, interest in developing chemical inhibitors of nitrification has stimulated research programs for many years. A variety of chemicals have been tested under prairie conditions, but none has achieved commercial success. For example, nitrapyrin (“N-Serve”) was recognized as a nitrification inhibitor in the early 1960s. Nitrification inhibitors have not been successful to date for various reasons, including cost, potential toxic effects on the soil environment and inconsistency.

Urease Inhibitors

The widespread adoption of direct seeding has resulted in interest in seed-placed fertilizer. While increasing seedbed utilization increases the safety of seed-placed fertilizer, the higher disturbance and draft is not always desirable. Openers with banding capability add cost and draft. Even with sidebanding, seedling damage can sometimes occur, particularly with wide row spacing, high rates of N application or insufficient separation between seed and fertilizer. Therefore, there is interest in “safening” urea fertilizer so that seedling injury is reduced, allowing more freedom for seedplaced N.

Agrotain (N-n-butyl-thiophosphoric triamide) “safens” urea by inactivating urease enzymes in the soil adjacent to the granule. This slows the breakdown of urea to ammonia, reducing the potential for seedling damage from seed-placed or sidebanded applications of urea or urea ammonium nitrate. As long as N remains in the urea form, the risk of damage is minimal. In addition, since Agrotain delays the release of ammonia, there is more time for the uncharged urea to move away from the seed-row in the soil water or with rainfall. Movement of the urea away from the seed, combined with a slower release of ammonia from the urea, will decrease the concentration of ammonia in contact with the germinating seedling, thereby reducing seedling damage.

Figure 16. Canola Seedlings with Sufficient N (left) Versus those with an N Deficiency

In field studies, Agrotain was effective in reducing seedling damage from side-banded urea and urea ammonium nitrate, where soil and environmental conditions led to seedling damage from the untreated fertilizer. The improved stand did not always lead to a higher crop yield because canola has the ability to compensate for reduced stands. The studies also showed that canola oil and chlorophyll content were often improved by using Agrotain.

While Agrotain appears effective for use with side-banded N applications, more information is needed to determine if the safening effect is great enough to allow for seed row placement of the full rates of urea or urea ammonium nitrate needed for a high-yielding canola crop.

Phosphorus (P)

Phosphorus is an important plant macronutrient, but it is required in smaller amounts than nitrogen. Western Canadian soils are commonly P deficient and fertilization usually increases yield and economic returns. Good P fertilizer management is important to optimizing canola production.

Role of Phosphorus in the Canola Plant

Phosphorus functions in the plant as a structural element and also in energy transfer. The structural components that rely on P include nucleic acids (the building blocks of DNA) and phospholipids (fats and oils), which are important membrane constituents.

Phosphorus plays a significant role in energy transfer in all living organisms. The P energy transfer compounds are phosphate esters—about 50 different esters have been identified. ATP (adenosine triphosphate) is the principal phosphate energy compound used for starch synthesis and nutrient uptake. Energy produced during respiration and photosynthesis is captured by these phosphate compounds, which then are transported to areas that are building plant tissue. The energy stored in the phosphate compound is released, and the molecule is recycled back to be “recharged.” This recycling of phosphate energy compounds is accomplished at extremely fast rates, and a small amount can satisfy the plant’s energy needs.

Figure 17. Progressively Less N Deficient Leaves (left to right)

Figure 18. N-sufficient (left) Flowering Plants and Deficient Plants

Figure 19. N-sufficient Raceme (left) and Deficient Raceme

Figure 20. N-sufficient Pods (left) Progressing to Deficient Pods

Characteristics of Phosphorus Uptake by Canola

The main P forms taken up by roots from the soil solution are the primary and secondary phosphate ions (H₂PO₄- and HPO₄^-2). These phosphate anions exist transiently in the soil solution due to rapid removal by roots and microbes, or reaction with other soil minerals. The P level is highest in young vegetative material and in the canola seed. Figure 21 illustrates the P uptake and level over the 1998 season at the AAFC Melfort, SK Research Centre. Canola seedlings take up P rapidly during early growth, but not as rapidly as N. Studies conducted in Manitoba in the 1960’s showed that canola P uptake in early growth stages was more rapid than oats, flax and soybeans. The P level remains fairly high in the leaves (0.3 to 0.4%) until late flowering when significant translocation occurs into developing pods and seeds. By maturity, 75 to 80% of the P in above-ground dry matter is in the seed. Canola seed contains 0.7 to 0.8% P, about double that of cereal grains. Canola stems and pods at harvest contain only 0.1 to 0.2% P.

Figure 21. Phosphorus Content (%) and Uptake by Canola Over the 1998 Growing Season, Melfort, SK

Canola is an efficient scavenger of soil P even though Brassica species are non-mycorrhizal (mycorrhizae are symbiotic associations between certain soil fungi and plant roots where the fungi contribute to the P nutrition of the plant). Many cereal crops can form these beneficial relationships. In spite of canola being non-mycorrhizal, research has shown that canola takes up more P than cereals. Canola has several mechanisms to achieve this efficient P uptake. Canola has abundant fine roots with the ability to branch and proliferate in zones of higher nutrient content such as around fertilizer bands or granules. In addition to root proliferation in fertilizer zones due to branching, canola roots can increase the root hair number and length in response to low P conditions.

The second mechanism in canola roots that enhances P uptake is solubilization of relatively insoluble mineral P forms. Canola has the ability to acidify the rhizosphere just behind the root tip near the zone of root hair formation. In a recent western Canada growth chamber experiment, the pH of the canola rhizosphere fell up to 0.8 units over five weeks compared to a drop of less than 0.4 units for wheat rhizosphere. Canola absorbed more of the relatively insoluble P forms than wheat. The acid generated by canola roots is predominantly caused by exudation of organic acids such as malic and citric acid. Canola roots also release enzymes (phosphatases) that mineralize phosphate from organic P pools. Cation-anion uptake imbalance may also contribute to rhizosphere acidification when the main form of N uptake is ammonium. However, under western Canadian field conditions, canola takes up the majority of N in the nitrate form.

After phosphate enters the root, there are three barriers to cross before reaching the xylem system that feeds aboveground growth. These barriers are the cell plasma membrane, vacuole membrane and the xylem “loading” site. The rate of phosphate transport across these membranes is affected by the plant’s P status. As the plant P content increases, the P transport rate decreases (feed back regulation). The P uptake rate is often more related to shoot than to root P level. This regulated transport system requires energy. Factors that influence root respiration will affect root P uptake. For example, cold soil or low oxygen content in a saturated soil reduces root respiration and consequently P uptake. There is competition for the phosphate transport system by arsenate. This can impact P nutrition in soils high in arsenate.

The xylem loading system is usually regulated separately from the systems at the plasma and vacuole membranes. Phosphate ions typically are rapidly transported from the roots to the shoots. Unlike N, P is absorbed and transported throughout the plant in the inorganic form (mainly H₂PO₄). Similar to N, phosphate is readily remobilized from aging tissue such as leaves to more active growing points. Phosphate stored in cell vacuoles can also be readily mobilized. Immature plants adequately supplied with P have 85 to 95% of the total inorganic phosphate stored in the vacuoles. In contrast, in P deficient plants, almost all the phosphate in leaves is found in active pools (cytoplasm and chloroplasts). By maturity, most of the plant P is stored in organic form as phytate in the grain. Phytate serves as a readily accessible P source for the germinating seedling. Animal nutritionists are interested in seed phytate (including canola meal) since these compounds interfere with absorption of minerals such as zinc, iron and calcium. Considerable attention has been given to reducing phytate levels in grains, including canola, and some success is being reported.

Phosphorus Effects on Canola Growth and Deficiency Symptoms

Canola plants suffering from strong P deficiency can experience slow leaf expansion, smaller and fewer leaves. Deficiency symptoms appear by the second week of growth since canola seedlings are able to obtain sufficient P from seed reserves for the first week of growth. Figure 22 (of field research results from five sites in western Canada in 1991) illustrates the significant increase in early season growth with P fertilization.

Phosphorus deficient leaves may have a dark green, bluish green to purplish colour since chlorophyll and protein formation are less affected than cell and leaf expansion. Under severe P deficiency, purple colouration arises from accumulation of anthocyanin pigments. Mildly deficient plants may look normal but are small. Above-ground plant P content at flowering should be above 0.24%.

Figure 22. Effect of P Fertilizer (P2O5) on Canola Dry Matter after Emergence

Root growth is less affected by P deficiency than shoot growth, leading to a typical decrease in the shoot-root ratio. With a more severe deficiency, root development is restricted, but not as dramatically as stem and leaf growth. Although overall root branching is restricted in P deficient soils, root hair length and density usually increase.

P deficiency affects the maturity and development of reproductive tissue. Even a mild P deficiency can result in maturity delays of several days compared to plants with adequate P. In addition to a flowering delay, a P deficiency can reduce the number of flowers and seeds per pod. Also, a P deficiency can cause leaves to die and drop early, which contributes to the overall yield loss.

Canola Response to Phosphorous Fertilizer

Most agricultural soils in Canada have inadequate P for producing canola crops. However, the canola yield response to P fertilizer on deficient soils usually is much less than the average response to N fertilizer on N deficient soils. Research in the 1960s showed that rapeseed often responded more to P fertilizer than wheat or flax. Subsequent research established that yield response could be predicted from soil test values. Central Alberta research in the 1980’s found that 23 of 48 sites responded to P fertilizer. A recent Alberta P study from 1991-1993 found a statistically significant response to P fertilizer at 42 site-years while 81 site-years had no response. Economic analysis of the results suggested that 70% of the canola sites responded to 7 kg (15 lb) P₂O₅/ac and 53% responded economically to 14 kg (30 lb)–given canola at $352/tonne ($8/bu)–and $0.75/kg ($0.34/lb) P₂O₅. Data from such fertilizer experiments are compiled into databases to predict fertilizer response. Figure 23 is an example of canola response to P fertilizer based on soil test P in Alberta.

Canola response to P fertilizer depends mainly on the amount of plant available P in the soil but is also influenced by moisture and temperature. In cold soil, P availability and movement is reduced. Canola response to P fertilizer is greater under these conditions. Phosphorus fertilization often slightly advances maturity of canola crops by one or two days. This slight difference may be important in short growing seasons.

Figure 23. Phosphate Fertilizer Recommendations for Canola on Medium to Fine Textured, Neutral Soil under Medium Moisture

Phosphorus Fertilizer Effect on Canola Quantity

P fertilization generally has negligible effects on canola quality. Experiments in western Canada have found that P fertilizer increased, decreased or did not affect oil content. Canola protein content has occasionally been slightly raised by P fertilization. A recent field experiment in Manitoba on two very deficient sites found that P fertilizer significantly increased both protein and oil content.

Phosphorus Cycle

Prairie soils contain significant amounts of total P—450 to 907 kg P/ac (1,000 to 2,000 lb P/ac). However, most of this soil P is relatively insoluble with limited availability to plants. Canola roots obtain P by absorbing phosphate dissolved in the soil water. Since the amount of phosphate dissolved in the soil water is very small at any given moment, there must be constant replenishment into the soil water from the insoluble forms. This replenishment of soil solution P around roots arises from slightly soluble minerals, P desorption from surfaces, organic P mineralization and fertilizer. Figure 24 depicts the “P cycle.”

Figure 24. The Soil Phosphorous Cycle

Both organic and inorganic P forms occur in soil, and both are important sources of plant available phosphate in soil water (soil solution P). Primary and secondary phosphate ions (H₂PO₄- and H₂PO₄^-2) can be present in soil solution, with H₂PO₄ - the major form at pH <7.2. This solution P has several possible fates: it may be absorbed by roots, adsorbed to mineral surfaces, precipitated with various cations such as Ca⁺², or immobilized into microbial biomass and soil organic matter. Soil phosphate supply is usually highest in the pH range of 6.5 to 7.0. At high pH levels (>7.5), calcium and magnesium cations can precipitate with phosphate to form salts with low solubility. In contrast, in acidic soils (pH<6), iron and aluminium cations react with the phosphate to form insoluble compounds. Phosphate is not a mobile nutrient in soil due to these soil constituent reactions.

The natural soil weathering process causes acidification and this encourages the eventual conversion of primary P to secondary minerals and unavailable forms (occluded P). This transformation to unavailable forms takes centuries.

As phosphate is removed from the soil solution, the lower level stimulates phosphate release from exchangeable and labile inorganic pools. As labile pools are depleted, nonlabile secondary P minerals slowly dissolve to maintain the labile and solution pools.

The organic P pool also contributes to the maintenance of phosphate in the soil solution. Organic P in prairie surface soil constitutes about 25 to 55% of total P and is a large pool of potential plant available P. Microbial processes drive the organic section of the P cycle. Phosphate from organic matter can be released through decomposition and then incorporated into new microbial biomass or enter into the soil solution. Most organic P compounds released during decomposition are quickly degraded and exist briefly in soil. Some organic P compounds can be stabilized in soil through adsorption to soil constituents or by physical isolation within aggregates. Tillage decreases the soil organic P content by exposing stabilized forms to new or more vigorous microbial attack. Organic P can be degraded to phosphate by enzymes (phosphatases) released by soil microbes and by canola roots.

Significant seasonal fluctuations occur in both organic and inorganic P pools. However, reports conflict on the direction and magnitude of the fluctuations. For example, several experiments reported decreases in organic P during the summer growing season and gains over the winter, while another experiment measured major declines over the winter. Inorganic P (soil test extractable P) can also vary widely and inconsistently between fall and spring.

Phosphorus Fertilizer Management for Canola

The majority of P fertilizer is not absorbed by canola in the application year. Instead, most of the fertilizer P reacts with soil constituents to form relatively insoluble salts or stabilized P compounds. Timing and placement are two management strategies used to maximize P fertilizer uptake and yield response.

Timing of Phosphorus Fertilization

Since phosphate reacts with soil constituents to form insoluble compounds over time, P fertilizer efficiency can be increased by limiting the time from application to crop uptake. However, the P fertilizer application date affects canola yield much less than placement method. Most growers apply P fertilizer at seeding, minimizing P availability losses by reducing reaction time. Research in western Canada has shown the effectiveness of fall and spring banding of P fertilizer is similar.

Phosphorus Fertilizer Placement

Phosphorus supply during the first two to six weeks of canola growth is critical to achieve optimal yield. Therefore, place P fertilizer to maximize early season access.

Seed row placement is an effective placement method when soil P levels are low to moderate and spring soil conditions are cold. Cold soil decreases phosphate solubility and diffusion in the soil solution, slowing P movement to roots and root uptake rates. This condition increases the likelihood of response to readily accessible seed row P (the “popup” or “starter” effect). Unfortunately, canola seedlings are sensitive to seed row fertilizer and this limits seed row P rates. The maximum safe rate of seed row P fertilizer for canola depends on seedbed utilization and soil moisture conditions. Pot experiment results (Figure 25) conducted at AAFC’s Beaverlodge, AB Research Centre in the 1960s illustrate canola’s sensitivity to seed-placed P fertilizer. The seedbed utilization in this experiment was very restricted (3%), even less than a double-disc press drill. The dry and moist soil corresponded to 30 and 50% of available water in a sandy loam.

Figure 25. Effect of Seed-Placed P Fertilizer on Canola Emergence

Subsequent field research has confirmed that excessive P fertilizer placed in the seed row can reduce plant populations and yield. High seed-placed P fertilizer rates have lowered plant populations in some cases but did not affect yield. However, lower plant populations due to excessive seed row fertilizer will increase yield variability and usually lowers high yield potential under optimal growing conditions.

Under dry soil moisture conditions with low seedbed utilization (such as disc opener), the maximum safe P₂O₅ seed-placed rate is approximately 22 kg/ha (20 lb/ac). The rate can be safely increased to 28 kg/ha (25 lb/ac) under good moisture conditions with low seedbed utilization. As seedbed utilization increases, proportionally increase seedplaced P fertilizer rates. Some research suggests that the larger seed of B. napus will tolerate slightly more seedplaced P than B. rapa. Due to the significant emergence and yield reductions caused by moderate to high rates of seedplaced P fertilizer, place these rates separately from the seed. This fertilizer/seed separation can be achieved by increasing the spread width in the seed row or by placing the fertilizer in bands away from the seed row.

Pre-plant band placement is an effective method since it reduces fertilizer contact with sensitive canola seed and with soil constituents that will fix P over time. Also, the deeper fertilizer placement tends to be more accessible to roots as they normally grow down to moist soil. Banding fertilizer prior to seeding reduces the fertilizer handled during seeding and can provide time and labour benefits. Pre-plant band placement is currently a common method for placing all fertilizer. Phosphorus fertilizer can be banded in late fall or spring prior to seeding.

Side-banding places fertilizer near the seed row during seeding. The fertilizer normally is banded 2.5 to 5 cm (1 to 2") below and beside the seed row. Several direct seeding machines use a mid-row or paired-row method of banding fertilizer. Air seeders with shovels or knives are used with shank spacings ranging from 20 to 36 cm (8 to 14"). The fertilizer is usually banded to a depth of 5 to 13 cm (2 to 5"). No consistent agronomic benefits accrue to banding deeper than 8 cm (3") and fuel costs increase significantly with deeper depths. In high P-fixing soils, place fall P bands deeper than subsequent tillage depths to avoid mixing the band with soil.

Split application methods refer to combinations of band and seed row placement. Split application takes advantage of the consistent benefit of seed-placed P fertilizer up to 22 kg/ha (20 lb P₂O₅/ac), and avoids seedling injury by placing the remainder of the P fertilizer in a band (usually with N and S).

Broadcast-incorporated placement involves spreading P fertilizer on the surface followed by cultivation to work it into the soil. This method is significantly less effective than seed-placed or banded P fertilizer due to increased contact between the P and reactive soil constituents. Application rates with broadcast-incorporated P fertilizer usually have to be two to four times seed-placed or banded rates to get an equal response. Therefore, broadcast-incorporated methods are less economical.

Research comparisons of P fertilizer placement methods at typical rates show that highest yields are frequently obtained with seed-placed and split applications, followed closely by pre-plant band methods. Broadcast-incorporated methods produce significantly lower yield responses.

Research by Westco Ag research illustrates the relative usefulness of various P fertilizer placements (Figure 26). Differences between placement methods are largest under conditions of low soil test P (such as after forage breaking), as well as cold spring soil. On typical prairie farms that have received P fertilizer for many years, canola yield response differences between seed-placed and banded methods tend to be minimal. Phosphorus fertilizer placement issues have been largely resolved by ground opener development with increased seed row spread or side-band capability. Split P application between a band and seed row appears to be the most consistent method due to reduced seed row toxicity, less P uptake interference from high N bands, and a dual location that hedges against poor access in either cold or dry surface soil conditions.

The application of both N and P fertilizer in a single band is called dual banding. At low to moderate rates of N, the uptake efficiency of P is sometimes increased from a dual band. At higher N rates—above 90 kg/ha (80 lb/ac)—the concentrated N in the band can reduce early season P uptake due to ammonia and nitrite toxicity that hinders root entry into the band. This P uptake interference appears to be strongest in recent band applications, and could be a problem with dual spring banded N+P fertilizer immediately before or during seeding.

Figure 26. Canola Yield Response to Different P Fertilizer Placements

Non-Traditional Sources of Phosphorus Nutrients

Phosphorus deficiencies on the prairies are normally corrected with annual applications of commercially refined P fertilizer—either dry blends of mono-ammonium phosphate (12-51-0) or liquid blends of ammonium polyphosphate (10-34-0). Manure also serves as a traditional source of P and other nutrients.

Rock Phosphate

Canola has an ability to absorb native soil P through acidification of the rhizosphere. Pot experiments have demonstrated that canola can utilize more rock phosphate than other crops, apparently due to the rhizosphere acidification. This has prompted promotion of rock phosphate as a viable alternative P fertilizer for canola. Rock phosphate is the relatively insoluble, gray-black powdery material that is refined by fertilizer manufacturing plants into soluble phosphate fertilizer.

Idaho is a common source of rock phosphate marketed in western Canada. Research on the prairies indicates that rock phosphates do not perform satisfactorily compared to fertilizer phosphate. The poor performance is due to poor solubility, lower P₂O₅ content, and the predominance of neutral, calcareous soils on the prairies. While high rates of rock phosphate do slightly improve canola yields on some soils, this is not cost effective compared to fertilizer phosphate. Typically, rock phosphate application rates need to be six to eight times that of fertilizer phosphate for equivalent yield response. Research by Alberta Agriculture Food and Rural Development at Ellerslie, AB illustrates the poor performance of rock phosphate compared to fertilizer P (Figure 27). All the P sources were seed placed and 112 kg N/ha (100 lb N/ac) was pre-plant banded.

Figure 27. B. Rapa Yield Response to Rock Phosphate and P Fertilizer

Another non-traditional means of P nutrition recently developed for canola is biologically based. Although canola does not form symbiotic associations with mycorrhizae that improve P nutrition in other crops, other rhizosphere microbes exist that increase P solubilization and subsequent plant P uptake. AAFC, Lethbridge, AB researchers identified an organism (Penicillium bilaii) that solubilized P minerals and improved the P uptake of cereals and canola. This organism was then commercialized as a seed inoculant (Provide®). Field experiments conducted on the prairies have shown that inoculating canola with Provide increases early season P uptake and vegetative growth, and results in higher yield with and without P fertilizer. The canola yield response to inoculation with Provide at 15 P-responsive sites in western Canada is summarized in Figure 28.

Figure 28. Canola Yield Response to Inoculation with Provide at P-Responsive Sites

On average, growers can expect to apply 11 kg less P₂O₅/ha (10 lb less P₂O₅/ac) when canola is inoculated with Provide. The adoption of Provide inoculation by canola growers has been limited, perhaps due to the inconvenience of inoculation, the short viable period after inoculation, inconsistency and cost relative to simply using more P fertilizer. In the future, biological fertility enhancing microbes will likely become more common.

Figure 29. P-sufficient Canola (left) Compared to P-deficient Canola

Photo Courtesy Adrian Johnston, PPI

Figure 30. P-sufficient Canola at Flowering (left) Compared to P-deficient Canola

Photo courtesy Adrian Johnston

Figure 31. A P-sufficient Canola Pod (top) Compared to a P-deficient Canola Pod

Photo courtesy Adrian Johnston

Potassium (K)

The macronutrient potassium (K) is required in large amounts by canola similar to nitrogen (see Table 3). In spite of the large requirement, canola yield responses to K fertilizer (potash) are infrequent, due to ample soil K reserves on the prairies, and canola’s strong ability to absorb K.

Role of Potassium in the Canola Plant

Potassium is different from most other essential nutrients since it does not become part of structural components in the plant. Instead, most of the K in plants remains dissolved in the cell sap and performs several major functions.

One major function for K is that of enzyme activation. Enzymes are protein complexes that catalyze chemical reactions. More than 60 enzymes need to be activated by K. This activation occurs when potassium cations (K+) bind to the enzyme surface, changing the enzyme shape, and allowing the enzyme’s active site to attach to its substrate more rapidly or accurately. For example, K stimulates the activity of an enzyme (starch synthase) that catalyzes starch formation from glucose. While other cations can also stimulate this enzyme, K+ is the most effective. In K deficient plants, the lack of stimulation of the starch synthase results in an accumulation of soluble sugars and N compounds, and a decrease in starch.

Another major function of K is in water relations. Potassium helps to maintain a favourable water status in plants in several different ways. Potassium cations dissolved in cell sap perform major osmotic functions. Osmosis is the tendency for water levels to equalize between different areas separated by a porous membrane. Dissolved ions such as K+ attract water and thus are osmotically active substances. Potassium is the major dissolved ion in cell sap and provides most of the osmotic “pull” that draws water into roots.

Potassium cations also maintain the water relations in plants through their crucial role in regulating water loss (called transpiration) from pores (stomata) in the leaves. Although the stomata must open to allow movement of carbon dioxide and oxygen in and out of the leaves, water loss also occurs. This transpiration creates a gradient that pulls water and nutrients up through the xylem to the leaves. However, plants cannot afford excessive water loss and need to regulate the stomata opening. For example, photosynthesis stops during darkness, and the need for nutrients and water decreases greatly during night. Plants have developed a system that closes stomata during the dark or during drought. Potassium cations, in combination with chlorine, calcium and certain hormones, are responsible for governing the opening and closing of the stomata. Upon receiving a “signal” induced by darkness, K+ and Cl- are pumped from the two guard cells surrounding the stomata, which causes a loss of turgidity of the guard cells and thus allows the pore to close. Potassium deficient plants often have higher transpiration rates and display wilting.

Potassium’s osmotic activity also provides the physical force that expands cells during growth. New cells accumulate K+ and associated anions like Cl- in the large central vacuole that occupy 80 to 90% of the cell volume. The K+ ions attract water and inflate the cell, stretching it to a new larger size. Potassium-deficient plants can exhibit low growth rates and small cells.

Energy relations in the plant are influenced by K. Potassium affects photosynthesis at several levels. K+ is the main ion that counterbalances the H+ flux during photosynthesis in the chloroplasts. Potassium also maintains a favourable pH gradient in the chloroplasts for making phosphate energy compounds. Potassium helps the translocation of photosynthate sugars by maintaining a high pH in phloem tubes needed for “loading”, and by maintaining osmotic gradients needed for sap flow.

Potassium is needed for N uptake and protein synthesis. K+ cations are the major counter ions that balance nitrate during transport and storage in vacuoles. Many steps of protein synthesis require high K+ levels.

The K level is highest in seedling canola, then declines steadily up to maturity as shown in Figure 32, B. napus canola grown at the AAFC Research Centre in Melfort, SK in 1998. Canola K uptake is rapid during the early growth stages and tapers off by the end of flowering. Under high K fertility and good growth, canola can absorb more K than apparently needed, a situation termed “luxury consumption.” As canola matures, the K level in leaves declines while the stem level increases. By harvest, the stem and straw material contain about 1 to 2% K. In contrast to N and P, the K content of the seed (0.8 to 1% K) is low relative to the stem. Unlike K+ in the vegetative parts, seed K is probably complexed with phytate as a salt.

Potassium Effects on Canola Growth and Deficiency Symptoms

Potassium deficiency reduces overall canola growth but to a lesser degree than N or P deficiency. Since K is mobile within the plant, deficiencies are first visible in older leaves. The edges and areas between veins of older leaves tend to turn pale green or yellow, followed by withering. The yellowing can occur first in middle leaves before older ones if observed at bolting to flowering stages. In severe cases, leaves die but remain attached to the stem. Small white spots can develop on leaves. Plants are prone to wilting during midday. Potassium deficiency symptoms in canola are rather nondistinct and can be easily confused with other problems. Fortunately K deficiency in canola is rare on the prairies.

Figure 32. Potassium Content (%) and Uptake by Canola over the Growing Season

Canola Response to Potassium Fertilizer

Although canola absorbs large amounts of K, responses to fertilizer K are rare on the prairies. In fact, canola or rapeseed responses to fertilizer K are infrequent around the world—a testament to the crop’s strong ability to absorb soil K.

Numerous fertilizer research studies on the prairies and in Ontario have established that canola rarely responds to applied K, even under conditions where cereals normally respond. Although the K soil test is adequate for cereals, the usefulness declines for canola. Critical levels are often stated to be around 280 kg K/ha (250 lb K/ac) or 112 ppm in the top 15 cm (6"), but research indicates that canola will not consistently or economically respond to fertilizer K unless the soil test is very low—78 to 112 kg K/ha (70 to 100 lb K/ac) or 35 to 50 ppm. Very sandy or peaty soils are the most likely soil types to have very low K soil test values. Other factors that increase the likelihood of K deficiency are:

• free lime in the rooting zone
• acid soil
• poor drainage
• cool temperatures
• soil compaction
• shallow root zone

Unlike cereal responses, potash applications have not been shown to help with canola disease resistance, lodging or seed quality (oil content or meal protein content).

Potassium Supply from the Soil

Western Canadian soils generally contain ample plant available K due to an abundance of K minerals (such as mica and feldspar) in the parent material (3 to 4% K). There is often 17,000 to 56,000 kg K/ha (15,000 to 50,000 lb K/ac) in the top 15 cm of prairie mineral (non-peat) soils. The weathering of these minerals slowly releases K+ held in crystal structures—typically only about 1% of total soil K is available for plant uptake. This available K is mostly (90%) exchangeable K+ adsorbed to clay surfaces and organic matter, while the other 10% is found dissolved in the soil solution. Approximately 10 to 20% of the total soil K is slowly available from smaller mica particles and certain clays. Figure 33 outlines these K pools. Losses due to leaching or erosion are ignored in this figure, as they are usually small.

Figure 33. Potassium Soil Cycle

Figure 33 shows that the various pools are in dynamic equilibrium. As K+ is removed by plant uptake and through leaching on sandy soils, additional K is released from the mineral soils to become available. Available K moves to plant roots by diffusion through the soil only up to 6 mm (1/4"). Therefore, the equilibrium process that repeatedly moves K from the slowly available to readily available pool is very important for K nutrition. The rate of movement from the slowly available to readily available pool varies among soils due to differences in minerals and clays. This variation in K dynamics creates problems for soil testing. An extractant that measures plant available K in soil solution and exchangeable K over a short time period does not assess the replenishment power. Unfortunately, tests that measure the replenishment power are time-consuming and cost-prohibitive.

Potassium Fertilizer Management for Canola

Potassium is relatively immobile in the soil since the K+ cations are readily adsorbed to the negative surface charges on clay particles and organic matter. Potassium can also be fixed into the clay lattice structure of certain clay types. Potassium is more mobile in sandy soil and thus can be leached in these soil textures. In most soils, K+ is much less mobile than nitrate but somewhat more mobile than phosphate. This relative immobility means that fertilizer placement will greatly affect uptake efficiency. Ensure application methods minimize contact with soil and increase root contact. Banding and seed-placed methods can achieve good uptake efficiency. Since canola responses to K fertilizer are rare on the prairies, there has been limited K placement research in this crop.

Seed-placed K fertilizer is an efficient application method but the high salt index of potash fertilizer limits the amount that can be safely applied near the seed. Canola has a much lower tolerance to seed-placed potash than cereals, and stands will be reduced if seed-placed K rates exceed 17 kg K20/ha (15 lb K20/ac) with drills that have low seedbed utilization (such as double disc drills). Higher rates of potash fertilizer can be safely seed placed as the seedbed utilization is increased. If other nutrients such as N or P are also seed placed, this reduces the safe rate of seed-placed K. Good seedbed moisture, higher clay and organic matter contents help reduce the severity of seedling damage from seedplaced K fertilizer. However, most K deficient soils are sandy, and are sensitive to seed-placed K.

Due to canola’s sensitivity to seed-placed K fertilizer, a band placement away from the seed row is more advisable. Sideband placement is an efficient method and the separation between fertilizer and seed reduces the risk of germination damage. Openers with side-band capability are becoming available, especially for direct seeding implements. Deep banding prior to seeding should also be an efficient and safe method of K fertilization. Potash fertilizer can be banded together with other nutrients. Banding efficiency should not differ greatly between fall and spring unless the soil is very sandy and subject to leaching loss under conditions of high snowmelt or spring rainfall.

The broadcast-incorporation application method is less efficient, and probably requires rates double that of banding to achieve a similar crop response. However, in situations where banding equipment is not readily available and seed placement is too risky, broadcast incorporation may be useful and not overly expensive due to the relatively low cost of potash fertilizer. The higher fertilizer rates necessary for broadcast K may also benefit subsequent crops with a higher K response than canola.

Figure 34. Leaf Edge Scorch Due to K Deficiency

Photo courtesy Adrian Johnston

Figure 35. Scorched Leaf Due to Severe K Deficiency

Photo courtesy Adrian Johnston

Figure 36. Severely K Deficient Pods (top) and Sufficient (bottom)

Photo courtesy Adrian Johnston

Figure 37. Sufficient Pod (left) and Severe K Deficient Pods

Photo courtesy Adrian Johnston

Sulphur (S)

Sulphur (S) is the fourth macronutrient, but ranks as the third most limiting nutrient on the prairies. Sulphur deficiency in western Canada was first identified in 1927 on Gray Wooded soils in Alberta. Canola is more sensitive than cereals to S deficiency and frequently responds to fertilizer S addition. Therefore, pay equal attention to N, P and S.

Role of Sulphur in the Canola Plant

As shown in Table 3, canola contains large amounts of S. Sulphur is part of structural and enzymatic components. Sulphur is a key component of two essential amino acids (cysteine and methionine) and is needed for protein synthesis. Chlorophyll synthesis also requires S. Both of these amino acids are also precursors for coenzymes and secondary plant substances. Glutathione, an important antioxidant in plants and animals, is synthesized from cysteine. Glutathione contents are higher in leaves than roots. It’s found primarily in the chloroplasts where its anti-oxidant ability is needed to detoxify free radicals generated during photosynthesis. Glutathione also functions as transient S storage, and a precursor of phytochelatins (compounds which detoxify heavy metals in plants). Thioredoxins, another important group of S compounds related to glutathione, help activate several enzymes in carbon metabolism. Sulphur also is part of several enzymes and coenzymes such as ferrodoxin, biotin (vitamin H), coenzyme A, urease, and thiamine (vitamin B1).

An important group of secondary plant S compounds in canola are glucosinolates. Plants contain over 100 different glucosinolate compounds. These secondary compounds, although not well understood, probably have a number of functions. Glucosinolates are stored in cell vacuoles, and can be broken down by an enzyme (myrosinase) to yield glucose, sulphate and volatile compounds such as isothiocyanate. Glucosinolates contribute to defence or attractant systems for certain insects and diseases. When plant cells are destroyed by insect feeding, glucosinolates are broken down, releasing various deterrents/attractants.

Glucosinolate levels are highest in growing points, roots, and youngest leaves, all of which are most vulnerable to insects and diseases. The role of glucosinolates as S reserves to maintain plant S during periods of high demand (such as bolting, flowering, podding and seed fill) is controversial. However, recent research in Europe showed that glucosinolates comprised a small S pool in leaves, and under induced S deficiency, sulphate (SO4-2) mobilization from storage in cell vacuoles was about 10 times greater than contributions from glucosinolates.

Sulphur is also a constituent of sulpholipids, which are membrane components.

Characteristics of Sulphur Uptake by Canola

The main S form absorbed by canola roots is sulphate. In industrial areas, atmospheric S compounds dissolved in rain can be absorbed by leaves. However, this amount is quite small and is decreasing with better air pollution control. sulphate absorption is accomplished with active transport systems across membranes. The uptake rate increases as the sulphate level increases in the soil water. Low plant S contents also increase the root uptake rate. Negative feedback signals for S uptake may be sulphate or glucosinolate levels in vacuoles, or the levels of organic S compounds such as cysteine, methionine or glutathione. Sulphate uptake faces competition from molybdenum and selenium. Therefore, soils high in these minerals can experience antagonism with S uptake.

The S level in canola plants is highest in the early seedling stage when young leaves comprise most of the dry matter (F