Crop Nutrition

Table of contents

    Basic Plant 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.

    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.

    Other factors such as improper management or pests can also result in 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 selectively absorb required elements for their growth and they also can take up elements not required for growth.

    The terms essential plant nutrient or essential mineral element describe the minerals needed by plants to grow and complete their life cycle. 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 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


    Plant nutrients are classified by the relative amounts needed for each macro and micronutrient. Macronutrients are needed in large amounts relative to micronutrients. Tables 1 and 2 show the relative amounts of nutrients contained in a typical canola crop. Some nutrients can also accumulate at higher levels than what is 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

    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, 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 in 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 negative charge 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 signaled 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 rapeseed in the vegetative state 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.

    The rate of nutrient uptake by canola varies by nutrient. For N and S, nutrient uptake is most rapid in earlier growth stages up to the middle flowering stage of the crop.  Phosphate uptake is more uniform throughout the growing season.  These uptake patterns show the importance of the 4R principle of right source at the right time at the right place and the right rate.  For instance, the mid-season demands for sulphur illustrate why in-season application of ammonium sulphate can correct an S nutrient deficiency.

    Table 3 shows the daily nutrient uptake of several macronutrients at different crop stages.

    Table 3. Nutrient uptake by crop development stage for canola at Melfort in 1998.
    Crop development stagelbs/day
    Emergence to 5-leaf 0.77 0.14 0.88 0.11
    5-leaf to first flower 3.85 0.66 5.53 0.68
    First flower to 50% flower 3.46 1.01 4.44 1.17
    50% flower to end of flower 0.96 0.57 0.83 0.69
    End of flower to 50% podded 0.92 0.48 0.77 0.65
    50% podded to full pod 0.05 1.08 1.81 0.50
    Full pod to harvest 0.05 0.14 0.17 0.05

    Source: Johnston et al., unpublished data

    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 directly or indirectly influence soil fertility.

    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")

    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. Nitrogen, on the other hand, is not a constituent of any soil mineral.

    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. Soil texture influences fertility by affecting moisture holding capacity, air exchange and CEC. Because weathering is very slow in prairie soils, mineralogical composition is fairly uniform and, therefore, texture can be a good indicator of CEC. 

    Adequate moisture is key to fertilizer response and achieving potential yield in canola in western Canada. Research on the Prairies found that seed yield could vary from 1.8 to 3.3 kg/mm, which is equivalent to 2 to 3.7 bu/inch of moisture. (Karamanos et al)

    Seed yield per unit of water
     Probability of precipitation, %
     kg mm-1
    Palliser Dry Plain 1.8 2 2.2
    Palliser Plain 2 2.2 2.5
    Parkland 2.5 2.7 2.9
    Moist Parkland 2.9 3.1 3.3
    Moist Parkland Transition 3.1 3.3 3.3
    Moist Parkland Wooded 2.9 3.1 3.1
    Humid Parkland 2.6 2.9 3.1
    Alberta Highlands 2.9 3.1 3.3
    Alberta Highlands West 2.9 3.1 3.3
    Dry Peace Country 2.9 3.1 3.3
    Peace Country 2.9 3.1 3.3
    Moist Peace Country 2.9 3.1 3.3
      Probability of precipitation, %
      75 50 25
    Palliser Dry Plain 2.0 2.2 2.5
    Palliser Plain 2.2 2.5 2.8
    Parkland 2.8 3.0 3.2
    Moist Parkland 3.2 3.5 3.7
    Moist Parkland Transition 3.5 3.7 3.7
    Moist Parkland Wooded 3.2 3.5 3.5
    Humid Parkland 2.9 3.2 3.5
    Alberta Highlands 3.2 3.5 3.7
    Alberta Highlands West 3.2 3.5 3.7
    Dry Peace Country 3.2 3.5 3.7
    Peace Country 3.2 3.5 3.7
    Moist Peace Country 3.2 3.5 3.7

    Source: Karamanos, R.E. and Henry, J.L. 2007.  A Water Based System for Deriving Nitrogen Recommendations in the Canadian Prairies.  Chapter 11, in. T. Bruuslema (ed.) Managing Crop Nitrogen for Weather. Proceedings of the Symposium “Integrating Water Variability into Nitrogen Recommendations” International Plant Nutrient Institute, Norcross, GA.

    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. A number of cations (e.g., K, Mg, Ca, Fe, Al) are part of the clay mineral structure and were ‘fixed’ when the soil was formed from its parent material.   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+). 

    All cations in soil are accompanied by a ‘hydration cell’, i.e., molecules of water, which affects the size and consequently the reactivity of cations.  For example, K has a smaller hydration cell than Ca or Na and reacts faster in soil.

    Soil negative charges arise due to substitutions in the mineral crystals by elements with smaller positive charge, and due to reactions at the edges. 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 extent of weathering. 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 particles also contain a much higher number of negative charges than clay colloids. 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. The CEC soil property allows a reservoir of nutrients to be stored then released to plant roots. There is a strong relationship between cations in the soil solution and those on the exchange phase of soil colloids; the more cations on the exchange phase, the more in the soil solution.  That is why most of western Canadian soils that are rich in K, Ca, and Mg minerals have ample supply of “available” forms of these elements (nutrients). A high CEC also means that fewer cations will be lost through leaching below 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 more prone to leaching loss.

    Soil organic matter (OM) plays an important role in soil fertility as a plant nutrient storehouse, especially since it is the only indigenous source of N in soils. 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. Mineralization also influences P availability with research showing that 50% of P availability comes from mineralization of organic matter. Mineralization of individual nutrients will be described in later sections. Soil OM also plays an additional role in soil fertility by improving physical properties such as water holding capacity, infiltration, aggregation (tilth) and buffering pH.

    Role of Nitrogen in Canola Plants

    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.

    Canola, like most crops, contains large amounts of N, which 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 1.

    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 and reaches the flowering stage, the overall N level declines due to leaf loss. By maturity, canola straw contains just 0.5 to 1.5% N while the seed contains 3.4 to 4% N.

    Figure 1

    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 2, 3, 4). 

    Figure 2

    Figure 3

    Figure 4

    Several western Canadian research studies have determined the seasonal plant growth and nutrient uptake of canola. In general, nutrient uptake increased with time at early growth stages and reached a maximum 4 weeks after emergence.  Total N accumulation in canola plants was highest 9 to 10 weeks after emergence. 

    Nitrogen 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 early in the season is necessary to produce a large, photosynthetically efficient leaf area that will support high numbers of flowers, pods and seed yield.

    Hybrid versus conventional canola

    Hybrid canola often demonstrates a greater yield response to N application compared to open-pollinated canola as shown in Figure 7.  Under an identical fertilizer regime, hybrid canola produced a 17% higher yield, indicating that hybrid canola is a more efficient scavenger of nutrients than open-pollinated canola. (Karamanos et al. 2005).  In contrast, other studies have not found statistically significant N response differences between hybrid and open-pollinated types (Mahli et al., 2007; Karamanos et al., 2007).  Almost all canola seeded in western Canada are now hybrid varieties.

    Nitrogen Cycle and Transformations

    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.

    Soil organic matter

    The decomposition rate is fairly slow and variable. As shown in Figure 8, organic N is first slowly changed to ammonium (NH4) by bacteria. Then different bacteria rapidly change the ammonium to nitrate (NO3), 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. The net amount of mineralization also depends on the level of organic matter.  The higher the soil OM, the greater amount of N will can be mineralized.  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, the breakdown of soil aggregates, and physical mixing that gives soil microbes access to new organic matter supplies.

    Long term no-till

    Long term no-till is beneficial for N cycling, and the increase in potentially mineralizable N under favourable climatic conditions.  A comparison of canola yields grown on adjacent fields found the area that had been in no-till for 31 years had 16% higher yield than the short term area consisting of a 9-year no-till field. These higher grain yields may be achieved without necessarily having to add additional crop inputs like fertilizer.

    There are several sources of plant available N to the soil system, in addition to soil organic matter decomposition and fertilizer additions (Figure 8). 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. Limited research on canola inoculation with associative N fixing bacteria 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 contributes to variable losses, and under good management practices, usually is less than 10% on Prairie soils. (Campbell and Rennie, 1984)

    As shown in Figure 8, 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.

    Considerable N denitrification losses occur during spring thaw. For example, research near Edmonton, AB found that 16 to 60% of the total amount of N lost annually to denitrification  was lost immediately following snowmelt (Note: This does not mean 60% of total soil N was lost, just that up to 60% of losses occurred in that time period). 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. 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 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.


    Immobilization is the second major N transformation that reduces the plant available N supply. Figure 8 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 (e.g. banding) and thus reduce immobilization losses.


    Leaching of nitrate can occur since this form is present in the soil solution 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, use ammoniacal forms of N such as anhydrous ammonia and try to avoid situations where fertilizer  applied to soils is likely to be exposed to wet conditions for long periods of time..

    Summerfallow is also a contributor to leaching losses of nitrate. However, research showed that 20 inches of water applied to a sandy soil that was summerfallowed was necessary to move N below the 24 inch depth. (Karamanos, 1979)


    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, place fertilizer into the soil. Growers can also consider the use of urease-inhibitors (Agrotain-type products), or slow-release fertilizers like ESN urea.


    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 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.

    Phosphorus (P)

    Phosphorus, although an important plant macronutrient, is only required in small amounts compared to 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.

    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.

    Canola seedlings take up P rapidly during early growth, but not as rapidly as N.

    Canola P uptake in early growth stages was more rapid than oats, flax and soybeans. Daily P uptake is highest in earlier growth stages.

    The P level remains fairly high in the leaves (0.3 to 0.4%) until late flowering when significant translocation to developing pods and seeds occurs. 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.

    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 Cycle

    Prairie soils contain significant amounts of total P -- 180 to 907 kg P/ac (400 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 12 depicts the P cycle.

    Figure 12. The Soil Phosphorous Cycle

    Figure 12. 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 aluminum 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, non-labile 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 breaking down soil aggregates and 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.

    Potassium (K)

    The macronutrient potassium (K) is required in large amounts by canola similar to nitrogen (see Table 2). In spite of the large requirement, canola yield responses to K fertilizer (potash) are infrequent, due to ample soil K reserves on prairie sols, 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. The release of K back to the soil is therefore quite rapid.

    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 and properly. 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.

    Canola K uptake is rapid during the early growth stages and tapers off by the end of flowering.  The K level is highest in seedling canola, then declines steadily up to maturity. (Malhi et al. 2007)

    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 (the principle P storage form) as a salt.

    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. Approximately 10 to 20% of the total soil K is slowly available from smaller mica and feldspar particles and certain clays. This is common in southwest Saskatchewan, and moving from SW to NE Saskatchewan, the feldspar and mica content declines, which is why K deficient soils can occur in NE Saskatchewan.

    Figure 15 outlines these K pools. Losses due to leaching or erosion are ignored in this figure, as they are usually small.

    Figure 15. Potassium Soil Cycle 

    Figure 15. Potassium Soil Cycle

    Figure 15 shows that the various pools are in dynamic equilibrium. As K+ is removed by plant uptake, 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 mineralogy and clay content. This variation in K dynamics creates problems for traditional 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.

    The Plant Root Simulator (PRS) from Western Ag Labs uses a negatively charged soil probe to estimate the plant available supply of K, and is another approach in estimating K soil fertility.

    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 2, 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 will antagonize S uptake.

    The S level in canola plants is highest in the early seedling stage when young leaves comprise most of the dry matter (Figure 16).

    Figure 16. Sulphur Content (%) and Uptake by B. napus over the Growing Season

    Figure 16

    As plants develop, the overall S level declines but not as dramatically as with N. By maturity, canola straw contains approximately 0.3 to 0.4% S while pod chaff contains slightly more S (0.5 to 0.6%). Canola seed contains about 0.4 to 0.6 % S. At harvest, canola straw and pod chaff contain roughly twice as much S per acre as that in the seed.

    Sulphur uptake increases rapidly after germination and peaks three to four weeks after emergence. (Malhi et al. 2007). This highlights the importance of early season S availability, but also indicates that S-deficiency can be corrected if top-dressing of ammonium sulphate occurs early enough, and before the canola bolts.

    There has been limited research on the complex S partitioning into the various compounds of different plant parts over the growing season. Most of the plant S ends up in protein and stored sulphate. As leaves senesce, protein-S is readily remobilized, while stored sulphate remobilization is slow and more limited. Therefore, overall, S has medium mobility.

    Sulphur Supply from the Soil

    The organic portion of the sulphur cycle in soil is closely tied to N due to their association in protein. Like N, the main S reserve in soil is in organic matter. Although there is considerable variability in the relative proportions of carbon, N and S (C:N:S) in soil organic matter, the ratios are quite similar for each soil group. In a study of Saskatchewan farm soils, the C:N:S ratio ranged from 58:6:1 in Brown soils, to 63:7:1 in Dark Brown, 83:8:1 in Black, 100:8:1 in Gray Black, and 129:11:1 in Gray soils.

    The soil S cycle is illustrated below in Figure 19.

    Figure 19. Soil Sulphur Cycle 

    Figure 19. Soil Sulphur Cycle

    A key component of the soil S cycle for plant growth is the mineralization path. Soil organic matter and plant residues are decomposed by soil microbes, releasing sulphate. The S mineralization rate is quite slow (much slower than N), and cannot match the uptake rate of growing plants. Also like N, the sulphate amounts released from residues will depend on the S content. When plant residues contain more than about 0.15% S (C:S ratio about 300:1), there will be a net release of sulphate through mineralization. Below 0.15% S, decomposition is slower and there will be immobilization of soil sulphate by soil microbes. The ability of soil to mineralize sulphate from organic matter has been found to be independent of the total amount of C, N or S, and of the C:N or N:S ratios in soils. However, research has also found that the initial amounts of sulphate mineralized from soil are closely correlated with the initial amounts of N mineralized in short-term incubation.

    Another important aspect of the soil S cycle is the oxidation path. In soils, sulphides, elemental sulphur and thiosulphate can be oxidized to sulphate by various soil microbes, but the main actors are bacteria from the genusThiobacillus. The oxidation of these inorganic S compounds produces considerable sulphuric acid. Sulphur oxidizing bacteria are most active under warm, moist, well aerated conditions. It is the oxidizing ability of these bacteria that permits the agricultural use of elemental S for crop growth.

    Although S reduction is shown in the soil S cycle diagram, it generally is not significant in aerated agricultural soils. In flooded soils, sulphate can be reduced by soil microbes to sulphides in a process analogous to denitrification. However, soil microbes will utilize nitrate, iron and manganese compounds before reducing sulphate.

    In many western Canadian soils, there is a subsoil salt (gypsum) and/or lime (calcium carbonate) layer. This subsoil layer contains considerable sulphate, often as coprecipitates with lime. Although this subsoil sulphate solubility is reduced, it still can contribute to plant needs if it exists within the rooting zone. However, the length of time that canola grows in S-deficient topsoil before rooting to the subsoil S will affect the yield response to fertilizer S. Also, the depth to subsoil S tends to vary greatly across the field. Total S amounts (organic and sulphate) generally increase from upper to lower slope positions.

    In most prairie soils, sulphate is not held by organic matter and clay particles since they are both negatively charged. Therefore, sulphate is vulnerable to leaching losses.

    Nitrogen:Sulphur Ratio

    A proper N:S balance is important for canola production. When N is in excess and S is deficient (high N:S ratio), there is insufficient S to combine with the N to make protein, and thus non-protein N accumulates.

    An outdated guideline is to add N and S fertilizer in a 7:1 ratio.  At the moderate to high levels of nitrogen fertilizer needed in areas with high crop yields, S sufficiency is reached far sooner than N sufficiency, and thus the fixed 7:1 ratio would overapply S. Further, the practice of balancing applied N and S in a fixed ratio on soils containing sufficient S levels appears unnecessary and wasteful. (Karamanos et al. 2007)

    Also, canola yield is highly dependent on N fertility. Optimum yields of canola are derived so long as individual N, P and S requirements are fulfilled. Nitrogen requirements to obtain optimum yield of hybrid canola cultivars are higher than that of conventional canola cultivars. Once an N or S deficiency is corrected, there appears to be little need for balancing N and S application rates at any particular ratio with hybrid canola, but there appears to be a N x S interaction with conventional canola. (Karamanos et al. 2005)

    There has been research into using the N:S ratio during tissue testing to determine S status. However, the N:S ratio of canola tissue has not proved reliable for predicting S status. The N:S ratio only indicates the relative proportions of N and S in the plant, and does not indicate their actual magnitudes. Therefore, if canola tissue tests show an optimal ratio of 7:1, there are three possibilities: both N and S levels are optimal, excessive, or deficient. At the rosette stage, tissue testing canola for S status should include several criteria to improve the reliability:

    • % S greater than 0.25%
    • N:S ratio of 10 or less
    • a sulphate:total S ratio (as indicated by hydriodic acid reducible S:total S) greater than 0.38

    Calcium  (Ca)

    Calcium is a macronutrient absorbed in relatively large amounts by canola (see Table 2). However, deficiencies in western Canada are rare due to ample soil reserves. Calcium is often referred to as a ‘secondary’ nutrient, probably due to uncommon deficiencies and non-specific roles in the plant.

    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

    Role of Calcium in the Canola Plant

    Calcium performs several roles in the plant. In contrast to other macronutrients, a high proportion of Ca is found as a structural component in cell walls. Calcium’s structural function is to provide stable but reversible molecular linkages. Pectins are calcium compounds in cell walls that strengthen the wall and contribute to tissue resistance against fungal and bacterial infections. Calcium also plays a fundamental role in membrane stability and maintains cell integrity. This membrane protection is important under low temperature or saturated soil stress. Calcium bound at membrane surfaces can be exchanged with other cations (such as K+, Na+ and H+). Calcium exchange with sodium (Na) at membrane surfaces is a main factor in salinity stress. Also, Ca replacement with Al+3 (or blocking of Ca channels) is a factor in aluminum toxicity in acid soil.

    Cell extension requires Ca. Rapidly growing parts are, therefore, most affected by Ca deficiency. Root extension, shoot elongation and pollen growth are dependent on adequate Ca. The secretion of mucilage by root caps (that help root tips penetrate through soil) also needs Ca. Downward root growth (gravitropic response) relies on adequate Ca in the root caps. Callose formation is another example of a process involving Ca. In response to injury, cells will produce callose instead of cellulose, which helps wounds to heal and reduce infection.

    Most plant Ca is present in leaf vacuoles where it likely contributes to the cation-anion balance. Calcium also stimulates a range of enzymes, but generally is not a constituent of enzymes. Calcium plays a key role in plants as a secondary messenger in turgor regulated processes such as stomata opening and closing.

    Characteristics of Calcium Uptake by Canola

    Canola roots mainly absorb calcium as the Ca+2 cation dissolved in the soil water. Plant available Ca also exists as exchangeable Ca adsorbed on soil organic matter, silt and clay surfaces. The amount of dissolved Ca+2 depends on the amount of Ca-containing minerals, the soil cation exchange capacity and soil pH.

    A survey of Ca levels of 1,220 western Canadian soils
    Soil type1N NH4OAc extractable Ca, (mg kg-1)
    Non-calcareous 3200 ± 1051
    Calcareous 6560 ± 3235
    All soils 4050 ± 5263

    Source: Karamanos et al. 2001.

    High pH soils (>7.5) usually contain the highest Ca due to significant amounts of precipitated Ca salts (lime and gypsum).

    Since Ca is absorbed out of the soil water, the dominant processes controlling the supply to roots are mass flow, diffusion and root interception. Therefore, Ca availability is dependent on adequate soil moisture.

    Unusual situations that can create calcium deficiency in canola in western Canada are Solonetzic soils (sodium induced Ca deficiency), acidic soils (hydrogen / aluminum induced Ca deficiency), and waterlogged soils (restricted root uptake of Ca inducing temporary Ca deficiency).

    The Ca content varies between different plant parts and ages, ranging from 0.2% to 5%. The highest Ca contents are found in old leaves. At maturity, only about 10% of plant Ca is found in the canola seed.

    Magnesium (Mg)

    Of all the macronutrients, magnesium is absorbed in the least amount by plants. (see Table 3). Magnesium deficiencies are rare on the prairies, similar to Ca. However, Mg has more specific roles in plant function than Ca.

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


    Role of Magnesium in the Canola Plant

    Magnesium is the central atom of the chlorophyll molecule, and depending on the Mg sufficiency level, up to 25% of the total plant Mg is bound to chlorophyll. Magnesium is also needed for protein synthesis, and to activate many enzymes such as glutathione synthase, carboxylases, phosphatases, and ATPases. Most of the plant Mg is contained in cell vacuoles where it serves as a reserve for the metabolic pool and contributes to cation-anion balance.

    Characteristics of Magnesium Uptake by Canola

    Magnesium uptake by canola is very similar to that of calcium. The plant available form is the Mg+2 cation that exists in the soil solution and as exchangeable Mg adsorbed on soil organic matter, silt and clay surfaces. The amount of dissolved and exchangeable Mg will depend on the extent of Mg containing minerals, soil pH and cation exchange capacity.

    A survey of Mg levels of 1,220 western Canadian soils
    Soil type1N NH4OAc extractable Mg, (mg kg-1)
    Non-calcareous 500 ± 490
    Calcareous 560 ± 880
    All soils 630 ± 314

    Source: Karamanos et al. 2001. Unpublished data.

    Soils slightly acidic to neutral in pH tend to have the highest available Mg levels. This is due to Mg being a weak competitor for exchange sites on soil colloids and root binding sites. At higher pH and in soils with free lime, Ca+2 will dominate the exchange sites, possibly inducing Mg deficiency. In contrast, at low pH, H+, Al+3, and manganese (Mn+2) under flooded conditions, will dominate the exchange sites, inducing Mg (and Ca) deficiency. High K+ or NH4+ levels in the root zone due to fertilization may also induce Mg deficiency, although it is often short term. High Mg levels relative to Ca can induce a Ca deficiency. This has been reported in barley grown on solonetzic soil in Alberta with high Mg/Ca ratios.

    Magnesium level is highest during early vegetative growth (about 0.5%), and declines with maturity. Stems and roots contain the least Mg. At harvest, straw Mg level is less than 0.2% while canola seed contains about 0.3% Mg. Roughly 1/3 to 1/2 of the above ground Mg is contained in the seed [about 7 or 8 kg/ha (6 or 7 lb/ac)].


    Micronutrients are those nutrients required in extremely small quantities (less than 100 ppm in plant dry weight). Unfortunately, the basic functions of micronutrients are less understood than macronutrients. Also, there is very limited knowledge about the forms and mechanisms of micronutrient transport in the xylem and phloem. Micronutrient deficiencies in canola are much less common than macronutrient deficiencies. However, canola yields can be severely depressed when micronutrient deficiency occurs. This section will review the various micronutrients and canola responses.

    Boron (B)

    Boron is a micronutrient that occasionally limits canola yield in western and eastern Canada. Unfortunately, current soil test methods using hot-water B extraction do not consistently predict economic responses to B fertilizer in canola. (Karamanos et al. 2003). B deficiency is rare but when it occurs, it usually is on sandy areas of fields – this may be due to aluminum toxicity to roots, which can occur in acidic sandy soils. (Blevins et al. 1998)

    Role of Boron in the Canola Plant

    Boron’s role in plant nutrition is the least understood of all the nutrients. Boron is not an enzyme constituent nor does it seem to directly affect enzyme activities. Most of our understanding about B arises from symptoms observed during deficiency. Possible roles for B include:

    • sugar transport and carbohydrate metabolism
    • cell wall synthesis and structure
    • RNA metabolism
    • respiration
    • hormone metabolism
    • stomatal regulation
    • membrane function

    Cell walls are dramatically affected by B deficiency. This shows up as cracked, hollow or corky stems. The cell wall diameter and proportion of plant dry weight increase under B deficiency. Most plant B is complexed with organic compounds in the cell walls, apparently serving a nonspecific structural role.

    One of the first plant responses to induced B deficiency is decreased root elongation, however, there is a lack of understanding how this occurs. Boron deficiency also restricts pollen tube growth. This is why B demand is higher during the reproductive stage than vegetative stage. Boron also affects fertilization and seed set by increasing pollen production and viability.

    Boron Supply from Soil and Uptake by Canola

    Boron uptake depends most critically on soil organic matter content, and then adequate soil moisture, pH and soil B level. Soil pH may not influence B availability as much as organic matter level and texture (Malhi et al 2003).

    As with most nutrients, B uptake rates are increased as the level in the soil water increases. The plant available forms dissolved in the soil water move to the root via mass flow and diffusion. Low soil temperature in the root zone decreases the sensitivity of canola to low soil B (Ye, et al. 2003). Under drought conditions, B deficiency can occur due to a combination of reduced mineralization from the soil organic matter, reduced mass flow to roots, and polymerization of boric acid. In contrast, under high rainfall conditions B can be leached in sandy textured soils.

    Soil pH is a major factor influencing B availability. Generally, B becomes less available as pH increases above 6.3 to 6.5. At higher pH, the borate anion is likely adsorbed to clay and organic particles.

    Daily B uptake is greatest around 50% flowering. (Karamanos et al. 2003 unpublished)

    Boron is classified as immobile within plants but mobility through phloem has been measured within canola plants and other species that transport sucrose as their primary photoassimilate (Stangoulis et al., 2010).

    Boron transporters within plants have been found to regulate B uptake based on B conditions.  Plants can sense internal and external B conditions and regulate deficiency or toxicity problems by regulating B transporters to maintain B balance within plant tissue. (Miwa et al. 2010).  In the future, identification of key genes and processes involved in B modulation within the plant may allow for the development of cultivars more tolerant of B stress.

    Boron uptake is affected by other nutrients. High levels of calcium and potassium have been shown to increase B deficiency symptoms but these antagonisms are not well understood.

    There is genetic variation for B efficiency in canola, with reports from China showing one major dominant gene (Shi et al, 2009) as well as multi-gene effects (Shi et al, 2012). Genetic differences in B efficiency were related to differences in root uptake or plant utilization. Research from Pakistan reported that Brassica napus was more sensitive to B deficiency than mustard (Brassica juncea), but needed relatively less B fertilizer for optimum grain yield. This area merits further research with western Canadian cultivars and conditions as many of these findings were observed on soils with less than 0.8% organic matter.

    Boron deficiency symptoms in canola first appear in new growth due to the intermediate mobility. Symptoms range from:

    • deformed, curled and rough skinned leaves with torn margins
    • yellow to brown spots in the interveinal areas of leaves
    • red to brown-purple coloured new leaves
    • early leaf drop
    • shortened stems
    • cracked stems
    • prolonged flowering
    • flower sterility
    • poor pod set and yield

    Canola response to boron

    On western Canadian Prairie soils, no significant yield increases were seen in 22 trials in seven separate experiments with soil- or foliar-applied B. In these experiments, B in canola tissue was correlated to the rate of B applied, but no yield response was observed. In addition, the hot-water B soil test extraction method was not an effective diagnostic tool for determining the B status of soils (Karamanos et al. 2003).

    On sandy soils in northeastern Saskatchewan, B fertilizer did not provide a consistent yield increase on soils thought to be deficient in B. The response to B depended on site, year, cultivar and B fertilizer rate, time and method of application – indicating that B deficiency in a field probably occurs in isolated patches. Boron fertilization had no impact on the amount of severity of sclerotinia stem rot, blackleg or alternaria pod spot.  (Malhi et al. 2003)

    Copper (Cu)

    Knowledge about copper fertility of western Canadian soils has increased over the past three decades. Previously, Cu deficiency was thought to be limited to organic or peat soils. More recent research has identified that Black, transitional Gray-Black and sandy Dark Brown soils may be Cu deficient for cereal production. Although copper deficiency and fertilizer response has been documented with cereals under field conditions, canola has only shown some response when soil test showed very low levels of Cu at less than 0.2 ppm. (Karamanos et al. 1986)

    Role of Copper in the Canola Plant

    Copper is a transition element that forms stable complexes in the plant and soil, and is capable of electron transfer (energy processes). Copper’s role in plant functions is mainly as a reactive constituent of enzymes that catalyze oxidation-reduction reactions. Some examples of Cu containing enzymes include:

    • plastocyanin (needed for energy capture through photosynthesis)
    • superoxide dismutase (needed for detoxification of oxygen radicals)
    • many different types of oxidases (enzymes that degrade or change compounds together with oxygen)

    One of the phenol oxidases is involved with lignin synthesis.

    Due to copper’s role in photosynthesis, deficiency leads to low carbohydrates levels, at least during the vegetative stage. The low carbohydrate content in Cu deficient plants contributes to impaired pollen formation and fertilization. The reduced lignification in Cu deficient plants also affects pollen fertility since lignification of anthers is needed to release pollen.

    Copper Supply from the Soil and Uptake by Canola

    Copper is a metallic nutrient that originates from minerals in the soil. The total Cu content in western Canada soils usually falls in the range of 5 to 50 ppm. Approximately 1/2 to 1/4 of the total Cu exists within minerals and is unavailable to plants. Copper associated with oxides and organic matter has been found to be an important source of plant available Cu, probably by replenishment of dissolved and exchangeable Cu. The oxide and organic fractions increase with the clay content, which explains why Cu deficiency is more likely on sandier textures. Exchangeable Cu ranges from 0.1 to 10% of total soil Cu. Only very small amounts of Cu exist as soluble Cu+2 in the soil water. Research on the prairies has found that DTPA extractable Cu is highly variable across cultivated and native fields. This means that larger numbers of soil samples are needed to obtain a precise estimate of the true soil average.

    Very little information exists on the mechanisms of Cu uptake by plant roots. The driving force for Cu uptake is the electrical chemical gradient across the root cell membranes. Since free Cu levels inside the cell are kept low to avoid harmful reactions, and the membranes have a large negative potential, this creates a large force for Cu uptake. Therefore, there is no need for active Cu uptake systems. It has been suggested that Ca channels likely also allow passage of other ions such as Cu+2.

    Within several weeks of emergence, daily Cu uptake rapidly rises, and peaks near mid-flowering.  (Karamanos et al 2003 Unpublished data)

    Copper remobilization is much higher in old leaves and is related to N remobilization. Despite the intermediate mobility of Cu, deficiency symptoms in sensitive crops during the vegetative stage first appear in new growth. However, canola does not display strong Cu deficiency symptoms. Pot experiments with extreme Cu deficiency have reported canola symptoms of:

    • interveinal chlorosis shortly after emergence
    • larger than normal leaves
    • wilting leaves
    • delayed flowering with a shortened flowering stem

    Evidence suggests that canola growth may be affected by imbalances between Cu, molybdenum (Mo) and manganese (Mn) levels. Manganese:copper ratios (DTPA extractable) greater than 15 may result in a Cu deficiency. Molybdenum also may antagonize Cu. However, the Mo antagonism is in turn affected by S levels. Sulphur additions were found to lower Mo contents in canola plants, reducing Mo antagonism with Cu, and Cu deficiency was alleviated without adding Cu fertilizer.

    Iron (Fe)

    Iron is one of the most abundant metallic elements in the earth’s crust. Western Canadian soils have developed from parent materials rich in Fe. Therefore, there have been no reports of Fe deficiency in field crops or responses to Fe fertilizer on the prairies. Also, there has been no work to calibrate soil test values for Fe on the prairies.

    Role of Iron in the Canola Plant

    Iron is a component of ferrodoxin, which acts as an electron transmitter in nitrate and sulphate reduction, nitrogen fixation, and energy production. Iron is needed for chlorophyll synthesis, and low chlorophyll contents of young leaves (interveinal ‘chlorosis’ or yellowing) is the most obvious visible symptom of Fe deficiency. In young growing leaves, about 80% of the Fe is located in the chloroplasts. Iron is also thought to be involved with protein synthesis and root tip growth.

    Iron concentration in whole plant tissue is highest in young canola plants, and declines in concentration as the plants mature. (Karamanos et al. 2003 Unpublished data)

    Manganese (Mn)

    Manganese is a metallic micronutrient that is occasionally deficient in western Canadian organic, high pH soils. Although oats can be affected by Mn deficiency in cold organic soils (gray speck of oats disease), there have been no documented deficiencies with canola in western Canada.

    Weathering of manganese containing soil minerals is the source of plant available Mn. The main Mn form that exists in soil solution or adsorbed to soil colloids is Mn+2, which is also the form absorbed by roots. Manganese availability decreases when pH increases above 6.2 in many soils. Low temperature and high organic matter can also decrease Mn availability. Rhizosphere microbes play a role in Mn availability by either oxidation or reduction. These microbes can either increase or decrease Mn availability. The rhizosphere acidification by canola roots likely increases Mn availability and makes this crop relatively tolerant of low soil Mn.

    Manganese concentration in canola tissue initially rises in the first 2 to 6 weeks after emergence, and then declines as canola matures. (Karamanos et al. 2033 Unpublished data)

    Plant Mn status is affected by Cu levels. High Mn:Cu ratios above 15 may lead to Cu deficiency while ratios below 1 may lead to Mn deficiency. Potash (KCl) has been shown to enhance Mn uptake by several crops. In some crops, seed Mn content has been shown to be important for initial Mn nutrition and plant growth, as well as disease resistance.

    Manganese toxicity

    Several cases of Mn toxicity were identified in northern Alberta in 2010 in areas of the field that had very strongly acidic soils.  The symptoms were similar to sulfur deficiency with chlorosis of leaf margins and cupping of leaves in canola. To correct the Mn toxicity problem, liming would be one tactic aimed at increasing soil pH.  However, the cost can be prohibitive especially with debatable returns given the lack of information on yield loss due to Mn toxicity in canola in western Canada. 

    Molybdenum (Mo)

    Molybdenum is a transition element needed in extremely low amounts; only nickel has a lower requirement. All the Brassica species appear to be sensitive to low Mo supply and can exhibit peculiar symptoms (for example ‘whiptail’ of cauliflower).

    Role of Molybdenum in the Canola Plant

    Only a few enzymes are known to contain Mo as a cofactor:

    • the enzyme that helps to change nitrate to organic N in the plant (nitrate reductase)
    • the major enzyme involved in nitrogen fixation in legumes (nitrogenase)
    • an oxidase/dehydrogenase enzyme involved in changing the products of N fixation
    • probably an enzyme involved in sulphate metabolism (sulphite reductase)

    Molybdenum functions are, therefore, closely related to N metabolism and N fixation.

    Some interaction occurs between Mo uptake and levels of P and S. Plant Mo uptake is usually enhanced by soluble P and decreased by sulphate. MoO4-2 and SO4-2 compete strongly for root uptake. Once absorbed by roots, Mo is readily mobile in both the xylem and phloem transport systems, probably as MoO4-2.

    Deficiencies in canola have not been documented in western Canada. If a Mo deficiency were to occur, there would be several options: seed treatment with Mo, soil or foliar fertilizer, and liming to raise soil pH. Molybdenum is unique among the micronutrients since there is a wide range between deficiency and toxicity.

    Zinc (Zn)

    Zinc is a metallic micronutrient that is commonly deficient in many countries, including Australia, China and India. In western Canada, sporadic Zn deficiencies have been identified in fields of alfalfa, flax, corn and beans, but not canola.

    Role of Zinc in the Canola Plant

    Zinc exists only in the Zn+2 form in plants, and is not involved with redox reactions. Zinc has an ability to form complexes with N, O and particularly S and performs catalytic and structural roles in enzymes. Many enzymes contain Zn as a structural, catalytic or cofactor component. Protein synthesis, hormone (auxin) and carbohydrate metabolism also require Zn. Membrane stability also relies on Zn, and the most obvious Zn deficiency symptoms (such as leaf chlorosis and inhibited stem elongation) probably arise from membrane breakdown. Pot experiments with canola have reported Zn deficiency symptoms ranging from purpling on new emerging leaves, brown spots on cotyledons, interveinal chlorosis and cupping of leaves.

    Uptake of Zinc by Canola

    Weathering of soil minerals is the primary source of plant available Zn+2. Weathering removes Zn faster than other metals except for Cu. Zn deficiency commonly occurs in acidic, highly weathered soils (typically tropical). Zinc deficiency may also occur in high pH, calcareous soils due to Zn adsorption to lime particles. Plant available Zn exists as exchangeable Zn+2, dissolved Zn+2 in soil water, adsorbed Zn to Mn oxide and organically bound Zn. Soil test labs often use a critical level of 0.5 ppm DTPA extractable Zn, but research in western Canada has found this level too high for predicting cereal response and that DTPA is an unsuitable extractant. No further work has been conducted to find a more suitable extractant and calibration for cereals and oilseeds on the prairies.

    The mechanism for Zn uptake by roots is not well understood and may involve both active and passive processes. Once absorbed by the roots, Zn is likely complexed with small organic molecules similar to other metallic micronutrients.

    Zinc availability increases as soil pH decreases (becomes more acidic). Copper and other cations compete for root Zn uptake. High P levels can induce Zn deficiency by inhibiting Zn translocation within the plant rather than affecting root uptake.

    Daily Zn uptake is greatest during canola flowering. (Karamanos et al. 2003 Unpublished data)

    Other Micronutrients

    There are other micronutrients and beneficial elements than those discussed above. However, deficiencies of the remaining nutrients are limited to certain plant species other than canola or deficiencies are infrequent anywhere in the world.

    Chlorine (Cl) is found in abundance in nature as chloride salts. Choride (Cl-) is highly mobile in soil and plants and is readily absorbed by roots. Chlorine is involved in photosynthesis, charge balance, enzyme activation, stomatal regulation and disease resistance. Cereal (winter wheat) responses to Cl fertilizer have occurred on the prairies, apparently due to disease suppression or improved water relations rather than Cl nutritional needs. Canola responses to Cl have not been reported.

    Nickel (Ni) has recently been established as an essential nutrient. In higher plants, urease is the only known Ni containing enzyme. Other Ni roles include Fe absorption, seed viability, N fixation and reproductive growth. The plant available form is Ni+2. Root uptake likely follows similar patterns as other micronutrient metals. Nickel appears to be readily mobile in both xylem and phloem. Ni deficiency in field grown crops has not been reported.

    Silicon (Si) is the second most abundant element in the earth’s crust and is a beneficial nutrient for a few wetland plant species such as rice. In non-wetland species, Si can counteract Zn deficiency induced by high P. Since Si is so abundant in nature, proving its essentiality is very difficult. Silicon may affect plant stability by influencing lignin biosynthesis as well as through deposition in cell walls. Increased leaf rigidity has been reported in cereal and cucumber crops. Silicon may also contribute to disease and insect resistance. Silicon may decrease toxicity from high levels of Mn, Fe and Al.

    Sodium (Na) is an essential nutrient for some plant species that use the C4 photosynthetic pathway. Canola uses the C3 pathway and, therefore, Na is not a beneficial nutrient for this crop.


    Allen, E.J. and Morgan, D.G. 1972.A quantitative analysis of the effects of nitrogen on the growth, development and yield of oilseed rape. J. Agric. Sci. Camb. 78:315-324.

    Bailey, L.D. 1986.The sulphur status of eastern Canadian prairie soils: sulphur response and requirements of alfalfa (Medicago sativa L.), rape (Brassica napus L.) and barley (Hordeum vulgare L.). Can. J. Soil Sci. 66:209-216.

    Bailey, L.D. 1990.The effects of 2-chloro-6 (trichloromethyl)- pyridine (N-Serve™) and N fertilizers on productivity and quality of Canadian oilseed rape. Can. J. Plant Sci. 70:979-986.

    Bailey, L.D. and Grant, C.A. 1990.Fertilizer placement studies on calcareous and non-calcareous chernozemic soils: growth, P-uptake, oil content and yield of Canadian rape. Commun. In Soil Sci. Plant Anal. 21:2089-2104.

    Bell, N.J. 1970.Fertilizer experiments with special crops. pp. 128-135 In: 14th Annual Manitoba Soil Science Society Proceedings. Winnipeg, Man.

    Bettany, J.R., Stewart, J.W.B., and Halstead, E.H. 1973.Sulfur fractions and Carbon, Nitrogen, and Sulfur Relationships in Grassland, Forest and Associated Transitional Soils. Soil Sci. Soc. Amer. Proc. 37:915-918.

    Blevins, D.G., and Lukaszewski, K.M. 1998.Boron in plant structure and function. Annual Review of Plant Physiology and Plant Molecular Biology. Vol. 49: 481-500.

    Bole, J.B. and Pittman, U.J. 1984.Availability of subsoil sulphates to barley and rapeseed. Can. J. Soil Sci. 64:301-312.

    Bolland, M.D.A. 1997.Comparative phosphorus requirement of canola and wheat. J. Plant Nutrition 20:813-829.

    Canola Council of Canada. Canola Production Centre Reports.
    Individual annual reports for 1997 to 2001.

    Campbell, C. A. and Paul, E. A. 1978.Effects of fertilizer N and soil moisture on mineralization, N recovery and A-values, under spring wheat grown in small lysimeters. Can. J. Soil Sci. 58: 39-51.

    Carter, M.R. and Webster, G.R. 1979.Calcium deficiency in some Solonetzic soils of Alberta. J. Soil Sci. 30:161-174.

    Cowell, L.E. and Doyle, P.J. 1993.Nitrogen use efficiency. pp. 49-109 In: Impact of Macronutrients on Crop Responses and Environmental Sustainability on the Canadian Prairies. Eds. D. A. Rennie, C. A. Campbell and T.L. Roberts. Published by Canadian Society of Soil Science.

    Cumbus, I.P. and Nye, P.H. 1982.Root zone temperature effects on growth and nitrate absorption in rape (Brassica napus cv Emerald). J. Exp. Bot. 33:1138-1146.

    Cumbus, I.P. and Nye, P.H. 1985.Root zone temperature effects on growth and phosphate absorption in rape Brassica napus cv. Emerald. J. Exp. Bot. 163:219-227.

    Doyle, P.J. and Cowell, L.E. 1993.Phosphorus. pp. 110-170 In: Impact of Macronutrients on Crop Responses and Environmental Sustainability on the Canadian Prairies. Eds. D. A. Rennie, C. A. Campbell and T.L. Roberts. Published by Canadian Society of Soil Science.

    Doyle, P.J. and Cowell, L.E. 1993.Potassium. pp. 171-201 In: Impact of Macronutrients on Crop Responses and Environmental Sustainability on the Canadian Prairies. Eds. D. A. Rennie, C. A. Campbell and T.L. Roberts. Published by Canadian Society of Soil Science.

    Doyle, P.J. and Cowell, L.E. 1993.Sulphur. pp. 202-250 In: Impact of Macronutrients on Crop Responses and Environmental Sustainability on the Canadian Prairies. Eds. D. A. Rennie, C. A. Campbell and T.L. Roberts. Published by Canadian Society of Soil Science.

    Dreccer, M.F., Schapendonk, A.H.C.M., Slafer, G.A. and Rabbinge, R. 2000.Comparative response of wheat and oilseed rape to nitrogen supply: absorption and utilisation efficiency of radiation and nitrogen during the reproductive stages determining yield. Plant and Soil 220:189-205.

    Foehse, D. and Jungk, A. 1983.Influence of phosphate and nitrate supply on root hair formation of rape, spinach and tomato plants. Plant and Soil 74:359-368.

    Fohse, D., Claassen, N., and Jungk, A. 1988.Phosphorus efficiency of plants. I. External and internal P requirement and P uptake efficiency of different plant species. Plant and Soil 110:101-109.

    Fohse, D., Claassen, N. and Jungk, A. 1991.Phosphorus efficiency of plants. II. Significance of root radius, root hairs and cation-anion balance for phosphorus influx in seven plant species. Plant and Soil 132: 261-272.

    Gleddie, S.C., Schlechte, D. and Turnbull, G. 1993.Effect of inoculation with Penicillium bilaji (Provide®) on phosphate uptake and yield of canola in western Canada. pp. 155-160 In: Proc. 30th Annual Alberta Soil Science Workshop, Edmonton, Alberta.

    Grant, C.A. and Bailey, L.D. 1993.Fertility management in canola production. Can. J. Plant Sci. 73:651-670.

    Grant, C.A., Flaten, D.N., Tomasiewicz, D.J. and Sheppard, S.C. 2001.The importance of early season phosphorus nutrition. Can. J. Plant Sci. 81:211-224.

    Grewal, H.S., Graham, R.D. and Stangoulis, J. 1998.Zinc boron interaction effects in oilseed rape. J. Plant Nutrition 21:2231-2243.

    Grewal, H.S., Stangoulis, J.C.R., Potter, T.D. and Graham, R.D. 1997.Zinc efficiency of oilseed rape (Brassica napus and B. juncea) genotypes. Plant and Soil 191:123-132.

    Gupta, U.C. 1993.Boron and its role in crop production. CRC Press, Boca Raton, Fla.

    Gupta, U.C. and Lipsett, J. 1981.Molybdenum in soils, plant and animals. Adv. in Agron. 34:73-115.

    Gupta, V.V.S.R., Lawrence, J.R. and Germida, J.J. 1988.Impact of elemental sulfur fertilization on agricultural soils. I. Effects on microbial biomass and enzyme activities. Can. J. Soil Sci. 68:463-473.

    Gupta, V.V.S.R., Lawrence, J.R. and Germida, J.J. 1988.Impact of elemental sulfur fertilization on agricultural soils. II. Effects on sulfur-oxidizing populations and oxidation rates. Can. J. Soil Sci. 68:475-483.

    Harapiak, J.T., Malhi, S.S., Campbell, C.A. and Nyborg, M. 1993.Fertilizer N application practices. pp. 251-313 In: Impact of Macronutrients on Crop Responses and Environmental Sustainability on the Canadian Prairies. Eds. D. A. Rennie, C. A. Campbell and T.L. Roberts. Published by Canadian Society of Soil Science

    Henry, J.L. and MacDonald, K.B. 1978.The effects of soil and fertilizer nitrogen and moisture stress on yield, oil and protein content of rape. Can. J. Soil Sci. 58:303-310.

    Herath, H.M.W. and Ormrod, D.P. 1971.Temperature effects on the response to sulphur of barley, peas and rape. Plant and Soil 35:635-646.

    Hocking. P.J., Kirkegaard, J.A., Angus, J.F., Gibson, A.H. and Koetz, E.A. 1997.Comparison of canola, Indian mustard and Linola in two contrasting environments. I. Effects of nitrogen fertilizer on dry-matter production, seed yield and seed quality. Field Crops Res. 49:107-125.

    Hocking, P.J., Randall, P.J. and DeMarco, D. 1997.The response of dryland canola to nitrogen fertilizer: partitioning and mobilization of dry matter and nitrogen, and nitrogen effects on yield components. Field Crops Research 54:201-220.

    Hoffland, E., Findenegg, G.R. and Nelemans, J.A. 1989.Solubilization of rock phosphate by rape. I. Evaluation of the role of the nutrient uptake pattern. Plant and Soil 113:155-160.

    Hoffland, E., Findenegg, G.R. and Nelemans, J.A. 1989.Solubilization of rock phosphate by rape. II. Local root exudation of organic acids as a response to P-starvation. Plant and Soil 113:161-165.

    Holmes, M.R.J. 1980.Nutrition of the oilseed rape crop. Applied Science Publishers Ltd. London. 158 pp.

    Huang, L., Hu, D. and Bell, R.W. 1995.Diagnosis of zinc deficiency in canola by plant analysis. Commun. Soil Sci. Plant Anal. 26:3005-3022.

    Huang, L., Ye, Z., and Bell, R.W. 1996.The importance of sampling immature leaves for the diagnosis of boron deficiency in oilseed rape (Brassica napus cv. Eureka). Plant and Soil 183:187-198.

    Jackson, G.D. 2000.Effects of nitrogen and sulfur on canola yield and nutrient uptake. Agron. J. 92:644-649.

    Janzen, H.H. 1990.Elemental sulfur oxidation as influenced by plant growth and degree of dispersion within soil. Can. J. Soil Sci. 70:499-502.

    Janzen, H.H. and Bettany, J.R. 1984a.Sulfur Nutrition of Rapeseed: I. Influence of Fertilizer Nitrogen and Sulfur Rates. Soil Sci. Soc. Am. J. 48:100-107.

    Janzen, H.H. and Bettany, J.R. 1984b.Sulfur Nutrition of Rapeseed: II. Effect of Time of Sulfur Application. Soil Sci. Soc. Am. J. 107-112.

    Janzen, H.H. and Bettany, J.R. 1987.Oxidation of elemental sulfur under field conditions in central Saskatchewan. Can. J. Soil Sci. 67:609-618.

    Janzen, H.H. and Karamanos, R.E. 1991.Short-term and residual contribution of selected elemental S fertilizers to the S fertility of two Luvisolic soils. Can. J. Soil Sci. 71:203-211.

    Johnston, A.M., Grant, C.A. and Clayton, G.W. 1999.Sulphur management of canola. In: GCIRC 10th Int. Congress Proceedings.

    Johnston, A.M., Johnson, E.N., Kirkland, K.J. and Stevenson, F.C. 2002.Nitrogen fertilizer placement for fall and spring seeded Brassica napus canola. Can. J. Plant Sci. 82: 15-20.

    Johnston, A.M., Johnson, E.N., Kirkland, K.J. and Stevenson, F.C. 2002.Nitrogen fertilizer placement for fall and spring seeded Brassica napus canola. Can. J. Plant Sci. 82: 15-20.

    Karamanos, R.E. 1979. Variations in natural 15N Abundance as an index of past nitrogen cycle stresses.  Ph.D. Thesis.  Saskatoon:  University of Saskatchewan, 165 pp.

    Karamanos, R.E. and Flore, N.A. 2001.Soil absorption of and plant response to applied calcium. In: Proc. Soil & Crops 2001, Saskatoon, Sask.

    Karamanos, R.E., Goh, T.B. and Flaten, D.N. 2007.  Nitrogen and sulphur fertilizer management for growing canola on sulphur sufficient soils.  Can. J. Plant Sci. 87:201-210.

    Karamanos, R. E., Goh, T. B. and Stonehouse, T. A. 2003.Canola response to boron in Canadian prairie soils. Can. J. Plant Sci. 83: 249–259.

    Karamanos, R.E. Goh, T.B. and Poisson, D.P. 2005.  Nitrogen, Phosphorus and Sulfur Fertilization of Hybrid Canola. J. Plant Nutr. 28: 1145 - 1161.

    Karamanos, R.E., Harapiak, J. and Flore, N.E. 2002.Fall and early spring seeding of canola (Brassica napus L.) using different methods of seeding and phosphorus placement. Can. J. Plant Sci. 82: 21-26.

    Karamanos, R.E., Hodge, N., and Stewart, J.W.B. 1989.The effect of sulphur on manganese and copper nutrition of canola. Can. J. Soil Sci. 69:119-125.

    Karamanos, R.E., and Janzen, H.H. 1991.Crop response to elemental sulfur fertilizers in central Alberta. Can. J. Soil Sci. 71:213-225.

    Karamanos R.E., Kruger, G.A. and Stewart, J.W.B. 1986. Copper deficiency in cereal and oilseed crops in northern Canadian prairie soils.  Agron. J., 78, 317-323.

    Karamanos, R.E., Poisson, D.P., and Tee Boon Goh. 2004.  Biomass and Nutrient Accumulation in Hybrid Canola. Proceedings Soils and Crops. Extension Division. University of Saskatchewan.

    Karamanos, R.E. and Henry, J.L. 2007.  A Water Based System for Deriving Nitrogen Recommendations in the Canadian Prairies.  Chapter 11, in. T. Bruuslema (ed.) Managing Crop Nitrogen for Weather. Proceedings of the Symposium “Integrating Water Variability into Nitrogen Recommendations” International Plant Nutrient Institute, Norcross, GA.

    Krogman, K.K. and Hobbs, E.H. 1975.Yield and morphological response of rape (Brassica campestris L. cv. Span) to irrigation and fertilizer treatments. Can. J. Plant Sci. 55:903-909.

    Kruger, G.A., Karamanos, R.E. and Singh, J.P. 1985.The copper fertility of Saskatchewan soils. Can. J. Soil Sci. 65:89-99.

    Kruger, G., Oliver, E., Balwin, A., and Polegi, J. 2001.Use of Agrotain to reduce the seedbed toxicity of urea. In: Soils & Crops 2001 Proceedings, Saskatoon, Sask.

    Kucey, R.M.N. and Leggett, M.E. 1989.Increased yields and phosphorus uptake by Westar canola (Brassica napus L.) inoculated with a phosphate-solubilizing isolate of Penicillium bilaji. Can. J. Soil Sci. 69:425-432.

    Lafond, G. P, Walley, F., May, W.E., and Holzapfel, C.B. 2011.Long term impact of no-till on soil properties and crop productivity on the Canadian Prairies. Soil & Tillage Research, Vol. 117: 110-123

    Liang, J., Karamanos, R.E. and Stewart, J.W.B. 1991.Plant availability of Zn fractions in Saskatchewan soils. Can. J. Soil Sci. 71:507-517.

    Liang, J., Stewart, J.W.B., and Karamanos, R.E. 1990.Distribution of zinc fractions in prairie soils. Can. J. Soil Sci. 70:335-342.

    Liang, J., Stewart, J.W.B., and Karamanos, R.E. 1991.Distribution and plant availability of soil copper fractions in Saskatchewan. Can. J. Soil Sci. 71:89-99.

    Liu, L., Shelp, B. J. and Spiers, G. A. 1993.Boron distribution and retranslocation in field-grown broccoli (Brassica oleracea var. italica). Can. J. Plant Sci. 73:587-600.

    Macduff, J.H., Hopper, M.J., Wild, A. and Trim, F.E. 1987.Comparison of the effects of root temperature on nitrate and ammonium nutrition of oilseed rape (Brassica napus L.) in flowing solution culture. I. Growth and uptake of nitrogen. J. Exp. Bot. 38:1104-1120.

    Macduff, J.H., Hopper, M.J., Wild, A. and Trim, F.E. 1987.Comparison of the effects of root temperature on nitrate and ammonium nutrition of oilseed rape (Brassica napus L.) in flowing solution culture. II. Cation-anion balance. J. Exp. Bot. 38:1589-1602.

    Malhi, S.S., Kryzanowski, L.M., Mumey, G., Nyborg, M. and Penney, D.C. 1993.Fertilizer N requirement for most economical yield of rapeseed as influenced by soil nitrate-N level. pp. 276-280 In: Proc. 30th Annual Alberta Soils Workshop. Edmonton, Alberta.

    Malhi. S.S, Kutcher, K., Johnston, A. and Leach, D. 2000.Feasibility of boron fertilization on canola in the Saskatchewan parkland. In: Proc. Soils & Crops 2000, Saskatoon, Sask.

    Malhi, S.S. and Leach, D. 2000.Influence of nitrogen rate on sulphur requirements for optimum yield of canola. In: Soils and Crops 2000 Proceedings, Saskatoon, Sask.

    Malhi, S.S. and Leach, D. 2001.Effectiveness of foliar applications of various sulphate-S fertilizers to correct sulphur deficiency on canola in the growing season. In: Soils and Crops 2001 Proceedings, Saskatoon, Sask.

    Malhi, S.S. and Leach, D. 2001.Effectiveness of Elemental S Fertilizers on Canola After Two Annual Applications. In: Soils and Crops 2001 Proceedings, Saskatoon, Sask.

    Malhi, S.S., Brandt, S., Ulrich, D., Lafond, G.P., Johnston, A.M. and Zentner, R.P. 2007.  Comparative nitrogen response and economic evaluation for optimum yield of hybrid and open-pollinated canola.  Can. J. Plant Sci. 87:449-460.

    Malhi, S.S., Nyborg, M., Walker, D.R. and Laverty, D.H. 1985.Fall and spring soil sampling for mineral N in north-central Alberta. Can. J. Soil Sci. 65:339-346.

    Malhi, S. S., Raza, M., Schoenau, J. J., Mermut, A. R., Kutcher, R., Johnston A. M. and Gill, K. S. 2003.Feasibility of boron fertilization for yield, seed quality and B uptake of canola in northeastern Saskatchewan. Can. J. Soil Sci. 83: 99–108.

    Manitoba Agriculture and Food. 2001.Soil Fertility Guide. Revised March 2001.

    Marschner, H. 1995.Mineral nutrition of higher plants. Academic Press Ltd., 889 pp.

    Maynard, D.G., Stewart, J.W.B. and Bettany, J.R. 1983.Use of plant analysis to predict sulfur deficiency in rapeseed (Brassica napus and B. campestris). Can. J. Soil Sci. 63:387-396.

    McAndrew, D.W., Loewen-Rudgers, L.A. and Racz, G.J. 1984.A growth chamber study of copper nutrition of cereal and oilseed crops in organic soil. Can. J. Plant Sci. 64:505-510.

    McKenzie, R.H., Dormaar, J.F., Schaalje, G.B. and Stewart, J.W.B. 1995.Chemical and biochemical changes in the rhizospheres of wheat and canola. Can. J. Soil Sci. 75:439-447.

    McKenzie, R.H. and Roberts, T.L. 1990.Soil and fertilizer phosphorus update. In: Proc. 27th Annual Alberta Soil Science Workshop, Edmonton, Alberta.

    McKenzie, R.H., Kryzanowski, L., Solberg, E., Penney, D., Coy, G., Heaney, D., Harapiak, J. and Flore, N.. 1994.Wheat, barley and canola responses to phosphate fertilizer in Alberta. pp. 136-145 In: Proc. 31st Annual Alberta Soil Science Workshop, Edmonton, Alberta.

    Miwa, K. and Fujiwara, T. 2010.Boron transport in plants: co-ordinated regulation of transporters. Annals of Botany 105: 1103 – 1108.

    Moorby, H., Nye, P.H. and White, R.E. 1985.The influence of nitrate nutrition on H+ efflux by young rape plants (Brassica napus cv emerald). Plant and Soil 84:403-415.

    N’dayegamiye, A., Simard, R.R. and Delisle, F. 1994.Evaluation of sulfur mineralization potential of meadow soils and availability to alfalfa. Can. J. Soil Sci. 74:259-265.

    Nuttall, W.F., Boswell, C.C., Sinclair, A.G., Moulin, A.P., Townley-Smith, L.J. and Galloway, G.L. 1993.The effect of time of application and placement of sulphur fertilizer sources on yield of wheat, canola and barley. Commun. Soil Sci. Plant Anal. 24:2193-2202.

    Nuttall, W.F. and Button, R.G. 1990.The effect of deep banding N and P fertilizer on the yield of canola (Brassica napus L.) and spring wheat (Triticum aestivum L.). Can. J. Soil Sci. 70:629-639.

    Nuttall, W.F. and Malhi, S.S. 1991.The effect of time and rate of N application on the yield and N uptake of wheat, barley, flax and four cultivars of rapeseed. Can. J. Soil Sci. 71:227-238.

    Nuttall, W.F., Moulin, A.P. and Townley-Smith, L.J. 1992.Yield response of canola to nitrogen, phosphorus, precipitation and temperature. Agron. J. 84:765-768.

    Nuttall, W.F., Ukrainetz, H., Stewart, J.W.B. and Spurr, D.T. 1987.The effect of nitrogen, sulphur and boron on yield and quality of rapeseed (Brassica napus L. and B. campestris L.). Can. J. Soil Sci. 67:545-559.

    Nyborg, M. 1961.The effect of fertilizers on emergence of cereal grains, flax and rape. Can. J. Soil Sci. 41:89-93.

    Nyborg, M. and Hennig, A.M.F. 1969.Field experiments with different placements of fertilizers for barley, flax and rapeseed. Can. J. Soil Sci. 49:79-88.

    Nyborg, M. and Hoyt, P.B. 1970.Boron deficiency in turnip rape grown on gray wooded soils. Can. J. Soil Sci. 50:87-88.

    Pageau, D., Lafond, J. and Tremblay, G.F. 1999.The effects of boron on the productivity of canola. In: GCIRC 10th Congress Proceedings.

    Penney, D.C., Goddard, T.W., Nolan, S.C., Skarberg, K. and McKenzie, R.C. 1995.Precision Farming: Yield, Terrain and Fertility Mapping with Global Positioning Systems (GPS). In: Soils & Crops Workshop, Saskatoon, Sask.

    Penney, D.C., Helm, J. and McKenzie, R.H. 1994.Intensive crop management systems for dryland and irrigated barley, wheat and canola production. Farming for the Future Final Report #87-0163.

    Qian, P., Schoenau, J.J., Greer, K.J., Liu,V. and Shen, J. 1995.Quick extraction and determination of potassium in fresh leaf sap and its use as a guide to potassium fertilization of canola, chickpea and dwarf sunflower. Commun. Soil Sci. Plant Anal. 26:2903-2912.

    Racz, G.T., Webber, R.J. and Hedlin, R.A. 1965.Phosphorus and nitrogen utilization of rape, flax and wheat. Agron. J. 57:335-337.

    Rashid, A., Rafique, E. and Bughio, N. 1994.Diagnosing boron deficiency in rapeseed and mustard by plant analysis and soil testing. Commun. Soil Sci. Plant Anal. 25:2883-2897.

    Raza, M., Mermut, A.R., Schoenau, J.J. and Malhi, S.S. 2000.Boron fractionation in some Saskatchewan soils. In: Proc. Soils & Crops 2000, Saskatoon, Sask.

    Roberts, T.L. and Bettany, J.R. 1985.The influence of topography on the nature and distribution of soil sulfur across a narrow environmental gradient. Can. J. Soil Sci. 65:419-434.

    Rood, S.B. and Major, D.J. 1984.Influence of plant density, nitrogen, water supply and pod or leaf removal on growth of oilseed rape. Field Crops Research 8:323-331.

    Rood, S.B., Major, D.J., Carefoot,V. and Bole, J.B. 1984.Seasonal distribution of nitrogen in oilseed rape. Field Crops Research 8:333-340.

    Schnug, E. and Haneklaus, S. 1994.Sulphur deficiency in Brassica napus - Biochemistry- Symptomatology- Morphogenesis. Landbauforschung Volkenrode, Sonderheft 144.

    Sheppard, S.C. and Bates, T.E. 1980.Yield and chemical composition of rape in response to nitrogen, phosphorus and potassium. Can. J. Soil Sci. 60:153-162.

    Shi, L., Wang, Y. H., Nian, F. Z., Lu, J. W., Meng, J. L. and Xu, F. S. 2009.Inheritance of boron efficiency in oilseed rape. Pedosphere. 19(3): 403–408.

    Shi, L., Wang, Y. H., Nian, F. Z., Lu, J. W., Meng, J. L. and Xu, F. S. 2009.Inheritance of boron efficiency in oilseed rape. Pedosphere. 19(3): 403–408.

    Singh, J.P., Karamanos, R.E., and Kachanoski, R.G. 1985.Spatial variation of extractable micronutrients in a cultivated and a native prairie soil. Can. J. Soil Sci. 65:149-156.

    Singh, J.P., Karamanos, R.E. and Stewart, J.W.B. 1987.The zinc fertility of Saskatchewan soils. Can. J. Soil Sci. 67:103-116.

    Solberg, E.D., Penney, D.C. and Nyborg, M.N. 1992.Factors affecting the effective use of elemental sulphur fertilizers in western Canada. In: Proceedings International Symposium on the Role of Sulphur, Magnesium and Micronutrients in Balanced Plant Nutrition. Chengdu, China.

    Soper, R.J. 1965.Effect of fertilizers on rape grown on stubble land. pp. 140-144 In: 9th Annual Manitoba Soil Science Meeting, Winnipeg, Man.

    Soper, R.J. 1971.Soil tests as a means of predicting response of rape to added N, P and K. Agron. J. 63:564-566.

    Stangoulis, J., Tate, M., Graham, R., Bucknall, M., Palmer, L., Boughton, B., and Reid, R. 2010.  The Mechanism of Boron Mobility in Wheat and Canola Phloem. Plant Physiology, 153: 876-881.

    Stewart, B.A., Porter, L.K. and Viets, Jr., F.G. 1966.Effect of Sulfur Content of Straws on Rates of Decomposition and Plant Growth. Soil Sci. Soc. Amer. Proc. 30:355-358.

    Strong, W.M. and Soper, R.J. 1973.Utilization of pelleted phosphorus by flax, wheat, rape and buckwheat from a calcareous soil. Agron. J. 65:18-21.

    Strong, W.M. and Soper, R.J. 1974.Phosphorus utilization by flax, wheat, rape, and buckwheat from a band or pellet-like application. I. Reaction zone root proliferation. Agron. J. 66:597-601.

    Strong, W.M. and Soper, R.J. 1974.Phosphorus utilization by flax, wheat, rape, and buckwheat from a band or pellet-like application. II. Influence of reaction zone phosphorus concentration and soil phosphorus supply. Agron. J. 66:601-605.

    Ukrainetz, H., Campbell, C.A., Biederbeck, V.O., Curtin, D. and Bouman, O.T. 1996.Yield and protein content of cereals and oilseed as influenced by long-term use of urea and anhydrous ammonia. Can. J. Plant Sci. 76:27-32.

    Welch, R.M. 1995.Micronutrient nutrition of plants. Crit. Rev. Pl. Sci. 14:49-82.

    Yang, Y., Xue, X., Ye, Z. and Wang, K. 1993.Responses of rape enotypes to boron application. Plant Soil 155/156:321-324.

    Ye, Z., Huang, L., Bell, R. W. and Dell, B. (2003).Low root zone temperature favours shoot B partitioning into young leaves of oilseed rape (Brassica napus). Physiologia Plantarum, 118: 213–220.

    Zhang, Q.Z., Kullmann, A. and Geisler, G. 1991.Nitrogen transportation in oilseed rape (Brassica napus L.) plant during flowering and early siliqua developing. J. Agron. & Crop Sci. 167:229-235.

    Zhao, F.J., Blake-Kalff, M.M.A., Riley, N., Hawkesford, M.J. and McGrath, S.P. 1999.Sulphur utilization efficiency in oilseed rape. In: GCIRC 10th Congress Proceedings.

    Zhao, F.J., Evans, E.J. and Bilsborrow, P.E. 1995.Varietal differences in sulphur uptake and utilization in relation to glucosinolate accumulation in oilseed rape. pp 271-273 In: GCIRC 9th Congress Proceedings.