Soil pH Cannot Be Used to Predict or Estimate Plant Nutrient Availability

By

Alfred E. Hartemink | Professor, Dept. of Soil Science, F.D. Hole Soils Lab, University of Wisconsin-Madison

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February 9, 2024

4-4-4 — Soil pH affects the availability of many nutrients, but the optimum pH for plant growth depends on which nutrient is the most limiting (photo courtesy Danny Klittich, Mission Produce, Inc.)

Soil acidity and soil alkalinity in relation to plant growth has been well-studied. Soil pH is often used as an indicator of the chemical fertility of the soil, and it is believed that most major and minor plant nutrients are best available around a slightly acid pH. This concept of soil pH-nutrient availability, the Achilles heel of soil fertility studies, was first developed in the 1930s and 1940s based on field trials, observation and various assumptions.

Early Conceptions
In 1936, a bulletin entitled, “A useful chart for teaching the relation of soil reaction to the availability of plant nutrients to crops” was published (Pettinger 1936) which stated, “…the effect of the degree of acidity or alkalinity on the availability of plant foods, or the relation between lime and fertilizers is one of the most widely discussed subjects in agriculture.” Soil reaction was perceived to be “…one of the pulses which indicates the state of health of the soil.” In the bulletin and diagram that came with it, Pettinger discussed the range of soil pH in relation to the availability of potassium, nitrates, magnesium, calcium, phosphates, iron, aluminium and manganese. A color diagram was presented that composed a series of bands representing the availability of plant nutrients in relation to a pH range of 4 to 10. The changes in width of the bands represent changes in the availability of the nutrient. It was stated that the diagram was designed to illustrate basic principles in the availability of nutrients in relation to soil reaction and did not “…portray the situation in a quantitative or absolute manner for any particular soil.” The diagram was considered only valid for well-drained soils of humid regions and not for alkali soils of arid regions or poorly drained or organic soils. The availability of some nutrients was directly affected by soil reaction whereas for other nutrients the availability was controlled by processes not related to the soil reaction. The bulletin noted, “…when the discovery of new evidence makes it necessary to discard present beliefs either wholly or in part, or when better methods of representing the facts are developed, the diagram will be revised and re-issued in improved form.”
The bulletin was not widely distributed, and it was received by Emil Truog at the University of Wisconsin-Madison who, by the 1930s, was a national leader in soil fertility and plant nutrition.

Relationship between soil pH and nutrient availability from Emil Truog’s 1946 paper in the Soil Science Society of America Proceedings (top), and a modern depiction of the relationship (bottom).

His work on the availability of plant nutrients emphasized the availability of plant nutrients was a relative matter and ‘available’ should be replaced by ‘readily available’, and ‘unavailable’ by ‘difficulty or slowly available’ (Truog 1937a), and that different cropping systems and crops have different levels of nutrient requirements and sufficiency levels.
Truog liked the soil pH-nutrient availability diagram, and considered it very useful and “…the subject of tremendous importance in connection with liming, fertilizing and soil management” (Truog 1946). He expanded the diagram to 11 nutrients and made it “…more simple in form but more complete in several aspects” (Truog 1946). The diagram illustrated the relation of the soil pH to plant nutrients in which the width of the band at any pH value indicates the relative availability of the nutrient. The band did not present the actual amount as that was affected by other factors such as the type of crop, soil and fertilization. For the 11 nutrients on the diagram, a pH of around 6.5 was most favorable but did not mean a satisfactory supply; it indicated as far as the soil reaction was concerned, the conditions were favorable. Likewise, it did not mean outside the favorable range that a deficiency would prevail. Nutrients outside the optimal range could be adequately supplied as other factors than the soil pH affected plant growth or as some plants had low requirements for a particular nutrient at a high or low pH (Truog 1946).

Previous research shows a direct effect of acidity on plant roots and on soil microorganisms, and pH at the root surface may differ from that of the bulk soil (photo courtesy Danny Klittich, Mission Produce, Inc.)

Limitations
The soil pH-nutrient diagram was presented as conceptual in 1937 and 1946 and contained several assumptions. It assumed the availability of nutrients was the same to all plants in all soils and it was best to have the soil around pH 6.5. However, many acid soils are highly productive as are some soils that have an alkaline pH. The diagram suggested deficiencies of micronutrients did not occur at low pH and there were no problems with the availability of potassium or sulfur at high pH (Blamey 2005). There are plants that require a high soil acidity such as tea, pineapple, blueberry and cranberry, and others that require a high soil pH (Hartemink and Barrow 2023).

There are numerous cases in the availability of plant nutrients that do not match the diagram, and some of them were already highlighted by Truog (e.g., the toxicity of copper and zinc in acid soils, and the fact that calcium may not be a limiting factor in acid soils, which is not uncommon). It was often found that despite the low availability of calcium at low pH, liming had limited effect as calcium was taken up from the subsoil, other nutrients were limiting (in particular phosphorus), or soil drainage was the problem (Truog 1937b). Improved crop performance with liming is often from the reduction in aluminum toxicity, and calcium deficiency is not always the major cause of poor growth (Blamey and Chapman 1982). Other exceptions to the diagram include manganese toxicity at low soil pH, iron toxicity on acid soils, boron deficiency in alkaline soils and sulfur deficiency on alkaline soils (Hartemink and Barrow 2023). Some of these exceptions to the pH-nutrient availability concept have been explained as “…simply due to methodology” (Penn and Camberato 2019).

Sources of soil acidity include urea- and ammonium-containing nitrogen fertilizers, sulfur soil amendments and biological soil processes (photo courtesy Danny Klittich, Mission Produce, Inc.)

The availability of phosphorus is often assumed to be problematic in low-pH soils where it is said to be fixed by iron and aluminium, or in soils with a high pH when phosphorus is precipitated by calcium. Of all the plant nutrients, this is probably the most widely accepted pH-availability relationship, and in a recent review it has been termed the “…the classic understanding of the effect of pH on P uptake from soils” (Penn and Camberato 2019). Barrow recently summed up the problems with this model: It makes wrong predictions, there is very little evidence for the existence of the separate postulated sinks for phosphate and it has no facility for explaining other aspects of the behavior of phosphates (Barrow 2017). There are different effects of pH on the P availability. When the pH is decreased from 6 to 4, the rate of uptake of phosphate by roots increases, the amount desorbed from soil increases and the amount sorbed by soil often also increases. The first two increase the P availability while the third effect decreases it. The pH-phosphorus availability diagram fails the most fundamental test of science and is difficult to understand why it persists (Barrow 2017).

Soil pH is a useful indicator of the soil condition, and it affects numerous soil chemical reactions and processes. But it cannot be used to predict or estimate plant nutrient availability, and different plants respond differently as nutrients interact which can be synergistic as well as antagonistic (Barrow and Hartemink 2023). Soil pH influences solubility, concentration in soil solution, ionic form, and mobility of most plant nutrients. Soil pH affects the availability of many nutrients, but the optimum pH for plant growth depends on which nutrient is the most limiting (Barrow 2017). Furthermore, the activity of microbial communities and a range of chemical reactions in soil are affected by fluctuating pH. The bulk pH of the soil (commonly measured in a soil-water ratio) may not reflect the pH in the rhizosphere where nutrients are taken up by the plant. The soil solution pH is relevant for soil and plant biogeochemical processes, and better a predictor of crop yields than the soil pH measured in a soil-water mixture. Too seldom have theories been tested by actually measuring the effects of pH on uptake of nutrients by plants growing in soil (Barrow and Hartemink 2023).

The influence of soil pH on bioavailability is indirect at best through the competition with cations for dissolved ligands or surface functional groups and through breakdown of minerals by the protons which may enhance the bioavailability of some metals. There is also a direct effect of acidity on plant roots and on soil microorganisms (Sposito 1989), and pH at the root surface may differ from that of the bulk soil . Some recent research highlighted the importance of root-induced changes in the rhizosphere pH. In soils with pH-dependent charge (e.g., ultisols, oxisols), pH increases tend to increase the P concentration in solution and its availability to plants, whereas in soils with permanent charge it is typically the other way around (Hartemink and Barrow 2023).

Truog believed the soil pH-nutrient availability diagram presented a fairly reliable picture, but he stressed it was generalized and tentative and partly based on assumptions as data were lacking. The 1946 paper “Soil reaction influence on availability of plant nutrients” provided no data and no references. The diagram has never received further investigation but ended up in many textbooks and popular soil books and continues to be used in textbooks, encyclopedias, extension bulletins and numerous papers. The diagram has many more usages, often without citation, which suggests it has been accepted as common knowledge. It has become a defining principle in soil fertility and plant nutrition.
Since the 1950s, a large amount of research work has been done on the solubility of nutrients, the biological transformations of nutrients in soils and the effect of soil pH on adsorption and plant uptake. None of that can possibly be summarized in a simple diagram. The relationship between soil pH and nutrient availability remains of interest as nutrient availability in acid and alkaline soils is unique for each soil, crop and climatic region.

References
Barrow, N.J., 2017. The effects of pH on phosphate uptake from the soil. Plant and Soil, 410(1): 401-410.
Barrow, N.J. and Hartemink, A.E., 2023. The effects of pH on nutrient availability depend on both soils and plants. Plant and Soil, 487(1-2): 21-37.
Blamey, F.P.C., 2005. Comments on a figure in “Australian Soils and Landscapes: An Illustrated Compendium” ASSSI Newsletter, 142.
Blamey, F.P.C. and Chapman, J., 1982. Soil amelioration effects on peanut growth, yield and quality. Plant and Soil, 65(3): 319-334.
Hartemink, A.E. and Barrow, N.J., 2023. Soil pH-nutrient relationships: the diagram. Plant and Soil, 486(1-2): 209-215.
Penn, C.J. and Camberato, J.J., 2019. A Critical Review on Soil Chemical Processes that Control How Soil pH Affects Phosphorus Availability to Plants. Agriculture, 9(6): 120.
Pettinger, N.A., 1936. A useful chart for teaching the relation of soil reaction to the availability of plant nutrients to crops. Virginia Agricultural and Mechanical College and Polytechnic Institute and the United States Department of Agriculture, Cooperating, Blacksburg.
Sposito, G., 1989. The chemistry of soils. Oxford University Press, New York.
Truog, E., 1937a. Availability of essential soil elements – a relative matter. Soil Sci. Soc. Am. Proc.(1): 135-142.
Truog, E., 1937b. A new soil acidity test for field purposes. Soil Science Society of America Proceedings, 1: 155-159.
Truog, E., 1946. Soil reaction influence on availability of plant nutrients. Soil Science Society of America Proceedings, 11: 305-308.

Useful Soil Maps in Microirrigated Orchards

By

Dennis L. Corwin | Research Soil and Environmental Scientist, U.S. Salinity Laboratory, USDA-ARS, Todd H. Skaggs | Research Leader, U.S. Salinity Laboratory, USDA-ARS and Elia Scudiero | Research Agronomist, Dept. of Environmental Sciences, UC Riverside, U.S. Salinity Laboratory, USDA-ARS

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February 9, 2024

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Figure1 copy — Figure 1. On the left panel, the Soil Survey Geographic Database (SSURGO) soil map for portion of the Agricultural Experiment Station at UC Riverside. A 3.5-acre field is zoomed in. Topsoil texture and soil available water capacity (AWC) information for the two mapping units within the field is reported at the bottom of the quadrant. On the right panel, a) soil apparent electrical conductivity (ECa) survey at the field and the soil sampling locations, b) topsoil (1 foot) sand content maps and c) topsoil AWC map.

Beyond planting and harvesting techniques, precision involves understanding and management of the very foundation of agriculture: the soil. Recent research led by UC Riverside and USDA-ARS U. S. Salinity Laboratory scientists (Scudiero et al. 2024; Corwin et al. 2022) offer some new insights on the use of soil apparent electrical conductivity (ECa) sensors to characterize the spatial variability of soil texture, soil moisture, salinity and related soil properties in microirrigated orchards. Please reach out to the corresponding author of this article if you would like a copy of the research papers discussed here.

Knowledge of Average, Variation and Spatial Patterns of Soil Properties is Key
In this context, where to sample soil and trees for nutrient level or water status, where to install soil moisture sensors and where to collect yield measurements. Soil maps from the USDA Natural Resources Conservation Service (e.g., SSURGO maps) are an invaluable resource for landscape-scale analyses. They can be accessed and explored at https://casoilresource.lawr.ucdavis.edu/gmap/. Unfortunately, these maps were not created to support agricultural management at the sub-field scale. When accurate, higher-resolution field-scale maps are needed and there are no funds to take and analyze hundreds of soil cores, geospatial soil sensors such as ECa can come to the rescue. The use of ECa to direct soil sampling and map soil properties is well established. However, ECa is not a direct measurement of any agronomically relevant soil property; it is a measure of how well a soil can conduct electricity. Dennis Corwin and colleagues (see Corwin and Scudiero 2020) have developed field and laboratory protocols to obtain reliable ECa measurements and soil maps. One of the key recommendations in the protocols is that effectiveness of ECa measurements peaks in uniformly wet fields. In flood- and sprinkler-irrigated fields, this condition is easily met, which contributed to making ECa arguably the most popular on-the-go near-ground sensor in the U.S. and globally. Figure 1, see page 14 compares the SSURGO mapping units information for one 3.5-acre field at UC Riverside with high-resolution soil maps obtained from hundreds of georeferenced ECa measurements and 30 soil cores and lab analyses. Linear regression statistical models were used to “calibrate” the ECa measurements to estimate target soil properties across the whole field. The ECa-derived soil maps reveal a different spatial pattern of soil properties compared to the SSURGO maps as well as a generally wider range of sand content and available water capacity (AWC). Most importantly, the AWC values mapped at the site are overall substantially lower than the values reported in SSURGO.

Figure 1. On the left panel, the Soil Survey Geographic Database (SSURGO) soil map for portion of the Agricultural Experiment Station at UC Riverside. A 3.5-acre field is zoomed in. Topsoil texture and soil available water capacity (AWC) information for the two mapping units within the field is reported at the bottom of the quadrant. On the right panel, a) soil apparent electrical conductivity (ECa) survey at the field and the soil sampling locations, b) topsoil (1 foot) sand content maps and c) topsoil AWC map.

Soil Moisture Rarely Uniform in Microirrigated Orchards
When done right, microirrigation delivers water only where desired and needed (i.e., where the tree roots are). At the beginning of irrigation, water content is usually very high near the emitters. Then, with time, water redistributes in depth and laterally. Lateral movement of water away from emitters is greater in finer-textured soils than in sandy soils. When salinity is present, salts are pushed downward and outward at the edges of the wetted soil volume. In contrast, soil in the alleyways (between the tree rows) is generally very dry unless it rains or during winter leaching irrigations. The typical ECa measurement setup has the sensor being dragged in a non-metallic sled behind a field vehicle. In orchards, that would therefore be in the middle of the alleyways, where the soil is generally too dry for making reliable ECa measurements.

What Happens if ECa is Surveyed Over Dry Alleyway Soils?
Chances are you will get an ECa survey that does not strongly correlate with the soil properties you intended to map. If the soil is too dry, electrical current may not find continuous pathways in a consistent manner through the soil (Scudiero et al. 2024). We took ECa measurements in a non-salt-affected drip-irrigated pistachio orchard (60 acres) in Lost Hills, Calif. both in the middle of the alleyways (around 10 ft away from the driplines) and along the driplines (1 foot away). The ECa measurements in the alleyways did not show reliable correlations with texture and water content. Conversely, ECa measured close to the driplines yielded significant correlations with soil moisture and texture in the topsoil and down to 5 ft.

Figure 2. The sled apparatus used by Scudiero et al. (2024) to measure soil apparent electrical conductivity (ECa) with electromagnetic induction sensors in microirrigated orchards.

If the goal is to map soil physical properties that do not change over time such as texture and available water capacity, one of the solutions to the dry alleyway problem is to carry out ECa surveys after rain events when the soil profile is close to field capacity. If the goal is to monitor more dynamic properties, such as salinity, water content or nutrient availability, ECa should be measured close to the water emitters under the tree canopy. We developed the rig shown in Figure 2, see page 16 for carrying out such surveys. The electromagnetic induction sensor that measures ECa is housed in a polyethylene utility sled. An adjustable arm allows the sled to be towed to the side of the utility vehicle so ECa can be measured under tree canopies, a position that is otherwise largely inaccessible to vehicle-mounted sensors. The sensor communicates via Bluetooth or cable to a datalogger in the field vehicle cabin. A GPS or GNSS receiver is placed on top of the field vehicle cabin to reduce geopositioning interference from dense tree canopies. It is important to note GPS coordinates and corresponding sensor locations do not coincide. To calculate the actual coordinates of the soil sensor based on a fixed offset, Scudiero et al. (2024) presented a computer script for geospatial ECa data post-processing, which is available at https://github.com/usda-ars-ussl/sensoff. A Microsoft Excel spreadsheet to calculate the coordinate offset is also available and can be requested from the corresponding author of this article.

Figure 3. a) Typical patterns of long-term salinity buildup on drip-irrigated/microirrigated orchards based on the work of Burt et al. (2003); b) salt efflorescence at the edges of the wetted soil in a microirrigated pistachio orchard in Fresno County (photo by E. Scudiero); and c) salinity changes from incremental radial distance (sideways and with depth) from drip emitters in a pistachio orchard in Lemoore, Calif. (data collected by Corwin et al. 2022).

We tested the rig in a 1-acre navel orange orchard in Riverside, Calif. where around 250 ECa (0 to 1.5 m) measurements were taken along with 20 topsoil (0 to 0.4 m) soil samples (Scudiero et al. 2024). The ECa was used to map soil particle size fraction. For example, the silt content was mapped with a R2 of 0.72 and a Mean Absolute Error of 1.55 %.

Using a Similar Rig to Map Soil Salinity
The short-scale changes of soil salinity from the microirrigated emitters and the edge of the wetted areas can be very large but are hard to capture with traditional soil sampling. Burt and colleagues (2003) characterized in detail the long-term salinity buildup on drip-irrigated/microirrigated trees in California (Figure 3a). Salinity buildup may often be seen on the soil surface at the edge of the areas wetted by the microirrigation emitters; see Figure 3b for an example in a drip-irrigated pistachio orchard in Fresno County. Corwin et al. (2022) presented a soil sampling protocol to reliably map soil salinity (ECe) from ECa surveys in microirrigated orchards. They present their recommended protocols using data from two pistachio sites in Lemoore, Calif. At each research site, they mapped ECa along the driplines. At 12 locations, they collected soil samples close to the dripline and out to 5 feet perpendicular from the dripline. Their survey was done days after irrigation and soil moisture was evenly distributed in the soil profile (Figure 3c). In contrast, ECe was low by the drip emitters and increased laterally and with depth (Figure 3c). Because of this short-scale variation in salinity, ECa was not correlated with the cores taken by the dripline (R2= 0.25; poor model performance). By averaging the soil cores collected by the dripline and 5 feet away from it, ECa’s correlation to soil salinity was much stronger (R2=0.73). Sampling by the dripline and 5 feet away from the dripline (i.e., at the edge of the root zone) provided the best “root zone” salinity average estimation. Mixing the two soil cores can make soil analyses cheaper.

Data Fusion Between ECa and Gamma-Ray Spectrometry
Researchers have utilized various geospatial sensor data, such as soil penetrometry, visible and near-infrared sensors and gamma-ray (γ-ray) spectrometry, either as independent sensing methods or in combination with ECa for a detailed understanding of soil variability and soil-plant interactions. Gamma-ray spectrometers detect radiation from soil, emanating from natural radioisotopes of elements like potassium, cesium, thorium and uranium (Figure 4). This technology has proven effective in creating accurate high-resolution maps of soil surface texture and clay mineralogy. While γ-ray spectrometry is generally employed for surveying the soil surface (usually the top few centimeters), under specific conditions, such as dry soil, it can assess soil profiles up to a depth of 1 meter. In dry soil conditions, which are less suitable for ECa measurements but ideal for γ-ray spectrometry, a 1% increase in soil moisture typically leads to an almost equivalent decrease in γ-ray emission from the soil. In microirrigated orchards, combining ECa data from the driplines with γ-ray data from these alleyways offers a promising approach to accurately assess soil spatial variability in such agricultural setups. We tested this fusion of ECa measured along the driplines and γ-ray total counts (TC) measured in the dry alleyways at a sandy loam citrus orchard in Riverside, Calif. (Scudiero et al. 2024). The study aimed to characterize field-scale soil particle size fraction spatial variability within the top 0.4 m of the soil profile. Significant Pearson correlation coefficients were found between sand and silt content with both ECa and TC. The results show a strong positive relationship between TC and clay content and negative with sand content. In particular, sand content was mapped with a very low mean absolute error (3.06%). These results indicated the ECa measurements obtained with the mobile platform were accurate and that both ECa and TC were effective predictors for soil texture spatial variability in non-salt-affected soils.

Figure 4. a) The gamma-ray spectrometry rig used by Scudiero et al. (2024) and b) the gamma ray total counts (measured in counts per second) they produced at the 1-acre citrus orchard in Riverside, Calif.

The integration of ECa and gamma-ray sensing technologies provides a robust and innovative approach to soil characterization in micro-irrigated orchard systems (Scudiero et al. 2024). If you can easily access the under-canopy space where the soil is moist, on-the-go ECa will be a good choice to map many soil properties. Otherwise, you may have to wait for abundant rains or resort to the use of γ-ray spectrometry in the alleyways. However, when salinity is a concern, there is no alternative to mapping the soil along the drip lines. Do not expect reliable soil salinity maps otherwise! The findings of Corwin et al. (2022) have significant practical implications for managing salinity in microirrigated orchards: ECa measurements should be calibrated with soil salinity measured in the entire rootzone, not just at a core by the dripline. Corwin et al.’s improved ECa-directed soil sampling guidelines offer a more accurate and representative measure of the soil salinity profile, which is critical for orchards with drip irrigation systems. We encourage crop consultants and practitioners in the field of precision agriculture to consider these advanced soil characterization techniques. Their adoption could significantly enhance the effectiveness of irrigation practices in microirrigated orchards, leading to more sustainable and productive agricultural outcomes.

References
Burt, C.M., Isbell, B. and Burt, L., 2003, November. Long-term salinity buildup on drip/micro-irrigated trees in California. In Proc. Irrigation Assoc. Tech. Conf (pp. 46-56).
Corwin, D.L., Scudiero, E., Zaccaria, D., 2022. Modified ECa – ECe protocols for mapping soil salinity under micro-irrigation. Agricultural Water Management 269, 107640. https://doi.org/10.1016/j.agwat.2022.107640.
Corwin, D.L., Scudiero, E., 2020. Field-scale apparent soil electrical conductivity. Soil Science Society of America Journal 84, 1405-1441. https://doi.org/10.1002/saj2.20153.
Scudiero, E., Corwin, D.L., Markley, P.T., Pourreza, A., Rounsaville, T., Bughici, T., Skaggs, T.H., 2024. A system for concurrent on-the-go soil apparent electrical conductivity and gamma-ray sensing in micro-irrigated orchards. Soil and Tillage Research 235, 105899. https://doi.org/10.1016/j.still.2023.105899.

Fertilizer Planning for Forage Crop Production

By

Karl Wyant | Ph.D., Director of Agronomy, Nutrien; Western Region Certified Crop Advisor Executive Board Member

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January 25, 2024

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Figure 1 alfalfa by Karl Wyant, Nutrien — Growing the Crop Consultant Industry One Reader at a Time

Forage crops represent a broad group and include crops specifically grown to be grazed on by livestock or conserved in some manner for later use (e.g., alfalfa and silage corn for this article). Forage preservation practices include the baling of hay or storing the forage under conditions that help prevent decomposition (e.g., silage). Although Western U.S. agriculture is famous for the production of tree nuts, citrus, wine grapes and vegetable crops, forage crops are also high on the list with Arizona and California growers consistently producing both high yields and high quality relative to national averages. In fact, the state of California alone has over 2 million acres of irrigated alfalfa and other forage crops spread across the state in multiple growing regions. A 2018 study estimated California alfalfa hay growers harvested 980,000 acres with a production value of $769.8 million. Additionally in 2018, California alfalfa hay was ranked as the state’s 11th most valuable commodity. Forage production remains a major crop category in Arizona as well, often ranking as the top five crops in several growing regions (Figure 1). In short, forage crops are a big deal!

Figure 1. A summer monsoon storm moves over an alfalfa field in the Sonoran Desert near Salome, Ariz. (photo by K. Wyant.)

Nutrient Needs for Silage Corn and Alfalfa
A unique characteristic of forage crops, relative to other crops, is the entire aboveground mass of stems, stalks, leaves, grain, etc. is harvested. As a result, large quantities of nutrients are exported off the field during a typical harvest (Figure 2). For most crops, only a portion is harvested (e.g., nut, fruit piece, cob, pod, etc.) and most of the plant remains behind on the field. Imagine if we removed the entire tree each time we harvested citrus? Sounds ridiculous, but whole-plant harvest for forage crops dictates a nutrient management plan that can 1) support the tremendous forage crop yields we can generate in Arizona and California and 2) replace the nutrients that were exported off the field to maintain long term soil fertility.

Figure 2. Silage corn (top) and alfalfa (bottom) nutrient uptake and removal estimates for 1 ton/acre of material (left) and fully expressed for a high yield goal (right). Source: IPNI Nutrien Removal Calculator; Nutrien-eKnomics.com

How Much NPK Do I Need?
Nutrient uptake and removal rates for high-yielding forage crops can be jarring to those unaccustomed to triple-digit export numbers. If improperly fertilized, forage crops may not hit estimated yield and quality goals, and subsequent soil fertility will decrease for the next crop in the rotation. However, there are many tools available for forage crop growers and their consultants to use to help match mineral fertilizer needs (e.g., urea, monoammonium phosphate, muriate of potash, etc.) and manage costs (Figure 3).

Figure 3. Full accounting of the NPK needed for high-yield forage systems (green box) can help determine proper input rates (e.g., manure and mineral fertilizers) and make sure the NPK in your soil and water are properly credited to avoid unnecessary expense (left side). On the harvest side of the nutrient budget (right size), we strive to ensure the crop uses the provided nutrients, which helps prevent losses to the environment.

A suite of soil and water tests can help determine what ‘free’ nutrients may already be available to the crop, and a good manure sample can be invaluable for determining exactly what was applied on the field as many forage cropping systems are near diaries, egg laying and broiler operations, and cattle feedlots. Manure can be a crucial source of NPK for forage crops, but lab tests are needed to help estimate NPK input rates (lbs/ac) given the observed ranges seen across the manure spectrum (Table 1). Upon quick inspection of the report, NPK ranges for each manure type will reveal how simple assumptions can quickly promote the development of underfertilized or overfertilized fields. Why the broad range? Manure are tricky as the NPK content can be impacted by animal species, diet, housing and bedding, manure storage and handling system, weather impacts on animals, etc.

Table 1. Not all manures are created equal, and lab tests will help focus your nutrient budget and avoid an overfertilized and underfertilized field (source: https://extension.umn.edu/manure-management/manure-characteristics#graph-summary-2317710).

Now that we know our various nutrient needs and have accounted for the NPK in the water, manure and soil, we can get a better estimate for what mineral fertilizer needs remain to drive optimal crop yields.

Total crop uptake (lbs/ac)
– Nutrients delivered in water (lbs/ac)
– Nutrients in soil (lbs/ac)
– Nutrients provided by manure application (lbs/ac)
= Nutrients needed by fertilizer program (lbs/ac)

Notes on Legumes, N Fixation and Fertilizer Programs
Nitrogen
Alfalfa production has a long history in Arizona and California, and its unique ability to ‘fix’ nitrogen and make its own N fertilizer is worth mentioning. This N fixation capacity (lbs N/ac) is a crucial part of the overall fertilizer budget for alfalfa. Due to the relationship between the crop and N-fixing bacteria, one might assume the plant does not need any additional N fertilizer to fuel crop growth (Table 2).

Table 2. Nitrogen fixing capacity of various legume crops Care must be taken when trying to supply alfalfa crops with extra N fertilizer as the fixation process is sensitive to excess N in the soil (source: Better Crops Vol. 83 #1 – 1999).

However, this assumption is not valid as it does not recognize the temporal dynamics inherent to the development of optimal rates of N fixation (Marschner 2012). High-yielding legume crops may need supplemental N fertilizer inputs to drive growth at a few key stages (Figure 4).

Figure 4. Total nitrogen status of a crop is a good predictor of crop performance and yield (top). In high-yielding alfalfa systems, crop demand for N can outstrip the capacity of the plant to generate it. For legumes, total N is related to two factors: the N produced by bacteria in the root nodules and the soil/fertilizer supply of N (source: Marschner 2012).

This includes early seedling stages when the N fixation relationship is not yet optimized and in later stages when crop growth rate is demanding N at a rate where fixation cannot keep up (Peoples et al. 1989). Care must be taken when trying to supply alfalfa crops with extra N fertilizer as the fixation process is sensitive to excess N in the soil. The traditional set of tests (manure, water and soil) can help determine if additional N is needed for alfalfa production.

Phosphate
The role of phosphorus in crop production is well established, and P also plays a key role in promoting N fixation. Under low P supply conditions, P deficiency limits plant root growth and the creation of ATP (biological currency) used to build sugars. Remember the critical relationship between plants and N fixing bacteria: No sugar equals no carbohydrates to pay for N fixation. In a study, Cassman et al. (1980) show that by increasing the P supply to a soybean crop, they were able to increase both root and nodule weight. This, in turn, drives an increase in aboveground yield as measured by shoot dry weight (Table 3).

Table 3. Increased phosphorus availability drives a substantial increase in nodule size and weight, which increases the nitrogen fixating capacity of the plant. This, in turn, increases the N content of the leaves and has the potential to influence yield (source: Better Crops Vol. 83 #1 – 1999).

Potassium
In general, potassium has been shown to increase rates of N fixation and overall yields via the following mechanisms (Better Crops 1998). K contributes to good root growth and has been shown to improve the number and size of nodules on roots. K serves as a cofactor for the action of an enzyme needed to transport carbohydrates across cell membranes and into the phloem. Remember the critical relationship between plants and N-fixing bacteria: No sugar equals no carbohydrates to pay for N fixation. In a study, Better Crops (1998), showed by increasing the K2O supply to a soybean crop, they were able to increase both nodule number and nodule weight. This, subsequently, led to an increase in aboveground yield and seed protein quality (Table 4).

Table 4. Increased potassium supply allows for the plant to support an increase of larger root nodules relative to the control. The increase in nitrogen fixing capacity leads to a ~2x increase in soybean yield and quality (source: Better Crops Vol. 82 # 3 – 1998).

Forage crop growers are facing multiple challenges including volatility in fertilizer prices, supply and logistics bottlenecks, and water supply constraints among others. These challenges come at a time characterized by a tandem increase in demand for animal-based products such as dairy, eggs and meat as well as a stricter regulatory environment. Proper nutrient management will help optimize the yield, quality and profitability of forage crops and help meet future demand for nutritious food. Furthermore, proper accounting and application of NPK from both manure- and mineral fertilizer-based sources will help prevent losses to the environment and ensure that regulatory conditions are met. Working with an experienced crop advisor can clarify the various sample reports (e.g., manure, water, soil tests), estimate total NPK uptake needs and, finally, help develop a sound nutrient management plan that meets the criteria of an increasingly complex production environment.

Resources
Better Crops/Vol. 82 (1998, No. 3) – LXXXII (82) 1998, No. 3 (ipni.net)
Better Crops/Vol. 83 (1999, No. 1) – LXXXIII (83) 1999, No. 1 (ipni.net)
Minnesota Extension Service – Manure Characteristics (umn.edu)
Understanding the Role of NPK in Promoting N Fixation in Legume Crops | Helena Agri-Enterprises, LLC
Alfalfa & Forage Industry – California Alfalfa & Forage Association (calhay.org)

LAMP as a New Tool for Testing Grapevine Red Blotch Virus

By

Joseph B. DeShields | Faculty Research Assistant, Oregon State University and Achala N. KC | Associate Professor, Oregon State University

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January 20, 2024

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Since its discovery in 2008, grapevine red blotch disease (GRBD) has negatively impacted the quality of wines due to reductions of sugar and color in the fruit. Its economic impact in the Western U.S. is estimated to range from $2,200 to $68,500 per vineyard depending on the growing region. Due to the presence of an insect vector capable of spreading the grapevine red blotch virus (GRBV), healthy grapevines can often become quickly infected during the growing season and symptoms can go unnoticed until the following season. Therefore, early detection of GRBV is even more crucial to preventing further transmission of the virus.

Symptoms of GRBD are often expressive in their characteristically red blotching patterns on leaves of red wine cultivars and likewise with yellow/yellow-white blotching on the leaves of white wine cultivars. However, symptoms on grapevines with established GRBV infections typically do not appear until after veraison. Consequently, molecular detection of GRBV can be critical for early determination of the infection status during early, non-symptomatic stages of infection.

Available GRBV Testing Strategies
There are multiple methods and strategies for diagnosing GRBD, including foliar symptom observation and monitoring, hyperspectral imaging, conventional polymerase chain reaction (PCR), quantitative PCR (qPCR), loop-mediated isothermal amplification (LAMP), plasmonic CRISPR and recombinase polymerase amplification (RPA). Among all these methods, PCR has remained the standard since 2014 due to its reliability, specificity and sensitivity; however, the PCR method creates technical, financial and infrastructure barriers for laymen due to the requirement for clean spaces, expensive instrumentation, complex troubleshooting and interpretation of results.

Other DNA-based methods such as LAMP and RPA, which are conducted at a stable reaction temperature throughout the procedure, do not require the same expensive equipment that PCR requires. The results from these two methods can be achieved much faster with reaction times as short as 20 or 30 minutes. In addition, LAMP and RPA are typically considered more sensitive to the DNA they target and less sensitive to impurities in the sample.

The pin-prick DNA extraction method being performed on grapevine leaves and dormant canes.

What is LAMP?
LAMP is a molecular tool used to detect DNA, commonly used as a diagnostic method for infectious diseases of plants and animals. Since its initial discovery by Notomi et al. (2000), the LAMP method has received much interest from private, academic and government sectors as well as from growers due to the low barriers to entry. In the past two decades, new formats for LAMP have been developed, making interpretation of results even more simplistic compared to the original method which had involved a gel electrophoresis system, additional chemicals and an advanced imaging system. These new formats allow the final interpretations to be done visually without any instrumentation. For example, the GRBV-positive reactions in some LAMP formats can create turbidity or cloudiness in the reaction tube, indicating GRBV was present in the sample, but more common is a change in color using pH indicators.

Recently, Romero Romero et al. (2019) published a LAMP method for the detection of GRBV along with a simplistic “pin-prick” DNA extraction method which consists of pricking leaf blades and petioles with a pipette tip and soaking them in water for 10 minutes to complete the extraction. Furthermore, these researchers paired this DNA extraction method with a colorimetric LAMP reagent that uses a pH indicator dye to determine whether the LAMP result was positive (yellow) or negative (pink) making interpretation faster and simpler.
We were interested in comparing its sensitivity and specificity to other more commonly used methods, such as PCR, qPCR and symptom monitoring.

The Experiment Design
We compared the four methods (LAMP, PCR, qPCR and visual symptom monitoring) at four different phenological time points per year for two years at a commercial vineyard in southern Oregon. We compared these methods using fully expanded, mature leaves sampled between berry set and harvest and using dormant shoot tissue during the winter. Both tissue types were collected at three different heights in the grapevine’s canopy: low-canopy (basal), mid-canopy and upper-canopy (apical). A tissue sample consisted of four leaves (one leaf from four shoots) or four dormant shoot segments and were collected for each canopy height and for each of the 40 vines used in this study. Vines were recorded for GRBD symptoms at the time of sample collection. Tissues were either subjected to a standard lab-based DNA extraction method and tested using PCR or qPCR or were subject to a simple, no-equipment-needed pin-prick DNA extraction method paired with LAMP.

Sampling of lower-canopy leaves for GRBV testing.

What Was Discovered
In leaf samples, the accuracy of all methods was reduced when samples were taken from higher positions within the canopy. Therefore, we will present the remaining results of this experiment from the data collected from basal samples only since this is already standard practice for most virus testing.

The sensitivity, or ability to detect a positive sample, of all four methods differed significantly at all time points and canopy heights. At berry set and veraison, both PCR and qPCR successfully detected GRBV in 98% GRBV-infected samples across both years whereas LAMP could only detect GRBV in 49% and 78%, respectively, of the same GRBV-infected vines. Only 31% of these same GRBV-positive grapevines expressed symptoms during veraison. At harvest, qPCR detected 100%, PCR detected 98% and LAMP detected 96% of GRBV-infected samples. At this stage, 94% of grapevines were symptomatic. At dormancy, where there are no leaves to observe GRBD symptoms, 96% of the dormant shoots tested positive using PCR and LAMP, and 95% tested positive using qPCR. There was no statistically significant difference in false-positive rates (the percentage of samples incorrectly testing positive) between methods.

Due to the nature of this virus and its vector, some of our GRBV-negative vines became infected some time into the two-year experiment. Among the eight new infections observed, seven vines tested positive at our earliest sampling timepoint, berry set, by PCR and qPCR whereas LAMP only detected one of these vines at berry set and the other six thereafter. The eighth newly infected sample tested positive by all methods, but only at the harvest sampling.

Sensitivity to GRBV compared across detection methods, grapevine phenology and canopy location.

The conclusion from this experiment was the accuracy of these three DNA-based methods very much depends on the location of the sample in the canopy. Use of lower-canopy leaf samples later into season increased the accuracy of GRBV diagnosis and reduced the variability in detectability. It is evidenced that testing with LAMP for GRBV later in the season (e.g., near commercial harvest) can yield comparable results to more standard methods such as PCR or qPCR.

More cost-effective and simple methods such as pin-prick DNA extraction and LAMP can offer a more accessible approach compared to external testing or the barriers and complexities of performing PCR in-house. While PCR and qPCR testing of GRBV remains the more accurate method when testing until veraison, this experiment suggests LAMP can serve as a useful tool for those who may be seeking alternatives to PCR testing. LAMP may be of interest for those wanting to test more routinely, closer to commercial maturity or during dormancy when foliar symptoms are absent.

2023 CCA of the Year Winner Seasoned Advisor Allan James Takes the Award at This Year’s Crop Consultant Conference

By

Taylor Chalstrom | Editor

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December 20, 2023

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This year's CCA of the Year winner was Allan James, technical services agronomist at Mid Valley Agricultural Services and CCA of eight years.

The 2023 Crop Consultant Conference, hosted on September 27 and 28 in a collaborative effort by JCS Marketing Inc. and Western Region Certified Crop Advisers (WRCCA), saw another year of record attendance and provided the opportunity for crop consultants, industry suppliers, researchers and others to network and learn.

In addition to CCAs, PCAs and growers receiving much-needed continuing education credits during the Conference’s established dual education track, WRCCA also presented its fourth-annual Crop Consultant of the Year award and Allan Romander Scholarship and Mentor Awards.

Karl Wyant, Ph.D., director of agronomy at Nutrien and WRCCA Board Chair, presented the awards.

CCAs, PCAs, growers and industry professionals congregated on the tradeshow floor during mornings and afternoons.

CCA of the Year
The CCA of the Year award recognizes a CCA in the western region (North Valley, South Valley, Coast and Desert) of the U.S. who has shown dedicated and exceptional performance as an advisor. The ideal candidate leads others to promote agricultural practices that benefit the farmers and environment in the western region. Selection criteria includes a peer nomination process, a scope of the CCA work, special skills and abilities, professional involvement and mentorship and community involvement.

This year’s CCA of the Year winner was Allan James, technical services agronomist at Mid Valley Agricultural Services. James is from Linden, Calif. and received his bachelor’s degree, master’s degree and doctorate at Iowa State University. He’s been consulting crops for over 35 years and a CCA since 2015.

“One of the great things about Allan is the expertise, having that experience really allows him to advise on crops, water and pest issues,” Wyant said.

James shared some words about what being a CCA means to him.

“I started out 43 years ago as a technical agronomist with a company out of Ripon, Calif.,” James said. “And I remember about three months after it started, I went home and told my wife, ‘This is the job I’ve been looking for.’

“I love the challenge of answering questions, solving the problem,” he continued. “Sometimes you say, ‘I can’t, I don’t know.’ And that’s really the joy of working in this business.

“Growers are exceptional, they have more backbone than anyone out there, our staff is exceptional. I couldn’t do what I do without them.”

Attendees had access to 8.0 DPR hours and 12.0 CCA hours as well as CDFA FREP and Arizona PCA hours.

Mentor Awards
Tracey Emmerick Takeuchi, plant science lecturer at California Polytechnic State University, Pomona; Richard Rosecrance, Ph.D., plant science professor at California State University, Chico; and Matthew Grieshop, Ph.D., director, Grimm Family Center for Organic Production and Research, California Polytechnic State University, San Luis Obispo were this year’s mentor award recipients. They were each nominated in the South Valley, North Valley and Coastal regions, respectively.

Takeuchi plans to use the award funds to support a student learning farm with both conventional and organic production through the purchase of a small, portable rototiller and high tunnel covers. Rosecrance plans to purchase temperature/light data, enabling students to conduct empirical investigations on dynamics of these within diverse orchard settings. Grieshop plans to support undergraduate and graduate student engagement in ongoing activities being managed by the Grimm Family Center at Cal Poly.

Western Region CCA Board Chair Dr. Karl Wyant posing with the 2023 CCA of the Year and Allan Romander Scholarship and Mentor Award recipients.

Scholarship Awards
Curtis Lefler of Hanford, Calif.; Matt Young of Modesto, Calif.; Sarah Ramirez of Morgan Hill, Calif.; and Isidro Lizarraga of Yuma, Ariz. were this year’s scholarship award recipients. All had excellent track records of awards, leadership and community service as well as internship experience. They were each nominated in the South Valley, North Valley, Coastal and Desert regions, respectively.

Lefler plans to become a CCA and achieve a master’s degree in an agronomy-related field. Young plans to become a CCA this year, achieve a master’s degree in soil science and become a Certified Professional Agronomist. Ramirez also plans to become a CCA this year and pursue a position as a soil conservationist with USDA-NCS in American Somoa. Lizarraga plans to achieve a master’s degree in agronomy.

Attendance for the Crop Consultant Conference broke records again with over 600 attendees.

The recipients of this year’s scholarship and mentor awards play a vital role in the development of CCAs in the western region and will continue to educate growers and prospective CCAs in the future.

On behalf of the JCS Marketing Inc. team and Progressive Crop Consultant magazine, the editor would like to thank all that attended this year’s Crop Consultant Conference in Visalia. The conference was a huge success with another record-breaking attendance year of over 600 that enjoyed the valuable seminars, exhibitors, food and entertainment.

An educational panel on DPR’s Pesticide Roadmap for the state was a crowd favorite at the conference.

Postharvest Fertigation in Trees and Vines Key to Root Development and Nutrient Storage

By

JW Lemons | CCA, CPAg, Verdesian Life Sciences

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December 20, 2023

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Figure 1_2 — Figure 1. Next season’s grape crop from budbreak to flowering relies solely on stored carbohydrates. Early shoot and root development, flowering and even fruit set are linked to those stored carbohydrates from the postharvest period (photo by Sean Jacobs, Agro-K.)

Studies and best practice examples corroborate it: When it comes to tree and vine postharvest fertilization, including fertigation is fundamental to ensure next season’ s crop success. It is that time of year to remind ourselves the season is not over after harvest.

Why Postharvest?
Fertigating at this stage is good for the roots. After the fruits have been collected, studies show, roots become the stronger sink for carbohydrates to fuel their growth, and access to readily available essential macronutrients and micronutrients such as nitrate and potassium can boost their development.

It may seem like nothing is happening with the grapevines after the grapes are harvested, the reality is different. After the harvest, vines continue to allocate resources; from the soil, grapevines are taking up nutrients and minerals, and with the process of photosynthesis, they create carbohydrate reserves and store them in permanent wood structures (roots and trunks). Therefore, the postharvest period is one of the most important periods for nutrient uptake, and carbohydrate reserves are used by vines for respiration during dormancy and for fueling new growth the following season. Next season’s grape crop from budbreak to flowering relies solely on stored carbohydrates. Early shoot and root development, flowering and even fruit set are linked to those stored carbohydrates from the postharvest period.

Figure 1. Next season’s grape crop from budbreak to flowering relies solely on stored carbohydrates. Early shoot and root development, flowering and even fruit set are linked to those stored carbohydrates from the postharvest period (photo by Sean Jacobs, Agro-K.)

Nutrient storage is important in all permanent crops. After heavy fruit and nut loads, the tree’s nutrient reserves are significantly reduced. Postharvest fertilizer, provided leaves are still photosynthetically active, will assure the tree can reload nutrient reserves to be well prepared to support next season’s early development. Tree crops grown in cooler climates with low temperatures during dormancy in winter will be faced with low soil temperature in early springtime and therefore limited root activity, even if ambient temperature is mild. In these conditions, tree crops and grape vines mainly rely on stored nutrients in the stem and roots.

Figure 2. Seasonal nitrogen uptake in a 13-year-old Nonpareil/Monterey almond orchard (cdfa.ca.gov/is/ffldrs/frep/FertilizationGuidelines/N_Almonds.html).

In the case of many tree fruit and nut crops, postharvest applications through foliar or fertigation could also reduce the “on-off” years incidence, where one year of heavy fruit load is followed by a year of low fruit yield. This phenomenon may be related to depleted nutrient stocks in the tree after heavy fruit load and nutrient export with harvested fruits from the orchard, rendering the tree crops unable to support a consecutive year of abundant fruit yield.

At the early bloom and fruit initiation stage, the tree fully depends on nutrient reserves, stored in the tree itself. The most important nutrients needed to top off at this period are nitrogen (N) and potassium (K), and up to 30% of total annual application of N and K should be applied. It is important to select readily available nutrient sources such as potassium nitrate, which will provide immediately available N in the form of nitrate, while tree crops need to be replenished with K as significant amounts of K are exported with the harvested fruits from the orchard.

Figure 3. Seasonal nitrogen distribution in a 13-year-old Nonpareil/Monterey almond orchard (cdfa.ca.gov/is/ffldrs/frep/FertilizationGuidelines/N_Almonds.html).

In the case of almonds, nitrogen can be applied any time after hull split up until a few weeks postharvest. In earlier harvested varieties and ‘Nonpareil,’ N can be applied shortly after harvest with the first postharvest irrigation. With later varieties like ‘Monterey’ or ‘Fritz,’ the application can be made post-hull split prior to harvest. This timing matches bud development that tends to occur about two weeks after ‘Nonpareil’ harvest for most varieties. Postharvest K applications may be a reasonable strategy if you are on soil that is able to hold the K. In sandy soils, K can be leached out of the rootzone, which may create a situation of deficiency in the following year.

In the case of grapes, the period after harvest but before leaf fall is one of the best times of the season for the uptake of N and K which the vine needs along with carbohydrates to provide for the period of rapid shoot growth in the spring after budbreak. This is encouragement to deliver these macronutrients after harvest when excessive growth and the K content of the fruit is not a concern. Replacing minerals is important as they are transported off-site in the crop. Even if some of these are recycled back into the soil like with leaves or canes, that recycling is slow and inadequate to provide the needed plant nutrients.

Figure 4. The period after grape harvest but before leaf fall is one of the best times of the season for the uptake of N and K.

In Research
There are two main stages of root growth. In a rhizotron study conducted in Chile in 1993 with two table grape varieties (Flame Seedless, Muscatel), it was shown that the first (and larger) peak root growth stage takes place from budbreak to petal fall/fruit initiation. The second (and smaller) peak root growth stage takes place after fruit harvest until leaf fall (postharvest). Root development is linked to the competition for carbohydrates between roots and developing fruits. Developing fruits are stronger sinks for carbohydrates produced in the leaf than roots. Therefore, root growth and development are suppressed during fruit development growth stages. Once the fruits have been harvested, roots become the stronger sink for carbohydrates to fuel their growth. Access to readily available essential macronutrients and micronutrients, applied with fertigation during postharvest, is equally essential to support root development. The recommended dose rate of nutrients in fertigation is to be decided by plant-soil-water diagnostics.

References
“Balanced soil fertility management in wine grape vineyards.” Grant, S. Practical Winery May/June 2002.
“Best Management Practices for Nitrogen Fertilization of Grapevines.” Peacock, B., Christensen, P. and Hirschfelt, D. University of California Cooperative Extension.
“Foothill Vineyard Post Harvest Activities: FERTILIZING: Information summarized from ‘Grapevine nutrition and fertilization in the San Joaquin Valley.’” Christensen, P., Kasimatis, A. and Jensen, F. UC ANR pub. 4087 (the “black book”) now out of print.” L.R. Wunderlich, UCCE Farm Advisor. Foothill Vineyard News, Issue 9, October 2013.
“Post-harvest Vineyard Management: Growers Guide for Riverina Vineyards.” Edited by Hackett, S. and Bartrop, K.. Riverina Wine Grapes Marketing Board. March 2011.
Resources
Almond Nutrients and Fertilization: fruitsandnuts.ucdavis.edu/crops/almond
Tree Fruit Soil Fertility and Plant Nutrition in Cropping Orchards in Central Washington: treefruit.wsu.edu/orchard-management/soils-nutrition/fruit-tree-nutrition/

Potassium and Potatoes: Understanding Fertilizer-Crop Interactions

By

Sarah Light| UCCE Agronomy Advisor, Sutter, Yuba and Colusa Counties and Dan Sullivan and Don Horneck | Oregon State University

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December 20, 2023

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Fresh,Organic,Potatoes,Isolated,On,White,Background.,Top,View,,Flat — Potassium and Potatoes: Understanding Fertilizer-Crop Interactions

Potatoes require high levels of potassium and nitrogen to achieve optimum yields. Potassium (K) is commonly applied as potassium chloride (KCl) while nitrogen (N) is applied both preplant and in season through fertigation. Crop N is managed in season through tissue testing of potato petioles for nitrate. Petiole nitrate levels are then used as a diagnostic tool for in-season N fertilizer applications based on extension recommendations.

Methods
This research was conducted in the Columbia Basin in Hermiston, Ore. Soils in this area are sandy, which increases leaching potential for nitrate and other anions, like chloride (Cl). Previous research found that petiole nitrate levels decreased as petiole Cl increased. There was concern that there was an antagonism in uptake of nitrate and Cl and that if in-season fertilizer recommendations based on petiole nitrate levels did not consider petiole Cl, growers might be applying more N than was required for crop demand. In other words, there was a concern that petiole nitrate levels might be lower not because of low soil N, but rather because of high Cl uptake following KCl application. This project was designed to understand Cl dynamics in potato production systems.

In this project, source and timing were included as treatments while rate and placement were consistent for all treatments. Three K sources (potassium chloride (KCl); sulfate of potash, K2SO4 (SOP); and K2SO4*2MgSO4 (KMag)) were applied at 200 lb K per acre at three different times in the season. The timing of the treatment applications were 206 days prior to planting (fall preplant), 14 days prior to planting (spring preplant) and 35 days after planting (layby). Russet Burbank potatoes were planted on April 11. After planting, beds were tilled by moving soil from between the rows into the potato rows. Potato plants had emerged but not closed canopy at the time of the layby fertilizer application.
Potato petioles were collected two times in the growing season (70 and 97 days after planting) to coincide with extension recommendations for in-season fertilizer management decisions. Soil samples were collected 68 to 70 days after planting. Aboveground whole plant biomass was collected 116 days after planting and potato tubers were harvested 21 days later. Potato yield and quality metrics including specific gravity were evaluated for all treatments. These plant and soil measurements were designed to understand how much Cl plants were taking up, where in the plant the Cl was going and how Cl uptake was impacting plant N levels.

Figure 1. Effect of K fertilizer source and timing on soil (0 to 8 inches depth) extractable Cl (a), SO4-S (b) and K (c) at 70 days after planting.

Source by Timing Interaction Revealed
Soil Cl concentrations revealed a source by timing interaction. When KCl was applied in the fall, soil Cl levels were similar to those in zero K control as well as KMag and SOP treatments; however, when KCl was applied in spring or during the growing season, soil Cl was greater with KCl compared to other treatments. Plant Cl measurements followed the same pattern as soil measurements. The most likely explanation for these results was greater leaching of Cl below rooting depth with fall fertilizer application as compared to spring or in-season fertilizer applications.

Despite being in a low-rainfall area, several factors favored overwinter leaching of Cl below the root zone. First, the soil texture is sandy loam with a low water-holding capacity. Second, the timing of field operations and bed preparation relative to fall fertilizer application facilitated Cl exclusion from beds. In the fall, fertilizers were applied to flat ground. Prior to potato planting in spring, beds were created from the top 2 to 4 inches of the soil present on flat ground. Therefore, overwinter leaching of Cl below a depth of approximately 4 inches would be sufficient to move it below the potato beds. Cumulative rainfall between fall preplant fertilizer application and planting in spring was 4.25 inches, though daily precipitation was never more than 0.4 inches. From February 1 to March 18 (date of spring preplant fertilizer application), cumulative rainfall was 2 inches as compared to cumulative estimated ET of 0.83 inch, which indicates a potential for leaching. In contrast, during the interval between spring preplant fertilizer application and planting (March 28 to April 11), cumulative rainfall was 0.3 inches as compared to cumulative estimated evapotranspiration (ET) of 0.43 inches. Therefore, little or no leaching was expected between spring preplant fertilizer application and planting. Following potato planting, irrigation plus precipitation did not leach out Cl.

Figure 2. Effect of K fertilizer source and timing on petiole NO3-N (a) and Cl (b) at 70 days after planting.

Petiole nitrate concentrations were similar for all treatments at both sampling dates. Higher Cl levels in petioles with spring preplant or layby KCl application did not reduce petiole nitrate. For example, at 70 days after planting, petiole Cl was increased twofold with spring preplant or layby KCl application as compared with fall application. In contrast, petiole nitrate concentrations were similar regardless of the timing of KCl application. At harvest, crop N concentrations were similar across all treatments.

K fertilizer source and timing did not impact total or marketable potato yield. There were no significant differences for specific gravity by treatment. However, among KCl treatments, the values for specific gravity and potato yield were as follows: Fall Preplant > Spring Preplant > Layby. This suggests that higher soil and plant Cl during the growing season may have delayed tuber initiation or growth. Typically, specific gravity increases with tuber maturity.

Figure 3. Effect of K fertilizer source and timing on petiole NO3-N (a) and Cl (b) at 97 days after planting.

This experiment was designed to minimize yield differences between treatments so that nutrient movement could be evaluated without regard to plant nutrient partitioning and physiological source-sink relationships. These results indicate potato plants accumulate large concentrations of Cl when available due to KCL being applied later in the growing season. Fall KCl applications, which allowed for overwinter leaching below the root zone, resulted in the lowest soil and plant Cl concentrations when compared to other application times.

In this research, aboveground biomass and tubers were collected three weeks apart and the Cl concentration in aboveground biomass was higher than in tubers. Peak nutrient uptake in potatoes occurs during times of significant aboveground growth. Photosynthates are then translocated into potato tubers during tuber bulking, which subsequently increases water uptake into tubers. As a result, Cl concentration in the tubers is diluted and decreases as a proportion of tuber weight during this bulking stage. Standard grower practice is to leave desiccated vines in the field after harvest. The majority of plant Cl is in the aboveground biomass at the end of the season and thus Cl is added back into the soil.
In this project, we did not measure an antagonism in crop update between nitrate and Cl. Our experimental design likely minimized the interaction between N and Cl because N was applied at lower rates during peak uptake to meet crop demand. Total N concentration in aboveground biomass and potato tubers as well as nitrate in petioles were generally unaffected by K source or time of K application. In this research, petiole Cl levels were affected by K source and time of application. Concentrations increased when KCl was the K source, and as KCl was applied closer to petiole sampling date. In this project, Cl levels were always higher in petioles collected later in the year, which indicates potatoes continue to accumulate Cl throughout the growing season. Petiole nitrate levels, by contrast, were consistently lower among all treatments for the second petiole collection date. Although Cl is an essential micronutrient, it is not metabolized into plant compounds, and high concentrations of Cl are maintained in aboveground plant tissue including petioles throughout the growing season.

Figure 4. Effect of K fertilizer source and timing on Cl concentration in aboveground biomass (tops) collected at 116 DAP (a) and in tubers at harvest (b). Aboveground biomass Cl uptake (kg ha-1; right axis in “a”) was estimated based on average biomass (2780 kg ha-1). Tuber Cl uptake (kg ha-1; right axis in “b”) was estimated based on average tuber dry matter (179 g kg-1) and average tuber total yield (64 Mg ha-1).

Though there were significant differences in petiole Cl concentrations by treatment, the antagonism in uptake between N and Cl in petioles that had been documented by other researchers was not measured in this study. In this project, N was applied weekly at a uniform rate to all treatments during periods of peak uptake throughout the growing season and was thus replenished and available for plant uptake. This application method likely allowed the Cl uptake and movement to be unaffected by N availability.

Potato plants can take up Cl when it is available, and that Cl accumulates in plant tissue (particularly aboveground biomass) until harvest. Higher concentrations of petiole Cl from preplant or in-season KCl application did not affect petiole nitrate when N was applied via fertigation throughout the growing season. Fall-applied Cl was not taken up by the crop because of the opportunity to leach out of the soil used to form potato beds prior to planting. Even in a low-rainfall area, sufficient leaching of Cl below the rootzone is possible if KCl is applied far in advance of planting.

This project was funded by United States Department of Agriculture: National Institute of Food and Agriculture and Compass Minerals. Dan Sullivan worked on data analysis and publication of results. Dr. Don A. Horneck proposed this research project and died before the completion of this project. He is missed.

Figure 5. Effect of fertilizer source on S concentration in petioles (pet), tubers and aboveground biomass (tops). Petioles sampled during tuber bulking growth stage (70 and 97 days after planting), aboveground biomass at 116 days after planting, tubers at harvest. Error bars indicate standard error of the mean (n=15).

A Native Plant Species as a Weedy Problem in Central Valley Orchards: The Case of Alkaliweed

By

James Schaeffer | UCCE Staff Research Associate, Fresno County, Kurt J. Hembree | UCCE Weed Science Advisor Emeritus and Anil Shrestha | Professor, Jordan College of Agricultural Sciences and Technology, California State University, Fresno

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December 20, 2023

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Fig3 — Figure 3. Reproductive structures of alkaliweed.

Alkaliweed (Cressa truxillensis) is a perennial herb (Figure 1a) native to California. Generally, it is found growing in natural areas, field margins and ditch banks. However, in recent years, it has been observed in agricultural areas. Alkaliweed was first reported as a problematic weed to the UCCE Fresno County in 2016. It is now being widely observed in tree nut orchards, agronomic crops, fallow fields, ditch banks and roadsides. It is more noticeable in young pistachio orchards in the southern Central Valley (Figure 1b). Standard orchard floor management practices such as between-row cultivation and herbicide applications have failed to control this species, and very little information is available on the biology, ecology and management of this species.

Figure 1. a) An alkaliweed plant and b) heavy infestation of alkaliweed in a Kings County pistachio orchard in late spring (all photos courtesy A. Shrestha.)

Common herbicides registered for use in orchards, such as glyphosate, glufosinate, salflufenacil, paraquat, 2-4 D, halosulfuron, carfentrazone, rimsulfuron and oxyfluorfen, only seem to suppress the plants for approximately 30 days before the plant starts regrowing. Treatments in pistachio orchards in Kings County showed the herbicide-treated plants regrew from new stems emerging from extensive underground root and/or shoot systems and from the aboveground parts of the treated plant. Similar observations were made in plants that had been cultivated with mechanical equipment. Such observations led us to suspect alkaliweed persists because of its dense aboveground and belowground plant parts (Figure 2a) and rhizome-like structures (Figure 2b).

Figure 2. a) Belowground plant parts and b) rhizome-like structures of alkaliweed.

While flowers (Figure 3) and seed are produced, very little is known about their contribution to invasion and spread. Also, very little is known about the germination ecology of its seeds, specifically in response to environmental stresses caused by soil pH, salinity and moisture that occur in the Central Valley. The populations in Kings County were observed in moderately alkaline and saline soils prone to summer drought.

Figure 3. Reproductive structures of alkaliweed.

Alkaliweed is stated to be a shade-intolerant plant (USDA-NRCS 2023). Our observations from 2016-18 showed it seemed to prefer and grow better in full sunlight than in shaded conditions. Plants were seen growing throughout young orchard floors that had very little canopy shading, but in older orchards they were observed mainly in the row middles and along unshaded areas of adjacent roadsides and canal banks. Therefore, we believed it was important to assess the shade tolerance ability of alkaliweed because such information could be beneficial in mitigating population spread and in the development of management strategies for this species. If flower and seed production are influenced by shaded conditions in this species, then more options become available in developing management practices to limit its sexual reproduction that contribute to seedbanks.

Although propagation of alkaliweed is primarily stated to be by seed (USDA-NRCS 2023), field observations have shown sprouting ability by underground plant parts that resemble rhizomes or stems as shown in Figure 2, see page 20. However, it is not known for certain what the reproductive potential of these structures are in alkaliweed and this needs to be studied, because limiting both seedbanks and bud banks may be necessary to effectively manage this species.

Therefore, we undertook a study on alkaliweed to 1) assess the germination of its seeds in response to environmental stresses such as pH, salinity, and moisture; 2) assess the impact of different levels of shade (full sunlight, 70% of full sunlight and 30% of full sunlight) on its aboveground growth and morphology; and 3) assess the effect of postemergence herbicides on its suppression.

Seed Germination
Seed germination experiments were conducted in a controlled-environment growth chamber at California State University, Fresno. Seed germination was tested under a range of pH solutions (5, 6, 7, 8 and 9) prepared using sodium hydroxide (NaOH) and hydrochloric acid (HCl) proportions. Germination was also tested in a range of salinity solutions measured by electrical conductivity (EC) of 0, 2.5, 5, 10, 15 and 20 dS/m (1 dS/m = 1 mmho/cm) that were prepared using laboratory-grade sodium chloride (NaCl). Furthermore, germination was tested under a range of water potential (ψ) solutions (0, -0.51, -1.88, -2.89, -4.12 and -5.56 MPa) prepared using polyethylene glycol (PEG 6000) to assess the drought tolerance of seeds for germination.

Effect of Shade on Alkaliweed Growth and Reproduction
An alkaliweed-infested roadside alongside the top of a ditch bank in Stratford, Calif. adjacent to a seven-year-old pistachio orchard infested with alkaliweed was used to study the effects of full sun and shade on alkaliweed morphology and growth. Shade tents (Figure 4) were constructed with PVC pipe frames and fitted with shade clothes representing 30% shade (70% of full sun) and 70% shade (30% of full sun). The shade tents were set up before alkaliweed plants emerged from the soil. Each treatment was replicated four times in a randomized complete block design. The photosynthetically active radiation (PAR) inside and outside the tents was taken every week between 11:00 a.m. and 1:00 p.m. using a hand-held quantum sensor. The number of plants and number of plants with flowers in the treatments were counted at the onset of flowering. The plants were harvested once five plants within a random area in each treatment plot had flowered. The plants were clipped at the soil surface, put into paper bags, and transported to the lab at Fresno State. In the lab, fresh weight of the five plants per treatment were taken, and the number of flowers and internodes on each plant were counted. The internode length on each plant was measured and recorded. The total leaf area on each plant was measured using a leaf area meter. The parts of the five plants were put together in separate paper bags for each treatment and placed in a forced-air oven and the dry weights were recorded after three days. The experiment was conducted from April to August 2019.

Figure 4. Layout of the shade study on the roadsides of the pistachio orchard in Stratford, Calif.

Postemergence Herbicides on Alkaliweed
The effect of postemergence herbicides on alkaliweed was tested by spraying actively growing plants on the edge of the road outside a pistachio orchard in Stratford on April 11, 2018. Since the plants had more of a semi-prostrate growth habit, the plants were approximately 1 to 1.5 inches tall and 6 inches in diameter at the time of the spray. The herbicide treatments included one-time applications, sequential applications and tank mixtures of herbicides (Table 1). Adjuvants were added to the herbicide treatments as recommended by each herbicide label and the solutions in all the treatments were buffered to a pH of 5.5 using BioLink® acidifier (Westbridge Agricultural Products, Vista, Calif). The herbicides were applied with a two-nozzle (Turbotwinjet 11004, Model TTJ60) spray boom using a CO2 backpack sprayer calibrated to spray 34.6 gallons/acre at 3 miles/hour. The spray pressure and spray height were maintained at 30 psi and 18 inches above the plant, respectively. Each treatment plot was 30 feet long and 3.3 feet wide. The experimental design was a randomized complete block with four replications of each treatment. The alkaliweed plants were evaluated at weekly intervals up to 28 days after treatment (DAT) for mortality on a scale of 0 to 100 where 0 was considered completely healthy without necrotic symptoms and 100 was considered entirely dead with no green tissue remaining.

Data for each of the experiments above were analyzed using analysis of variance (ANOVA) and when the analysis showed a significance at α=0.05, the means were separated using Fisher’s Least Significant Difference tests. Regression analysis was also used for the germination studies.

Effect of Environmental Factors on Seed Germination
Alkaliweed seeds were moderately drought-tolerant as germination was 68.5% at -0.51 MPa and 12% of the seeds germinated at a fairly high negative water potential of -1.09 MPa. There was no germination beyond water potential values of -1.09 MPa. The water potential level that reduced seed germination by 50% was estimated as -0.78 MPa. For comparison, in field bindweed (Convolvulus arvensis), another plant of the Convolvulaceae family, the water potential that reduced germination by 50% was estimated as -0.4 MPa (Tanveer et al. 2013). Therefore, it appears alkaliweed seeds are more tolerant to water potential stress than field bindweed during germination, enabling it to germinate in the semiarid regions of the Central Valley under fairly dry conditions.

Alkaliweed seeds were very tolerant to salinity (sodium chloride) stress at germination as approximately 24% of the seeds germinated at a very high EC level of 20 dS/m, although percent germination was significantly lowered at EC levels higher than 10 dS/m. The salinity level that reduced seed germination by 50% was estimated as 15.3 dS/m. Again, there are no studies that have reported the effect of salinity levels on alkaliweed, but in field bindweed Tanveer et al. (2013) reported similar tolerance to salinity stress. Therefore, alkaliweed can germinate in the high-salinity regions of the westside of the Central Valley.
Unlike water potential and salinity, germination of alkaliweed seeds was not affected by the range of the pH levels tested. Germination was similar at all pH levels (5 to 9) and ranged from 76% to 84%, indicating this species has a wide range of adaptation to pH levels during germination. Although more than 80% of the seeds germinated at a pH level of 5, it is not known if the plants would grow and reproduce at this pH level because the optimum pH range for growth of this species is reported as 6.8 to 9.2 (USDA-NRCS 2023). Again, there are no published studies on the germination of alkaliweed seed in response to pH, but Tanveer et al. (2013) reported that field bindweed germinated at a pH range of 4 to 9 but the optimum pH was 6 to 8. Therefore, it appears that alkaliweed can germinate in the alkaline and acidic soils of the Central Valley.

Effect of Shade on Growth of Alkaliweed
Shade levels influenced the morphology, growth and reproductive potential of alkaliweed plants. The PAR taken close to noon in the treatment plots during the experiment, on average, ranged from 1700 to 2000, 1000 to 1300 and 500 to 700 µmol m-2 s-1 in the full sun, 30% shade and 70% shade, respectively. Although the total aboveground biomass and number of internodes on the stem were not affected by shade level, other morphological characters showed that this species was not a shade-tolerant plant and that plants exhibited efforts to adapt to shade. For example, the length of internodes on the stem increased as the level of shade increased. The average length of the internodes in the plants growing in the full sun was approximately 1.3 inches whereas at 70% shade it was approximately 3.5 inches. Similarly, the total leaf area per plant was higher in the plants grown in the shade than in the full sun. Leaf thickness was not measured in the study. However, it is suspected that leaves on the plants growing in the shade were comparatively thinner than on those growing in the full sun because thin leaves are an adaptation strategy of shade-intolerant plants when growing under shade (Glime 2017). Similar results were reported in field bindweed where internode length and total leaf area were lesser in plants grown in the full sun than those grown under shade (Gianoli 2001). However, field bindweed is more of a viny plant than alkaliweed.

Although the plants growing in the shade had similar aboveground biomass as those in the full sun, none of the plants in either the 30% or the 70% shade treatments produced any flowers for the duration of this study. Therefore, it appears that reproduction can be severely inhibited in alkaliweed by shade, and use of shading strategies may be an effective management method to reduce sexual reproduction of this species.

Effect of Postemergence Herbicides on Alkaliweed
Plant mortality differed between the herbicide treatments at each evaluation date (Table 2). Initially, saflufenacil (alone) and the tank mix of glyphosate + saflufenacil + glufosinate looked promising as more than 80% of the plants showed injury symptoms at 7 DAT and the plants appeared to be dying. However, by 28 DAT, the plants regrew, and the mortality was reduced to 50% and 53%, respectively, for the two treatments. This level of control would not be considered acceptable to a grower managing alkaliweed. None of the herbicide treatments provided acceptable control as the mortality rate was 20% or less in most cases. It was expected that the sequential application treatments would work better but it was not the case. Sequential applications of glyphosate and carfentrazone provided 43% control whereas that of glyphosate and paraquat provided only 15% control. Although appropriate adjuvants were applied in all the treatments, the postemergence herbicides failed to control the plants which is perhaps due to reduced contact, retention and absorption of the herbicides because of the hairiness of the alkaliweed plants as observed under a microscope (Figure 5). Even the leaves of small plants were observed to be hairy. Also, it is not known if the regrowth is because of adequate levels of stored carbohydrates or growth of new stems (aboveground or belowground) as discussed earlier. Therefore, alkaliweed control may not be feasible using postemergence herbicides alone. It has been reported that 2,4-D can provide some level of control but in-season use of this herbicide in orchards could be a concern due to phytotoxic effects on the trees.
Our study showed alkaliweed seeds could germinate in moderate drought and high salinity conditions under a range of soil pH. The species was not very shade-tolerant, and no reproductive structures were observed in the plants growing in the 30 and 70% shade levels. None of the postemergence herbicides provided adequate control of the plants. Therefore, an integrated management plan which includes multiple tactics needs to be developed for managing alkaliweed in Central Valley orchards. Complete details on this study are available at mdpi.com/2392752.

Figure 5. Microscopic images showing the presence of hairs on the leaves of alkaliweed.

We would like to thank Mr. Kevin Brooks, PCA, for the wealth of information provided on alkaliweed.

References
Gianoli E. 2001. Lack of differential plasticity to shading of internodes and petioles with growth habit in Convolvulus arvensis (Convolvulaceae). Int J Plant Sci 162:1247–1252. https://doi.org/10.1086/322950.
Glime J. 2017. Light: Adaptations for Shade. In: Glime, J. M. Bryophyte Ecology. Volume 1. Physiological Ecology. 9-2-1, http://digitalcommons.mtu.edu/bryophyte-ecology.
Tanveer A, Tasneem M, Khaliq A, et al. 2013. Influence of seed size and ecological factors on the germination and emergence of field bindweed (Convolvulus arvensis). Plant daninha 31(1) https://doi.org/10.1590/S0100-83582013000100005.
USDA-NRCS, 2023. Conservation plant characteristics, Cressa truxillensis Kunth, spreading alkaliweed, CRTR5. https://plantsorig.sc.egov.usda.gov/java/charProfile?symbol=CRTR5.

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