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The Mechanics of Soil Fertility: Use of Sugar in Field Crops
Jolee Derrick, Precision Nutrient Management Ph. D. Student
Grace Williams, Soil Microbiology Ph. D. Candidate
Brian Arnall, Precision Nutrient Management Specialist
Recently, there has been increased interest in adding sugar to spray tank mixes, whether for post-emergence weed control or foliar nutrient applications. While there is limited work on impact of sugar inclusion in herbicide applications, some papers have posed potential enhancement (Devine and Hall, 1990). But since this is coming from a soil science group, we will only focus on soil impact. Following up the last blog, unlike humic substances, which represent more complex and relatively stable carbon forms, sugar is a highly labile carbon source. This rapid utilization of simple carbon sources is well documented to stimulate microbial activity and growth (Kuzyakov and Blagodatskaya, 2015). The general idea of utilizing sugar applications is that sugar has the capacity to improve spray performance, stimulate biological activity, increase organic matter mineralization, and ultimately result in improved yields.
Sugar additions can influence soil processes differently depending on system conditions. In systems with higher residual nitrogen and organic matter, responses may differ from those observed in Oklahoma production environments, where soils are typically lower in organic matter and microbial activity can occur for much of the year. Understanding how sugar functions in these systems requires a basic discussion of carbon dynamics. Sugar itself is almost entirely carbon and is readily consumed by microbes. It’s a simple molecule, which allows it to dissolve easily in water and be quickly utilized in the soil system. Crop residues, like wheat straw, are also carbon-rich but much more complex. They contain cellulose, hemicellulose, and lignin which are long carbon chains that take time to break down because microbes need specialized enzymes to access them.
For the sake of simplicity, we can group carbon into two key pools: labile carbon and particulate organic matter (POM). Labile carbon includes easily decomposed materials, which include the previously mentioned simple sugars that microbes can metabolize rapidly. These pools differ in turnover time and microbial accessibility, with labile carbon driving short-term microbial responses (Cotrufo et al. 2013). POM breaks down more slowly and serves as a longer-term nitrogen source through residue breakdown.
Soil microorganisms require both carbon and nitrogen to grow and maintain biomass, typically at a ratio of approximately 24 parts carbon to 1 part nitrogen. When readily available carbon is abundant, but nitrogen is limited, microbes increase their nitrogen demand and begin scavenging nitrogen from the surrounding soil. This process, better known as nitrogen immobilization, temporarily reduces nitrogen availability to crops. Additions of readily available carbon sources have consistently been shown to increase microbial nitrogen immobilization in soil systems (Recous et al. 1990).
In systems where sufficient nitrogen is present, microbial populations can expand rapidly. Fast-growing microbial species may dominate, continuing to immobilize nitrogen within their biomass. Eventually, when nitrogen becomes limiting, microbial populations decline to levels the system can support. This boom-and-bust cycle can disrupt nitrogen availability during critical stages of crop growth. These rapid shifts in microbial population and activity following carbon inputs are commonly observed in soil systems receiving easily decomposable substrates (Blagodatskaya and Kuzyakov, 2008).
This dynamic becomes especially relevant when considering residue management practices common in Oklahoma. Under no-till or limited-tillage systems, the crop residues have wide carbon-to-nitrogen (C:N) ratios, creating conditions where nitrogen immobilization can occur during the growing season.
Table 1 provides approximate C:N ratios for several crops commonly grown in Oklahoma. When additional carbon is introduced into these systems without accompanying nitrogen, the likelihood of microbial immobilization increases. While immobilization is not bad, it does create a question mark as Oklahoma’s variable climate means the following release of nutrients will be unpredictable.
Table 1. Table depicting the range of C:N ratios for residues of commonly utilized crops in Oklahoma. Ratios were obtained from Brady, N. C., & Weil, R. R. (2017). The Nature and Properties of Soils (15th ed.)
Now consider conventional tillage systems. In Oklahoma, no-till systems typically contain 2 to 3 percent organic matter, which is relatively high given our climate and extended periods of microbial activity. Conventional tillage systems often fall between 0.75 and 2.25 percent organic matter. Because soil organic matter is approximately 58 percent carbon, this represents a substantial difference in the soil carbon pool.
Tillage can temporarily enhance microbial access to both previously mentioned carbon pools. When tillage exposes previously protected carbon, microbial activity increases rapidly. This initial flush can temporarily increase nitrogen mineralization as organic nitrogen is converted to plant-available forms. However, this phase is short-lived. As microbial populations expand, nitrogen demand increases, leading to immobilization and reduced nitrogen availability.
Hypothetically, increased microbial growth and activity would rapidly mineralize organic matter, trigger a surge in NO₃⁻, deplete soil organic matter, and as resources become limiting and the environment can no longer sustain elevated microbial populations, this boom would be followed by a population crash. This relationship is ultimately driven by the soil C:N ratio, which introduces an interesting additional complexity of residue. Different residues bring very different carbon-to-nitrogen balances into the system, and microbes respond accordingly. High carbon residues give microbes plenty of energy but very little nitrogen, so they pull N out of the soil to meet their needs. Residues with lower C:N ratios (soybean, alfalfa, etc.) do opposite, releasing nitrogen as they break down. Now the real question becomes where the critical point sits, and when does management push the system from the threshold of immobilization and mineralization.
These hypotheses form the foundation for new research currently underway through the Precision Nutrient Management Program. Initial proof-of-concept work has already been completed, providing a necessary steppingstone to address these questions.
Figure 1. Graph depicting the different concentrations of nitrate leached corresponding to applied treatments in the proof-of-concept work
The preliminary work (Figure 1) evaluated different sugar sources applied alongside a high-nitrogen product to assess the extent of nitrogen immobilization. Although these studies were conducted using potting soils, clear trends were apparent. Treatments containing sugar consistently showed greater nitrogen immobilization compared to treatments without sugar. This response is consistent with studies showing that additions of simple carbon substrates stimulate microbial growth and increase nitrogen immobilization (Dendooven et al. 2006). Building on this work, an active field-based research project is underway to evaluate how sugar additions influence nitrogen availability and microbial dynamics under real-world Oklahoma production conditions.
From an agronomic standpoint, sugar functions primarily as a readily available carbon source that stimulates microbial growth. In nitrogen-limited systems, this response increases the likelihood that nitrogen will be incorporated into microbial biomass rather than remaining immediately available for crop uptake.
Finally, we conclude with a conceptual consideration. If increased OM mineralization leads to greater plant biomass, this process may partially offset losses of OM. Greater biomass production could return more residues to the soil, contributing to the OM pool in the upper soil profile. Therefore, the system may compensate for OM mineralization through the rebuilding of organic matter via plant inputs. However, the stabilization of this carbon depends on microbial processing and physical protection within the soil matrix (Cotrufo et al. 2015)
However, while the underlying logic is sound, this concept has not been extensively studied within Oklahoma cropping systems. This blog does not address the impact of sugar applications on residue breakdown, and the potential impact of such. Future research through the Precision Nutrient Management Program will further investigate the mineralization process to better understand carbon dynamics within these systems.
Take Home:
- Oklahoma production systems generally have lower residual N and high carbon residues, creating conditions conducive to N immobilization
- Adding sugar increases microbial growth, creating population booms that will momentarily increase mineralization, but then immediately immobilize residual nitrogen.
- Tillage can amplify the negative effects of sugar by exposing more carbon and reducing soil organic matter
- Proof-of-concept work shows sugar triggered a net nitrogen immobilization in a carbon heavy environment
- Proof-of-concept work also suggests that when additional nitrogen is present, sugar additions may shift the system toward net mineralization rather than immobilization.
Work Cited:
Blagodatskaya, E., & Kuzyakov, Y. (2008). Mechanisms of real and apparent priming effects. Biology and Fertility of Soils, 45, 115–131.
Brady, N. C., and R. R. Weil. “The Nature and Properties of Soils, 15th Edn (eBook).” (2017).
Cotrufo, M. F., Wallenstein, M. D., Boot, C. M., Denef, K., & Paul, E. (2013). The Microbial Efficiency-Matrix Stabilization (MEMS) framework. Global Change Biology, 19, 988–995.
Cotrufo, M. F., Soong, J. L., Horton, A. J., Campbell, E. E., Haddix, M. L., Wall, D. H., & Parton, W. J. (2015). Formation of soil organic matter via biochemical and physical pathways of litter mass loss. Nature Geoscience, 8(10), 776–779.
Dendooven, L., Verhulst, N., Luna-Guido, M., & Ceballos-Ramírez, J. M. (2006). Dynamics of inorganic nitrogen in nitrate- and glucose-amended alkaline–saline soil. Plant and Soil, 283(1–2), 321–333.
Devine, M. D., & Hall, L. M. (1990). Implications of sucrose transport mechanisms for the translocation of herbicides. Weed Science, 38(3), 299–304.
Kuzyakov, Y., & Blagodatskaya, E. (2015). Microbial hotspots and hot moments in soil: Concept & review. Soil Biology and Biochemistry, 83, 184–199.
Recous, S., Mary, B., & Faurie, G. (1990). Microbial immobilization of ammonium and nitrate in cultivated soils. Soil Biology and Biochemistry, 22, 913–922.
Mechanics of Soil Fertility: Understanding Humic and Fulvic Acids
Brian Arnall, Oklahoma State University, Precision Nutrient Management Extension Specialist
Oliver Li, Oklahoma State University, Soil Chemistry
Interest in humic and fulvic acid products has increased substantially in agricultural production systems during the past two decades. These materials are frequently promoted as tools for improving soil biology, increasing nutrient availability, enhancing fertilizer efficiency, and stimulating plant growth. Because humic substances are known to be important components of soil organic matter, it is reasonable to ask whether adding humic or fulvic products to soil can meaningfully influence soil fertility.
As with many soil fertility questions, the answer depends on understanding two key factors: the mechanism involved and the magnitude of that mechanism relative to the soil system. Soil processes operate within large natural pools of organic matter, nutrients, and microbial activity. Therefore, evaluating the potential effects of humic products requires examining both how these compounds function chemically and biologically and how their application rates compare with the soils organic matter.
What Are Humic and Fulvic Acids?
Humic substances are heterogeneous organic compounds formed during the decomposition and transformation of plant and microbial residues. Historically, soil scientists have divided these materials into three operational fractions based on their solubility behavior: humic acid, fulvic acid, and humin (Stevenson, 1994; Tan, 2014). Humic acids are relatively large molecules that are insoluble under acidic conditions but dissolve in alkaline solutions. Fulvic acids are smaller molecules that remain soluble across the entire pH range, which allows them to move more freely in soil solution.
Both humic and fulvic acids contain numerous functional groups, particularly carboxyl and phenolic groups, which carry negative charge. These functional groups allow humic substances to interact with metal ions and nutrient cations and contribute to several important soil properties, including cation exchange capacity, buffering capacity, and metal complexation (Stevenson, 1994; Lehmann and Kleber, 2015). Because these materials originate from decomposed organic residues, they represent one portion of the complex mixture that collectively makes up soil organic matter. The distribution of the soil organic matter fractions varies among soil types and land uses, but fulvic acids and humic acids are each typically estimated to comprise approximately 10–35% of total soil organic matter (Guimarães et al., 2013).
Nutrient Retention and the Role of Cation Exchange
One of the most commonly cited mechanisms associated with humic substances is their ability to retain nutrients through cation exchange. The negatively charged functional groups present on humic molecules attract positively charged ions in soil solution. Through this electrostatic attraction, humic materials can retain several plant nutrients, including ammonium, potassium, calcium, magnesium, and certain micronutrients such as zinc and copper (Stevenson, 1994; Tan, 2014). This mechanism functions in the same manner as cation exchange on clay minerals. Of course, negatively charged surfaces do not retain negatively charged ions. As a result, nutrients such as nitrate are not held by humic substances and remain mobile in soil solution.
Laboratory measurements indicate that humic materials may possess relatively high cation exchange capacity on a mass basis. Reported values commonly range from approximately 300 to 600 cmolc kg⁻¹ depending on the source material and extraction method (Stevenson, 1994; Tan, 2014). These values demonstrate that humic substances can retain a large amount of cationic nutrients. A question that can be posed, however, is how this capacity compares with the nutrient retention already provided by soil organic matter.
Understanding the magnitude of humic additions requires comparing product application rates with the organic matter already present in soil. Calculations based on typical cation exchange values suggest that one pound of humic material with a CEC of 300–600 cmolc kg⁻¹ could theoretically retain approximately 0.04 to 0.08 pounds of ammonium-nitrogen. When viewed in isolation this number may appear meaningful. However, agricultural soils already contain large quantities of organic matter. An acre furrow slice, representing approximately the upper six inches of soil, weighs roughly two million pounds. Soil containing one percent organic matter therefore contains about 20,000 pounds of organic material per acre (Brady and Weil, 2016). Humified organic matter typically has cation exchange capacities ranging between 150 and 300 cmolc kg⁻¹ (Stevenson, 1994), meaning that the exchange capacity associated with native soil organic matter is already substantial. To put this into perspective, one pound of humic material can retain roughly 0.04 to 0.08 pounds of cation charge. Ammonium and potassium carry a single positive charge, while calcium carries two, meaning two ammoniums can be held for every two calcium. To provide contrast to the application of a humic substance, increasing soil organic matter by just 0.1% equivalent to about 2,000 pounds of additional organic material per acre can provide the capacity to retain approximately 40 to 80 pounds of cation charge or 40 to 80 pounds of ammonium.
The key point is not that humic materials cannot retain nutrients. They clearly can. Rather, the scale of material already present in soil is extremely large compared with the few ounces or pounds of humic products typically applied in agricultural systems. Consequently, the nutrient retention capacity associated with soil organic matter overwhelmingly dominates the soil system.
Micronutrient Complexation
Humic and fulvic substances are also known to interact with micronutrients through metal complexation reactions (also known as ‘chelation’). Carboxyl and phenolic functional groups can coordinate with metal ions such as iron, zinc, copper, and manganese to form organic complexes (Stevenson, 1994; Tan, 2014). These complexes can influence micronutrient mobility and availability in soils.
Fulvic acids are particularly effective at forming soluble complexes because they remain dissolved across the full range of soil pH. In some cases, these complexes may increase micronutrient mobility and transport within the soil solution. This mechanism has been well documented in soil chemistry research and may explain some responses observed in systems where micronutrient availability is limited.
Effects on Plant Physiology
In addition to soil chemical interactions, humic substances may influence plant growth through physiological mechanisms occurring in the rhizosphere. Several studies have shown that humic substances can stimulate root development, including increases in root elongation, lateral root formation, and root hair production (Nardi et al., 2002; Canellas and Olivares, 2014).
Research suggests that these responses may involve interactions with plant hormonal pathways and membrane transport processes. Humic substances have been shown to activate plasma membrane H⁺-ATPase enzymes, which are involved in proton pumping and nutrient uptake across root membranes (Canellas et al., 2002; Trevisan et al., 2010). Activation of these transport systems can enhance nutrient absorption and influence root architecture.
These physiological effects appear to occur primarily at the root–soil interface, where dissolved organic molecules interact directly with plant tissues. As a result, the responses observed in plant growth experiments are often attributed to rhizosphere signaling processes rather than large changes in bulk soil fertility.
Microbial Responses to Humic and Fulvic Compounds
Soil microorganisms respond strongly to carbon availability, and different carbon sources can produce very different microbial responses. Simple carbohydrates such as glucose and sucrose are readily metabolized by soil microbes and therefore produce rapid increases in microbial respiration and biomass. Humic substances, in contrast, consist of chemically complex and partially oxidized organic compounds that decompose much more slowly (Lehmann and Kleber, 2015).
Experimental studies comparing carbon sources consistently show that microbial respiration increases dramatically when simple sugars are added to soil, whereas humic substances produce smaller responses (Blagodatskaya and Kuzyakov, 2008). This difference reflects the relative degradability of these compounds as microbial energy sources.
Carbon Inputs from Humic Products Compared with Natural Soil Carbon
Soil microbial activity is largely driven by carbon supplied from plants through root exudation, residue decomposition, and organic matter turnover. The carbon pools already present in soil are therefore important for understanding the potential influence of humic product additions. A soil containing one percent organic matter holds approximately 11,600 pounds of carbon per acre (Brady and Weil, 2016).
Research on plant–soil carbon cycling indicates that living roots release significant quantities of organic carbon into soil each growing season through root exudation and rhizodeposition (Kuzyakov and Domanski, 2000). These plant-derived carbon inputs commonly amount to hundreds of pounds of carbon per acre and serve as a major energy source for soil microbial communities. Viewed in this context, humic product applications represent extremely small additions to the soil carbon pool. Consequently, microbial stimulation in agricultural soils is dominated by carbon inputs from plant residues and root exudates rather than by small additions of humic materials.
Building Organic Matter in the Central Plains
Increasing soil OM in the central Great Plains is achievable, but the magnitude of change is governed primarily by carbon inputs and water availability rather than any single management practice. Systems that combine no-till, increased residue return, diversified crop rotations, and where feasible cover crops or manure inputs are the most effective because they simultaneously increase carbon inputs and reduce decomposition losses (Lyon et al., 2007; Mikha et al., 2013; Nielsen et al., 2016). In semi-arid systems, realistic rates of OM increase are modest: over a 5-year period, changes are often small, approximately +0.05 to 0.1% OM, but significant in relation to the system which is often at total OM levels between 0.7 and 1.25 prior to establishment of conservation practices. The increase is confined to the top inch of the soil surface (Mikha et al., 2013; Saha et al., 2024). Mechanistically, these gains occur through greater residue and root-derived carbon inputs, reduced soil disturbance which slows microbial oxidation, and improved aggregation that physically protects organic matter from decomposition (Six et al., 2002; Lehmann and Kleber, 2015). However, as emphasized throughout this discussion, the scale of change is small relative to the large existing organic matter pool, and meaningful increases require long-term, system-level management focused on maximizing biomass production rather than relying on small external carbon additions such as commercial products.
Take-Home Points
- Humic and fulvic acids can retain cations, chelate micronutrients, and influence plant and microbial processes.
- Typical application rates are small relative to existing soil organic matter, so whole-soil impacts are limited.
- Most observed effects are localized in the rhizosphere, not broad changes in soil fertility.
- Evaluating both mechanism and scale is key to understanding their role in nutrient management.
References
Blagodatskaya, E., & Kuzyakov, Y. (2008). Mechanisms of real and apparent priming effects and their dependence on soil microbial biomass and community structure. Biology and Fertility of Soils, 45(2), 115–131.
Brady, N. C., & Weil, R. R. (2016). The nature and properties of soils (15th ed.). Pearson.
Canellas, L. P., Olivares, F. L., Okorokova-Façanha, A. L., & Façanha, A. R. (2002). Humic acids isolated from earthworm compost enhance root elongation and lateral root emergence in maize. Plant Physiology, 130(4), 1951–1957.
Canellas, L. P., & Olivares, F. L. (2014). Physiological responses to humic substances as plant growth promoters. Chemical and Biological Technologies in Agriculture, 1, 3.
Guimarães, D. V., Gonzaga, M. I. S., Silva, T. O., Silva, T. L., Dias, N. S., & Matias, M. I. S. (2013). Soil organic matter pools and carbon fractions in soil under different land uses. Soil and Tillage Research, 126, 177–182.
Kuzyakov, Y., & Domanski, G. (2000). Carbon input by plants into the soil: Review. Journal of Plant Nutrition and Soil Science, 163(4), 421–431.
Lehmann, J., & Kleber, M. (2015). The contentious nature of soil organic matter. Nature, 528(7580), 60–68.
Lovley, D. R., Coates, J. D., Blunt-Harris, E. L., Phillips, E. J. P., & Woodward, J. C. (1996). Humic substances as electron acceptors for microbial respiration. Nature, 382, 445–448.
Lyon, D. J., Stroup, W. W., & Brown, R. E. (2007). Crop production and soil water storage in long-term winter wheat–fallow tillage experiments. Soil and Tillage Research, 94(2), 387–397.
Mikha, M. M., Vigil, M. F., Benjamin, J. G., & Sauer, T. J. (2013). Cropping system influences on soil carbon and nitrogen stocks in the Central Great Plains. Soil Science Society of America Journal, 77(2), 702–710.
Nardi, S., Pizzeghello, D., Muscolo, A., & Vianello, A. (2002). Physiological effects of humic substances on higher plants. Soil Biology and Biochemistry, 34(11), 1527–1536.
Nielsen, D. C., Lyon, D. J., Hergert, G. W., Higgins, R. K., Calderón, F. J., & Vigil, M. F. (2016). Cover crop mixtures do not use water differently than single-species plantings. Agronomy Journal, 108(3), 1025–1038.
Saha, D., Kukal, S. S., & Bawa, S. S. (2024). Long-term impacts of conservation agriculture practices on soil organic carbon and aggregation. Soil Science Society of America Journal.
Six, J., Conant, R. T., Paul, E. A., & Paustian, K. (2002). Stabilization mechanisms of soil organic matter: Implications for C saturation of soils. Plant and Soil, 241(2), 155–176.
Stevenson, F. J. (1994). Humus chemistry: Genesis, composition, reactions (2nd ed.). Wiley.
Tan, K. H. (2014). Humic matter in soil and the environment. CRC Press.
Trevisan, S., Francioso, O., Quaggiotti, S., & Nardi, S. (2010). Humic substances biological activity at the plant–soil interface. Plant Signaling & Behavior, 5(6), 635–643.
For any questions or commments please feel free to reach out to Brian Anrall, b.arnall@okstate.edu
How do you handle your soil sample….
It’s that time of year where wheat producers are in all stages of prep across the state, the graze out folks are wondering when the rains will hit to get the dusted in wheat up and the grain only folks are prepping grain drills or dealing with summer crop harvest. While this blog is focused on the wheat producers it should be acknowledged as a work of caution for anyone who pulls soil samples in environments where the temps during sampling can be above the mid 80’s.
Following some fantastic work by our Wild Cat Soil Fertility Counterparts (see blog) we dug into the same question of how does soil sample handling post collection impact the results. Dr. Ruiz-Diaz results on the impact of storage on nitrogen values was not surprising. But we wanted to go the next step and add in sample bag type.
Our Project
We collected a soil ground and homogenized. Placed in three types of bags Ziploc, Commercial (resin lined paper that is closed) and SWFAL bag (breathable material). Placed samples in a mini-van (Field) and brought a group onto campus (Office). With both sets of samples we had temperature loggers collecting hourly data. Every three days we collected four samples from each location determined moisture content and ran a full spectrum soil analysis including CO2 burst. The soil we used had a OM of 1.1% and soil pH of 6.1
Now let’s back up. Why was the KSU data not surprising. Well for those certified and honorary nitrogen ninjas we understand that the N-cycle processes of mineralization, immobilization, and nitrification are biological reactions which are significantly impacted by soil moisture and soil temperature. But the short and quick version is that the microbes that convert organic N to mineral N (NH4) and convert NH4 to NO3 like warm conditions with good soil moisture. So, when you pull soil samples, hopefully there is some moisture in the soil and then you place it in a bag and seal it. This creates an effective greenhouse environment where moisture cannot escape and is the perfect place for microbes to microbial things. Therefore, you expect the organic cycle and nitrogen cycle to move and move quickly through the processes of mineralization, ammonification, and nitrification.
Back to the data.
First, we have the average daily temperatures from the sitting vehicle and the lab. On days 2-5 the average temp of the van was over 100°, after that the temps dropped. One thing to remember is the temp of the van is an average of a wide-ranging highs and lows between morning and midday while the office temperature being stable throughout the entire day.
What is the most interesting and revealing data may be the soil moisture of the samples. Unfortunately, day 3 samples went straight to the oven, but you can see by day 6 the SFWAL bags (yellow line) had dried significantly while it wasn’t until day 27 did the Commercial bags in the van (Black line w/ triangle) show signs of drying. It’s also important to note that while there are two lines (Field and Office) for the Commercial bags and Ziploc bags, but only one for the SWFAL bags. That is because for all variables measured there was never a significant difference between the SWFAL Field value and the SWFAL Office value.
We are going to start off with the variable that changed the most, NO3-N. This is also the form of N that is measured in most soil test. With a starting point of 3.4 lbs. N ac-1 (based on a six inch soil sample depth) by day 6 the Field samples had jumped to 15.6 lbs. The office samples had much slower increase in NO3 with it taking 9 days to reach 10 lbs. You can also see that the error bars start showing up in the samples after about two weeks. The bars show the range in the sample results of a treatment. The interesting thing we saw was that the location within the van was significant. We set each replication of samples in a different spot in the van, the front seat, middle seat, and rear. The thermometers set with each replications showed that each area of the van had slightly differing temperature, which in turn affected the samples.
Ammonium (NH4-N), which is only reported by a few labs is also impacted by the storage location and bag type, however not to the sample level as NO3-N. Effectively the SWFAL bags and all samples taken to the office maintained consistent NH4 levels. The Com bag and Ziplocs left in the van however saw a significant increase in NH4 over time, basically a 2 lb per day increase.
Of the other measured parameters such as potassium, sulfur, and CO2 burst data were effected. These will be discussed more in depth in a peer reviewed publication and factsheet.
Take Home’s from the Work
While the majority of the nutrients were relatively un affected by the bag type or storage those that are more heavily influenced by organic matter and biological activity where, this includes many of the soil health parameters. The use of SWFAL soil bags resulted in data that was not impacted by storage or time. While most clients of the OSU lab use these bags, more than one samples been brought into the county via a sealed Ziploc baggie. However the majority of the soil samples collected are placed into the resin lined paper bags and sealed. This data set is also important to those who use laboratories that run the Haney test which includes measures of NO3, NH4, Organic N, CO2 respiration and H2O extracted Organic C. It is critical that when using these type of bags that the samples be brought into climate controlled facilities as soon as possible. Its good to remember when looking at the data that the soil we used had a OM of 1.1%, initial NO3 of 3.4 lbs. and NH4 of 14 lbs. A soil with a higher OM% could have even greater change. Another viable is if the soil sample depth is greater than 6”. If your sample depth is 8” then the value is 33% higher.
Questions or comments feel free to reach out via email or social media.
Brian Arnall, Precision Nutrient Management b.arnall@okstate.edu
Soil sample handling practices can affect soil nitrate test accuracy
From Guest Authors,
Bryan Rutter, PhD student and Soil Testing Lab Manager, Kansas State University
Dr. Dorivar Ruiz Diaz, Soil Fertility Specialist, Kansas State University
The accuracy of a soil test is limited, in part, by the quality of the tested sample. For this reason, strong emphasis is placed on ensuring representative samples are collected in the field. However, these samples must also be handled properly after they have been collected.
Soils are home to a diverse population of microorganisms, many of which help decompose crop residue and cycle nutrients in soils. This nutrient cycling is crucial for crop production, but can skew soil test results if it continues in soil samples after they have been collected.
Microorganisms drive the soil nitrogen cycle
The nitrogen (N) cycle in soils is particularly complex and is strongly influenced by microbial activity and, therefore, temperature and soil moisture conditions. Bacteria and fungi consume organic material and use carbon as an energy source. During this process, N contained in the organic matter undergoes several transformations, ultimately converting it to ammonia. This conversion from organic-N to inorganic-N (NH4+, ammonium) is called “mineralization.” Plants can then take up the ammonium (NH4+), or converted to nitrate (NO3–) by certain bacteria through a process known as “nitrification”.
The microbial activity requires moisture and heat, and the processes described above happen more quickly in warm, wet soils than in cold, dry soils. Microbial activity does not stop just because a sample has been collected and put in a bag. This activity continues as long as the environmental conditions are favorable. As a result, soil tests for plant-available N have the potential to change substantially if samples are not handled properly. This is an important consideration for growers because these soil test results are used to determine the profile-N credit and, ultimately, adjust N fertilizer recommendations.
Research study on soil sample storage
A recent study at the K-State Soil Testing Lab illustrates what can happen if sample submission is delayed. For this study, soil was collected from the Agronomy North Farm (Manhattan, KS) and thoroughly mixed/sieved to homogenize the material. This soil was then placed into sample bags, which were randomly assigned to different combinations of storage temperature and duration. One set of samples was kept in a refrigerator while the other set was kept in a cargo box in a truck bed. To monitor changes in soil test levels over time, three sample bags were removed from the refrigerator and truck box every two days (48 hours) and tested in the lab.
Figure 1. Change in soil test nitrogen parameters over a 14-day storage period. Samples stored in an unrefrigerated cargo box are indicated by purple points. Samples stored in a refrigerator (38F) are indicated by grey points. Graphs by Bryan Rutter, K-State Research and Extension.
Take home points from the K-State Soil Testing Lab study:
- Mineralization and nitrification led to more than a 3x increase in soil test nitrate in the undried and unrefrigerated “Truck Cargo Box” samples (purple points in Figure 1).
- Soil test nitrogen did not change substantially in refrigerated samples.
- Profile-N credits calculated from soil test N results were nearly 100 lbs of N/acre higher for the unrefrigerated samples (Figure 2).
- Improper handling and storage of soil samples can dramatically reduce soil test accuracy and may lead to under or overfertilizing crops.
K-State Soil Testing Lab Recommendations
- Submit soil samples to the lab as soon as possible, ideally on the same day they were collected.
- If same-day submission is not possible, samples should be air-dried or placed in a refrigerator set at 40 degrees F or less.
Please see the accompanying article “The challenge of collecting a representative soil sample” for guidance on field soil sampling practices.
For detailed instructions on submitting soil samples to the K-State Soil Testing Lab, please see the accompanying article “Fall soil sampling: Sample collection and submission to K-State Soil Testing Lab”.
For detailed information on how N credits are calculated please see the MF-2586 fact sheet: “Soil Test Interpretations and Fertilizers Recommendations”.
Bryan Rutter, PhD student and Soil Testing Lab Manager
rutter@ksu.edu
Dorivar Ruiz Diaz, Soil Fertility Specialist
ruizdiaz@ksu.edu
The original article can be found on the KSU Agronomy E-update site
https://eupdate.agronomy.ksu.edu/article_new/soil-sample-handling-practices-can-affect-soil-nitrate-test-accuracy-511-4
Phosphorus decisions, Is it worth cutting P?
With the current conditions and input cost many wheat producers are considering cutting back on inputs. I can’t disagree with the plan, but I would caution against what you cut. If you have read any of my past blogs, or seen me speak, you should know I’m all for cutting back on pre-plant nitrogen (N). Based on some recent trials I would not argue cutting the potassium (K) side, but phosphorus (P) that’s another story that we will walk through in this blog.
First and foremost, soil testing is the key to P management. If your soil test is below the critical threshold for the test you use, 32.5 for Mehlich 3 (M3P), then you need to add phos. We have enough work that shows current recommendations work for P in wheat. Reeds paper Evaluation of incorporated phosphorus fertilizer recommendations on no-till managed winter wheat Link to Paper goes over soil test recommendations in no-till and the recent double crop soybean project Double Crop P and K Blog highlights the importance of P fertility, regardless of yield level. Also if your soil test is below a 5.5 and you haven’t limed (Liming is the best solution, Band-aids not so cheap Blog ), then the next best option is adding additional P to alleviate the aluminum toxicity Band-aids for low pH Blog. In-short if the fields soil test P and or pH is below optimum you should not forgo P application.
But the primary reason I am writing this blog is for those looking at fields with composite soil test that is right around the critical thresholds, and they are trying to make the call on to apply P or not to apply P. Even on fields with soil test values in the good level, I am usually in favor of banding in-furrow fertilizer wheat, but not because of the same reasons I am for corn. With corn you are planting in cool soils and the availability of nutrients like P is lower in cool wet soils. For wheat cold soil isn’t the concern until we reach the end of the planting window. It will serve as a bit of a “pop-up” as the crop comes out of dormancy in the spring. I have also seen little to no value of N applied in furrow. I see same response to DAP (18-46-0), MAP (11-52-0), and TSP (0-46-0) when all applied at same rate of P. Meaning it was the P not N making the difference.
For me the reason I still recommend getting a little phosphate out even when the soil test comes back is that the great majority of fields have a large range of variability. Looking at a set of 650 grid sampled fields across Oklahoma and Kansas it showed on average soil pH 6.0 and M3P was 34 ppm. Both pH and P are at adequate/optimum levels. However, the average is usually somewhere between the low and high point and in this data set and the range of soil pH was 1.8 units and the range in M3P was 67 ppm. That meant on average of the 648 field with pH values the average difference between low pH and high pH was 1.8 units and the difference between low P and high P was 64 ppm.
The field below is from Kingfisher county and was sampled at a resolution of 10 acres per sample. This is a fairly course resolution for grid sampling but provides a great view of how variable our soils can be. The field average pH is 5.3, which is below optimum but our aluminum tolerant wheats would be able to handle fairly well. For the P the average is 22 ppm which needs about 18 lbs of P2O5 to max yields. If the farmer applied a flat rate of 20 lbs there would be significant forage loss on about 65% of the field, for grain only about 45% of the field due to underapplication of P. Note that low P and low pH are not correlated well, meaning the areas low in pH are not always low in P.
Banding P makes it more efficient because it slows the rate of tie. However, we have plenty data that says broadcast applied P is still a great option, even after planting. So what are my take homes from this blog?
First: If you are grazing wheat get down 40-50 lbs of N pre. But I have plenty of data the pre-plant N on grain only wheat is not needed. I have the same amount of data that shows the only value of in-furrow N for grain only is that it forces you to plant more seeds, because it just lowers stand.
Second: When it comes to wheat pay attention to Phosphorus and soil pH. Even our acid tolerant wheats preform better in neutral soil pHs, especially forage wise.
Third: A composite soil sample is an AVERAGE of the field. If your average is right at the ok level (pH of 5.6ish and M3P of 30 ppm), then half of your field is below optimum and will benefit from P.
Fourth: If you can band P great, but if you cant broadcast is still a viable option. Do Not Skip P when soil test says there is a need.
Questions or comments please feel free to reach out.
Brian Arnall b.arnall@okstate.edu
Soil Acidity and Cotton Production
Raedan Sharry, Precision Nutrient Management Ph.D. student
Brian Arnall, Precision Nutrient Management Extension Specialist
Cotton production in Oklahoma has expanded the past decade into areas which other production systems such as continuous wheat may have traditionally dominated. Fields that have been managed for continuous wheat production may have become acidic in response to management practices, such as the use of ammoniacal nitrogen fertilizers. In response to the acidification of these soils it may be important to recognize and understand the potential impacts of soil acidification on cotton production.
To better document the impact of soil pH on cotton lint yield and quality a study was conducted over the 2019 and 2020 growing seasons at two locations: Stillwater, OK (EFAW farm) and Perkins, OK. Soil pH in these experiments were adjusted to a depth of approximately 6 inches using aluminum sulfate (acidifying) and hydrated lime (alkalinizing). All three locations were planted to two varieties; NexGen 3930 and Deltapine 1612 at a rate of approximately 35,000 seeds per acre, into plots with soils ranging in pH from 4.0 to 8.0.
In season measurements taken included stand count, plant height, node count and NDVI (normalized difference vegetative index. All four in season measurements demonstrated a significant critical threshold in which soil pH negatively impacted crop performance. Stand was significantly decreased at a soil pH of 5.3 or lower. This trend is displayed in Figure 1. Plant height and node count depicted in figures 2 and 3 respectively were both significantly decreased when soil pH dropped below 5.3 and 4.9, while NDVI began to deteriorate around a pH of 5.1 (Figure 4). Response to soil pH level was also visually observable as shown by Figure 5 from the Perkins 2019 location.
Yield levels across this experiment ranged from 0 to 1284 lbs. of lint per acre. In this work yield is reported as relative yield. To calculate relative yield the yield of each plot is normalized to the average of the three highest yields for that site. This method of reporting yield response allows this work to be applied across a range of yield environments.
When all sites and cultivars were combined relative yield reached a plateau at approximately 73% of yield with a critical threshold observed at a pH of 5.4. Below this critical threshold yield decreased at a rate of 37% per point of pH decline. This equates to 15% yield loss at a pH of 5.0, 33% yield loss at a pH of 4.5, and 52% yield loss at a pH of 4.5. This relationship is depicted in Figure 6. It is important to note that yield was 0 when pH was 4.3 in two plots. This represents the possibility for total crop failure when planting into very acidic conditions.
Differences in relative yield between cultivars were insignificant. However, further investigation may produce a significant difference. Critical threshold for the DP 1612 and NG 3930 cultivars were 6.1 and 5.2 respectively. This suggests that there may be value in further examining the influence of genetics on cotton response to soil acidity.
SUMMARY
The influence of soil pH level on cotton productivity is confirmed by this study. All three sites evaluated provided a strong correlation between soil pH and lint yield, as well as the in-season growth parameters measured. While this study is likely to be expanded to another location to provide a more robust evaluation of the potential impact of soil acidity on cotton, the current dataset provides ample evidence to conclude that soil acidity is likely to be detrimental to cotton production in the southern plains. Soil pH levels below 5.5 appear to provide the greatest opportunity for yield loss as depicted above. Lint quality measurements taken in the study (micronaire, length, uniformity, and strength) showed no consistent trends in the relationship between quality parameters and soil pH suggesting that soil acidity had a limited influence on lint quality characteristics. Cotton response to soil pH is likely to be influenced by the environment of a specific location and growing season. This underlines the importance of understanding the soil properties that negatively impact productivity, such as the presence of toxic forms of aluminum or manganese. It is also important to highlight the possibility of acidic conditions significantly affecting the ability of the crop to access important nutrients such as phosphorus. Under acidic conditions cotton productivity is likely to be significantly decreased unless soils are neutralized using a soil amendment such as lime.
Project Supported by the
Oklahoma Cotton Council
Cotton Incorporated
Oklahoma Fertilizer Checkoff Program
The challenge of collecting a representative soil sample
Guest Author, Dorivar Ruiz Diaz, Nutrient Management Specialist Kansas State University
At first glance, soil sampling would seem to be a relatively easy task. However, when you consider the variability that likely exists within a field because of inherent soil formation factors and past production practices, the collection of a representative soil sample becomes more of a challenge.
Before heading to the field to take the sample, be sure to have your objective clearly in mind. For instance, if all you want to learn is the average fertility level of a field to make a uniform maintenance application of P or K, then the sampling approach would be different than sampling for pH when establishing a new alfalfa seeding or sampling to develop a variable rate P application map.
In some cases, sampling procedures are predetermined and simply must be followed. For example, soil tests may be required for compliance with a nutrient management plan or environmental regulations associated with confined animal feeding operations. Sampling procedures for regulatory compliance are set by the regulatory agency and their sampling instructions must be followed exactly. Likewise, when collecting grid samples to use with a spatial statistics package for drawing nutrient maps, sampling procedures specific to that program should be followed.
Figure 1. The level of accuracy of the results of a soil test will depend, in part, on how many subsamples were taken to create the composite sample. In general, a composite sample should consist of 15 or more subsamples. For better accuracy, 20-30 cores, or subsamples, should be taken and combined into a representative sample. F
Regardless of the sampling objectives or requirements, some sampling practices should be followed:
- A soil sample should be a composite of many cores to minimize the effects of soil variability. Take a minimum of 12 to 15 cores from a relatively small area (two to four acres). Taking 20-30 cores will provide results that are more accurate. Take a greater number of cores on larger fields than smaller fields, but not necessarily in direct proportion to the greater acreage. A single core is not an acceptable sample.
- Use a consistent sampling depth for all cores because pH, organic matter, and nutrient levels often change with depth. Match sampling depth to sampling objectives. K-State recommendations call for a sampling depth of two feet for the mobile nutrients – nitrogen, sulfur, and chloride. A six-inch depth is suggested for routine tests of pH, organic matter, phosphorus (P), potassium (K), and zinc (Zn) (Figure 2).
- When sampling a specific area, a zigzag pattern across the field is better than following planting/tillage pattern to minimize any past non-uniform fertilizer application/tillage effects. With a GPS system available, recording of core locations is possible. This allows future samples to be taken from the same locations in the field.
- When sampling grid points for making variable rate nutrient application maps, collecting cores in a 5-10 foot radius around the center point of the grid is preferred for many spatial statistical software packages.
- Avoid unusual spots obvious by plant growth and/or visual soil color/texture differences. If information on these unusual areas is desired, collect a separate composite sample from these spots.
- If banded fertilizer has been used on the previous crop (such as strip tillage), then it is suggested that the number of cores taken should be increased to minimize the effect of an individual core on the composite sample results, and to obtain a better estimate of the average fertility for the field.
- For permanent sod or long-term no-till fields where nitrogen fertilizer has been broadcast on the surface, a three- or four-inch sampling depth would be advisable to monitor surface soil pH
Figure 2. Consistency is sampling depth is particularly important for immobile nutrients like P. Stratification of nutrients and pH can be accentuated under reduced tillage.
Soil test results for organic matter, pH, and non-mobile nutrients (P, K, and Zn) change relatively slowly over time, making it possible to monitor changes if soil samples are collected from the same field following the same sampling procedures. However, there can be some seasonal variability and previous crop effects. Therefore, soil samples should be collected at the same time of year and after the same crop.
Soil test results for organic matter, pH, and non-mobile nutrients (P, K, and Zn) change relatively slowly over time, making it possible to monitor changes if soil samples are collected from the same field following the same sampling procedures. However, there can be some seasonal variability and previous crop effects. Therefore, soil samples should be collected at the same time of year and after the same crop.
Soil testing should be the first step for a good nutrient management program, but it all starts with the proper sample collection procedure. After harvest in the fall is good time for soil sampling for most limiting nutrients in Kansas.
For instructions on submitting soil samples to the K-State Soil Testing Lab, please see the accompanying article “Fall soil sampling: Sample collection and submission to K-State Soil Testing Lab” found in this eUpdate issue.
For any questions Contact.
Dorivar Ruiz Diaz, Nutrient Management Specialist
ruizdiaz@ksu.edu
Soil sampling for pH and liming in continuous no-till fields
Quest Author, Dorivar Ruiz Diaz, Nutrient Management Specialist Kansas State University
One question that commonly comes up with continuous no-till operations is: “How deep should I sample soils for pH?” Another common question is: “How should the lime be applied if the soil is acidic and the field needs lime?”.
Sampling depth in continuous no-till
Our standard recommendation for pH is to take one set of samples to a 0-6 inch depth. On continuous no-till fields where most or all of the nitrogen (N) is surface applied, we recommend taking a second sample to a 0-3-inch depth. We make the same recommendation for long-term pasture or grass hayfields, such as a bromegrass field that has been fertilized with urea annually for several years.
Nitrogen fertilizer is the primary driving force in lowering soil pH levels, so N application rates and methods must be considered when determining how deep to sample for pH. In no-till, the effects of N fertilizer on lowering pH are most pronounced in the area where the fertilizer is actually applied. In a tilled system, the applied N or acid produced through nitrification is mixed in through the action of tillage and distributed throughout the tilled area.
Where N sources such as urea or liquid UAN solutions are broadcast on the surface in no-till system, the pH effects of the acid formed by nitrification of the ammonium will be confined to the surface few inches of soil. Initially this may be just the top 1 to 2 inches but over time, and as N rates increase, the effect of acidity become more pronounced, and the pH drops at deeper depths (Figure 1). How deep and how quickly the acidity develops over time is primarily a function of N rate and soil CEC (cation exchange capacity), or buffering capacity.
Where anhydrous ammonia is applied, or liquid UAN banded with the strip-till below the surface, an acid zone will develop deeper in the soil. As with long-term surface applications, these bands will expand over time as more and more N fertilizer is placed in the same general area. The graphic below (Figure 1) illustrates the effect of repeated nitrogen and phosphorus application with strip-till in the same area in the row middle on a high CEC soil for more than 12 years.
Figure 1. Soil pH stratification after 25 years of no-till and surface nitrogen fertilizer application, and the effect of repeated fertilizer application with strip-till in the same area after 12 years.
Liming application methods in continuous no-till
Where do you place the lime in continuous no-till?
If you surface apply N, then surface apply the lime. That’s a simple but effective rule. But remember that surface-applied lime will likely only neutralize the acidity in the top 2-3 inches of soil. So if a producer hasn’t limed for 20 years of continuous no-till and has applied 100 to 150 pounds of N per year, there will probably be a 4-5 inch thick acid zone, and the bottom half of that zone may not be neutralized from surface-applied lime. So, if a producer is only able to neutralize the top 3 inches of a 5-inch deep surface zone of acid soil, would that suggest he needs to incorporate lime? Not really. Research has shown that as long as the surface is in an appropriate range and the remainder of the acid soil is above pH 5, crops will do fine.
Liming benefits crop production in large part by reducing toxic aluminum, supplying calcium and magnesium, and enhancing the activity of some herbicides. Aluminum toxicity doesn’t occur until the soil pH is normally below about 5.2 to 5.5 and KCl-extractable (free aluminum) levels are greater than 25 parts per million (ppm). At that pH the Al in soil solution begins to increase dramatically as pH declines further. Aluminum is toxic to plant roots, and at worse the roots would not grow well in the remaining acid zone.
This implies that the acid zones from ammonia or banded UAN are probably not a major problem. We have monitored ammonia bands in the row middles of long-term no-till for many years and while the pH dropped very low, we never saw any adverse impacts on the crop that would justify liming and using tillage to incorporate the lime. In fact, some nutrients such as zinc, manganese, and iron can become more available at low pH, which can be an advantage at times.
Yield enhancement is not the only concern with low-pH soils, however. Herbicide effectiveness must also be considered. The most commonly used soil-applied herbicide impacted by pH is atrazine. As pH goes down, activity and performance goes down. So in acidic soils, weed control may be impacted. We do see that happen in corn and sorghum production.
Liming products for no-till
When choosing a liming product, is there any value to using dolomitic lime (which contains a large percentage of magnesium in addition to calcium) over a purely calcium-based lime product?
Most Kansas soils have high magnesium content. So as long as we maintain a reasonable soil pH, there normally is enough magnesium present to supply the needs of a crop. Calcium content is normally significantly higher than magnesium, so calcium deficiency is very, very rare in Kansas. The soil pH would need to be below 4.5 before calcium deficiency would become an issue. Before calcium deficiency would occur, aluminum toxicity or manganese toxicity would be severely impacting crop growth. So producers really don’t have to worry about a deficiency of calcium or magnesium on most Kansas soils.
What about the use of pelletized lime as a pH management tool on no-till fields?
The idea has been around for a while to use pel-lime in low doses to neutralize the acidity created from nitrogen and prevent acid zones from developing. . Pel-lime is a very high-quality product, normally having 1800 to 2000 pounds of effective calcium carbonate (ECC) per ton, and can be blended with fertilizers such as MAP or DAP or potash easily. Therefore, if you apply enough product this can be an excellent source of lime. Lime can be from various sources and with different qualities. Consecutively, to ensure a standardized unit of soil-acidity neutralizing potential, we use units of ECC.
Summary
Applying N fertilizer to soil will cause the soil to become acidic over time. Placement of the applied N and the level of soil mixing done through tillage determine where the acid zones will develop. Make sure your soil testing program is focused on the area in the soil becoming acidic, and apply the lime accordingly.
For any questions Contact.
Dorivar Ruiz Diaz, Nutrient Management Specialist
ruizdiaz@ksu.edu
Precision Nutrient Management in Forage Systems
Published in Progressive Forage http://www.progressiveforage.com/ 9.1.2016
First, let’s agree the term “precision” is relative. Forage is a diverse system with an even more diverse set of management strategies. Regardless, every manager should be constantly striving to improve the precision in which nutrients are managed. The ultimate goal of any precision nutrient management tool should be this: producing the highest quality output (in this case forage) with the least amount of input – ultimately, optimizing efficiencies and maximizing profits. Within this readership there are those who are soil sampling at a 1-acre resolution and others who have likely not pulled a soil sample in the past decade. For both spectrums we can make improvements – let’s start basic and move forward.
A soil sample should the basis for all nutrient management decisions. Is soil testing a perfected science? No, far from it. However, there must be a starting point. A soil sample is that first bit of information we can start with and the basic data collection for precision ag to make improved management decisions. When fertilizer is applied without a recent soil sample, it is done based upon pure guesswork. How many other management decisions are made on a farm or ranch by a guess?
The composite soil sample is a great start, but it is just that – a start. While there are some soils that are very uniform most are extremely variable. In a survey of 178 fields in the southern Great Plains on average the soil pH was 6.12; phosphorus (Mehlich 3 phosphorus [M3P] and Bray 1 phosphorus [B1P]) was 28 ppm while soil test potassium averaged 196 ppm. So on the average the primary components of soil fertility were okay. However, on average the 178 fields had a range in soil pH of 1.8 units, M3P and B1P both had range of a 52 ppm and STK had a range of 180 ppm.
Table 1 shows the minimum and maximum soil test values for the 178 fields.
| Average | Range | Min | Max | |
| Soil pH | 6.12 | 1.77 | 5.23 | 7.01 |
| Phosphorus | 28 | 52 | 2 | 54 |
| Potassium | 197 | 180 | 107 | 287 |
| Sulfur | 15 | 24 | 3 | 27 |
| Organic matter | 1.9 | 1.2 | 1.3 | 2.5 |
This data helps support the concept that we should find ways to increase the resolution or decrease the number of acres represented by a single soil sample. Increasing soil sample resolution is typically done using one or two methods – zone or by grid.
Zone sampling
The basis of a zone sample is creating a smaller field. The biggest question with zones is how to draw the lines. There are dozens, if not hundreds, of possible methods, each having their own reasons and benefits. My basic recommendation is that before lines are drawn goals have to be established. For example, if phosphorus or soil pH management is important, the basis for the lines should be soil based. This could be based on soils map, soil texture, slope and on and on. If the target is improved nitrogen management, then the reason for drawing lines should be yield based. This could be based on yield maps, aerial images, historic knowledge or many soil parameters.
Why does it matter? Two reasons: First, across the broad spectrum of soils and environments two nutrients are hardly spatially correlated, which means the zone that is best at describing phosphorus variability does an extremely poor job describing potassium variability. Second, more theoretically the demand for nutrients are driven by different factors. Phosphorus (a soil immobile nutrient) fertilizer need is driven by the soil P concentration (look up Brays Sufficiency Concept). Many use yield as a parameter for phosphorus application, but this is not a plant need or even a yield maximizing practice. Fertilizing based on removal is done to prevent nutrient mining. However, nitrogen (a nutrient mobile in the soil) fertilizer need is based on yield and crop removal. Hence, the common Land Grant University N and sulfur recommendations are yield goal based.
Grid sampling
To be honest even the experts disagree on the hows, whys and ifs of grid sampling. I like data, therefore I naturally lean towards grid sampling if the field warrants it. For me, the biggest benefit of grid over zone sampling is that soils data from zone samples are biased to whatever parameter was set for the zone and therefore any resulting map for all nutrients must reflect the original zones. In a grid, each data point is independent therefore the maps of each nutrient can be independent, and (the science tells us) in most cases nutrients are independent of each other.
Ideally two pieces of information are available for determining whether a field is grid sampled or not. The first piece of information is a yield map from any previous crop. If yield is fairly uniform, I question the need for variable rate management, much less the expense of grid sampling. Regardless of the sampling method zone or grid, the discussion is moot if spatial variability does not exist across the field. However, many forage producers may not have access to this kind of data.
One of the most useful decision aid tools for grid sampling is the composite soil sample. The reason is simple statistics: A composite sample should be a representative average of the field. If the data is normally distributed, that means half of the field is above and half the field is below the sample average. So the optimum fields to grid are those in which an input falls at the point in which the benefit of applying is in question, because it suggests that approximately half the field needs the inputs while the other half likely does not. It is in this scenario that the return on investment can be greatest. As with pH, for example, fields with a very low value should have a flat broadcast application and should be sampled again at a later date. Fields with a composite pH well above 6.0 will unlikely have enough acres needing lime to warrant sending out an applicator.
Is grid sampling a lifelong activity? No. The initial activity of grid sampling will provide both an indicator of the variability level and overall needs of the field. From that point, decisions can be made and actions taken. Identify the greatest limiting factor in the field based on the samples, and focus on impacting change upon it. Zone sampling in subsequent years can be utilized to document change. When that issue is resolved, move to the next factor. It may require grid sampling again or using the original grid to develop new management zones. For instance, if the greatest issue first identified on the field is soil acidity then after the soil pH is neutralized the field should be grid sampled again. The reason is for this is that changing soil pH will influence many nutrients and the amount of change is not consistent but dependent upon many other factors.
In precision ag we tend to look at layers, yield, soil, etc. However, none of these tell the whole story independently. An area in a field may have moderate soil fertility and be under producing. Using the data collected the decision may be made to increase inputs; yet, the issue is a shallow restrictive layer limiting production. Therefore, the extra inputs will be of no benefit and could even further reduce production. It is at this point I like to bring out the importance of “getting dirty.” There is no technology that can take the place of “boots on the ground” agronomy.
For producers who have historically preformed intensive soil sampling there is still room for improvement. Soil testing and nutrient management is not an exact science; in fact, it was originally built for broad sweeping, statewide recommendations. As technology advances and inputs can be applied at sub-acre resolutions, all of the environment (weather, soil) by genotype inactions becomes more evident.
The next step in precision ag is to develop recommendations by upon site specific crop responses. This is where nutrient response strips can further improve nutrient use efficiencies and crop production. In Oklahoma, nitrogen-rich strips are applied across fields (grain and forage) to determine in-season nitrogen needs. Having a strip in the field with 50 to 100 extra units of N acts as a management tool which takes into account soil, environment and plant need. If the strip is visible the field or zone needs more N, if it is not visible then the crop is not deficient and at that point in the season does not need more N. Producers have taken this approach for N and adopted it for P and K with strips across the field with a zero and high rate of either nutrient. After a few seasons, responsive and non-responsive zones are developed and P and K applications are managed accordingly.
One misconception of precision ag is that the end result should be a field with uniform yield from one corner to the other. This is often not the case; in fact, in many cases the variability in production across the field can be increased. Theoretically, precision ag is applying inputs at the right rate in the right place. This means areas of the field which are yield limited due to underlying factors which cannot be managed have a reduction in inputs with no effect on yield. Other areas of the field have not been managed for maximum production therefore an increase inputs result in increasing yield widening the gap between the low and high yield levels.
Regardless of where a producer currently sets on the technology curve, there are potential ways to increase productivity and efficiency. There is nothing wrong with taking baby steps; it is often the simple things that lead to the greatest return.
Now may not be the time for Replacement
For phosphorus (P) and potassium (K) fertilizer management there are three primary schools of thought when it comes to rate recommendations. The three approaches are Build-up, Maintenance/Replacement, and Sufficiency. There is a time and place for each one of the methods however the current markets are making the decision for the 2016-16 winter wheat crop a very easy one. The OSU factsheet PSS-2266 goes in-depth on each of these methods. For the rest of the blog I will use P in the conversation but in many scenarios K should/could be treated the same.
Build-up is when soil test is below a significant amount of fertilizer, about 7.5 lbs P2O5 per 1 ppm increase, is added so that soil test values increase. This method is only suggested when grain price is high and fertilizer is relatively cheap. Given the market, this is a no go. The two most commonly used methods of recommendation are Replacement and Sufficiency. In the replacement approach if the soil is at or below optimum P2O5 rate it based upon replacing what the crop will remove. The sufficiency approach uses response curves to determine the rate of P that will maximize yield. These two values are typically quite different. A good way you boil the two down is that replacement feeds the soil and sufficiency feeds the plant.
Oklahoma State Universities Soil, Water, and Forage Analytical Lab (SWFAL) provides recommendations utilizing sufficiency only while many private labs and consultants use replacement or a blended approach. Some of this is due to region. Throughout the corn belt many lease agreement contain clauses that the soil test values should not decrease otherwise the renter pays for replacement after the lease is over. For the corn belt both corn and soybean can be expected to remove 80 to 100 pounds of P per year. Conversely the Oklahoma state average wheat crop removes 17 lbs P a year. In areas where wheat yields are below 40 bushel per acre (bpa) using the sufficiency approach for P recs can increase soil test P over time.
This conceptual soil test response curve is divided into categories that correspond with below opti-mum, optimum and above optimum soil test values. The critical level is the soil test level, below which a crop response to a nutrient application may be expected, and above which no crop response is expected. At very high soil test levels crop yield may decrease.
*Rutgers Cooperative Extension Service FS719
Back to subject of this blog, consultants, agronomist, and producers need to take a good look at the way P recs are being made this year. Profitability and staying in the black is the number 1, 2, and 3 topic being discussed right now. The simple fact is there is no economic benefit to apply rate above crop need, regardless of yield level. The figures above demonstrate both the yield response to fertilizer based upon soil test. At the point of Critical level crop response / increase in yield is zero. What should also be understood is that in the replacement approach P fertilizer is still added even when soil test is in Optimum level. This also referred to as maintenance, or maintaining the current level of fertility by replacing removal. If your program is a replacement program this is not a recommendation to drop it completely. Over a period of time of high removal soil test P levels can and will be drawn down. But one year or even two years of fertilizing 100 bpa wheat based on sufficiency will not drop soil test levels. On average soils contain between 400 and 6000 pounds of total phosphorus which in the soil in three over arching forms plant available, labile, and fixed. Plant available is well plant available and fixed is non plant available. The labile form is intermediate form of P. When P is labile it can be easily converted to plant available or fixed. When a plant takes up P the system will convert labile P into available P. When we apply P fertilizer the greatest majority of was is applied makes it to the labile and fixed forms in a relatively short period of time. For more in-depth information on P in the soil you can visit the SOIL 4234 Soil Fertility course and watch recorded lectures Fall 2015 10 26-30 Link .
How to tell if your P recs have a replacement factor, not including calling your agronomist. First replacement recs are based on yield goal, so if you change your yield goal your rate will change. The other and easier way is to compare your rates to the table below. Most of the regional Land Grant Universities have very similar sufficiency recs for wheat. Another aspect of the sufficiency approach is the percent sufficiency value itself. The sufficiency can provide one more layer in the decision making process for those who are near the critical or 100% level. Response and likelihood of response to P is not equal. At the lowest levels the likelihood of response is very high and the yield increase per unit of fertilizer is the greatest. As soil test values near critical (32.5 ppm or 65 STP) the likelihood of response and amount of yield increase due to fertilizer P decreases significantly. At a STP of 10 the crop will only produce 70% of its environmental potential if P is not added while at a STP of 40 the crop will make 90% of its potential. The combination of % sufficiency and yield goal can be used to determine economic value of added P.
*From Oklahoma State University Soil Test Interpretations. Fact Sheet PSS-2225
*Based on Mehlich 3
*PPM (parts per million) is used by most labs
*STP (soil test P) is a conversion used by some Universities. Equivalent to pounds per acre.
* for a 0-6” in soil sample PPM * 2 = STP.
This data is available from OSU in multiple forms from the Factsheet PSS-2225, the SWFAL website, Pete Sheets quick cards, and the Field Guide App.
This year with margins tight soil testing is more important than ever before. Knowing the likelihood of response and appropriate amount of fertilizer to apply will be critical maximizing the return on fertilizer invest while maximizing the quality and amount of grain we can produce. Visit with your consultant or agronomist to discuss what the best approach is for your operation. Lets ride this market out, get the most out of every input and come out of this down cycle strong.
Feel free to contact me with any questions you may have.
Brian
b.arnall@okstate.edu
Down and Dirty with NPK 