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OSUNPK Fertilizer Market Outlook — July 27, 2026
Produced by Chat GPT Work, reviewed by Brian Arnall, OSU Precision Nutrient Management.
Oklahoma fertilizer markets moved in two different directions during the latest reporting period. Nitrogen prices generally declined, while phosphate and potash remained nearly steady. That local stability does not mean the broader market is free of risk. International production costs, sulfur availability, trade flows and large import programs continue to influence the prices Oklahoma producers may encounter later in the season.
The USDA’s July 24 Oklahoma report placed urea at an average of $672.29 per ton, down $14.42 from the previous report. UAN 28 declined $16 to $472.60, UAN 32 fell $5 to $548.86, and anhydrous ammonia declined $29 to $947. One reported urea quote fell by approximately $300 per ton, while an NH₃ quote moved below $900 and a UAN 28 quote dropped below $400. Those individual movements contributed to unusually wide ranges and should not be treated as representative statewide prices without verification.
The USDA attributed part of the softer market to reduced fertilizer demand during triple-digit heat. Seasonal demand also appears less urgent following major spring and early-summer applications. However, the spread between low and high quotes remains substantial: $365 per ton for urea, $232 for UAN 28 and $225 for NH₃. Those differences make product basis, location and service charges particularly important when comparing offers. (USDA Oklahoma Production Cost Report)
International nitrogen conditions remain uncertain. Fertilizer-market analyst Josh Linville recently identified the resumption of vessel traffic through the Strait of Hormuz and the possibility of renewed Chinese urea exports as potentially bearish influences. Either development could improve global availability. However, fertilizer remains expensive relative to grain prices, and freight, energy or geopolitical disruptions could quickly change the cost calculation. (Josh Linville market commentary)
Oklahoma phosphate prices showed little immediate movement. DAP averaged $952.78, an increase of $2.22, while MAP held at $942.88. The local numbers appear quiet, but upstream phosphate conditions deserve attention. Linville has noted renewed strength in international phosphate values as limited sulfur availability and high production costs constrain output.
Sulfur matters because it is used to produce sulfuric acid, a critical input in manufacturing phosphoric acid and finished phosphate fertilizers such as DAP and MAP. Ammonia is another important production input. Consequently, higher sulfur or ammonia costs can support phosphate prices even when Oklahoma retail quotations have not yet responded. This is an upstream warning signal rather than a prediction that local prices must rise. (Josh Linville on phosphate and sulfur, World Bank fertilizer-market analysis)
Potash remained the quietest major nutrient in Oklahoma. The reported average increased only 75 cents to $491.55, with a range of $460 to $550. Global potash supply is geographically broader than the supply chain for several nitrogen and phosphate inputs. Expanded or recovering shipments from major producing regions may help moderate market pressure, although trade restrictions, transportation constraints and regional demand can still affect delivered costs.
International buying also remains important. India reported importing more than 3.2 million metric tons of urea and DAP during its first fiscal quarter and is pursuing longer-term overseas supply agreements. Large import programs can affect competition for available urea and phosphate cargoes, especially when export availability is already uncertain. (Times of India fertilizer-import report)
For Oklahoma producers, the current report offers a somewhat more favorable nitrogen signal, but not a uniform one. Wide quote ranges mean that comparison shopping may be as important as the regional average. Phosphate prices are locally stable, while sulfur availability and production costs remain meaningful external risks. Potash is comparatively steady.
Before acting on any quotation, confirm the nutrient analysis, cash-price basis, pickup or delivery terms, quote expiration and whether application or blending charges have been separated. UAN quotations should always be identified as either 28% or 32% before comparison.
Market Sources
Don’t Let the Heavy Rains Fool You – Scouting for Sorghum Aphids and Chinch Bugs in Sorghum
Ashleigh Faris, Cropping Systems Entomologist and IPM Coordinator
Depending on where you are in Oklahoma, you may have had anywhere between just under 0.5” or over 7.5” of rain in the last 7 days. If you were on the higher end, you may think that the rainfall has helped to prevent or wash off any insect pests building up in your summer crops. Unfortunately, this isn’t case; this morning sorghum aphids and chinch bugs were detected in research plots in northern Payne County. With rain out of the forecast and typical summer temperatures kicking in these insects, and other pests, can start to take off. Both pests have the potential to inflict severe economic damage if left unchecked, particularly during vulnerable crop stages or near recently harvested wheat. Scout now, and if needed, get together a plan to help you keep these economically damaging pests in check.
Sorghum Aphid, Melanaphis sorghi
Formerly known as the sugarcane aphid, the sorghum aphid can infest sorghum at any growth stage but causes the most severe damage from the boot to soft dough stages. The aphids form dense colonies on the undersides of leaves and produce heavy amounts of honeydew that can negatively affect plant growth and severely impede harvest.
Identification: Sorghum aphids are small aphids with light tan to pale yellow bodies, black feet, black antennae, and black cornicles (the two “tailpipes” on the back of the abdomen) (Figure 1).They almost exclusively reside on the underside of the leaves, often nestled against the midrib (Figure 2).
Figure 1. Sorghum aphids are light tan to yellow in color, have black legs, black antennae, and black cornicles (two tailpipe-like appendages at the tip of their abdomen. Photo credit: Oklahoma State University IPM.
Figure 2. Sorghum aphid nymphs and adults along the midrib of a sorghum leaf. Photo credit: Ashleigh M. Faris, Oklahoma State University Extension.
How to Scout Sorghum Aphids: Begin scouting once a week and increase to twice a week once the aphid is detected. There are two primary ways to scout:
- Option A: The Glance-N-Go App (Recommended) Download the SCA Glance-N-Go App (available on Apple and Google Play). Enter your control cost (
/bushel). Sample two leaves per plant until the app tells you whether to treat or not. It tracks the threshold automatically based on your specific economic variables.
- Option B: The Pen-and-Paper Method
- Walk 90 feet into the field. Inspect the top and bottom leaves of 3 consecutive plants, walk 5 rows over, and sample 3 more plants. (This is 1 “stop” = 6 plants).
- Walk 30 feet in an inverted “U” shape to your next stop and repeat.
- Complete 9 total stops (54 plants total).
- Record the number of plants that have 50 or more aphids.
Sorghum Aphid Economic Threshold: If using the manual method (Option B), the threshold to treat is when 25% of the plants are infested with 50 to 125 aphids per leaf. Do not spray before the threshold is met, as premature spraying can disrupt the natural enemies (lady beetles, lacewings, parasitic wasps) that naturally keep sorghum aphid populations in check.
Sorghum Aphid Management: Sivanto 200 SL and Transform WD are recommended insecticides as both are soft on beneficials and provide good residual. Follow label rates and directions. Coverage is critical so apply with high water volume (5+ gallons/acre by air; 10+ gallons/acre by ground). Avoid pyrethroids. Using pyrethroids to control secondary pests (like headworms once at grain fill) even when aphid numbers are low will wipe out beneficial insects, frequently causing sorghum aphid populations to explode.
Chinch Bug, Blissus leucopterus
Newly seeded sorghum (think double-crop and late planted sorghum for this time of the year) is especially at risk for chinch bug damage. Chinch bugs frequently migrate into sorghum fields looking for a new food source once adjacent wheat fields mature and are harvested.
Identification: Adult chinch bugs are about 1/8 inch long, black body with white wings folded over their back. The wings have distinctive black, triangular markings (Figure 3). Chinch bug nymphs (immatures) are bright red/orange right after hatching (Figure 4). They gradually darken to brown/black as they mature. They do not have wings but do have a light-colored band across their abdomens.
Figure 3. Chinch bug nymphs photographed with a microscope. Nymphs are wingless, early nymphal stages are orange to red in color with a white band, later nymphal stages are darker in color. Photo credit: Jacie Guerrero and Nikolai Thielepape, Department of Entomology and Plant Pathology, Oklahoma State University Extension.
Figure 4. Chinch bug adults have black bodies and white wings that give a triangular shape to the chinch bug’s back when folded over. Photo credit: Ashleigh M. Faris, Oklahoma State University Extension.
How to Scout Chinch Bugs: Focus on border rows adjacent to harvested wheat or thin stands. Double-cropped sorghum planted directly into wheat stubble is at the highest risk, but we have detected chinch bugs in full-season planted sorghum throughout North Central Oklahoma. Chinch bugs pierce the plant and suck the sap, injecting toxins as they feed. Look for stunting, wilting, reddish discoloration on the stalk, or poor root development (Figure 5).Check the leaf blades, but importantly, look under the leaf collars/sheaths and down at the base of the plant near the soil line where the chinch bugs hide (Figure 6).
Figure 5. Chinch bug damage and chinch bug nymph on sorghum stalk. Photo credit: Ashleigh M. Faris, Oklahoma State University Extension.
Figure 6. Chinch bug nymphs and damage to sorghum stalks found by peeling back lower sheaths of damaged sorghum stalks.Photo credit: Ashleigh M. Faris, Oklahoma State University Extension.
Chinch Bug Economic Threshold: For sorghum seedlings (less than 6 inches), treat if 2 or more chinch bugs are found on 20% of the plants.Larger plants (over 1 foot), treat if you find 10 or more chinch bugs per plant.
Chinch Bug Management: The best defense is an insecticide seed treatment (e.g., Gaucho, Poncho, or Cruiser), which provides about two to three weeks of early control. Avoid planting sorghum directly adjacent to poor wheat stands if possible.If migrating populations breach the threshold, foliar pyrethroids are the primary option (e.g., lambda-cyhalothrin or zeta-cypermethrin). Because chinch bugs hide behind leaf sheaths and at the soil level, high water volume is essential to achieve proper penetration and coverage on small plants. Spot treatments on border rows are often enough if the migration is caught early and chinch bugs have not spread into the field.
Dr. Ashleigh Faris’s Cropping Systems Entomology IPM team is currently conducting chinch bug insecticide efficacy trials. Stay tuned for the results!
For detailed insecticide rates and grazing/post-harvest intervals, always consult the product’s label guidelines and read the label carefully before application.
Banding P for Acidic Soils: Its not the time to be paying for poor practice.
I am bringing this topic back to the surface now with the current outlook on phosphorus fertilizer. If you have heard its not only becoming more expensive but the supply is short and will likely stay short through summer into the fall, which wont help prices. So this year’s wheat crop, we need to be prepared to be smart with Phosphorus, and applying an extra 30lbs to band aid for soil acidity should not be in the cards. Look at it this way, if the phosphorus was at $0.66 a lb that $20 that could be spent on a ton of lime. That lime will last 3-5 years, while that P needs to be added every year. Not only that, but the lime will help root growth (better when we dry up), produce significantly more biomass, and make the phosphorus you’ve applied in the past available again for plant uptake. So make the plans now to soil sample as soon as this crop is off, you can get a soil test recommendation and plan for the lime trucks. This is also not the year to just apply phosphorus for the sake of applying. Soil tests are inexpensive relative to buying excess fertilizer.
Current quotes on 4.24.26 are at $0.54 + per lbs P2O5 with DAP at $830 a ton.
Quick Fertilizer Price Calculation:
Urea at $860 a ton means N is $0.93 a lbs.
DAP at $830 has $334 worth of $0.93 nitrogen and $495 of phosphorus at $0.54 a lb.
Banding P as a band-aid for soil acidity, not so cheap now.
Original Blog Posted in 2021
Whoi Cho, PhD student Ag Economics advised by Dr. Wade Brorsen
Raedan Sharry, PhD Student Soil Science advised by Dr. Brian Arnall
Brian Arnall, Precision Nutrient Management Extension.
In 2014 I wrote the blog Banding P as a Band-Aid for low-pH soils. Banding phosphate to alleviate soil acidity has been a long practiced approach in the southern Great Plains. The blog that follows is a summary of a recent publication that re-evaluated this practices economic viability.
Many Oklahoma wheat fields are impacted by soil acidity and the associated aluminum (Al) toxicity that comes with the low soil pH. The increased availability of the toxic AL3+ leads to reduced grain and forage yields by impacting the ability of the plant to reach important nutrients and moisture by inhibiting root growth. Aluminum can also tie up phosphorus in the soil, further intensifying the negative effects of soil acidity. More on the causes and implication of soil acidity can be found in factsheet PSS-2239 or here (https://extension.okstate.edu/fact-sheets/cause-and-effects-of-soil-acidity.html). The acidification of many of Oklahoma’s fields has left producers with important choices on how to best manage their fields to maximize profit.
Two specific management strategies are widely utilized in Oklahoma to counter the negative impacts of soil acidification: Lime application and banding phosphorus (P) fertilizer with seed. While banding P with seed ties up Al allowing the crop to grow, this effect is only temporary, and application will be required every year. The effects of liming are longer lasting and corrects soil acidity instead of just relieving Al toxicity. Historically banding P has been a popular alternative to liming largely due to the much lower initial cost of application. However, as P fertilizers continue to increase in cost the choice between banding P and liming needed to be reconsidered.
A recent study by Cho et al.,2020 compared the profitability of liming versus banding P in a continuous wheat system considering the impacts that lime cost, wheat price and yield goal has on the comparison. This work compared the net present value (NPV) of lime and banded P. The study considered yield goal level (40 and 60 bu/ac) as well as the price of P2O5 fertilizer and Ag Lime. The price of P2O5 used in this study was $0.43 lb-1 while lime price was dictated by distance from quarry, close to quarry being approximately $43 ton-1 and far being $81 ton-1. For all intents and purposes these lime values are equivalent to total lime cost including application. Wheat prices utilized in the study were $5.10 bu-1 and $7.91 bu-1. It is important to note that baseline yield level was not considered sustainable under banded P management in this analysis. This resulted in a decrease in yield of approximately 3.2 bu ac-1 per year. This is attributable to the expected continued decline in pH when banding P is the management technique of choice.
The analysis in this work showed that lime application is cost prohibitive in the short term (1 year) when compared with banding P regardless of lime cost, yield goal level, and wheat value (within the scope of this study). This same result can be seen over a two-year span when yield is at the lower level (40 bu ac-1). While in the short-term banding P was shown to be a viable alternative to liming, as producers are able to control ground longer lime application becomes the more appealing option, especially when producers can plan for more than 3 years of future production. In fact, under no set of circumstances did banding P provide greater economic return than liming regardless of crop value, yield, or liming cost when more than 3 years of production were considered and only under one scenario did banded P provide a higher NPV in a 3-year planning horizon.
While historically banding P was a profitable alternative to lime application for many wheat producers the situation has likely drastically changed. At the time of writing this blog (09/17/2021) Diammonium Phosphate (DAP) at the Two Rivers Cooperative was priced at $0.78 lb-1. of P2O5. This is a drastic increase in P cost over the last year or so since Cho et al. was published in 2020. With P fertilizer prices remaining high it will be important for producers to continue to consider the value of liming compared to banded P. This is particularly crucial for those producers who can make plans over a longer time frame, especially those more than 3 years.
Addendum: As fertilizer prices have continued to rise a quick analysis utilizing the $0.78 lb-1 of P2O5was completed to consider the higher P fertilizer cost. Under this analysis an estimated decrease in NPV of approximately $38 an acre for P banding occurred. When considering this change in NPV, lime application becomes the more profitable option for alleviation of soil acidity symptoms even in the short term (assuming lime price values are equivalent to the previous analysis). This underlines the fact that it is imperative to consider the impact on profitability of the liming vs. banding P decision in the current economic climate for agricultural inputs.
Link to the Open Access Peer Reviewed publication “Banding of phosphorus as an alternative to lime for wheat in acid soil” https://doi.org/10.1002/agg2.20071
Mechanisms of Soil Fertility: Looking at Biologicals and MOA
Brian Arnall, Oklahoma State University Precision Nutrient Management
The use of biological products in commercial agriculture has expanded rapidly, with large corporations entering a space once dominated by smaller groups. This has created an arms race, with nearly every company offering a biological product. Over the past twenty years, I have had the opportunity to test products from the biggest groups with billions in backing, to solutions raised in stock tanks delivered in Braums milk jugs. It is critical to understand what is in the jug and the biological function it is expected to perform. Like herbicides, knowing the mode of action determines whether the product fits the intended purpose. No different than herbicides and knowing mode of actions. It’s important to know and understand that if you are trying to kill ryegrass 2.4-D, a broadleaf herbicide is not the right answer.
So what are we working with that’s in these products?
My approach has been to classify the products by operation not by species or genre. Doing so I have grouped products into five biological classifications and a sixth group, which is often in concluded in conversations.
Decomposers / Organic Matter Mineralizers
Nitrogen Fixers (Symbiotic and Associative)
Symbiotic Root Associations (Mycorrhizae, PGPR)
Nutrient Solubilizers
Biological Pest Control
Plant Growth Regulators (Hormonal Effects)
So, let’s dig into each of the mechanisms.
Decomposers / Organic Matter Mineralizers
Decomposition is carried out by a diverse group of organisms including fungi (e.g., Trichoderma, Aspergillus), bacteria (e.g., Bacillus, Pseudomonas), and actinomycetes (e.g., Streptomyces), each contributing to the breakdown of organic materials through different enzymatic pathways. This process of decomposing organic matter releases the nutrients tied up into plant available forms. The release of nitrogen is usually first thought, but this process adds significant amounts of potassium, calcium, and magnesium.
The process occurs both in the soil and on the soil surface. While it seems simple in application though this is a complex process. Let’s start with the soil pool, triggering decomposition of a system where the previous crop was wheat is significantly different than following corn. Following wheat, the carbon nitrogen ratio will be very high (see sugar blog), so while decomposition will release cations such as potassium and calcium, it is very likely to immobilize and residual nitrogen in the system. However, in fields that previously had corn the carbon to nitrogen ratio is much closer and the probability of seeing nitrogen release is much higher (Kuzyakov & Blagodatskaya, 2015). The process is similar for surface residues, but the rate is heavily controlled by rainfall. While both the soil and surface systems require moisture for the process to progress, the surface moisture is much more dynamic with frequent wetting and drying. Rain or irrigation is also needed to move the nutrients into the root zone.
One aspect of increasing decomposition of OM that I do not have a handle on is the long-term impact of expediting OM breakdown in and on the soil, especially in the central plains. As mentioned in the sugar blog, you would hope that the increase in nutrients from OM decomposition would increase plant growth enough to replenish the OM that was burned up. One caveat to this is that the decomposition would have to add nutrients that are deficient. Otherwise, there is no increase in plant growth and hypothetically the system is not net negative on OM. When it comes to decomposing surface residue, I have always been a bit hesitant in Oklahoma as I see having surface coverage to preserve soil moisture typically has a greater value than the nutrients from the residue.
Nitrogen Fixers (Symbiotic and Associative)
Nitrogen fixation is carried out by both symbiotic organisms such as Rhizobium and Bradyrhizobium, which form nodules on plant roots and supply significant nitrogen, and associative organisms such as Azospirillum and Azotobacter, which reside in the rhizosphere and contribute smaller, more variable amounts of nitrogen. Symbiotic nitrogen fixation, such as we have come to expect from legumes, is tightly regulated by the plant, with carbon supplied to the microbe in exchange for fixed nitrogen, making it one of the most efficient biological nitrogen inputs in agriculture.
Associative nitrogen fixation is not directly coupled to plant demand, and nitrogen contributions are typically limited by carbon availability and environmental conditions (Kennedy et al., 2004). While these organisms possess the ability to fix atmospheric nitrogen, the magnitude of nitrogen contribution, particularly from non-symbiotic systems, is highly variable and often limited under field conditions. We know that in soybean nodulation is greatly reduced when excess nitrogen is present in the soil, basically the plant does not need rhizobia, so it does not trigger symbiosis. I expect that as we move symbiotic fixation out of legumes that this mechanism does not change. Finally fixed N is no different than fertilizer N, if you add more then the crop needs, its lost. Therefore, if I am planning to use a N fixer, I would significantly reduce the amount of fertilizer N apply well below crop demand. Otherwise, the money spent on the N fixer is a waste. The only argument I have heard for this is the security blanket, making sure that if more is needed than normally the system is covered. But I circle back to the question about a system with high levels of residual N and rhizobium nodulation.
Symbiotic Root Associations (Mycorrhizae, PGPR)
Symbiotic root associations include arbuscular mycorrhizal fungi (e.g., Rhizophagus, Funneliformis) that extend the effective root system and improve nutrient uptake, particularly phosphorus, as well as plant growth-promoting rhizobacteria (e.g., Pseudomonas, Bacillus) that influence root development and plant signaling through multiple biochemical pathways (Smith & Read, 2008). In my visits with soil microbiologist, I have been left with the understanding that these relationships are not generic, but quite specific. There is significant influence of genotype and environment. And even more interesting is that the majority expect that the plant needs to signal for this relationship to happen.
The effectiveness of these associations is highly dependent on soil conditions, existing microbial communities, and nutrient availability, with responses often diminishing in systems where nutrients are not limited or where native populations are already established. I was able to follow along with some work down at OSU a few years back that was working with sorghum looking for symbiotic relationships to improve water and nutrient uptake specifically phosphorus. The work was successful, the researchers were able to identify a AMF that created a symbiotic relationship with sorghum, with a few caveats. First land race cultivars had a much higher incidence of symbiosis. For the landraces it worked well in extremely nutrient depleted soils and any additions of N or P reduced forage yield over the none. In the end the researchers were able to show improved the grain yield in landraces above fertilized, but these yields did equal fertilized hybrids. This work had great impact on small holders in developing counties with limited resources.
Nutrient Solubilizers
Nutrient solubilization is carried out by organisms such as Bacillus, Pseudomonas, and Aspergillus, which increase nutrient availability through mechanisms including organic acid production, proton release, and chelation, allowing nutrients like phosphorus and micronutrients to become more accessible in the rhizosphere.
Phosphorus-solubilizing fungi, such as Aspergillus and Penicillium, function similarly to bacterial solubilizers but are often more effective at producing strong organic acids. These acids can lower pH in localized zones and release phosphorus from mineral-bound forms, particularly in soils with high fixation capacity. Fungal systems can operate across a wider range of environmental conditions and may play a larger role in longer-term phosphorus cycling. However, as with bacterial systems, these effects are generally localized and dependent on soil chemistry (Richardson et al., 2009). I tend to see these having the greatest benefits in systems that have historically received manures or long-term applications of fertilizer P. I do not believe this is a good fit for soils with limited available phosphorus, as it is trying to focus the soil into something, it does not want to do or have too spare.
Potassium-solubilizing organisms, including species such as Bacillus mucilaginosus and Frateuria aurantia, contribute to the release of potassium from primary minerals like feldspars and micas. These microbes facilitate mineral weathering through acidification and chelation processes that slowly break down mineral structures. While the mechanism is well understood, the rate of potassium release is typically slow relative to crop demand. As a result, these organisms are more influential in long-term soil development than in short-term fertility management (Sheng & He, 2006).
Micronutrient-mobilizing organisms, particularly Pseudomonas and Bacillus species, enhance availability through the production of siderophores and other chelating compounds. These molecules bind metals such as iron and zinc, increasing their solubility and facilitating uptake in the rhizosphere. This process is especially important in soils where micronutrients are present but not readily available due to chemical constraints. However, the impact is typically limited to the immediate root zone and depends on both microbial activity and soil conditions (Ahmed & Holmström, 2014).
Biological Pest Control
Biological pest control organisms, including species such as Bacillus, Pseudomonas, and Trichoderma, function by suppressing pathogens through several well-documented mechanisms. These include the production of inhibitory compounds, competition for space and nutrients, direct antagonism of pathogens, and the activation of plant defense systems through induced systemic resistance. While these mechanisms are well established under controlled conditions, their effectiveness in field environments is highly dependent on environmental conditions, pathogen pressure, and the ability of the organism to persist and colonize the soil or plant surface (Lugtenberg & Kamilova, 2009).
I’ve been working with a lot of folks from Brazil who historically make four to six nemacide applications in soybean, but utilizing Pseudomanas they have been able to reduce that number by half or more. The caveat, as I understand, the application rates needed are significantly higher than anything I have seen in the US. If you look through the literature, you are seeing more and more documentation of this such as Li et al. 2022. But as Spescha et al. (2023) documented, different biological control agents operate through complementary mechanisms, including infection, toxin production, and host targeting. However, effectiveness depended on environmental conditions and interactions among organisms, reinforcing that biological control outcomes are system-dependent rather than universally consistent.
Plant Growth Regulators (Hormonal Effects)
This group differs slightly, as the primary effect is not direct nutrient cycling but modification of plant physiological response. This group is one I hold the greatest expectations for. I mean we have been using PGRs in crop production for decades, we just did not have an inkling of how many PGRs exist.
Plant growth regulator effects are associated with organisms such as Azospirillum, Bacillus, and Pseudomonas, which can influence plant development through the production of phytohormones and related compounds. These microbes produce substances such as auxins, cytokinins, and gibberellins that alter root architecture and plant growth patterns, and in some cases reduce stress responses through enzymes like ACC deaminase. Rather than supplying nutrients directly, these organisms modify how plants respond to their environment and utilize available resources. However, just like everything previously discussed the magnitude of response is often subtle and highly dependent on environmental conditions and crop system interactions (Glick, 2012).
Final thoughts.
There is one situation that pops up that I do not agree with, based upon my limited understanding of soil microbiology. Its adding more of what is already there. The soil system is a dynamic system. While there are population booms and bust, it supports what it is able to. Adding more of what is already there is like dropping a million rabbits into a prairie that has rabbits already. The current population is where it is because that is what the system can support. Adding means one of two things, a lot of rabbits die immediately, or they overwhelm the system and another animal species dies off due to lack of resources. Also, most microbiologists tell me the system is amazing at signaling and finding what it wants. It may take a season, but it will be there, in the quantities that soil needs, just given time.
So, the final slide in all my biological additives talks ends with this statement. My experiments show one thing. The impact of adding these products on crop yields is very consistently inconsistent. I’ve had many show a significant positive response, once. I have struggled to ever get repeated successes. It is my belief that I will have more success improving the soil biome by managing the soil (no-till, crop rotation, cover crops) than I will ever have with adding a product.
Final comment, Read the label. Many of the biological products I have tested are not singularly pure species. There are many blends of species and organisms which encompass many of the modes. A lot of these blends also contain extras such as humics, fulvics, carbohydrates, and sugars, see previous blogs.
Take-Home Messages
- Biological products function through specific mechanisms, not as broad “boosters,” and understanding that mechanism is critical to proper use.
- The presence of a biological function does not guarantee a yield response, outcomes are driven by soil, crop, and environmental conditions
- Decomposers and carbon-driven systems can immobilize or mineralize nitrogen, depending largely on residue quality and system balance
- Mycorrhizae and PGPR improve access to existing nutrients, not total nutrient supply
- Nutrient-solubilizing organisms mobilize nutrients already present in the soil
- Plant growth regulators influence plant signaling and development
- Adding biological organisms to soil does not guarantee establishment or persistence, as soil systems can regulate microbial populations.
- Management practices such as no-till, crop rotation, and cover crops are effective at improving soil biological function
- Across all biological products, mechanism exists, but response depends on the system
Any questions or comments please reachout to me @ b.arnall@okstate.edu
Citations
Ahmed, E., & Holmström, S. J. M. (2014). Siderophores in environmental research: Roles and applications. Microbial Biotechnology, 7(3), 196–208.
Glick, B. R. (2012). Plant growth-promoting bacteria: Mechanisms and applications. Scientifica, 2012, 963401
Kennedy, I. R., Choudhury, A. T. M. A., & Kecskés, M. L. (2004).
Non-symbiotic bacterial diazotrophs in crop-farming systems. Plant and Soil, 266, 65–79.
Kuzyakov, Y., & Blagodatskaya, E. (2015).
Microbial hotspots and hot moments in soil. Soil Biology and Biochemistry, 83, 184–199.
Lugtenberg, B., & Kamilova, F. (2009). Plant-growth-promoting rhizobacteria. Annual Review of Microbiology, 63, 541–556.
Richardson, A. E., Barea, J. M., McNeill, A. M., & Prigent-Combaret, C. (2009). Acquisition of phosphorus and nitrogen in the rhizosphere and plant growth promotion by microorganisms. Plant and Soil, 321(1–2), 305–339.
Sheng, X. F., & He, L. Y. (2006). Solubilization of potassium-bearing minerals by a wild-type strain of Bacillus edaphicus and its mutants and increased potassium uptake by wheat. Canadian Journal of Microbiology, 52(1), 66–72. https://doi.org/10.1139/w05-117
Smith, S. E., & Read, D. J. (2008).
Mycorrhizal symbiosis. Academic Press.
Spescha, A., Weibel, J., Wyser, L., Brunner, M., Hess Hermida, M., Moix, A., Scheibler, F., Guyer, A., Campos-Herrera, R., Grabenweger, G., & Maurhofer, M. (2023). Combining entomopathogenic Pseudomonas bacteria, nematodes and fungi for biological control of a below-ground insect pest. Agriculture, Ecosystems & Environment, 348, 108414.
Ye S, Yan R, Li X, Lin Y, Yang Z, Ma Y and Ding Z (2022) Biocontrol potential of Pseudomonas rhodesiae GC-7 against the root-knot nematode Meloidogyne graminicola through both antagonistic effects and induced plant resistance. Front. Microbiol. 13:1025727. doi: 10.3389/fmicb.2022.1025727
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
One Well-Timed Shot: Rethinking Split Nitrogen Applications in Wheat production
Brian Arnall, Precision Nutrient Management Specialist
Samson Abiola, PNM Ph.D. Student.
Nitrogen is the most yield limiting nutrient in wheat production, but it’s also the most unpredictable. Apply it too early, and you risk losing it to leaching or volatilization before your crop can use it. Apply it too late, and your wheat has already determined its yield potential; you’re just feeding protein at that point. For decades, the conventional wisdom has been to split nitrogen applications: put some down early to get the crop going, then come back later to apply again. But does splitting actually work? And more importantly, when is the optimal window to apply nitrogen if you want to maximize both yield and protein quality? We spent three years across different Oklahoma locations testing every timing scenario to answer these questions.
How We Tested Every Nitrogen Timing Scenario in Oklahoma Wheat
Between 2018 to 2021, we conducted field trials at three Oklahoma locations, including Perkins, Lake Carl Blackwell, and Chickasha, representing different soil types and growing conditions across the state. We tested three nitrogen rates: 0, 90, and 180 lbs N/ac, applied as urea at five critical growth stages based on growing degree days (GDD). These timings were 0 GDD (preplant, before green-up), 30 GDD (early tillering), 60 GDD (active tillering), 90 GDD (late tillering, approximately Feekes 5-6), and 120 GDD (stem elongation, approaching jointing). We also compared single applications at each timing against split applications, where half the nitrogen (45 lbs N ac-1) went down preplant, and the other half was applied in-season (45 lbs N ac-1).
The Sweet Spot: Yield and Protein at the 90 lbs N/ac Rate
Across all site-years, at the 90 lbs N/ac rate, timing had a significant impact on both yield and protein. The highest yields came from the 30 and 90 GDD timings, producing 62 to 66 bu/ac, with 60 GDD reaching the peak (Figure 1). Protein at these early timings stayed relatively modest at 13%. The 90 GDD timing delivered 62 bu/ac with 14% protein matching the yield of the 30 GDD application but pushing protein a percentage higher (Figure 2). The real problem appeared at 120 GDD. Delaying application until stem elongation dropped yields to just 49 bu/ac, even though protein climbed to 15%. That’s a 13 bushel penalty compared to the 90 GDD timing. At current wheat prices per bushel, that late application may cost farmers over $100 per acre in lost revenue. By 120 GDD, the crop has already determined its yield potential tillers are set, head numbers are locked in and nitrogen applied at this stage can only be directed toward protein synthesis, not building more yield components.
More Nitrogen Does not lead to high yield
Doubling the nitrogen rate to 180 lbs N/ac revealed something critical, more nitrogen doesn’t mean more yield. The yield pattern remained nearly identical to the 90 lbs N/ac rate. The 60 GDD timing produced the highest yield at 68 bu/ac, followed closely by 30 GDD at 67 bu/ac. The 90 GDD timing yielded 62 bu/ac, and the 120 GDD timing again crashed to 51 bu/ac. The only difference between the two rates was protein concentration (Figure 2). At 180 lbs N/ac, protein levels increased across all timings: 13% at preplant, 15% at both 30 and 60 GDD, 15-16% at 90 GDD, and 16% at 120 GDD. This confirms a fundamental principle: once farmers supply enough nitrogen to maximize yield potential, which occurred at 90 lbs N/ac in these trials, additional nitrogen only increases grain protein. It does not build more bushels. Unless farmers are receiving premium payments for high-protein wheat, that extra 90 lbs of nitrogen represents a cost with no yield return.
Should farmers split their nitrogen application?
Now that timing has been established as critical, the next question becomes: should farmers split their nitrogen applications, or is a single application sufficient? The conventional recommendation has been to split nitrogen apply part preplant to support early growth and tillering, then return with a second application later in the season to boost protein and finish the crop. But does the data support this practice? We compared three strategies at each timing: applying all nitrogen preplant, applying all nitrogen in-season at the target timing, or splitting nitrogen equally between preplant and in-season timing. The goal was to determine whether the extra trip across the field will deliver better results.
Our findings revealed that splitting provided no consistent advantage. At 30 GDD, all three strategies preplant, in-season, and split performed identically, producing 62-65 bu/ac with 12-13% protein (Figure 3 and 4). No statistical differences existed among them. At 60 GDD, similar pattern was held. Yields ranged from 61 to 66 bu/ac and protein stayed at 12-13% regardless of whether farmers applied all nitrogen preplant, all at 60 GDD, or split between the two. At 90 GDD, the single in-season application actually outperformed the split. While yields remained similar across all three methods (61-64 bu/ac), the in-season application delivered significantly higher protein at 13.7% compared to 12.4% for preplant and 12.5% for split applications. This suggests that concentrating nitrogen at 90 GDD, rather than diluting it across two applications, allows more efficient incorporation into grain protein. The only timing where splits appeared beneficial was 120 GDD, where the split application yielded 59 bu/ac compared to 51 bu/ac for the single late application. But this is not a win for splitting, it simply demonstrates that applying all nitrogen at 120 GDD is too late and putting half down earlier salvages some of the yield loss. Across all timings tested, splitting nitrogen into two applications offered no agronomic advantage over a single well-timed application, meaning farmers are making an extra pass for no gain in yield or protein.
Practical Recommendations for Nitrogen Management
Based on three years of field data, farmers should target the 90 GDD timing (late tillering, Feekes 5-6) for their main nitrogen application to achieve the best balance between yield and protein. This window typically falls in late February to early March in Oklahoma, though farmers should monitor crop development rather than relying solely on the calendar apply when wheat shows multiple tillers, good green color, and vigorous growth. A rate of 90 lbs N/ac maximized yield in these trials; higher rates only increased protein without adding bushels, so farmers should only exceed this rate if receiving premium payments for high-protein wheat. Splitting nitrogen applications provided no advantage at any timing, meaning a single well-timed application at 90 GDD is sufficient for most Oklahoma wheat production systems. The exception would be sandy soils with high leaching potential, where splitting may reduce nitrogen loss. Farmers should avoid delaying applications until 120 GDD or later, as this timing consistently resulted in 15-25 bushel per acre yield losses even though protein increased. For farmers specifically targeting premium protein markets, a two-step strategy works best: apply 90 lbs N/ac at 90 GDD to establish yield potential and baseline protein, then follow with a foliar application of 20-30 lbs N/ac at flowering to push protein above 14% without sacrificing yield. Finally, weather conditions matter hot, dry forecasts increase volatilization risk and reduce uptake efficiency, so farmers should consider moving applications earlier if low humidity conditions are expected.
Split Application Caveat * Note from Arnall.
The caveat to the it only takes one pass, is high yielding >85+ bpa, environments. In these situation I still have not found any value for preplant nitrogen application. I have seen however a split spring application is valuable. Basically putting on 30-50 lbs at green-up, with the rest following at jointing (hollowstem). The method tends to reduce lodging in the high yielding environments.
This work was published in Front Plant Sci. 2025 Nov 6;16:1698494. doi: 10.3389/fpls.2025.1698494
Split nitrogen applications provide no benefit over a single well timed application in rainfed winter wheat
Another reason to N-Rich Strip.
Yet just one more data set showing the value of in-season nitrogen and why the N-Rich Strip concept works so well.
Questions or comments please feel free to reach out.
Brian Arnall b.arnall@okstate.edu
Acknowledgements:
Oklahoma Wheat Commission and Oklahoma Fertilizer Checkoff for Funding.
What OK Cotton Growers Should Know about the Two-Spotted Cotton Leafhopper, aka Cotton Jassid
Ashleigh M. Faris, OSU Extension Cropping Systems Entomologist Department of Entomology and Plant Pathology Oklahoma State University
Jenny Dudak, OSU Extension Cotton Specialist, Department of Plant & Soil Sciences Oklahoma State University
Maxwell Smith, OSU IPM for Cotton Extension Specialist Department of Entomology and Plant Pathology, Oklahoma State University
Published 8.29.25
The two-spotted cotton leafhopper, also known as the cotton jassid (Amrasca biguttula), has not been detected in Oklahoma cotton production as of August 27, 2025. However, this insect pest has been confirmed to be causing significant injury in cotton fields that have high populations of the jassid throughout the southeastern United States (Figure 1). Although it is not known to currently be in Oklahoma cotton acres, our cotton growers and consultants are encouraged to keep an eye out for this pest and to report sightings of the insect or related damage to the OSU Cotton Extension team for confirmation.
Two-Spotted Cotton Leafhopper Identification & Injury
Cotton jassids are small, sap-sucking insects that are pale green insects with yellowish-green wings. They can be distinguished from most native North American species by the pair of black spots head and black spots on the tip of each wing (Figure 2). The immature nymph stages are even smaller than the adults; they are wingless but light green in color like the adults. (Figures 3 and 4).
The cotton jassid feeds on the underside of leaves, causing hopperburn—a rapid yellowing, reddening, and browning that can quickly weaken plants (Figure 5). Initially, injury symptoms may look like nutrient deficiency with slight yellowing along the leaf tips and margins. Injury may also present as upward curling/cupping of the leaves. Once early symptoms are visible, leaves decline rapidly, turning red and brown. Because late-stage hopperburn can resemble a spider mite infestation, growers and consultants should scout for the pest causing injury prior to initiating control. Cotton jassid populations and hopperburn symptoms usually begin on field edges before moving into the cotton field.
Suggested Threshold, Scouting, and Chemical Control Guidance
Current recommendations for the suggested threshold and chemical control options are based on insecticide trials conducted by University of Florida, University of Georgia, Clemson University, and the University of Auburn Extension Entomologists. The threshold at which treatment is recommended for the cotton jassid is 2 nymphs per leaf with injury present anywhere in the field. To scout, inspect the underside of the main stem leaf on the third, fourth, or fifth node from the top of the cotton plant. Sample a minimum of 25 leaves across plants and average counts across the number of leaves sampled.
In terms of management, multiple products seem to provide good efficacy. Bidrin (6 oz/a) has been consistent in multiple trials across Florida, Georgia, and South Carlina, but may be difficult if whiteflies are an issue. Centric (2 oz/a) and Transform (1.5 oz/a) seem to be consistent as well. One product that should not be used is Bifenthrin, which has proven not to be effective across multiple trials. Information on suggested chemical control options, their efficacy, and general IPM guidance will be forthcoming as more is learned about this pest in U.S. cotton systems.
If you detect the cotton jassid in Oklahoma cotton, please contact
IPM for Cotton Extension Specialist Maxwell Smith,
PRE-EMERGENT RESIDUAL HERBICIDE ACTIVITY ON SOYBEANS, 2025
Liberty Galvin, Weed Science Specialist
Karina Beneton, Weed Science Graduate Student.
Objective
Determine the duration of residual weed control in soybean systems following the application of Preemergent (PRE) herbicides when applied alone and in tank-mix combination.
Why we are doing the research
PRE herbicides offer an effective means of suppressing early-season weed emergence, thereby minimizing competition during the critical early growth stage. However, evolving herbicide resistance and the need for longer-lasting weed suppression underscore the importance of evaluating multiple modes of action and their residual properties alone and tank-mixed.
Field application experimental design and methods
Field experiments were conducted in 2022, 2023, and 2024 growing seasons in Bixby, Lane, and Ft. Cobb FRSU Research Stations across Oklahoma. Each herbicide (listed in Table 1) was tested individually, in 2-way combinations, 3-way mixtures, and finally as 4-way combinations that included all active ingredients listed at the label rate.
Soybeans were planted at rates between 116,000 and 139,000 seeds/acre from late May to early June, depending on the year and location. The variety used belongs to the indeterminate mid- maturity group IV, with traits conferring tolerance to glyphosate (group 9 mode of action), glufosinate (group 10), and dicamba (group 4). Not all soybean varieties have metribuzin tolerance. Please read the herbicide label and consult your seed dealer for acquiring tolerant varieties. Row spacing was 76 cm at Bixby and Lane, and 91 cm at Fort Cobb. PRE treatments were applied immediately after planting at each experimental location.
POST applications consisted of a tank-mix of dicamba (XtendiMax VG® – 22 floz/acre), glyphosate (Roundup PowerMax 3®- 30 floz/acre), S-metolachlor (Dual II Magnum® – 16 floz/acre), and potassium carbonate (Sentris® – 18 floz/acre). Applications were made on different dates, mostly after the first 3 weeks following PRE treatments. These timings were based on visual weed control ratings, particularly for herbicides applied alone or in 2-way combinations, which showed less than 80% control at those early evaluation dates. The need for POST applications also depended on the species present at each site, with most fields being dominated by pigweed, as illustrated in the figure below.
Results
Tank-mixed PRE herbicide combinations generally provided superior residual control compared to a single mode of action application (Shown in Figure 1). Timely post-emergent (POST) herbicide applications helped sustain high levels of weed suppression, particularly as the effectiveness of residual PRE declined.
Residual control of tank-mixed PRE
Some herbicides applied alone or in simple 2-way mixes, such as sulfentrazone + chloransulam- methyl and pyroxasulfone + chloransulam-methyl required POST applications within 20 to 29 days after PRE, indicating moderate residual control.
In contrast, 2-way combinations containing metribuzin, such as sulfentrazone + metribuzin and pyroxasulfone + metribuzin, extended control up to 50 days after PRE in some cases, highlighting metribuzin’s importance even in less complex formulations.
Furthermore, 3-way and 4-way combinations including metribuzin provided the longest-lasting control, delaying POST applications up to 51–55 days after PRE.
Injury of specific weeds
Palmer amaranth (Amaranthus palmeri) control in Bixby was consistently high (≥90%) at 2 weeks after PRE in 2022 and 2024 across all treatments. At 4 WAPRE, treatments containing metribuzin alone or in combination maintained strong control (90% or greater).
Texas millet (i.e., panicum; Urochloa texana) and large crabgrass (Digitaria sanguinalis) were effectively managed with most treatments delivering over 90% control early in the season and maintaining performance throughout. In 2024, control remained generally effective, though pyroxasulfone alone showed a temporary lack of control for Texas millet, and single applications declined in effectiveness against large crabgrass later in the season. These reductions were likely due to continuous emergence and the natural decline in residual herbicide activity due to weather. The most consistent late-season control for both species came from 3- and 4-way herbicide combinations.
Morningglory (Ipomoea purpurea) control reached full effectiveness (100%) only when POST herbicides were applied, across all years and locations. Their late emergence beyond the residual window of PRE herbicides reinforces the importance of sequential herbicide applications for season-long control.
Take home messages:
- Incorporating PRE and POST herbicides slows the rate of herbicide resistance
- Tank mixing with *different modes of action* ensures greater weed control by having activity on multiple metabolic pathways within the plant.
- Tank mixing with PRE herbicides could reduce the number of POST applications required, and
- Provides POST application flexibility due to residual of PRE application
For additional information, please contact Liberty Galvin at 405-334-7676 | LBGALVIN@OKSTATE.EDU or your Area Agronomist extension specialist.
Laboratory evaluation of Liquid Calcium
Liquid calcium products have been around for a long time. The vast majority of these products are either a calcium chloride or chelated calcium base which is now commonly found with the addition of a humic acid, microbial, or micronutrient. Many of these make promises such as “raises your soil pH with natural, regenerative, liquid calcium fertilizers that correct soil pH quickly, efficiently, and affordably!”. From a soil chemistry aspect the promise of adding 3 to 5 gallons of a Ca solution, which is approximately 10% Ca, will raise the soil pH is impossible on a mass balance approach. In this I mean that to increase the pH of an acid soil {soil pH is the ratio of hydrogen (H) and hydroxide (OH) in the soil, and having an acid soil means the concentration of H is greater than that of OH} requires a significant portion of the H+ that is in solution and on soil particle to be converted to OH, or removed from the system entirely.
The blog below walks through the full chemical process of liming a soil but in essence to reduce the H+ concentration we add a cation (positively charged ion) such as Ca or magnesium (Mg) which will kick the H+ of the soil particle and a oxygen (O) donator such as CO2 with ag lime or (OH)2 which is in hydrated lime. Each of these O’s will react with two H’s to make water. And with that the pH increases.
However regardless of the chemistry, there is always a lot of discussion around the use of liquid calcium Therefore we decided to dig into the question with both field and laboratory testing. This blog will walk through the lab portion.
This was a laboratory incubation study. The objective was to evaluation the impact of the liquid Ca product (LiqCa**) on the soil pH, buffer capacity, Ca content and CEC of two acidic soils. LiqCa was applied at three rates to 500 g of soil. The three rates were equivalent to 2, 4, and 6 gallon per acre applied on a 6” acre furrow slice of soil. One none treated check and two comparative products were also applied. HydrateLime (CaO) as applied at rate of Ca equivalent to the amount of Ca applied via LiqCa, which was approximately 1.19 pounds of Ca per acre. Also AgLime (CaCO3) was applied at rates equivalent to 1, 2, and 4 ton effective calcium carbonate equivalency (ECCE). The Ag lime used in the study had a measured ECCE of 92%. The two soils selected for both acidic but had differing soil textures and buffering capacities. The first LCB, had an initial soil pH (1:1 H2O) of 5.3 and a texture of silty clay loam and Perkins had a initial pH of 5.8 and is a sandy loam texture. Both soils had been previously collected, dried, ground, and homogenized. In total 10 treatments were tested across two soils with four replications per treatment and soil.
Project protocol, which has been used to determined site specific liming and acidification rates, was to apply the treatments to 500 grams of soil. Then for a period of eight weeks this soil wetted and mixed to a point of 50% field capacity once a week then allowed to airdry and be mixed again. At the initiation and every two weeks after soil pH was recorded from each treatment. The expectation is that soil pH levels will change as the liming products are impacting the system and at some point, the pH reaches equilibrium and no longer changes. In this soil that point was week six however the trail was continued to week eight for confirmation. See Figures 1 and 2.
ANOVA Main effect analysis showed that Soil was not a significant effect so therefore both soils were combined for further analysis. Figure 3 shows the final soil pH of the treatments with letters above bars representing significance between treatments. In this study all treatments were significantly greater than the check with exception of LiqCal 2 and CaO 6. Neither LiqCal or CaO treatments reached the pH level of Aglime, regardless of rate.
Summary
The incubation study showed that application of LiqCal at a rate of 4 and 6 gallons per acre did significantly increase the soil pH by 0.1 pH units and 6 gallons per acre increased the Buffer index above the check by 0.03 units. Showing the application of LiqCal did impact the soil. However the application of 1 ton of Ag lime resulted in significantly great increase in soil pH, 1.0 units by 8 weeks and a buffer index change of 0.2 units. The Aglime 1 was statistically greatly than all LiqCal treatments. Ag lime 2 and 4 were both statistically greater than Ag lime 1 with increasing N rate with increasing lime rate. Given the active ingredient listed in LiqCal is CaCl, this result is not unexpected. Ag lime changes pH by the function of CO3 reacting H+ in large quantities. In a unsupported effort a titration was performed on LiqCal, which show the solution was buffered against pH change. However it was estimated that a application of approximately 500 gallons per acre would be needed to sufficiently change the soil pH within a 0-6” zone of soil.
Results of the field study.
https://osunpk.com/2025/06/02/field-evaluation-of-lime-and-calcium-sources-impact-on-acidity/
Take Home
The application of a liquid calcium will add both calcium and chloride which are plant essential nutrients and can be deficient. In a soil or environment suffering from Cl deficiency specifically I would expect an agronomic response. However this study suggest there is no benefit to soil acidity or CEC with the application rates utilized (2, 4, and 6 gallon per acre).
** LiqCal The product evaluated was derived from calcium chloride. It should be noted that since the completion of the study this specific product used has changed its formulation to a calcium chelate. This change however would not be expected to change the results as the experiment did include a equivalent calcium rate of calcium oxide.
Other articles of Interest
https://extension.psu.edu/beware-of-liquid-calcium-products
https://foragefax.tamu.edu/liquid-calcium-a-substitute-for-what/
Any questions or comments feel free to contact me. b.arnall@okstate.edu
Down and Dirty with NPK 