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Certified Crop Adviser Conservationist of the Year Award

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Multiple industry groups and federal agencies came together to recognize the CCA Conservationist of the Year Award.

In early December 2023, Russell Taylor received the 2023 Certified Crop Adviser Conservationist of the Year Award in a ceremony in Washington, D.C., hosted by USDA. The CCA Conservationist of the Year Award recognizes a crop advisor who has demonstrated leadership in conservation within the agriculture industry.

Taylor is a CCA and vice president of Live Earth Products, Inc., which mines and manufactures humic and fulvic acid-based products. To him, conservation in ag means doing more with less.

2023 CCA Conservationist of the Year Award recipient Russell Taylor speaking at the award ceremony in Washington D.C. hosted by USDA (all photos courtesy R. Taylor.)

“Agriculture is going to deal with the human population doubling by 2050,” Taylor said. “And how will we do that with the same amount of resources? That’s going to be using the same amount of water and the same amount of land, or even less land due to loss to development, to feed that growing population. So as somebody who’s looking at conservation, we need to maximize our output and make better use of our inputs.
“Conservation is also reshaping the rules that prevent farmers from accessing products and information that aid in conservation,” he said. “So, getting CCAs’ and researchers’ expanded knowledge is essential to accomplish the goal of helping [growers].”

Taylor has contributed to the conservation movement for years through various educational avenues. He advocates for using humates to reduce fertilizer loss and add organic matter to the soil to improve conservation. He’s also worked with states and the federal government to change rules restricting label claims and products growers can use.
“Some of the rules we’re working under are 1950s laws,” Taylor said. “And through agricultural advancement, we have new technologies that these old laws don’t cover. So, the laws were written for DDT and pesticides, and that was it. No other beneficial crop inputs like plant biostimulants were included.

“The efforts that both me and many individuals within the regenerative agriculture and the biostimulant communities have worked hard toward is just changing these rules.”
While receiving the award, Taylor got the chance to speak with multiple leaders in the ag industry about conservation, including USDA NRCS’ national nutrient management specialist and national agronomist as well as members of the Senate and House Ag committees. Reflecting on these conversations, Taylor agreed that in terms of conservation, the ag industry is headed in the right direction.

“You see the grind on the news about Congress and how things aren’t getting done,” Taylor said. “Then you go meet with those in the Ag committee, and it’s a lot of positivity because we’re making significant changes.

“There’s a lot of positivity going on… we’re doing great things.”
USDA-Natural Resources Conservation Service, Agricultural Retailers Association, American Society of Agronomy, CropLife America, Crop Science Society of America, National Association of Conservation Districts, National Association of State Departments of Agriculture, Soil Science Society of America and The Fertilizer Institute all came together to recognize the CCA Conservationist of the Year award.

“You’ve got multiple industry groups and federal agencies working together to recognize this award, so it is a big deal,” Taylor said. “The Conservationists Award is a collaboration of the leading conservation groups in agriculture.”

Taylor encouraged others within the ag industry to support the conservation movement where possible, highlighting the Plant Biostimulant Act as an important measure. The Plant Biostimulant Act, authored by U.S. Representative Jimmy Panetta (D-CA-20) and introduced with Rep. Jim Baird (R-IN-4), “would create a uniform process for approving commercial plant biostimulant use and require more federal research on the technology’s benefits for soil health,” according to a press release on Congressman Panetta’s website.
“The No. 1 thing a grower could do is contact a congressman and say, ‘I support this. Please get it in the farm bill,’” Taylor said.

Cover Crop Use in California and Measurable Outcomes

Trials suggest best cover crop benefits for almonds in the Central Valley come from late termination (photo by J. Kratt.)

According to the Sustainable Agriculture Research and Education (SARE), “a cover crop is a plant that is used primarily to slow erosion, improve soil health, enhance water availability, smother weeds, help control pests and diseases, increase biodiversity and bring a host of other benefits to your farm.”

Why Use Cover Crops?
The first answer I give: Nature fights against bare ground. Andre Lu wrote, “Bare ground is the best way to encourage weeds as most weeds are pioneer species. They rapidly germinate to cover disturbed and bare ground. Nature always regenerates disturbed soil by rapidly covering it with plants. Weeds are nature’s way of healing disturbed soil. Living plants feed the soil microbiome with the molecules of life so they can regenerate healthy soil.”

The way nature regenerates bare ground is through a process called plant succession, which is the change in the species structure of an ecological community over time. A great example of this is the area around Mt. Saint Helens the past 43 years since its eruption. A plant community gradually or rapidly replacing another can result from developmental changes in the ecosystem itself or from disturbances such as wind, fire, volcanic activity, insects and disease, or harvest. Different plants require different ratios of fungi and bacteria based on their successional growth traits. Cover crops are a way to dramatically speed up this process on the farm.

Different plants require different ratios of fungi and bacteria based on their successional growth traits (photo courtesy Earthfort, LLC.)

In my opinion, the more species you have in your cover crop from different plant families, the better it performs. The regenerative agriculture department at CSU Chico wrote: “Dr. Christine Jones lays out a strong case for the importance of nurturing the biodiversity in the soil using multispecies cover crops. The most diverse mix produces the best results both for the soil microbiome and for the productivity and resiliency of crops grown together in that soil.”

Side-by-side blocks on the same farm, same day, 24 hours after a heavy spring rain showing cover crops’ ability to improve water infiltration (photos by J. Kratt.)

I’ve had the benefit of meeting Dr. Jones twice in person and sat through her lectures on how cover crops can signal something called “Quorum Sensing,” which is when a very diverse mix of plant species enable the soil microbiome to become more active, which then exponentially speeds up soil health processes. According to Dr. Jones, multispecies mixes that induce Quorum Sensing need to include species from different functional plant groups (grasses, brassicas, legumes and broadleaves).

Measurable Outcomes
The first thing we saw was dramatically larger and more diverse microbial communities using a variety of soil biology tests, including Direct Microscopy, MicrobiometerTM, and BiomeMakers DNA testing. We also saw physically improved soil structure and water infiltration rather quickly. Often, our growers report seeing more earthworms and a diversity of arthropods in the soil.

After initial stunted growth at planting and through spring, whole orchard recycling cover-cropped almond trees (top) eventually outperform older conventional plantings (bottom) (photos by J. Kratt.)

In a Wasco, Calif. almond trial, samples collected June 2023 showed an orchard farmed regeneratively the past three years had 5.5x more fungi, 1.7x more bacteria, a 3.3x higher fungi-to-bacteria ratio, 3x more biologically active carbon, 2x more biologically active nitrogen, and a 1.5x higher carbon-to-nitrogen ratio than the conventional almond orchard. In addition, the regenerative orchard used to be full of several diseases, is over 22 years old, yielded 1800 to 2200 lbs. per acre the previous five years and was ready for the bulldozer, but last year and this year yielded 3600 and 3000 lbs. per acre, respectively, vs the conventional orchard which averaged 2700 and 2300 lbs. per acre. Both orchards were in the same soil series, and both were Nonpareil/Monterey on Hansen hybrid rootstock, and the conventional orchard is nine years old. The regenerative orchard used cover crops, biologicals made by Earthfort and 60% less conventional N compared to the conventional orchard which had no cover crop, no biologicals, a full NPK program per UC Davis guidelines and bare orchard floors using herbicides.

In 2021, Fenster et al. published a study titled, “Regenerative Almond Production Systems Improve Soil Health, Biodiversity and Profit” in Frontiers in Sustainable Food Systems. The study included another orchard we work with in the Wasco area. One of the more interesting findings was the water infiltration rate of regenerative orchards in California versus conventional orchards (each conventional orchard was adjacent to each regenerative orchard to control for soil type variances.) The conventional orchards averaged 0.04 ml water infiltration per second, whereas the regenerative orchards averaged 0.8 ml/s, and the orchard in Wasco we work with averaged 1 ml/s. We believe the combination of having a 16-species diverse cover crop mix, keeping it alive until June and using fungal-dominant biologicals were the reasons why this orchard (which used to have sealed ground prior to becoming regenerative) had superior water infiltration.
As mentioned before, we see improved disease suppression when using cover crops, primarily due to improved water infiltration, but we also see a dramatic increase in microbial diversity including microbes that are predatory against bacterial and fungal soil pathogens.

After mowing in late spring or early summer, cover crop residue acts as an armor to protect against water evaporation and can keep soil microbes alive during hot summer months (photo by J. Kratt.)

Plant-parasitic nematode suppression is one of the more impressive benefits we have seen. In past research, wild mustard reduced a wide range of parasitic nematodes, velvetbean reduced root-knot nematodes, and hairy indigo and joint vetch with mulched cowpea maintained low populations of B. longicaudatus and M. incognita nematodes. In a grower trial we did in Paso Robles, Calif. in wine grapes, a combination of cover crops and Earthfort biologicals were used to try and improve vineyard health and productivity. In 2021, there were 2.8 times more plant parasitic nematodes than beneficial nematodes. After three years on our regenerative program, there were 85.5x more beneficial nematodes than plant-parasitic nematodes. This was a 196% increase in beneficial nematode species and a 98.7% decrease in parasitic species.

One of the most popular reasons almond growers in California have been using cover crops is to have a food source for bees. This is important, but typically in the southern end of the Central Valley there aren’t many (or any) flowers open at the time of almond bloom due to the lateness of the planting after orchard operations or the lateness of winter rains. Then, many growers mow it all down in March either from fear of frost damage or fear that if they let the cover grow too large it will interfere with harvest operations. In contrast, we have found our best benefits come from late termination as this allows for more beneficial insects to assist with mite control, and it also helps with improved water infiltration because the roots were allowed to go much deeper.

This brings us to the most frequently stated benefit we hear: improved water infiltration. This past spring when much of the Central Valley was flooded, orchards with cover crops were able to resume orchard activities sooner than those without. Cover crops have been widely shown to reduce erosion and runoff, and in many cases we find water penetration is improved so much that gypsum is no longer needed.

After mowing in late spring or early summer, cover crop residue acts as an armor to protect against water evaporation and can keep soil microbes alive during hot summer months. Some growers and irrigation GSAs are fearful that cover crops increase water use, but recent studies by UC Davis point to no net increase of water inputs and in some cases measurable decreases in water use.

Cover crops might assist in the breakdown of wood chips in whole orchard recycling (WOR). A Kern County almond grower who did WOR in November 2022 and planted trees in January 2023 seeded a cover crop immediately after. Initially, the trees were significantly stunted from a lack of N tied up by the excess C (wood). The cover crop grew incredibly well due to the heavy spring rains. The trees were fed monthly with amino N and a microbial package from Earthfort, and the cover crop was disc-incorporated in July 2023. By the end of September, the trees outgrew the neighbor’s trees that were from the same nursery and rootstock and planted two months earlier in fumigated ground, kept bare with herbicides and fertilized with UN32. Our trees didn’t have long whippy growth either; they had thicker caliper trunks and more lateral branches than the neighbor’s. Further work needs to be done in this area to determine how cover crops and biologicals can best be used in WOR plantings.

In the final analysis, with water and weather issues continuing to be problematic, cover cropping in California is proving to be a practice that growers can’t afford not to do.

References
Lu, Andre; Growing Life: Regenerating Farming and Ranching. ACRES USA, 2021.
Frontiers in Sustainable Food Systems, Volume 5, Article 664359, August 2021.
Bending and Lincoln 1999
Rodriguez-Kabana et al. 1992
Rhoades and Forbes 1986
Resources
https://www.csuchico.edu/regenerativeagriculture/ra101-section/cover-crop-biomass.shtml
https://www.plantsciences.ucdavis.edu/news/Mitchell-Gaudin-cover-crops-video
https://www.csuchico.edu/regenerativeagriculture/blog/water-use-and-cover-crops.shtml

The Difference Between Leaf Tissue and Sap Analyses

A side cut of a leaf. Sap analysis analyzes available nutrients from the xylem and phloem. Tissue testing analyzes available and unavailable (total) nutrients from the entire leaf (photo by Jose Aburto, NEWAGE Labs.)

Utilizing lab data successfully comes down to knowing what is being measured and why. Take for example soil testing; there is a broad range of analyses. A Total Nutrient Digest uses strong acids to provide a combined (available and unavailable) total nutrient assay, whereas a Saturated Paste test utilizes no acid and provides insight on nutrients that are water soluble in solution. Both are soil tests but provide drastically different data based on the type of lab analyses performed. The relationship between the two types of soil tests is analogous to the differences between leaf tissue and sap analysis for plant nutrition assessment.

Sap extractions ready to be analyzed. Increased understanding of nutrient mobility from sap analyses can often identify nutrient deficiency, excess or toxicity long before any symptoms become visible (photo by Sierra Wall, NEWAGE Labs.)

Leaf tissue testing (whole leaf or petiole) quantifies the total accumulated nutrients that are both available and unavailable inside the plant. Unavailable nutrients have been taken up and used to build the leaf and its cellular structures. Because this test is measuring the nutrient content of the entire leaf, the results are mostly illustrating what the nutrient status was during the growth and development of those tissues. Sap analysis is measuring the liquids actively flowing in the vascular tissues (xylem and phloem) and provides a near real-time assessment of the nutrients currently available in the plant.

Leaf tissue analysis is carried out by drying, heating and pulverizing tissue for consistency in size and then employing ashing and strong acids to create a total nutrient assay solution. Sap analysis only employes linear pressure to extract the sap but keep the integrity of the leaf with no leaf mastication, for a liquid sap extraction, no heat, acids or solvents are used in preparation of sap for analyses. Leaf sap analysis identifies mostly available nutrients located inside the xylem and phloem. Nutrients within the vascular bundle are not yet incorporated into the leaf structure. The proprietary sap extraction process used at NEWAGE Laboratories in South Haven, Mich. uses differing pressures for different crop types so as not to violate the integrity of the leaves and cellular structures.

Table 1. Nitrogen parameters from a sap report showing three nitrogen parameters and nitrogen conversion efficiency comparison.

With these two different types of leaf tests also comes differing sampling protocols. Leaf tissue samples are collected from a singular age of leaf and placed in a paper bag to facilitate dehydration. Sample dehydrating is a key component for tissue analysis but in so doing one loses the ability to assess nitrogen species such as nitrate or ammonium, as well as sugars, brix, pH and electrical conductivity. With sap analysis, it is critical to maintain the samples’ integrity from the moment leaves are removed from the plant. Protocols for sampling and handling are available to help ensure leaves maintain a condition of stasis (the least amount of moisture lost and low respiration until received by the lab). To alleviate this issue, NEWAGE provides an overnight shipping program and protocol to keep samples fresh all the way to the lab.

Sap samples are collected from both new and old age leaf sources and placed in separate plastic zip style locking bags in a cooler. Collecting a new yet fully formed leaf plus petiole and an old yet still viable leaf plus petiole sample set allows for plant nutrient uptake and mobility to be observed. Nutrient mobility assessment is something Leaf Tissue testing cannot offer though an important feature of sap analysis.

How to Analyze and Interpret Nutrient Mobility from Sap
Increased understanding of nutrient mobility from sap analysis can often identify nutrient deficiency, excess or toxicity long before any symptoms become visible. As an example, N, being highly phloem-mobile, will produce a deficiency symptom initially in the old leaves if the plant has been nitrogen deficient for a prolonged period. Old leaves start to turn yellow due to lack of chlorophyll synthesis. On a sap report, N deficiency is identified, many times long before visual symptoms occur, when higher amounts of N are measured in the new leaf than the old leaf, meaning the plant is remobilizing N from old leaves to meet the high demand in the new leaves when not enough N uptake is available. An excess of N is commonly identified in sap results as well. In an excess N condition, the sap report shows more N in the old leaves than in the new. When an excess of N is persistent in sap, this points to an area where fertilizer rates can be potentially decreased and production problems with pests and diseases associated with excess N can be avoided. This comparison of new vs old leaves is applicable to identify deficiencies and excesses of all phloem-mobile nutrients which include phosphorus, potassium, magnesium, chloride, sodium, molybdenum, nickel and, of course, N. The unique N information available from sap analysis also includes N conversion efficiency analysis, something unique to NEWAGE Labs’ sap analysis and again something not available from leaf tissue analysis.

A new and old leaf sap set. Sap sampling is normally taken in pairs (photo by Finn Telles, Penny Newman.)

Nitrogen Conversion
Consideration with N fertilization is the type of N being applied and how to measure it. Sap reports from NEWAGE have four N parameters to provide an enhanced picture of N uptake and its conversion, ultimately to more complex forms like amino acids and proteins. NEWAGE measures Total Nitrogen (all species of N in the plant), N in nitrate and N in ammonium and compares all three. To compare Total Nitrogen to the N in nitrate and ammonium, the oxygen in NO3 and hydrogen in NH4 need to be removed from the equation so they are directly comparable in terms of N concentration.

Now that the different forms of N have been normalized for the analysis, this apples-to-apples N comparison can be translated into an algebraic equation to solve for ‘x’. Total Nitrogen is the product, N in NH4 and N in NO3 are known values and x is the amino acids and proteins portion of this Nitrogen Conversion equation. NEWAGE Labs has labeled this as Nitrogen Conversion Efficiency % (NCE%). The NCE% provides insight on how N is converted inside the plant. If the NCE% is at or above 90%, it means 90% of total N being taken up by the plant is being converted to amino acids and/or proteins and 10% is staying in NO3 and/or NH4 forms. A 90% or higher conversation rate is the target. As seen in Table 1, see page 30 NCE% in new and old leaves are less than 80%, and there is more Total Nitrogen in the old leaves by >80%; therefore, the color code in the report is blue to indicate an excess condition. The plants from this example are potentially being overfertilized, and the nitrogen isn’t transforming well inside the plant, which can ultimately lead to increased diseases and pests, reduced grain or fruit quality (and fruit shelf life) and an undue burden on your fertility budget.

Quality of Data
When deciding which sap lab to work with, here are some laboratory attributes to evaluate that will have a direct bearing on the quality of the reports. These laboratory attributes to evaluate include:

  • Report turnaround time
  • Quality training
  • Support of sample collection staff to ensure quality samples are collected, handled and shipped properly
  • Sample transit time to the lab
  • Report interpretation support and training

Easy-to-fill-out sample documentation is also important as a practical nutrient management tool. NEWAGE’s sample collection sheet is simple, capturing the essential information of collection date and time, field name, and who the sample was collected by. All this information is then transferred onto a NEWAGE report. With sap sampling it’s imperative the samples are kept cool to prevent N volatilization and degradation of the sample. If samples get too warm, usually by poor shipping, results of the 25 nutrient parameters NEWAGE analyses can be adversely affected. NEWAGE Labs’ overnight shipping decreases turnaround time to approximately 48 hours. Same-day sampling and shipping is recommended. If samples are held over to the next day for shipping, keep samples in refrigerator or cooler and let the air out of the plastic bags. Make sure samples do not freeze. When cellular structures burst, sap analysis is no longer a viable testing method. Leaves freeze mainly when samples are stored right up against a frozen gel back without a barrier between them like a paper towel or thin bubble wrap. Request a crop-specific sampling guide as how and when to sample are of great importance and impact data quality.

Leaf sap analysis is a tool to help growers make informed in season decisions by looking at nutrient uptake in greater detail using nutrient mobility and N conversion efficiency. Go to www.newagelaboratories.com or contact Info@newagelaboratories.com for more information.

Ecology, Monitoring and Management of Carpophilus Beetle: A New Invasive Pest of Tree Nuts in California

Figure 2. Adult Carpophilus truncatus (blue circles) inside of a hull-split almond (photo by J. Rijal.)

Growers and PCAs should be on the lookout for a new pest called carpophilus beetle (Nitidulidae: Carpophilus truncatus) (Figures 1 and 2).

Figure 1. Adult Carpophilus truncatus as seen from the (a) dorsal, (b) ventral, (c) left lateral and (d) anterior end (photos by Sarah Meierotto, UC Riverside.)

Damage occurs when adults and larvae of this pest feed directly on the developing kernel, causing reductions in both crop yield and quality. This pest was initially found infesting almonds and pistachios in the northern and central part of the San Joaquin Valley, and a broader survey is now underway to verify the extent of its spread in California.

Figure 2. Adult Carpophilus truncatus (blue circles) inside of a hull-split almond (photo by J. Rijal.)

The carpophilus beetle is recognized as one of the top two pests of almond production in Australia, where growers typically experience 2% to 5% infestation, but it can be closer to 30% to 40% in more extreme cases (Madge 2022). In addition to feeding on new crop nuts, the carpophilus beetle can also likely facilitate the spread of Aspergillus fungi that can lead to the production of aflatoxins, which are known human carcinogens that are heavily regulated in key markets.

Biological and chemical control options are very limited or unknown. As such, the key to managing carpophilus beetle is crop sanitation since this pest overwinters on remnant mummy nuts in the orchard like navel orangeworm (Pyralidae: Amyelois transitella) (NOW). Growers and PCAs can monitor carpophilus beetle by directly inspecting mummy or new crop nuts, although there are no known economic thresholds for this pest.

Global Distribution and Initial Detections in California
Carpophilus beetle has been well-established in Australia for over 10 years, where it is considered a key pest of almonds. More recently, the beetle was reported from walnuts in Argentina and Italy. Carpophilus truncatus is a close relative to other beetles in the genus Carpophilus, such as the driedfruit beetle (C. hemipterus) that is known primarily as a postharvest pest of figs and raisins in California.

In California, populations of carpophilus beetle were first detected in September 2023 in almond and pistachio orchards in Madera and Kings counties, respectively. Pest identification was subsequently confirmed by CDFA. A broader survey of orchards throughout the San Joaquin Valley is now underway to determine the extent of the outbreak as well as confirm additional hosts such as walnuts and pecans. So far, almond or pistachio orchards infested by carpophilus beetle have been confirmed in Stanislaus, Merced, Madera and Kings counties, suggesting the establishment of this new pest is already widespread. In fact, some specimens from Merced County were from collections that were made in 2022, suggesting the pest has been present in the San Joaquin Valley for at least a year already.

Host Plants
While in Australia the carpophilus beetle is primarily found in and around almond orchards, alternative host plants have been identified that the beetle can use for feeding and reproduction. These include Brazil nuts (Lecythidaceae: Bertholletia excelsa), candlenuts (Euphorbiaceae: Aleurites moluccanus), cashews (Anacardiaceae: Anacardium occidentale), pistachios (Anacardiaceae: Pistacia vera), quandong seeds (Santalaceae: Santalum acuminatum), walnuts (Juglandaceae: Juglans regia), acacia seeds (Fabaceae: Acadia spp.), pine nuts (Pinaceae : Pinus spp.), desiccated coconut flesh (Arecaceae: Cocos nucifera), sunflower seeds (Asteraceae: Helianthus annuus) and granulated or powdered pollen (Madge 2022). While adults have also been shown to survive on dried apricots (Rosaceae: Prunus armeniaca), dates (Arecaceae: Phoenix dactylifera) and cup mushrooms (Pezizaceae: Peziza spp.), the larvae are unable to complete development on these hosts (Madge 2022). In California, this pest has so far been confirmed feeding on almonds, pistachios and walnuts.

Seasonal Ecology
Carpophilus beetles overwinter in remnant nuts (i.e., mummy nuts) that are left in the tree or on the ground following the previous year’s harvest. In contrast to NOW, carpophilus beetles tend to prefer remnant nuts on the ground, likely due to increased moisture, and they generally tend to forage closer to ground level (Madge 2022). That said, carpophilus beetles can also be found in tree mummies as well.

The beetles become active when temperatures and photoperiod increase in the spring. They can have multiple generations per year, although there is no information available on the upper and lower temperature thresholds for this pest, much less degree-day requirements. While more research is still needed to better characterize beetle activity in the spring prior to hull split, they likely make use of mummy nuts as a reproductive host during that early part of the season, similar to NOW.

At hull split, carpophilus beetles will move onto new crop nuts, although some fraction of the population does remain on mummies all year. Prior to egg deposition on new crop nuts, the adult beetles will chew through the shell (Figure 3) and/or feed on the kernel, which is thought to both facilitate larvae access to the developing kernel, as well as inoculate the nut with a symbiotic gut-associated yeast that likely help the larvae feed (Madge 2022). Recent studies in Australia have demonstrated adults prefer to oviposit onto nuts previously fed upon by other carpophilus beetles and/or inoculated with yeasts (Madge 2022). While carpophilus beetles have been found infesting nuts throughout the tree, they tend to concentrate in the lower canopy (Madge 2022). This contrasts with NOW, which tend to initially infest nuts higher in the tree canopy.

Figure 3. Prior to laying eggs, adult carpophilus beetle can chew a hole through the shell (yellow circle) (photo by J. Rijal.)

Following egg deposition, the larvae that emerge feed on the developing kernels, leaving the almond kernel packed with a fine powdery mix of nutmeat and frass that is sometimes accompanied by an oval-shaped tunnel (Figures 4 and 5).

Figure 4. Almond with fine powdery frass due to infestation by Carpophilus truncatus (photo by H. Wilson.)

Figure 5. Carpophilus truncatus feed directly on the nut kernel, which can sometimes result in a distinct oval-shaped tunnel (photo by H. Wilson.)

Damage from carpophilus beetle may be confused with NOW and/or ants, but these can be differentiated. For example, NOW tends to feed all over the kernel (rather than tunneling) and produces a darker and larger type of frass (i.e., excrement) along with webbing (Figure 6).

Figure 6. NOW damage to almonds results in larger, darker frass as well as webbing (photo by Jack Kelly Clark, courtesy UC Statewide IPM Program.)

In contrast, ants tend to chew through the skin of the kernel and feed primarily on the white nutmeat, leaving the papery skin behind (Figure 7).

Figure 7. While ant damage to almonds can also produce a fine white powder, damage to the nut tends to be broad and superficial in contrast to carpophilus beetles which tunnel into the nut (photo by Jack Kelly Clark, courtesy UC Statewide IPM Program.)

Ant feeding is also associated with the presence of a fine white powder (similar to carpophilus beetle damage) that can be seen while sampling in the field but disappears in the hulling/shelling process prior to the inspection of processed kernels (Figure 8).

Figure 8. Damage from ants can result in a fine white powder like what is produced by carpophilus beetles (photo by D. Haviland.)

Monitoring
Monitoring for carpophilus beetle is currently limited to direct inspection of hull-split nuts for the presence of feeding holes and/or larvae or adult beetles. Given their tendency to infest nuts lower in the canopy, this is a good area to focus on for initial inspection of new crop nuts. That said, to get an idea of average overall infestation levels, it is best to take a sample from freshly shaken new crop nuts on the ground at harvest, since this is more representative of the entire tree canopy.

Over the winter, mummy nuts can also be inspected for the presence of carpophilus beetles. Initial studies from Australia suggest that monitoring points should be at least 200 yards apart (Madge 2022). While no specific economic thresholds have been developed, summer infestations on new crop nuts tend to reflect the distribution of infested mummy nuts during the winter.

A lure based on male-produced aggregation pheromone from three related species of Carpophilus spp. was previously developed in Australia and used for monitoring as well as an attract-and-kill strategy in stone fruit orchards (Hossain 2018). With the arrival of C. truncatus in Australia, this monitoring and management strategy was tested in almond orchards. Unfortunately, in its current form, this trap and lure system is not very attractive to C. truncatus, and so it has no utility for either monitoring or as a control strategy (Hossain 2018).

As such, Australian researchers are now working to develop a pheromone lure that is specific to C. truncatus as well as one or more co-attractants based on host plant volatiles and/or gut-associated yeasts. Preliminary studies have shown a lot of promise, and this new lure may soon provide a better monitoring tool for growers, PCAs and researchers, but it is not yet commercially available (Madge 2022). Initial studies with the new species-specific pheromone lure have demonstrated that carpophilus beetles appear to forage mostly at ground level, and beetle catch with the candidate pheromone was improved when traps were moved from the tree canopy (i.e., hung at about 5 ft height) to ground level (Madge 2022).

Biological Control
While there are certainly parasitoids and predators that attack Nitidulids and species in the genus Carpophilus in particular, very little is known about the specific natural enemies of C. truncatus, much less their efficacy in an orchard setting. Much more information exists on the parasitoids of C. hemipterus, the driedfruit beetle, a key pest of figs and stone fruit in California. Key parasitoids of C. hemipterus include the Encyrtids Zeteticontus spp. and Cerchysiella spp. as well as a Proctotrupid Brachyserphus abruptus, all of which attack the larvae. Additional parasitoids include the Braconid Microctonus nitidulidis and the Bethylid Pseudisobrachium flavinervis, which attack the adult and pupal life stages, respectively. These parasitoids have a wide host range and may attack C. truncatus, but more research will be necessary to confirm this as well as their efficacy for population control.

Documented predators of Carpophilus spp. include the Staphylinid Atheta coriaria and the Reduviid Peregrinator biannulipes. Other generalist predators commonly found in tree nut orchards (e.g., spiders, lacewings) may also contribute to biological control of C. truncatus, but again more research is needed to further characterize this, much less determine their impacts on C. truncatus specifically.

In the absence of specialist parasitoids or predators, researchers in Australia have focused on the use of entomopathogenic fungi (EPF) for carpophilus beetle, particularly the EPF Beauvaria bassiana. While preliminary studies found that in some cases B. bassiana could cause up to 70% mortality of C. truncatus larvae (and to a lesser extent adults), the use of this EPF under field conditions is still being developed (Madge 2022).

Finally, vertebrates like birds and rodents may provide some degree of control by consuming or damaging remnant mummy nuts infested by carpophilus beetle, but the impacts of this remain unclear in California orchards.

Chemical Control
The ability to control carpophilus beetle with insecticides is limited, primarily due to challenges with spray coverage. Most of the beetle’s life cycle is spent protected inside the nut, with relatively short windows of opportunity available to spray the adults while they are exposed. In Australia, the use of bifenthrin for control of carpophilus beetle has produced inconsistent results, and experiments to improve coverage with various adjuvants did not lead to improved control (Madge 2022). Furthermore, the continued presence of carpophilus beetles on remnant mummy nuts throughout the season presents an additional challenge to control with insecticides.

Cultural Control
In the absence of clear chemical or biological control strategies, the most important tool for managing carpophilus beetle is crop sanitation. In Australia, this is currently the primary method for managing this pest. While carpophilus beetle can be found overwintering on remnant mummy nuts both in the tree canopy and on the ground, they tend to prefer ground mummies, likely due to elevated moisture conditions. After removing remnant mummy nuts from trees, it is critical all ground mummies be thoroughly broken apart and destroyed. Simply disking mummies under the soil will not be effective, since research in Australia demonstrated even when mummies are buried as deep as 3 ft down, adult carpophilus beetle can still survive and crawl up to the soil surface (Madge 2022). As such, make sure to use a flail mower to thoroughly destroy mummy nuts. It might need a double pass to ensure all nuts are shredded.

Carpophilus beetle is a new pest in California that will need to be addressed by both researchers and growers alike. Within the research community, new research and extension activities are being developed by UCCE personnel in collaboration with their counterparts in Australia. Until more is known about this pest in California, growers are advised to monitor for its presence and follow the Australian model of focusing on winter sanitation as the primary means for its control.

If you suspect that you have this beetle in your orchard, please contact your local UCCE Farm Advisor (https://ucanr.edu/About/Locations/), County Agricultural Commissioner (https://cacasa.org/county/) and/or the CDFA Pest Hotline (https://www.cdfa.ca.gov/plant/reportapest/) at 1-800-491-1899.

Selected References
Hossain, M. 2018 “Final Report – Management of Carpohilus Beetle in Almond” Project Code AL15004, Horticulture Innovation Australia, North Sydney, Australia.
https://www.horticulture.com.au/growers/help-your-business-grow/research-reports-publications-fact-sheets-and-more/al15004/
Madge, D. 2022 “Final Report – An Integrated Pest Management program for the Australian almond industry.” Project Code AL16009, Horticulture Innovation Australia, North Sydney, Australia.
https://www.horticulture.com.au/growers/help-your-business-grow/research-reports-publications-fact-sheets-and-more/al16009/

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

Useful Soil Maps in Microirrigated Orchards

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

Fertilizer Planning for Forage Crop Production

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

LAMP as a New Tool for Testing Grapevine Red Blotch Virus

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

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

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

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

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

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

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

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

Sampling of lower-canopy leaves for GRBV testing.

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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


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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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