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Foliar Feeding of Plant Nutrition

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According to studies from the United Nations, the world population will increase by 1 billion people over the next decade, reaching an estimated 8.6 billion people by 2030. By 2050, Earth will have 9.8 billion inhabitants. This enormous population growth will be coupled with a corresponding increase in demand for food. Projections from the FAO have indicated that agricultural production worldwide will have to increase by an estimated 50% by 2050 to meet the needs of a growing population.

Proportionately, agricultural land is becoming scarce. Therefore, one of the biggest challenges is to improve the yield, quality and shelf life of crops by using the best fertilizers, applied in very precise doses and with proven methods. One of the best procedures to reach those goals is foliar application of nutrients in customized doses based on critical timing for each crop.

Foliar feeding is the entry of small amounts of liquid fertilizer through the surface of plant tissue. This can allow for rapid nutrient utilization by the plant. Foliar feeding provides the consultant and applicator the ability to blend the fertilizer with other products such as pesticides and micronutrients.

Current formulations of liquid fertilizers are believed to penetrate mostly the transcuticular pores on foliage, which are open virtually all the time compared to stomata. Nutrients also enter stomata, but these often are closed due to environmental stresses and darkness. Most stomata are located underneath leaves away from fertilizer spray patterns. Drawbacks to foliar feeding include the inability to apply large amounts of N, phosphorous (P) and potassium (K) without potentially burning the foliage and possibly scarring the fruit. Therefore, frequent applications of the right fertilizer and fertilizer blends at a low volume are required to maintain optimum tissue levels of key nutrients, resulting in consistent plant growth and functions like the production and transference of assimilates.

 

Foliar Nutrient Applications

Selecting the right fertilizers will continue to be an important part of the equation to improving yield, quality and shelf life of crops. How we apply the nutrient is also vital. Let us examine foliar spraying more closely. Crops treated with foliar spray tend to have a higher nutritional value. Improved Total Soluble Solids (TSS) have contributed to increased sugar, vitamins, minerals and proteins in the harvested crops. It is not enough just to grow more crops, but those crops must have better nutritional value for us as well.

We can see very positive effects by examining just one mineral nutrient such as K. K regulates plant functions and increases nitrogen efficiency. By increasing the saline concentration of the cells, the plants are better able to resist frosts. Applied at the right time and in the correct amount, K promotes the development of a stronger cell structure, which allows the crops to better withstand drought, disease and pests. A better internal structure promotes a prolonged shelf life. In many of our soils, we lack adequate available K to meet the crop demands. One way to meet those phenological growth stages is to supplement K sprays. This could affect the production levels and quality of numerous crops such as tuber fill and skin condition in potato, nut meat density in almond and even oil production in mint.

Originally, it was thought that using foliar nutrient applications was only appropriate when a nutritional deficiency was present. This is definitely not the case. Ongoing scientific research has consistently shown that periodic applications of certain nutrients in various crops can have a positive effect on both quantity and quality of fruits, nuts, vegetables and grain crops. Although most supplemental nutrients are absorbed through the root system, it is also important to note that leaves (and to a lesser degree stems,) flowering plant tissues and even fruit surfaces can absorb limited amounts of nutrients. It is crucial to understand which nutrients can be supplied effectively by foliar applications if the soil-supplied nutrients are inadequate or impaired in some way.

In agriculture, our scientists have identified many cases where nutrient supplementation using a foliar application may be considered more effective than an in-crop topdressing application. One such situation that has had success is during late stages of fruit development where Ca sprays applied through the season to apples has shown ability to combat Ca deficiency. This deficiency is directly connected with the physiological disorder called bitter pit. The application of Ca making direct contact with the fruit has shown good results in controlling this disorder. Also, Ca being sprayed on cherries preharvest amounted to less cracking in fruit during late maturation stages.

Research trials done in almonds have also demonstrated where a foliar-applied nutrient can have a huge impact on a crop. Knowing that fruit set is crucial in almond trees, it has been proven that micronutrients, zinc (Zn) and especially boron (B), have a significant impact on fruit set as well as on fruitlet abscission. In several fruit trees, it has been reported that even foliar spraying of one or both elements has improved productivity. In one experiment, Nonpareil trees were sprayed at full blossom with boric acid at 0.2%, Zn-50 at 0.3% and, with the combination of these micronutrients as a separate treatment, compared with a non-sprayed control. Fruit set of the B treatment was 27.7% and Zn was 22.2%. However, the B + Zn combination produced a significantly higher fruit set (38.1%) (Sotomayor et al., 2002).

 

Additional Benefits

As consultants, we must understand that at times only a small amount of a specific nutrient is required, and due to low soil mobility of the nutrient, it is actually more efficient to supply the small amount of needed nutrient as a foliar application. This can be the case for both macronutrients and micronutrients.
There can also be other conditions that cause a nutrient to be less available to crop roots. One example is cool, excessively wet conditions on an alkaline soil (e.g. pH >8.0) where iron is less available to certain crops (e.g. iron chlorosis of citrus trees) in heavy wet clay. Even if supplemental iron fertilizer has been applied to the soil before or at planting, it may not be available through the roots. This can also be true for numerous other crops. By monitoring the trees and observing patterns, we can offset the chlorosis in our crop with a supplemental iron foliar application. Spray applications of elements such as iron, zinc, copper and manganese may have to be repeated every three weeks when we get deficiencies of these nutrients.

Many studies include yield data collected after applying foliar sprays of various fertilizer solutions, not only to nutritionally deficient crops, but even to crops that have adequate levels of nutrients such as potassium. Crops including wheat, almonds, tomatoes, citrus, cucurbits, pome fruits and rice among many others react positively to certain nutrient sprays even when adequate soil nutrient levels are present.

A good reference to the benefits of foliar spraying is in citrus research. After years of trials addressing HLB or Huanglongbing citrus greening disease in Florida, it was determined that a foliar nutrition approach to the disease was a very viable option (www.citrusbr.com.br/download/Foliar_nutrition_forHLB.pdf). The nature of the disease restricts the ability of the plant to adequately take up nutrients though its damaged root system. Finding an alternative way to increase plant nutrition and prolong the life and productivity of the infected tree was crucial. Numerous combinations and timings were trialed.

Trials show that in the case of one nutrient combination, potassium nitrate feeding increased yields in citrus. Foliar applications with potassium nitrate have proven to be highly efficient in fulfilling the potassium requirements for many crops. The combination of potassium and nitrate in this fertilizer has been found to be beneficial in improving fruit size, dry matter, color, taste and integrity as well as resistance to biotic and abiotic stresses for citrus and tomato fruit. Moreover, the integration of potassium nitrate in routine management or in specific growth stages resulted in remarkably positive benefits to cost ratio.

 

Take Caution

In our management plan, caution should be used when foliar application might be more efficient but not practical. When a foliar application is relatively effective, but we cannot supply the needed nutrient in one application, multiple applications would be needed, spaced out sufficiently and timed as often as once a week. Multiple applications in field crops can be expensive due to fuel, equipment and labor costs, or where there may not be sufficient time to apply enough of the needed nutrient. We must measure the economics and limitations when determining our fertilizer plan.

In the case of a severe P deficiency, there may not be time to apply sufficient applications of low rates of foliar P to be effective. The application costs can become excessive based on the return in production. In this situation, it may be better to realize there is not much that can be done in the current season to correct the issue. Recognizing the deficiency, the preferred course of action is to apply sufficient P fertilizer to the soil prior to the planting of subsequent crops to correct the P deficiency.

If we continue to use phosphorus P as an example, we also need to understand foliar P efficiency and when it does have a fit in our fertilizer program. Foliar-applied nutrients have the benefit of being 4 to 30 times more efficient, and there is no risk of groundwater contamination. Studies using labeled P on apple, cherry, corn, tomato, potato and bean crops have shown that as much as 12 to 14% of the total P can be supplied by multiple foliar sprays. Since P can be very immobile in the soil, foliar applications can be up to 20 times more effective than soil applications.

Recognizing another nutrient use as a foliar, we can see that some foliar N sprays compared to soil applications of N include lower application rates and the ease of obtaining timely, uniform applications. With attention to best-use guidelines, the efficiency of foliar-applied N may be optimized at nearly 95 to 100%. Based on the foregoing information, if the recovery of soil-applied N can be impaired to as low as 15 to 62%, it can then be concluded by the method of estimation that foliar-applied N has an efficiency of 1.3 to 1.6 times soil-applied N at the low end and seven times at the upper end. If foliar-applied N can be up to seven times more efficient than soil-applied N, then on a pound-for-pound basis, it makes sense that we could use this information to prevent N loading to ground water. In crops such as citrus, cherries and wine grapes, we may be able to achieve 100% of the needed N by foliar application. This is not always the case, but understanding your N efficiency in your soils may prompt the need for a supplemental foliar N application. Mature, low-N-requiring crops could acquire 40 to 50 pounds N with foliar application alone. This in no way warrants replacing all of our soil applied N in all crops by using foliar.

 

Conclusion

In summary, it should be pointed out that we have barely scratched the surface of all the features and benefits of foliar nutrition sprays. Get to know your crop and soils as well as the correlation between total nutrients and available nutrients. Understand if your crop has shown positive response to foliar treatment even when adequate nutrient levels are in the soil. Ask yourself: when do soil conditions prevent adequate nutrient uptake? What conditions should I be aware of to optimize foliar applications (rain, temperature, wind, sprayer capability, pH of my solution, humidity, physiological growth stage, activity of plant parts like stomate, additives such as pesticides or adjuvants and antagonist as well as synergistic reactions with other nutrients.) Study absorption rates and what affects them. Like all other things in agriculture, foliar nutrition is a tool.

 

References

Sotomayor, C., Silva, H. and Castro, J. (2002). Effectiveness of boron and zinc foliar sprays on fruit setting of two almond cultivars. Acta Hortic. 591, 437-440, https://doi.org/10.17660/ActaHortic.2002.591.67.

Impact of Some Biostimulants in Improving Strawberry Yields

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Microbial and botanical biostimulants have been used for promoting plant growth and to help plants withstand various pests, diseases or environmental stressors. While macro and micronutrients are necessary for plant growth and optimal yields, biostimulants play multiple roles by increasing the bioavailability of nutrients, improving nutrient and water absorption, and protecting plants from pestiferous organisms, either through direct antagonism or by triggering plants’ defense mechanisms (Berg, 2009). In addition to improving health and yields, biostimulants are also known to increase nutritional quality (Parađiković et al., 2011; Fierentino et al., 2018).

Multiple field studies in California demonstrated the potential of biostimulants and soil amendments in improving yields in tomato (Dara, 2019a; Dara and Lewis, 2019) and strawberry (Dara and Peck, 2018; Dara, 2019b). As the knowledge of biostimulants and their potential for sustainable agriculture is expanding, there has been a steady introduction of biostimulant products in the market warranting additional studies. A study was conducted to evaluate the potential of different biostimulant materials on strawberry growth, health and fruit yields.

The study measured the impact on growth, health and yield parameters for each treatment.

 

Methodology

This study was conducted in an experimental strawberry field at the Shafter Research Station during 2019-2020. Cultivar San Andreas was planted on October 29, 2019. No preplant fertilizer application was made in this non-fumigated field, which had both Fusarium oxysporum and Macrophomina phaseolina infections in the previous year’s strawberry planting.

Each treatment was applied to a 300-foot-long bed with single drip tape in the center and two rows of strawberry plant on either side. Sprinkler irrigation was provided immediately after planting along with drip irrigation, which was provided one or more times weekly as needed for the rest of the experimental period. Each bed was divided into six 30-foot-long plots representing replications with 18-foot buffers in between. This study included both biostimulant and nutrient supplements, but this article presents data from the biostimulant treatments only. Treatments were applied either as fertigation through the drip system using a Dosatron, or sprayed over the plants with a handheld garden sprayer.

 

Treatments

The following treatments were evaluated in this study:
Grower Standard (GS): Between November 6, 2019 and May 9, 2020, 1.88 qt of 20-10-0 (a combination of 32-0-0 urea ammonium nitrate and 10-34-0 ammonium phosphate) and 1.32 qt of potassium thiosulfate were applied 20 times at weekly intervals through fertigation. This fertilizer program was used as the standard for all treatments except for the addition of biostimulant materials.

GS + Abound: Transplants were dipped in 7 fl oz of Abound (azoxystrobin) fungicide in 100 gal of water for four minutes immediately before planting. Transplant dip in a fungicide is practiced by several growers to protect from fungal diseases and is considered as another standard in this study.

GS + Str10 + Rhizolizer: Str10 (Wickerhamomyces sp.) was applied at 5 fl oz/ac with molasses at 10 fl oz/ac immediately after planting and Rhizolizer (Trichoderma harzianum and Bacillus amyloliquefaciens) at 3 fl oz with a food source blend at 10 fl oz two weeks after Str10 application through the drip system. The same pattern was repeated starting from mid-February 2020. From February to May, 6 fl oz/ac of Rhizolizer was applied with 20 fl oz/ac of food source once a month. Str10 is an unregistered product with yeast that is expected to help with nutrient uptake and phosphorous mobilization for improved plant vigor and yield. Rhizolizer is expected to solubilize soil nutrients and improve crop growth and yield.

GS + ON-Gard: 32 fl oz of ON-Gard was applied every two weeks through the drip system from planting until canopy development and then sprayed in 50 gpa. ON-Gard is expected to increase the nutrient use efficiency and decrease abiotic stress to the plants.

GS + ON-Gard + RootShield Plus: 32 fl oz of ON-Gard (soy protein-based) was applied every two weeks through the drip system from planting until canopy develops and then sprayed in 50 gpa. RootShield Plus WP (T. harzianum and T. virens) was also applied at 2 lb/ac through drip immediately after planting with 1 lb/ac at the end of November and again at the end of December 2019. RootShield is a biofungicide expected to protect strawberry from phytopathogens and improve water and nutrient uptake.

GS + CropSignal: CropSignal was applied at 10 gpa six days before planting and at 5 gpa 30 days after transplanting through the drip system. CropSignal is a carbon-based nutrient formula containing botanical extracts along with cobalt, copper, manganese and zinc, and is expected to support the growth and diversity of beneficial aerobic soil microbes for improved soil structure, water retention, nutrient cycling and plant protection.

Parameters observed during the study included canopy growth (area of the canopy) in January, February and March; first flower and fruit count in January; leaf chlorophyll and leaf nitrogen (with chlorophyll meter) in January, February and May, fruit sugar (with refractometer) in March and May; fruit firmness (with penetrometer) in March, April and May; severity of gray mold (caused by Botrytis cinereae) and other fruit diseases (mucor fruit rot caused by Mucor spp. and Rhizopus fruit rot caused by Rhizopus spp.) 3 and 5 days after harvest (on a scale of 0 to 4 where 0=no infection; 1=1-25%, 2=26-50%, 3=51-75% and 4=76-100% fungal growth) in March and May; sensitivity to heat stress (expressed as the number of dead and dying plants) in May; and fruit yield per plant from 11 weekly harvests between March 11 and May 14, 2020. Data were analyzed using analysis of variance in Statistix software and significant means were separated using the Least Significant Difference test.

 

Results and Discussion

The impact of treatments varied with different measured parameters. The interactions among plants, beneficial and pathogenic microorganisms in the crop environment, the influence of environmental factors and how all these biotic and abiotic factors respond to various biostimulant inputs can be very complex. The scope of this study was only to measure the impact on growth, health and yield parameters, not to investigate those complex interactions.

The canopy size does not always correspond with yields but could be indicative of stresses and how the plant is responding to them in the presence of treatment materials. Plants in some treatments had significantly larger canopy size in January and February, but the grower standard plants were significantly larger than the rest by March. Leaf chlorophyll and nitrogen contents were significantly different among treatments only in January where the grower standard plants had the lowest and the plants that received CropSignal had the highest. When the counts of the first onset of flowers and developing fruits were taken in January, plants that received ON-Gard alone had the highest number followed by the CropSignal and Abound treatments.

A similar trend was also seen for the average fruit sugar content. There was no statistically significant difference in the average fruit firmness among the treatments, although the value was numerically higher for the fruits in the CropSignal treatment. Severity of the gray mold, which occurred at low levels during the observation period, did not statistically differ among the treatments, but it was numerically higher in fruits from the grower standard plots. However, the severity of other diseases was significantly different among various treatments with the highest level in fruits from the grower standard and the lowest in fruits from plants that were treated with Abound. Temperatures were unusually high during the last week of May, and several plants exhibited heat stress and started to die. The number of dead or dying plants on May 28 was the highest grower standard and the lowest in Str10 + Rhizolizer and Abound treatments, although the differences were not statistically significant.

Table 1: The trial showed significant differences in marketable and unmarketable fruit yields among treatments.
There were significant differences in marketable and unmarketable fruit yields among treatments (see Figure 1.) Highest marketable yields were seen in ON-Gard and CropSignal treatments followed by the grower standard, ON-Gard + RootShield and Str10 + Rhizolizer. Transplant dip in a fungicide seems to have a negative impact on fruit yields as observed in the current study or earlier studies (Dara and Peck, 2017 and 2018; Peck unpublished data). While the grower standard had the highest amount of unmarketable fruits, the Str10 + Rhizolizer treatment had the lowest in this study. Fruit yield and some of the observed parameters appeared to be better in the grower standard with no fungicide or biostimulant

treatments, which has also been seen in some earlier strawberry studies.

Compared to the grower standard, marketable fruit yield improvement was seen only in ON-Gard and CropSignal treatments. However, marketable fruit yield was higher in all treatments compared to the grower standard with Abound. Sometimes, natural balance of the nutrients, organic matter and microbial community in the soil might result in optimal yields in the absence of pathogens or other stressors. However, it is very common to use fungicidal treatments or add biological or synthetic amendments to protect from potential threats and improving yields. These results help understand the impact of various biostimulant materials and warrant the need to continue such studies under various environmental, crop and soil conditions.

Field assistant Tamas Zold measures fruit weight to further determine impacts on treatments.

 

References

Berg, G. 2009. Plant-microbe interactions promoting plant growth and health: perspectives for controlled use of microorganisms in agriculture. Appl. Microbiol. Biotechnol. 84: 11-18.

Dara, S. K. 2019a. Effect of microbial and botanical biostimulants with nutrients on tomato yield. CAPCA Adviser, 22(5): 40-45.

Dara, S. K. 2019b. Improving strawberry yields with biostimulants: a 2018-2019 study. UCANR eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=31096.

Dara, S. K. and D. Peck. 2017. Evaluating beneficial microbe-based products for their impact on strawberry plant growth, health, and fruit yield. UCANR eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=25122.

Dara, S. K. and D. Peck. 2018. Evaluation of additive, soil amendment, and biostimulant products in Santa Maria strawberry. CAPCA Adviser, 21 (5): 44-50.

Dara, S. K. and E. Lewis. 2019. Evaluating biostimulant and nutrient inputs to improve tomato yields and crop health. Progressive Crop Consultant 4(5): 38-42.

Fiorentino, N., V. Ventorino, S. L. Woo, O. Pepe, A. De Rosa, L. Gioia, I. Romano, N. Lombardi, M. Napolitano, G. Colla, and Y. Rouphael. 2018. Trichoderma-based biostimulants modulate rhizosphere microbial populations and improve N uptake efficiency, yield, and nutritional quality of leafy vegetables. Frontiers in Plant Sci. 9: 743.

Parađiković, N., T. Vinković, I. V. Vrček, I. Žuntar, M. Bojić, and M. Medić-Šarić. 2011. Effect of natural biostimulants on yield and nutritional quality: an example of sweet yellow pepper (Capsicum annuum L.) plants. J. Sci. Food. Agric. 91: 2146-2152.

(The author would like to thank BioWorks, Inc., Fauna Soil Production and Locus Agricultural Solutions for the financial support of the study and Marjan Heidarian Dehkordi and Tamas Zold for their technical assistance.)

Assessing the Accuracy and Precision of Commercial Ag Laboratories

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Soil chemical analysis is the cornerstone of an effective nutrient management program. Without a reliable soil test, significant miscalculations in fertilizer recommendations can occur, leading to drastic effects on profitability and the environment as a result of under and overfertilization and soil amendment recommendations.

Despite the large number of analytical commercial laboratories serving California agriculture, deciding which laboratory to send a sample to can be a daunting task. Unfortunately, there are no public data reporting the accuracy of the analysis performed by agricultural laboratories, and there isn’t a “true” certification program in the U.S. Although a lab may participate in a proficiency program such as the Agricultural Laboratory Proficiency (ALP) program or the North American Proficiency Testing (NAPT), these programs are not mandatory, nor do they certify lab quality. Therefore, laboratories are chosen based on “word of mouth” and prices, which can vary significantly.

Because of the absence of data, growers, farm managers, consultants, environmentalists and even researchers are left without a reliable means by which to select a testing laboratory. A study was conducted in 2019 to assess the performance of soil testing laboratories.

 

Accuracy and Precision Assessment

Four reference soil samples from the ALP program were submitted to eight commercial Ag laboratories in the Western U.S. (seven in California and one in Idaho) for typical fertility analyses. In most cases, fertility “package analyses” offered by the laboratories were chosen in order to optimize project funds. The same four reference soil samples were resubmitted two more times totaling three rounds, approximately three months apart each round, in order to assess the precision of each laboratory.

Methods and materials used for the laboratory accuracy assessment (photo courtesy A. Biscaro.)

Standard reference soil samples were selected from the ALP program archives, each previously analyzed by a minimum of 30 credible laboratories, in triplicate for each soil sample (totaling 90+ analyses per reference soil.) The median and median absolute deviation (MAD) of these 90+ analyses per reference soil were used to assess the accuracy and precision of the eight laboratories assessed in this study (method based on ALP consensus statistics.) While the accuracy assessment is focused on contrasting each analysis with the ALP medians, the precision assessment is focused on the variability of the analyses across the three rounds (same reference soils analyzed at different times.) Sample IDs were modified and submitted to each laboratory so they wouldn’t be aware of the objectives of the study. Names of laboratories are not disclosed to follow university policy; laboratories are referred as #1 to #8 for discussion purposes.

Each reference soil was analyzed for NO3, P, extractable K, Na, Ca and Mg, SO4-S, electrical conductivity (ECe), Cl, Ca, Mg, Na and B, pH and five micronutrients. Some labs provided additional analyses in their fertility package, such as soil organic matter, estimated and measured CEC and others, however, these were not used in this study since they were not performed by all laboratories. Nineteen analyses performed on four reference soils by eight laboratories three times equals a total of 1,824 analyses, or 228 per laboratory. While that is a rich dataset, trying to create a performance rank for the laboratories across all analysis types is quite challenging since there are multiple types of soil analyses, extraction methods and units. For that reason, performance standards used by the ALP program were applied to this project in order to assess the accuracy and precision of the analysis performed by the laboratories. Eight analyses were chosen for this assessment: Olsen P, extractable K, Ca and Mg, ECe, pH, sodium adsorption ratio (SAR) and DTPA Zn. For the purpose of accuracy assessment, each of these analyses were attributed a pass or failure score, totaling 96 scores per laboratory (8 analysis types, 4 reference soils and 3 rounds). The precision assessment was based on the relative standard deviation of each analysis across the 3 rounds.

Although all labs presented certain inaccuracy and imprecision, some stood out. Laboratories #2 and #8 were consistently inaccurate and imprecise, while laboratories #1 and #7 were the most accurate and precise. Laboratory #8 in particular presented the poorest performance for both accuracy and precision. Laboratories #3, #4, #5 and #6 presented varying accuracy and precision. These patterns of accuracy and precision are illustrated in the three graphs below.

 

Laboratory Results

Figure 1 illustrates the results for saturated paste pH for soil C. Listed is the pH median and the Median Absolute Deviation (MAD), with results for each lab and each round. Labs #1 and #7 were the most accurate over all rounds. Labs #2 and #8 had high bias, and lab #2 was imprecise (inconsistent). Due to funding limitations, only two rounds of samples were submitted to lab #7.

Figure 1. Soil pH analysis by the saturated paste method performed by eight commercial laboratories for soil C.

Results for exchangeable potassium analysis by ammonium acetate for soil C (Figure 2) illustrates a common occurrence of accuracy levels observed across most reference soils used in this study. Generally, labs #1 and #7 consistently reported results near the median.

Figure 2. Exchangeable potassium analysis by ammonium acetate extract performed by eight commercial laboratories for soil C (SRS-1604).

Results for Zn extractable by DTPA for soil A (Figure 3) show a general trend of all eight labs reporting higher Zn values relative to the median for this standard reference soil of 0.9 ppm. Labs #1, #4, #5 and #7 generally reported equivalent Zn concentrations for each round. Labs #2, #3, #6 and #8 were inconsistent across the three rounds. Lab #6 in particular reported values that varied by 300% across the three rounds.

Figure 3. Zinc analysis by the DTPA method performed by eight commercial laboratories for soil A (SRS-1809).

 

Need for Consistency

Besides the accuracy and precision parameters assessed in this study, it seems like consistency is an overall challenge for the lab industry: consistency of methods used for certain analyses, of how methods and units reported, and of the interpretation of the results (e.g. graphs illustrating sufficiency and deficiency ranges). Although it is the responsibility of the client to verify the methods used and request the most pertinent information for their application, many growers and farm managers are not familiar with the intricacies of soil analyses and nutrient management. Hence, providing an electrical conductivity analysis in 1:1 or 1:2 extraction instead of the standard saturated paste extract (ECe) can lead to misleading conclusions and inappropriate management decisions since the literature for most salinity thresholds for crop yields were defined with the saturated paste extract method. Another observation in regard to the analysis type is about the phosphorus extraction method used for soils with different pH, where some labs used the Olsen extraction method for soils with pH below 6.0, and others utilized the Bray P1 method.

Soil testing lab users in California and in the Western U.S. could benefit tremendously from a certification program designed to certify the accuracy and precision of all labs on a regular basis. Please feel free to contact the lead author directly for more detailed information about this study at (805) 645-1465.

The author wishes to acknowledges the following contributors to this article: Robert Miller, ALP Program Director, former Extension Soil Specialist UC Davis; Dirk Holstege, Director, UC Davis Analytical Laboratory (retired); UCCE Advisor Steve Orlof, Siskiyou County (in memorian); Tim Hartz, UCCE Vegetable Crops Specialist, UC-ANR (retired); Ben Faber, UCCE Advisor, Ventura County; Anthony Luna, UCCE Advisor, Ventura County; and Eryn Wingate, Agronomist, Tri-Tech Ag Products.

Fusarium Wilt of Tomato Race 3

Fusarium oxysporum f. sp. lycopersici (Fol) race 3 causes Fusarium wilt, a disease currently affecting most tomato-producing counties in California. Fol is divided into groups called races based on the ability to overcome plant resistances. Fol race 3 is the most recent race, which broke resistance to race 2. Fol race 3 was long restricted to the Sutter Basin, but began spreading in the early 2000s and is now present in every county with large-scale tomato production, making this one of the greatest economic threats to the industry.

Growers are seeking solutions for this damaging soil-borne disease. In this article, an issue overview as well as the latest information on Fusarium wilt race 3 spread, control and prevention is provided. The focus is on current research that is shining a light on new prospects for management of Fusarium wilt in tomato.

 

Key Characteristics

Bright yellow foliage on one or several shoots on an otherwise normal plant are the earliest symptoms, starting as early as 45 days after planting but typically occurring at about 60 days. The one-sided yellowing of a branch or whole plant can help distinguish this disease from other wilt pathogens (e.g. Verticillium) and other causes of chlorotic conditions (e.g. nutrient disorders) (Figure 1A). From time of initial symptoms to harvest, disease symptoms progress from shoot yellowing to branch death which leads to partial or entire canopy collapse (Figure 1B). Fruit in this exposed canopy develop sunburn and may rot. Another important diagnostic feature of Fusarium wilt of tomato is the presence of chocolate-brown vascular discoloration in the plant stem (Figure 1C). Vascular discoloration is also a symptom of Verticillium wilt, which can lead to misdiagnosis. At advanced stages of Fusarium wilt, the general canopy collapse is similar to other tomato diseases such as southern blight, bacterial canker and Fusarium crown rots. Because of the potential for misdiagnosis of Fusarium wilt, even by experienced scouters, it is prudent to submit plant samples to a diagnostics laboratory prior to making management decisions.

Figure 1. Symptoms of Fusarium wilt in tomato plants, shown here as a shoot with bright yellow and dying foliage (also known as “yellow flagging”) on an otherwise healthy plant (A), collapse of the vine (B) and chocolate-brown discoloration inside a stem (C) (photos by K. Paugh.)

 

Survival and Spread

Fusarium wilt race 3 occurs across the Central Valley from the Sacramento Valley region (Colusa, Sutter, Solano, Yolo and Sacramento) to the central San Joaquin Valley (San Joaquin, Stanislaus, Merced, Fresno and Kings) and, most recently, to the southern end of the San Joaquin Valley (Kern) (Figure 2). The pathogen is thought to move locally from field to field by hitching a ride in the soil and plant debris that cling to farm equipment. Hence, the increased movement of farm equipment across processing tomato regions may have facilitated spread of this disease.

Figure 2. California counties with documented cases of Fusarium wilt race 3, as highlighted in red.

Once present in the field, Fol can persist in dead tissue in the soil. Ongoing studies suggest that this pathogen can persist for at least two years in infested tomato tissue after incorporation. In addition, although Fol can only cause disease in tomatoes, it can infect many different non-tomato crops, including melons, pepper and sunflower, without causing any symptoms and survive in non-tomato crop residue in soil. Thus, Fol can feasibly be introduced into a field that has never had tomato, propagate on these silently-infected crops and cause severe losses in the first year the field is planted to tomato. Of note, there are also many Fusarium wilt diseases of rotation crops (e.g. Fusarium wilt of melon, watermelon and lettuce,) but these Fusarium wilts are all caused by completely different pathogens. Therefore, if you have Fusarium wilt of melon in your field, this does not mean you will get Fusarium wilt in your tomatoes.

 

Management

Overview of IPM for Fusarium wilt
The most effective tool for Fusarium wilt management is preventing pathogen introduction. If introduced into a field, the disease can usually be successfully managed with resistant cultivars (F3 cultivars), although there are some caveats in F3 efficacy. If F3 cultivars are not available for management, pathogen-tolerant cultivars and early-season chemical management options are also available. Crop rotation can help reduce pathogen pressure and reduce risk that an F3 resistance-breaking race will emerge (race 4).

Effective management of Fusarium wilt requires accurate diagnosis. As noted above, there are many diseases that look like Fusarium wilt, and at present there is no way to differentiate these diseases in the field. Before developing a Fusarium wilt management plan, it is critical to submit samples for analysis by a diagnostic lab. Ideally, growers could have their soils tested for Fusarium wilt prior to planting. Although there are no commercial soil testing services available, there are preliminary soil testing tools under development at UC Davis (contact C. Swett for more information.)

Management with genetic resistance
Fol race 3 resistant cultivars, called F3 cultivars, typically develop no disease and are an excellent management tool. The tomato industry has worked hard to overcome challenges in F3 cultivar quality, yield and seed availability. In addition, certain F2 cultivars are “tolerant” of Fusarium wilt race 3 in that their yield does not appear to be significantly impacted in infested fields. Fusarium wilt tolerance is not a listed trait for existing commercial cultivars, but this information is available through seed dealers.

In some cases, F3 cultivars develop Fusarium wilt due to Fol race 3. This is typically attributed to either the presence of off-types (when incidence is below 2%) or environmental stresses (when incidence is higher.) Abiotic and biotic stresses appear to play a role in influencing stability of resistance, and recent studies have demonstrated that salt stress can compromise F3 resistance, leading to Fusarium wilt development in up to 30% of F3 plants in a field. While the role of various stresses in mediating F3 resistance is not well-characterized, research in this area is ongoing. Management of these stresses can help maintain the efficacy of host resistance against Fol race 3.

Chemical control pre- and post-planting
Fusarium wilt race 3 is notoriously difficult to control once established in soil. Although host resistance is the gold standard for management, F3 cultivars are not always available. Chemical management may function as a short-term alternative. Recent studies have shown promising results for pre-plant fumigation with K-Pam HL (AMVAC Corporation) at 30 gal/A or higher (maximum rate of 60 gal/A) for optimal efficacy (Figure 3, see page 6).

Figure 3. Results for a 2019 small plot trial at the UC Davis Plant Pathology research farm on the efficacy of the drip-applied fumigant, K-Pam HL, against Fusarium wilt of tomato.

Crop rotation
Crop rotation is a common recommendation for management of host-specific pathogens like Fusarium wilt because non-host crops suppress pathogen propagation and survival. However, the efficacy of this method relies on the inability of the pathogen to infect rotation crops. It has been discovered that Fol race 3 can infect many crops without causing symptoms, which may reduce the effectiveness of crop rotation. Several rotation crops were found to be poor hosts and were suppressive to pathogen build up in soils; these include cotton, bean crops (i.e. garbanzo, fava, lima and green bean), grass crops, including wheat and potentially corn and rice (poor hosts, not field tested), and onion (Figure 4, see page 6). These appear to be good crops to grow right after tomato and the year before planting to tomato. Pathogen-enhancing crops should be avoided when possible; these include pepper, melons, pumpkins, and sunflower.

Figure 4. Preliminary results for the effect of rotation crops on development of Fusarium wilt (FW) in tomato. Plots at the UC Davis Plant Pathology research farm were previously planted to a summer or winter rotation crop (or tomato) or left in chemical or unmanaged fallow (=weeds) in summer 2019 (A) or winter 2019-2020 (B), respectively

No free rides for pathogens
The most effective tool for Fusarium wilt management is preventing pathogen introduction. On farms where Fusarium wilt is not present, this is best achieved by only using equipment that remains within the farm. However, as this is not an option for most producers, a second option is to use a sanitation regime for shared equipment. There is limited information on which equipment is the most important to target for sanitation, but Fusarium has been found at high levels on harvesters which retain large amounts of plant debris (Figure 5). An assessment of critical control points on harvesters indicates that areas which only have contact with fruit have lower levels of microbes, whereas the chipper and other areas which come into contact with whole plant material have higher levels. Robust analysis of effective sanitation methods is lacking, but preliminary data indicates some efficacy of current practices such as physical removal of contaminants using scrapers, combined with chemical and steam treatment.

Further management options understudy
Compost amendments are commonly used for soil fertility management, and recent studies at UC Davis suggest that composts may also suppress Fusarium wilt in soil. Pathogen-infested tomato residue decomposes more rapidly in soil with long-term inputs of poultry manure compost. Preliminary studies also indicate that cover crops such as hairy vetch may be suppressive to Fusarium wilt, as has been seen for Fusarium wilt of watermelon, but results are inconsistent.

 

Where to Go Next

There are no documented cases of Fol race 4 in California. However, given the history of this pathogen, it is almost inevitable that a new race (race 4) will emerge that overcomes race 3 resistance. As a result, race 4 monitoring continues to be a top priority across the state.

F3 resistance can be compromised by stress, and Fol race 3 has been documented causing disease in multiple F3 tomato fields every year. Knowing which types of plant stress affect host resistance would help growers prioritize management strategies. Long-term studies of Fol race 3 survival in soil are important for knowing optimal durations for rotating out of tomato.

There are several common rotation crops whose risk status is still unknown, including safflower, alfalfa, potato and hemp. Furthermore, there is clearly a need for soil testing tools for Fol race 3. The UC Davis Vegetable Pathology program is working to develop more rapid molecular tools for both soil detection and diagnosis in plants. Industry innovations in effective sanitation, such as full room steam or fumigation sanitation stations, could provide a breakthrough in slowing spread of Fusarium wilt as well as other soil-borne pathogens.

While the aforementioned management approaches have primarily focused on Fusarium wilt alone, in reality, Fusarium wilt also occurs as part of disease complexes. Developing strategies that minimize damage wrought by these soil-borne complexes is a critical next step in achieving healthy tomato production in California.

For more information, contact Cassandra Swett at clswett@ucdavis.edu.

Weed Control in Lettuce

Economical and successful weed control in lettuce can be accomplished by utilizing key cultural practices, cultivation technologies and herbicides. Planting configurations vary from 40-inch wide beds with two seedlines to 80-inch wide beds with 5 to 6 seedlines. Recent studies of weeding costs for lettuce ranged from $454 to $623/A for 80-inch wide beds with 5 seedlines of head and 6 seedlines of romaine hearts lettuces, respectively (see coststudies.ucdavis.edu/en/current/commodity/lettuce/).

Weeding costs included the following: Herbicide applied in 4-inch wide bands over the seedlines, cultivation, auto thinning using a fertilizer to kill unwanted lettuce plants and hand weeding/double removal. The costs for auto thinning also include fertilizer costs, which can satisfy the need for the first fertilizer application.

Significant weed control is accomplished by practices that occur before the crop is planted. For instance, weed pressure is affected by prior crop rotations and how much weed seed was produced in them. The weeding costs given above are rough averages. If weed pressure is light, weeding costs can be lower, but if weed pressure is high, weeding costs can be much higher. In the Salinas Valley, good management of weeds is possible with rotational crops such as baby vegetables (spinach, baby lettuce and spring mix) because they mature in 25 to 35 days and don’t allow weeds to set seed. Long-season crops such as pepper and annual artichokes allow multiple waves of weeds to germinate and which are difficult to see and remove once the plants get bigger.

Preirrigation is standard practice to prepare the beds for planting. It stimulates germination of a percentage of weed seeds in the seedbank, and they are subsequently killed by tillage operations. Studies have shown that preirrigation followed by tillage lowers weed pressure to the subsequent crop by about 50%. In organic production, pregermination is one of the most powerful practices for reducing weed pressure, and if time allows, it can be repeated to further reduce weed pressure.

 

Preemergence Herbicides

There are three pre-emergence herbicides available for use in lettuce production: Balan, Prefar and Kerb. Balan and Prefar provide good control of key warm season weeds such as lambsquarters, pigweed and purslane, as well as grasses (Table 1). Kerb is better at controlling mustard and nightshade family weeds such as shepherd’s purse and nightshades. Balan is mechanically incorporated into the soil and Prefar and Kerb are commonly applied at or post planting and incorporated into the soil with germination water.

Table 1. Weed susceptibility to registered preemergent herbicides.

Kerb is more mobile in water than Prefar which can lead to issues with its efficacy. Often 1.5 to 2.0 inches of water are applied with the first irrigation to germinate the crop which can cause Kerb to move below the zone of germinating weed seeds, especially on sandy soils. For instance, Kerb is capable of controlling purslane however, its efficacy can be low on sandy soils due to its movement below the zone of germinating weed seeds with the first germination water. Prefar does not leach as readily as Kerb and that is why these two herbicides are often mixed in the summer to control purslane (Figure 1).

Figure 1. On left: Kerb at 3.5 pints/A applied at planting; On right Kerb at 3.5 pints/A + Prefar at 1.0 gallon/A applied at planting. The main weed is common purslane which was not controlled by Kerb because it was pushed below the zone of germinating weed seeds by the germination water (photo courtesy R. Smith.)

In the desert, the use of delayed applications of Kerb has been used for many years. Due to the large amounts of water that are applied to keep the seeds moist and cool, Kerb is applied in the 2nd or 3rd germination water, approximately 3 to 5 days following the first water, just prior to the emergence of the lettuce seedlings. The amount of water applied in the second and third irrigation is less than the first application and therefore does not push the Kerb as deep in the soil.  Although the Salinas Valley is cooler than the desert, evaluations here have also found delayed applications to improve the efficacy of Kerb (Figure 2).  These data illustrate the loss of control of purslane by Kerb when applied before the first germination water, as well as the improvement in efficacy that results when applied after the first germination water. It also illustrates the role that Prefar plays in the control of purslane when the efficacy of Kerb is reduced by being pushed too deep. Clearly, there is benefit from applying the Kerb in the 2nd or 3rd germination water because it helps to keep it in the zone where weed seeds are germinating.

Figure 2. Efficacy of Kerb applied at 3.5 pints/A at planting or in the 3rd germination water; crop was romaine. Note that applying the Kerb after the first heavy application of germination water greatly improved its effectiveness.

The use of single use drip tape injected 3 inches deep in the soil has become popular in the Salinas Valley. The uniformity of using new tape with each crop has allowed growers to consider using drip irrigation to germinate lettuce stands. Although the same amount of water may be applied to germinate the stand with drip irrigation as with sprinklers, the water tends to move upward with drip irrigation. In drip germinated lettuce, Kerb is sprayed on the soil surface and is solubilized by the upward movement of the drip applied water which allows it to move just deep enough in the soil to control germinating weeds, but not too deep to reduce its efficacy (Table 2). Interestingly, drip germination alone resulted in fewer weeds than sprinkler irrigation.

Lettuce is typically planted with 4-5 times more seed than is needed in order to assure a good stand. At about 3 weeks after the first irrigation, lettuce is thinned. Traditionally lettuce has been thinned by hand, but increasingly growers are using auto thinners which spray an herbicide (Shark) or concentrated liquid fertilizer (e.g. AN 20, 28-0-0-5, and others) to kill the unwanted plants and achieve the desired plant spacing. In the process of thinning by hand or by auto thinning, a significant portion of weeds in the seedline is also removed.

Table 2. Effect of Kerb application (at 3 pints/A) method (surface applied, drip injected or untreated) and irrigation method (surface tape, buried tape or sprinkler) on weed densities, lettuce stand and visual injury.

 

Automated Thinning and Weeding

About 10 to 14 days after thinning, hand weeding is carried out to remove weeds from the seedline and any double lettuce plants that were not removed in the thinning operation. An increasing number of Salinas Valley growers are using autoweeders prior to hand weeding.  There are several autoweeders available: Robovator (Denmark), Steketee (Netherlands), Ferrari (Italy) and Garford (England). These machines use a camera to capture the image of the seedline and a computer that processes the image and activates a kill mechanism (a split or spinning blade) to remove unwanted plants. The machines were originally designed for use with transplanted vegetables. We tested auto weeders and found that they remove about 50% of the weeds in the seedline and reduced the subsequent hand weeding times by 35%. In order to safeguard the crop plants, the auto weeders leave an uncultivated safe zone around the crop plants where weeds can survive. As a result, auto weeders do not remove all the weeds in the seedline, but they help to make subsequent hand weeding operations more efficient and economical.

Depending on the weed pressure, some lettuce fields are hand weeded one more time a week or so prior to harvest. Given the practices just outlined, perennial weeds are not problems in the typical lettuce rotations in the Salinas Valley. The rapid turnaround of the crop (55 to 70 days during the summer) and the frequent use of cultivation does not allow enough time for weeds like field bind weed or yellow nutsedge to build up root reserves or nutlets before they are cultivated or disced out. In the summer, purslane is the biggest concern because it can build up high populations in the seedbank and, because of their fleshy tissue, can set seed even after being cut by the cultivator knives. As a result, if it is not effectively controlled in prior rotations, it can result in high hand weeding costs. Growers address purslane issues by making bedtop applications of the combination of Prefar and Kerb, as well as by a combination of other practices outlined above.

Although there have been no new herbicides registered for use on lettuce in many years, there have been significant technological developments that have improved efficiency of weed control in lettuce. The increasing use of single use drip tape and new automated thinning and weeding technology have recently contributed greatly in this regard.

Making Sense of Biostimulants for Improving your Soil

Biostimulants…bio what??? You may have heard or read this phrase several times over the past year as this product category gains traction in the agricultural marketplace. Confused about what exactly constitutes a biostimulant? You are not the only one! A biostimulant includes “diverse substances and microorganisms that enhance plant growth” or helps “amend the soil structure, function, or performance.” Got it? No? That is ok, please read on for more information.

 

Market Confusion

The exact definition of what a biostimulant is, and what it is not, can be confusing and leave some folks scratching their head on what to expect regarding product performance (See Figure 1). A biostimulant tends to be an “environmentally friendly alternative to synthetic products” and can have multiple impacts on the crop or soil, although the exact definition of the category is vague and open-ended. This uncertainty has received increased attention by regulators, and we should expect to see more precise definitions soon.

Figure 1: Biostimulants can impact a crop in many ways depending on the active ingredient applied (graphic courtesy Ute Albrecht, Southwest Florida Research and Education Center).

As it stands, there are many active ingredients in this arena, and some growers have struggled to find the right fit for their farm. This confusion is regrettable given the increasing popularity of the category and the forecasted sales growth rates. For example, the global market for biostimulants was valued at $2.19 billion in 2018 and is projected to have a compound annual growth rate of 12.5% from 2019 to 2024.

 

Matching Clear Goals

Biostimulants can be derived from a laundry list of different materials, with studies listing roughly eight major classes of active ingredients or more, each with unique properties and modes of action. However, my experience in the field suggests that many of us have unfortunately lumped the various products in this category into one largeother” bucket for simplicity, regardless of the difference in how the product works or what outcome should be expected.

Below I help clarify the role of several active ingredients to allow you to better understand and also mix and match the desired characteristics you are looking for (See Table 1). This reference table will allow you to determine which features you want to put to work into your biostimulant blend based on your crop production method, application equipment, and comfort level. The biostimulant categories listed complement an agronomically sound fertilizer and irrigation program and should be included as a part of a comprehensive crop management program. Caveat: I do not have enough space to list all possible modes of action, but instead I limit the table to the materials that have an impact on the soil.

Table 1: Biostimulants are sorted by their active ingredient (left side), a description of how they work (center) and some general handling notes (right side).

Understanding the Nuances

The biostimulant category offers many exciting opportunities to growers and can deliver new functionality to common fertilizers when used in a blend. Before jumping into this ‘other’ category, start with the following question “What features am I looking for?” This honest query will help you pick the correct ingredient needed and bring clarity to the nuances of the biostimulant category. Getting your product blend right from the get-go can help improve the soil on your farm and help jumpstart your 2020 yield and quality goals. Please consult with your local sales representatives to help pick the right active ingredient for the job and be sure to jar test any new blend ideas you have prior to tank mixing for compatibility concerns.

Furthermore, running a pilot or test study can be a great way to learn which biostimulant product is right for your crop and production system. Keeping good records of your observations will help jog your memory about product performance as the season wears on and will help you formulate the right blend for the job. A good pilot or trial plan can go a long way with helping you keep track of important information on how your biostimulant blend is impacting your crop.

Hungry for more information about biostimulants and what they can do for you? Many trade publications, such as the one you are reading now, have begun to cover this category in more detail and there are several good articles out there that are worth reading. Below I provided some recommended reading to help get you started along with some online resources that are worth a look.

 

About the Author

Dr. Karl Wyant currently serves as the Director of Ag Science at Heliae® Agriculture where he oversees the internal and external PhycoTerra® trials, assists with building regenerative agriculture implementation, and oversees agronomy training. Prior to Heliae® Agriculture, Dr. Wyant worked as a field agronomist for a major ag retailer serving the California and Arizona growing regions. To learn more about the future of soil health and regenerative agriculture, you can follow his webinar and blog series at PhycoTerra.com.

 

Further Resources

 

References

Albrecht, Ute. (2019). Plant biostimulants: definition and overview of categories and effects. IFAS Extension HS1330.

Calvo Velez, Pamela & Nelson, Louise & Kloepper, Joseph. (2014). Agricultural uses of plant biostimulants. Plant and Soil. 383. 10.1007/s11104-014-2131-8.

Drobek, Magdalena & Frąc, Magdalena & Cybulska, Justyna. (2019). Plant Biostimulants: Importance of the Quality and Yield of Horticultural Crops and the Improvement of Plant Tolerance to Abiotic Stress—A Review. Agronomy. 9. 335. 10.3390/agronomy9060335.

Rouphael, Y., Colla, G., eds. (2020). Biostimulants in Agriculture.  Lausanne: Frontiers Media SA. doi: 10.3389/978-2-88963-558-0

Detection of Marked Lettuce and Tomato by an Intelligent Cultivator

Weeds are difficult to control in lettuce and tomato due to labor shortages, increasing costs of hand weeding and limited herbicide options. Lettuce is very sensitive to weed competition, plus there is no tolerance for contamination of bagged lettuce salad mixes with weeds; therefore, weeds must be controlled if lettuce is to be harvested.

Consequently, mechanical weed control is an important part of an integrated weed management program in conventional and organic vegetable crops. Traditional inter-row cultivation, however, only removes weeds between crop rows and leaves the weeds within the crop row. The removal of in-row weeds requires hand weeding, a time-consuming and expensive process.

 

Vegetable Weed Control Costs

Weed control costs for conventional head lettuce production in California are estimated at $216 to $319 per acre, while weed control costs in organic leaf lettuce are $489 per acre, on average, at current labor rates. In conventional processing tomatoes, weed control costs are about $225 per acre or 12% of production costs. Additionally, hand weeding costs have increased due to labor shortages, changes in California overtime regulations and increasing minimum wages as well as decreased labor immigration from Mexico. The result is greater vulnerability of growers to crop losses due to weeds.

Automation of weed removal may be a method to contain or reduce weed control costs in vegetable crops. Intelligent intra-row cultivators (IC) provide an alternate weed management option to standard inter-row cultivation. Previous results have shown that IC can reduce the need for hand weeding compared to standard cultivators and may reduce weed control costs.

The Robovator® cultivator evaluated by Lati et al. (2016) relied on pattern recognition of the rows and crop plants within the rows based on the expected crop spacing within the rows. When these spatial cues are unavailable, as can occur in an organic field with a high weed density, this approach cannot differentiate between crops and weeds, and thus it relies on a size difference between crops and weeds, as well as a low to moderate weed population to function accurately.

Intelligent Cultivation. Intelligent intra-row cultivation requires three technologies; a machine-vision system that detects crop plants and weeds, image classification and decision algorithm that differentiates between crop plants and weeds, and an automated weed removal mechanism that controls the weed while protecting the crop. Precision guidance systems, decision algorithms, and precision in-row weed control devices are commercially available or are at an advanced level of development. Accurate crop detection and differentiation from weeds, at normal cultivation speeds, would allow for greatly improved intra-row cultivators.

Weed/Crop Differentiation. The main challenge for intelligent intra-row cultivation is to differentiate between crops and weeds using digital imagery and processing at field operation speeds of at least 1 mph in high weed density fields with travel speeds above 2 mph required for economic acceptability for low to moderate weed loads.

A new method of crop and weed differentiation called “crop signaling” is presented in the research “Crop Signaling for Automated Weed/Crop Differentiation and Mechanized Weed Control in Vegetable Crops” by Raja et al. 2019 out of UC Davis. It is based on the idea that the identity of the crop is known with certainty when it is planted, whether transplanted or seeded. Thus, if the crop has a marker or signal that an IC can reliably detect, then the IC would recognize the signal and protect the crop. Plants without the signal, i.e., weeds, would not be protected and would be removed by the IC. The objective of this work was to test a crop signaling system for crop detection accuracy and weed control efficacy by an IC in lettuce and tomato.

Marking System Descriptions. Two methods of plant signaling were tested, physical plant markers and topical markers. Biodegradable straws coated with a fluorescent marker were used as the plant markers in this study (Figure 1). The straws were then placed next to tomato seedlings in the planting trays and then transplanted together (Figure 2).

Figure 2. Holland transplanter with butterfly transfer fingers used for transplanting plant labels and tomatoes together.

The topical marker used on plant foliage was green or orange fluorescent water-based paint (Figure 3a,b). A paint sprayer was used to apply the topical marker to lettuce foliage and tomato seedlings prior to planting, while they were in trays. Another method was to spray the marker onto tomato stems as they were transplanted (Figure 4).

Figure 3. (left) Topical marker on lettuce plants, (right) Spray application of topical marker on crop plants.
Figure 4. Topical marker sprayed on tomato transplants by applicator mounted on the transplanter during the process of transplanting.

Intelligent Cultivator. The IC used in this research was developed at the University of California, Davis. It uses a machine vision system specifically designed to detect the physical labels and topical markers on the crop (Figures 5&6). Weed control was done by mechanical knives, which the IC opens (Figure 6b) to avoid the marked crop plants and closes (Figure 6a) to uproot weeds in the intra-row space.

Figure 5. Image of a tomato plant with a green label taken (a) under normal light plus UV light, and (b) under UV light only. Note the reflections of the green label in the six mirrors, and the actual label in the center of the image.
Figure 6. The actuator device used in this project: (a) Weed knives closed – uprooting weeds in crop row, (b) Weed knives open avoiding tomato plant.

 

Field Trials

Eight field trials in tomato at Davis, Calif., and six in lettuce at Salinas, Calif., were conducted during 2016-2018.

Tomato. Field trials in processing tomatoes were located on a silt loam soil on the UC Davis vegetable field crops research station near Davis. The tomatoes were seeded in trays and kept in a greenhouse for 45 to 60 days until they were about 10 inches tall. Tomatoes were transplanted into 60-inch beds at 15-inch spacing in a single center row. Two tomato trials were carried to yield.  Plant labels were added to seedling trays prior to transplanting (Figure 1) or the topical marker was applied to trays of tomato seedlings as described above (Figure 4). Tomato transplants were marked with paint 4 inches above the soil line. About three weeks after planting, all plots were cultivated with a standard mechanical cultivator which only removed weeds outside the plant line. The standard cultivator left a 7-inch non-cultivated band centered on the crop row.

Weed densities by species were measured before and after cultivation in four 7-inch-wide (centered on crop row) by 20-foot-long sample areas randomly placed along the length of the plots. The time required by a laborer to hand weed the 20-foot areas was recorded. Two tomato trials were maintained until harvest so that marketable yield data could be collected.

Lettuce. Field trials using Romaine lettuce were conducted in a sandy loam soil at the USDA research station in Salinas, Calif. Four weeks after seeding, the whole experiment was cultivated with a standard mechanical cultivator. The standard cultivator left a 6-inch non-cultivated band centered on the crop row (Figure 7). The IC operated within .75 inches of the lettuce plants on all sides. Pre-cultivation weed counts were measured the day before cultivation and post-cultivation weed counts were taken the day after cultivation. Weed densities were measured in a 6-inch band centered on the crop row in each of two 20 -foot-long samples in the field. Weeds that were uprooted were considered dead. After cultivation, hand weeding was performed and timed as described for the tomato trials. The time spent by a laborer to hand weed with a hoe was recorded.

Figure 7. The plant layout used in the lettuce plantings: (a) Single crop row of lettuce on 1 m beds. The control rows are with no crop signal visible, (b) physical labels in lettuce row two weeks after transplanting.

The 2017 lettuce trials were maintained until commercial maturity and number of marketable heads and weight of marketable heads were recorded. The 2018 trial was conducted at a commercial lettuce field near Salinas, Calif.

Statistical Analysis. RStudio Version 1.1.383 was used for statistical analysis. Differences between pre- and post-cultivation weed counts determined weed removal effectiveness. The most efficacious treatments removed the greatest proportion of weeds.

The difference in weed densities between pre and post cultivation were analyzed using analysis of co-variance, to measure the effect of cultivator type on weed density. Analysis of variance (ANOVA) was performed on the hand-weeding time data to measure the effect of the cultivators.  Weights were determined for both lettuce and tomato yields, and in lettuce, the number of heads was also determined.

Weed Control. The IC was more effective than the standard cultivator at removing weeds from the inter-row space. The data were pooled separately for tomato and lettuce. In tomato seed lines, 1 weed per square foot remained after IC while 10.5 weeds per square foot remained after standard cultivation. This is a 90% reduction in the number of weeds remaining after cultivation (P<0.05).  In the lettuce trials, 1.7 weeds per square foot remained in the seed line after intelligent cultivation while 5 weeds per square foot remained after standard cultivation, which is a 66% reduction in weeds remaining after cultivation (see Table 1).

Table 1: Effect of cultivator type on in-row weed densities after cultivation, time to hand weed and marketable yield in tomatoes and lettuce.

Handweeding in the tomato trials required 7.8 hours/A following the IC while the standard cultivator required 14.9 hours/A which is a 48% reduction (P<0.05). Hand weeding of lettuce required 16 hours/A following cultivation the IC while 29 hours/A was required for the standard cultivator, a 45% reduction in time (P<0.05).

The time-spent hand weeding after IC cultivation was a notably smaller percentage reduction than it was for weed densities, i.e. 48% vs. 90% in tomato. This is because the IC consistently removes the readily accessible weeds that are more than an inch from the crop; while the remaining weeds after IC cultivation are typically close to the crop plants and take more time for the field crew to remove than weeds further from the crop plant. The IC did not remove all the weeds it passed over due to some algorithmic uncertainty in the precise location of the crop’s main root and a risk-averse control strategy. Thus, weed control in close proximity to crop plants may still require some hand weeding. However, significant reductions in manual labor were achieved while maintaining effective weed control.

Crop Yields. There was no difference between the cultivators in their effect on tomato fruit yield in 2017 (P>0.05) (Table 1).  The 2018 tomato yields had marketable fruit yields in the IC and standard cultivator treatments of 44,045 and 50,217 lbs./A, respectively (P>0.05). Similarly, there were no differences between the cultivators in their effect on lettuce yields (P>0.05) (Table 1). Yield data were analyzed both as the number of marketable lettuce heads per acre and fresh weights.  

Weed/Crop Differentiation.  One of the biggest challenges for automated intra-row cultivation is to enable a computer and vision system to differentiate between crops and weeds at normal field travel speeds. The commercially available IC ‘Robovator®’ uses pattern recognition to recognize the crop row and can perform intra-row weeding at speeds of 1 mph (Lati et al. 2016). However, this requires a distinct crop pattern best found such as in a transplanted field where the crop is much larger than the weeds and the crop stand is consistent. Further, when high weed densities obscure the 2-dimensional crop row pattern, the intra-row weeding program does not work.

Two types of crop signals were tested, physical plant labels and topical markers. The methods have very low false positive error rates and the classification accuracy achieved for both techniques approaches 100%. The crop signaling technique appears to be effective in creating a reliable method for automatic detection of crop plants in vegetable fields with high weed densities. Crop signaling technology could facilitate development of automated weed control robots that are as accurate in crop/weed differentiation as human workers are.

A recommendation for future work is to develop a commercially viable marking method that is machine readable, yet does not contaminate harvested produce or the field soil and subsequent rotational crops. For transplanted stem crops like tomato, a biodegradable machine-readable tag attached to each stem as the transplanter sets the plants should be explored for commercial potential. Lettuce will probably require a machine-readable label attached to the first true leaves or a machine-readable label on the fiber-coated plant plug as it is set in the soil as is done with the Plant Tape® (www.planttape.com) system of vegetable transplanting.

Regardless of the technology used for crop weed differentiation, development of intelligent weed removal technology has improved weed control programs for horticultural crops that continue to rely on a limited number of herbicides and hand weeding. However, there is much more to do to improve vegetable weed control.

Acknowledgments. Thanks to the USDA Institute of Food and Agriculture Specialty Crop Research Initiative (USDA-NIFA-SCRI-004530) the California Tomato Research Institute and the California Leafy Greens Research Program for financial support.

 

References

Lati, R.N., M.C. Siemens, J.S. Rachuy, and S.A. Fennimore. 2016. Intra-row Weed Removal in Broccoli and Transplanted Lettuce with an Intelligent Cultivator. Weed Technology 30:655-663

Raja R, Slaughter DC, Fennimore SA, Nguyen TT, Vuong V, Sinha N, Tourte L, Smith RF, Siemens MC (2019) Crop signaling: a novel crop recognition technique for robotic weed control.  Biosystems Engineering 187:278-291.

Virus Pathogens: Challenges to the Health of Vegetable Crops

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Farmers and other field professionals producing vegetable crops face a bewildering array of challenges. Insects and mites feed on, disfigure, and eat away at produce quality. Weeds compete with the vegetables for precious resources and can require extensive labor to be removed. Fertilizer and water inputs can be costly. The economic cycle of planting, growing, harvesting, and marketing can be a “black hole” that engulfs company resources while offering few guarantees of profits. Another group of challenges is embodied by the many plant pathogens that cause diseases of vegetable crops. One particular group of pathogens of interest are the viruses that infect plants.

 

Virus Pathogens of Plants

Viruses that infect plants are similar, in shape and constitution, to the viruses that infect insects, animals, and yes, people. A virus consists of a piece or two of genetic material (either DNA or RNA) that is surrounded and protected by a protein coat or covering. In the grand scheme of biology, such a nucleic acid + protein structure is extremely simple and basic. This entity is also extremely tiny. Since a virus is composed of two types of chemicals, it is much smaller than a plant cell and cannot be observed with a regular microscope. Only with the use of electron microscopes can the body of the virus be observed. The outer protein coat gives the virus a distinctive shape, and plant viruses can look like long flexible threads, short rigid rods, or spherical, geometric polyhedrals.

Different viruses have different shapes and appear as long threads, rigid rods, or geometric spheres. Left, tomato chlorosis virus (photo courtesy K. Schlueter, USDA) and, right, cucumber mosaic virus (photo courtesy M. Kim, USDA)

Plant viruses, like all viruses, do not function or operate outside of their hosts. To become active the virus must be introduced into a living plant cell, after which the virus mechanism activates and highjacks the cell’s processes, forcing the host cell to produce more virus RNA or DNA and virus proteins. These components are assembled into new viruses which are then translocated throughout the plant by being carried in plant fluids that stream into stems, leaves, flowers, and fruits.

 

Diseases Caused by Viruses

As with viruses that infect people and animals, plant pathogenic viruses at first show no evidence of their initial incursion into the host. There is a latent period or lag-time during which the virus is steadily orchestrating the manufacture of additional virus nucleic acids and proteins. At a certain critical point, the virus population causes enough physiological and metabolic disruption so as to cause visible symptoms, which collectively we call the disease.

Disease symptoms caused by viruses can vary greatly and are influenced by the vegetable variety, age of plant when first infected, the strain of the virus, and environmental conditions under which the crop is grown. In general, vegetable crops infected with viruses will show one or more types of foliar symptoms. Leaf color changes with the development of yellow or brown spots, light and dark green patterns (mosaic, mottling), concentric ring patterns (ringspot), and yellow or white blotches and streaks. In some cases, the entire foliage of the plant turns yellow, orange, or red. Some viruses cause a curious reaction where only the veins of the leaf become yellow or brown. Leaves can be misshapen in various ways, from simple curling, to unusual elongation (strap leaf), to severe twisting and deformation. Internodes along the stem become abnormally shortened, resulting in tight bunching of leaves. Flowers also change appearance with streaks of color in the petals (color break). For fruiting vegetables, the fruit may show only subtle color breaks and patterns, or alternatively become grossly deformed. Overall plant growth can be stunted and crop development can be delayed.

All vegetable crops suffer from at least one virus pathogen, while some crops are subject to a dozen different ones. Table 1 lists selected vegetable crops and some of the viruses affecting these crops in the U.S. Like fungal and bacterial pathogens, virus pathogen occurrence and importance vary with geographic region. A virus that is important on California lettuce may be incidental or lacking on lettuce in Florida. Likewise, the set of viruses that North American tomato growers must deal with will be different than tomato viruses occurring in South America or Asia.

Table 1. Selected vegetable crops, virus pathogens, and means of virus dispersal.

The economic impact of a particular crop-virus interaction depends on the inherent aggressiveness of the virus, incidence of the disease, and the susceptibility of the crop. Regarding the crop, a critically important factor is the type of harvested commodity. For example, leafy commodities such as lettuce and spinach will be especially vulnerable to viruses that cause leaf symptoms. The viruses of pepper that cause fruit malformations are more important than the pepper viruses that only cause mild mosaics in the foliage. For celery grown in California, cucumber mosaic virus (CMV) causes some leaf mosaic and mottling but rarely causes any symptoms on the celery petioles and, therefore, is of little concern. However, a different virus, Apium Virus Y, can cause celery petioles to turn brown, making the celery unmarketable.

In lettuce, Impatiens necrotic spot virus results in distorted plants and brown leaf lesions (photo by S. Koike.)

Detecting and Diagnosing Viruses

Confirmation of a virus requires testing. We acknowledge that experienced growers and field personnel, who have looked at virus diseases of a particular crop for many years, can develop a good diagnostic sense for such problems. However, to be scientifically sound and accurate, diagnosing virus diseases cannot be achieved without clinical testing. Virus disease symptoms pose particular challenges to diagnosticians because the wide range of virus-like symptoms can also be caused by other factors.

Symptoms caused by viruses can also be caused by genetic disorders, nutritional imbalances, environmental extremes, phytotoxicity from pesticides and fertilizers, and other factors (see Table 2.) Fortunately, diagnostic labs have the tools that can identify most of the commonly occurring viruses in vegetables. Such tests rely on either serology (using antibodies that detect the antigens of virus proteins) or molecular biology (using probes that recognize nucleic acid sequences of the virus.)

Table 2. Symptoms caused by viruses and other factors that can create similar symptoms

Epidemiology of Virus Diseases

Development of virus diseases of plants involves several factors. In contrast to some human viruses, plant viruses are not moved around in the air or deposited on surfaces waiting to come into contact with a plant. Rather, plant pathogenic viruses typically originate from a living source or “reservoir.” (Factor 1) The reservoir is often an infected weed that is near the site where the vegetable crop will be planted, or the reservoir can be an infected volunteer crop plant in the field. Vectors (Factor 2) are the insects, mites, and nematodes that have fed on a virus-infected plant, ingested virus particles, and now are capable of injecting the viruses into the next plant that is fed upon. For the great majority of viruses that infect vegetables, the viruses are moved by vectors from reservoir hosts to healthy crops (Factor 3). Aphids are the most common vectors (See Table 1.) Other insects (thrips, leafhoppers, beetles) also carry viruses, as do a few soilborne nematodes and one soilborne fungus.

Apium virus Y causes disfiguring brown lesions on celery petioles (photo by S. Koike.)
A number of virus pathogens cause damage to the fruits of some vegetable crops (photo by S. Koike.)

The epidemiology, or progress of disease spread, depends on the complex interaction of the three factors mentioned above.

Factor 1 Reservoir: What is the nature of the virus reservoir? Which weed species are present? Are there high numbers of virus-infected weeds or volunteer plants in the area? A virus with a broad host range, such as Tomato spotted wilt virus (TSWV), may be present in dozens of weeds and numerous volunteer plants on a particular ranch.

Factor 2 Vector: Which vectors are in the vicinity? What are their populations and dispersal patterns? How do wind patterns and geographic features influence dispersal? What is the extent of vector increase within the crop, which can result in plant-to-plant spread within that planting?

Factor 3 Vegetable Crop: What is the crop diversity in the area being considered and which viruses affect these crops? For example, could CMV, which has a broad host range, spread between different vegetables? If the region is widely planted to one crop, such as lettuce, will a particular virus affect many lettuce plantings? Too much of the same crop, densely cropped in one region, could result in rapid virus spread and disease epidemics. In contrast, if a region has only one onion field among many non-allium crops, a narrow host-range pathogen such as Iris yellow spot virus will infect only the onions. The answers to these and other questions have significant bearing on the management of virus diseases.

 

Managing Virus Diseases

Diagnosis: The first step in disease management is accurately identifying the precise pathogen involved. Molecular and serological assays are available for most of the major virus pathogens affecting vegetables. Knowing which virus is involved enables one to know the reservoir plants harboring the virus, the vectors involved, and the potential target crops.

Exclusion: Prevent the virus from entering the production system. For lettuce, cucurbits and tomato, some viruses are carried in the seed; therefore, use seed that has been tested or certified to not harbor the pathogen. For crops started as transplants, employ IPM practices to prevent infection at the transplant stage. Note that for the few vegetable crops propagated by cuttings or plant divisions (example: artichoke), viruses will be readily spread if infected propagative material is used to plant new fields.

Reservoir host eradication: Remove the initial sources of the virus, which are infected weeds and volunteer crop plants. Plant viruses are present mostly in living plants and generally not in soil, water, equipment surfaces, or the air. Controlling weeds and other reservoir plants is therefore a critical part of virus control.

Manage the vectors: Use IPM practices to control the virus vectors. The great majority of vegetable-infecting viruses only reach a crop via an insect vector. Complete control of an insect pest is rarely possible, so strategies should attempt to manage the insects as best as possible. Keep in mind that the vectors are also present on the reservoir weeds and plants outside of the field. Once a virus is introduced into the crop, intra-field, plant-to-plant spread will be achieved only through movement of the vector.

Destruction of the old crop: Once a crop has been harvested, the passed-over plants and shoots growing from remaining crop roots can serve as virus reservoirs if they are infected. Old vegetable fields should, therefore, be disked and plowed under in a timely manner.

Resistant cultivars: If available, growers should select cultivars that are bred to be resistant to the virus pathogens. Note, however, that the usefulness of such genetic plant resistance may not last. Researchers found that the use of tomato and pepper cultivars resistant to TSWV has allowed for the development of “resistance breaking” (RB) strains of the virus. Through mutation and selection, these new strains of TSWV can cause disease in the previously resistant cultivars.

Chemicals or pesticides: Currently there are no chemical treatments that can be applied to plants that would prevent infection from viruses or prevent development of virus disease.

Carrot fields severely infected with viruses become noticeably yellow to orange in color (photo by S. Koike.)

 

Cover Crops in California Agriculture: An Overview of Current Research

Growers throughout the country and around the world plant a wide range of cover crops for a variety of reasons. Cover crops can reduce soil compaction, improve water infiltration, improve soil structure, and feed soil microbes: they encourage a healthier and more diverse soil ecosystem.

Researchers in California are analyzing the best ways to incorporate cover cropping into the state’s diverse agricultural systems, from high-value vegetable production on the central coast to the cotton, tomato, and almond fields of the central valley.

 

Cover Crops on the Central Coast

Researchers working with central coast vegetable growers have devised innovative ways to use cover crops to reduce nitrate leaching and agricultural runoff, thereby improving both local ecosystems and soil health.

Eric Brennan and his team at the USDA Agricultural Research Service started the Salinas Organic Cropping Systems trial in the Salinas Valley in 2003 to understand the long-term impacts of various cropping systems and soil amendments. This trial focuses on organic lettuce and broccoli, two of the high-value crops grown in the area known as the nation’s salad bowl.

To maintain soil organic matter and provide nutrients to their crops, organic vegetable growers in this area prefer applying compost instead of planting cover crops. The amount of time that cover crops require for incorporation and decomposition can shorten the growing season for these high-value crops (Brennan & Boyd, 2012.) To make this practice more feasible for growers in the area, this group of researchers has developed three strategies for integrating cover crops into the vegetable cropping systems of the Central Coast.

Option 1: Plant the cover crops only in furrow bottoms, not the entire field. After 50 to 60 days of growth, the grower can spray the cover crops and then do the usual tillage necessary to prepare the ground for planting the cash crops. By planting time, the cover crop residue has already decomposed. This method reduces runoff and erosion but does not reduce nitrate leaching, so this is best for fields with runoff problems but without high nitrate levels. However, this method makes controlling weeds during a wet winter difficult and costs more than simply leaving the field bare (Brennan, 2017.)

Option 2: Plant non-legume cover crops on the vegetable beds and mow the cover crops repeatedly throughout the growing season. This maximizes nitrate scavenging while minimizing the amount of residue that needs to decompose right before planting. The ideal cover crop for this practice would be a grass, like cereal rye. Repeated mowing would reduce the amount of water lost to evapotranspiration from the cover crop but still enable the rye to scavenge nutrients that could otherwise be lost to leaching (Brennan, 2017.)

Option 3: Turn the cover crop residues into a highly nutritious juice and compost. To do this practice, a grower would plant a non-leguminous cover crop in October and allow it to grow until mid-December, at which point it will have scavenged most of the nitrogen that it will use. The grower then harvests the cover crop, leaving as little residue behind as possible. They can then feed the residue into a screw press, which will separate the liquids and solids. The liquid component has a relatively low nitrogen concentration and can be applied to the vegetable crop to fulfill some of the crop’s nutrient needs. The solid residues can be composted and applied at a convenient time, to provide organic matter to the soil (Brennan, 2017.)

Researchers are still working on refining these strategies, but they could allow central coast vegetable growers to reap the rewards associated with cover crops while maintaining a profitable enterprise.

Field day at the West Side REC in 2010, discussing cover cropping and conservation tillage (photo courtesy Jeff Mitchell, UCCE.)

 

Annual Systems in the Central Valley

For the past 20 years, Jeff Mitchell and his team at UC Cooperative Extension have studied the effects of reduced tillage and cover crops on a tomato-cotton rotation at the UC’s West Side Research and Extension Center. This study measures the efficacy of these practices in reducing air pollution and increasing soil organic matter. Reduced tillage and cover cropping have resulted in less dust emissions compared to conventionally managed fields (Mitchell et al., 2017.) They found that cover cropping increased soil organic matter more than conservation tillage alone did (Veenstra et al., 2006.) Overall, these practices have improved soil health by increasing aggregate stability, water infiltration, and soil organic matter while maintaining similar yields to the conventional system (Mitchell et al., 2017.) This study has allowed researchers to see the long-term effects of conservation tillage and cover cropping on tomato and cotton systems in the San Joaquin Valley.

Another UC research team in the Central Valley, led by Kate Scow at the Russell Ranch near UC Davis, examined the long-term effects of cover cropping on organic tomatoes and corn. These researchers found that cover cropping encouraged the proliferation of diverse types of beneficial fungi known as arbuscular mycorrhizal fungi (Bender & Bowles, 2018). Under optimal environmental conditions, cover cropping was correlated with higher tomato yields. In contrast, corn did not enjoy the same benefits from organic management that the tomatoes did and had lower yields compared to fields without cover crops (Bender & Bowles, 2018). These studies have found important benefits to including cover crops in annual systems, but growers will need to further refine the practice to fit their needs.

 

Perennial Systems in the Central Valley

Amélie Gaudin and her team from UC Davis and UC Cooperative Extension are quantifying and communicating the benefits and tradeoffs of planting winter cover crops in almond orchards. They established trials throughout the Central Valley. Planting cover crops in almonds increases bee forage, improves soil health, and encourages resiliency. The researchers have found that cover crops resulted in increased water infiltration. Despite the common concern that cover crops would increase frost risk, they found that cover cropping did not affect ambient air temperatures 3 and 5 feet above the ground. Moreover, the ground cover worked as a buffer, keeping temperatures more stable than bare ground did (Gaudin, 2020.)

Other benefits included a decrease in sodicity, improved trafficability in the wintertime, and an increase in aggregation. The soil microbial ecosystem showed increased biomass. Bees enjoyed a more diverse, varied diet, contributing to better bee health. Finally, cover crops reduced weed diversity and growth. They did not reduce germination since both the cover crops and the weeds emerged at the same time. All these benefits start to outweigh the costs of implementation after about 10 years (Gaudin, 2020). Many of these soil and ecosystem benefits are not unique to almond orchards, and could also benefit other perennial cropping systems in the Central Valley.

Mustard cover crops in a table grape vineyard, March 2020 (photo by S. Shroder.)

 

Funding Options

UC and USDA researchers have found benefits to cover cropping in diverse agricultural systems throughout California, from almond orchards to lettuce and tomato fields. These include reducing erosion, compaction, and nutrient leaching, along with improving soil aggregation and providing habitat for beneficial insects. Cover crops may improve the soils upon which your crops depend and increase your operation’s resiliency in the face of a changing climate.

The California Department of Food and Agriculture’s Healthy Soils Program and the USDA NRCS EQIP provide incentives for planting cover crops. Check out cdfa.ca.gov/oefi/healthysoils/IncentivesProgram to learn more about the CDFA’s program. There are 10 technical assistance providers working throughout the state who can help you select your cover crop species, apply for the program, and implement your practices. Go to ciwr.ucanr.edu/Programs/ClimateSmartAg to find your closest climate smart specialist.

Community Education Specialist Alli Fish and a daikon radish cover crop in December 2019 (photo by Rose Hayden-Smith.)

 

Works Cited

(2010). [Field day at West Side Research and Extension Center] [Photograph]. California Agriculture. http://calag.ucanr.edu/Archive/?article=ca.v070n02p53

Bender, S.F & Bowles, T.M. (2018). Effects of AMF diversity and community composition on nutrient cycling as shaped by long-term agricultural management. Russell Ranch 2018 Annual Report. https://asi.ucdavis.edu/sites/g/files/dgvnsk5751/files/inline-files/RRSAF%20Progress%20Report_2018.pdf

Brennan, E. B. (2017). Can we grow organic or conventional vegetables sustainably without cover crops? HortTechnology27(2), 151-161.

Brennan, E. B., & Boyd, N. S. (2012). Winter cover crop seeding rate and variety affects during eight years of organic vegetables: I. Cover crop biomass production. Agronomy Journal104(3), 684-698.

Gaudin, A. (2020, February 4). What do cover crops have to offer? [PowerPoint slides]. University of California Agriculture and Natural Resources. https://ucanr.edu/sites/calasa/files/319850.pdf

Mitchell, J. P., Shrestha, A., Mathesius, K., Scow, K. M., Southard, R. J., Haney, R. L., … & Horwath, W. R. (2017). Cover cropping and no-tillage improve soil health in an arid irrigated cropping system in California’s San Joaquin Valley, USA. Soil and Tillage Research165, 325-335.

Veenstra, J., Horwath, W., Mitchell, J., & Munk, D. (2006). Conservation tillage and cover cropping influence soil properties in San Joaquin Valley cotton-tomato crop. California Agriculture60(3), 146-153.

Lettuce Dieback: New Virus Found to be Associated with Soilborne Disease in Lettuce

Lettuce dieback is a soilborne virus disease known to cause significant losses for lettuce production throughout all western growing regions. The disease was originally described in the Salinas Valley in the late 1990s following severe flooding along the Salinas River but has now been found throughout coastal and inland lettuce production regions of California, the winter production region in southwestern Arizona and Imperial Valley, California.

The disease is most prevalent on romaine lettuce but is known to occur on all non-crisphead (iceberg) lettuce types. Most modern crisphead lettuces are resistant, and an increasing number of romaine cultivars now carry resistance as well. Symptoms of lettuce dieback include yellowing and necrosis of outer leaves, stunted growth and death of affected plants (Fig. 1). Plants infected young may fail to develop beyond the 8 to 10 leaf stage, but symptoms can develop at any point in the growing season, and fields often exhibit a range of plant sizes with some plants appearing healthy and maturing normally, while others become stunted and never fully develop (Fig 2).

Figure 2. Romaine lettuce plants in a field showing variation in severity typical of lettuce dieback including stunted growth, as well as yellowing and necrosis of outer leaves.

Initial symptoms begin with yellowing and necrosis (death) of small veins in outer leaves, with the necrosis expanding into larger areas within and between veins. Inner leaves of the head usually retain their color, but some romaine varieties may also exhibit bright chlorotic flecks within veins of leaves at the center of the head that resembles tiny stars. These are most visible when affected leaves are held up to a light source (Figure 3).

Figure 3. Romaine lettuce leaf from the inner portion of a head showing star-shaped chlorotic flecking in veins characteristic of lettuce dieback disease on romaine.

This vein-flecking symptom is not always present on infected romaine, but when observed it is an excellent diagnostic indicator. The vein flecking symptom is less common on other types of lettuce and is more difficult to observe on red lettuce. Losses resulting from lettuce dieback can range from a few plants to complete loss of crop. In most severely affected fields lettuce heads are not harvested because the plants will not meet quality standards. Symptoms of the disease are frequently found in low lying areas with poor drainage, in areas near rivers, on recently flooded land, and in areas where soil has been dredged from a river or ditch and spread onto adjacent fields.

Symptoms of lettuce dieback can be mistaken for those of other diseases, particularly lettuce drop, a disease caused by a fungus, and symptoms of two viruses transmitted by thrips. It is fairly easy to differentiate lettuce drop from lettuce dieback because lettuce drop, caused by fungi in the genus Sclerotinia, results in a soft rot, outer leaves often flatten against the ground, and heads easily separate from the root, whereas with lettuce dieback the root remains firmly attached to the head. The two thrips-transmitted viruses, impatiens necrotic spot virus (INSV) and tomato spotted wilt virus (TSWV), also cause necrotic (dead) patches on leaves of infected lettuce plants that resemble symptoms of lettuce dieback, and therefore it can be difficult to differentiate the two diseases. Diagnostic tests can be used to differentiate lettuce plants infected with these viruses from those with lettuce dieback disease. Serological detection methods including commercially available immunostrips that can be used in the field to determine infection with INSV or TSWV, but immunostrips are not available for the viruses associated with lettuce dieback disease. Therefore, confirmation of lettuce dieback requires laboratory testing, which can include both molecular biology and serological methods. In some cases, lettuce plants may be infected by multiple pathogens simultaneously and this may complicate diagnosis.

Lettuce dieback is probably a very old disease of crisphead (iceberg) lettuce that disappeared for many years before reemerging with a new name as a disease of other lettuce types. In the 1930s a disease known as brown blight devastated lettuce production in California with symptoms that closely resembled those of lettuce dieback based on descriptions and illustrations at the time.

Iceberg lettuce was the main type of lettuce grown in the 1930s, and it suffered severe losses from brown blight for many years until a source of resistance was identified by a USDA scientist, Ivan Jagger. This source of resistance was eventually bred into all subsequent iceberg lettuce types, beginning with the variety Imperial, and this eliminated the threat from brown blight. In the early 2000s, after the appearance of lettuce dieback, USDA scientists identified a source of resistance to lettuce dieback from the crisphead lettuce variety Salinas, and through genetic studies found that the source of resistance to lettuce dieback is also present in the brown blight-resistant lettuces developed by Jagger over 70 years earlier, but was not in earlier susceptible lettuce varieties. In other words, only crisphead lettuce varieties that predate the variety Imperial could develop symptoms of lettuce dieback. This suggests the two diseases may actually be the same. The resistance to lettuce dieback has been incorporated into several romaine lettuce varieties, as well as some leaf and butter lettuce varieties, but there remain many lettuces that are susceptible to lettuce dieback disease.

Since the late 1990s, lettuce dieback has been believed to be caused by infection of lettuce plants with either of two viruses from the genus Tombusvirus; tomato bushy stunt virus (TBSV) and Moroccan pepper virus (MPV). These viruses are absent from healthy lettuce but have been found regularly in association with lettuce dieback disease. However, there have been numerous situations in which neither virus was found in association with obvious disease symptoms. Furthermore, it has not been possible to consistently and easily reproduce disease symptoms when lettuce is inoculated with either virus in a laboratory setting, raising the possibility that an additional virus may contribute to causing lettuce dieback disease.

In an attempt to identify a possible additional virus contributing to lettuce dieback disease, high throughput sequencing (HTS) was used on several lettuce plants exhibiting dieback symptoms, which led to the identification of a new virus consistently associated with diseased plants but not with healthy lettuce plants. This novel virus was most closely related to a recently identified and poorly characterized virus from watermelon in China, watermelon crinkle leaf associated virus, which was found using the same HTS approach.

The newly identified lettuce virus, tentatively named lettuce dieback associated virus (LDaV) shares an extremely low genetic relationship with the watermelon virus, which suggests that although the two viruses are related, they are very distantly related to one another. Using a combination of HTS and traditional DNA sequencing the genome of the new virus, LDaV, was assembled and methods were developed to allow rapid detection of the virus from lettuce leaf extracts using RT-PCR, a routine laboratory diagnostic method. LDaV has now been found not only in lettuce showing dieback symptoms collected recently, but it has also been found in older archived samples of lettuce nucleic acid collected from plants showing dieback symptoms over the past 20 years, including many that also contained MPV or TBSV. To date, LDaV has not been found in healthy lettuce plants. Interestingly, genetic comparison showed that LDaV isolates collected from coastal California production regions are closely related to one another, and desert isolates from Arizona and Imperial Valley, California also are closely related to one another. However, coastal and desert isolates differ genetically from one another, suggesting perhaps some regional adaptation of the virus to plants grown under the different climatic conditions.

Further research will clarify the role of LDaV in lettuce dieback disease and how it relates to the two tombusviruses, MPV and TBSV, that have long been linked to the disease. Studies to date, however, strongly suggest a role for LDaV in lettuce dieback disease development, and research is in progress to clarify the ability of LDaV to produce lettuce dieback symptoms when inoculated to lettuce plants, as well as whether or not the new virus can infect lettuce plants carrying a gene for resistance to lettuce dieback.

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