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Enhancing Diamondback Moth Management with Mating Disruption

By
Surendra K. Dara
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Brussels sprouts field in Santa Maria (photo by S.K. Dara.)

Brassica crops such as broccoli, brussels sprouts, cabbage, canola, cauliflower, collards, kale, kohlrabi, turnip and mustards are important vegetable or oilseed crops. The value of brassica vegetables, also known as cole crops, is more than $1.2 billion in California, which is the leading producer of these crops. Among various arthropod pests that attack brassica crops, the diamondback moth (DBM), Plutella xylostella (Lepidoptera: Plutellidae), is of significant importance. Thought to be of European origin, now with worldwide distribution, DBM exclusively feeds on cultivated and weedy crucifers. DBM can have up to 12 generations per year, especially under warmer climate.

Female moths deposit 150 eggs on average. Four larval instars feed on foliage and growing parts of young plants or bore into heads or flower buds, resulting in skeletonization of leaves, stunting of the plants or failure of head formation in some hosts. Pupation occurs on the lower surface of leaves or in florets. Adult moths are grayish-brown, and when at rest, a light-colored diamond-shaped pattern can be seen on the upper side of the wings.

Farmers typically rely on synthetic and biological insecticidal applications for controlling DBM. Multiple species of parasitoids and predatory arthropods also provide some control. Due to a heavy reliance on insecticidal control, DBM resistance to several insecticides is a common problem. Resistance of DBM to Bacillus thuringiensis (Ferré et al., 1991), abamectin (Pu et al., 2009), emamectin benzoate, indoxacarb and spinosad (Zhao et al., 2006), pyrethroids and other insecticides (Leibee and Savage, 1992; Endersby et al., 2011) have been reported from around the world. Excessive use of any kind of pesticide leads to resistance problems (Dara, 2020) to an individual pesticide or multiple pesticides.

Diamondback moth larva and adult (photos by Jack Kelly Clark, UC IPM.)

Integrated pest management (IPM) strategy encourages the use of various control options for maintaining pest control efficacy and reducing the risk of resistance development (Dara, 2019). Regularly monitoring pest populations to make treatment decisions, rotating pesticides with different modes of action, exploring the potential of biocontrol agents, and other non-chemical control approaches such as mating disruption with pheromones are some of the IPM strategies for controlling the DBM. While sex pheromones are effectively used to manage several lepidopteran pests and are proven to be a critical IPM tool, mating disruption is not fully explored for controlling DBM. A study was conducted in Brussels sprouts to evaluate the efficacy of a sprayable pheromone against the DBM and to enhance current IPM strategies.

 

Methodology

The study was conducted on a 10-acre Brussels sprouts field in Santa Maria. Cultivar Marte was planted in early July for harvesting in December 2020. A typical diamondback control program includes monitoring DBM populations with the help of sticky traps and lures and applying various combinations of biological and synthetic pesticides at regular intervals. This study evaluated the efficacy of adding CheckMate DBM-F to the grower standard practice of monitoring the DBM populations with traps and lures and applying pesticides. Treatments included 1.) grower standard pesticide program (See Table 1) grower standard pesticide program with two applications of 3.1 fl oz of CheckMate DBM-F on August 9 and September 11. Treatment materials were applied by a tractor-mounted sprayer using a 100 gpa spray volume and necessary buffering agents and surfactants. Each treatment was five acres and divided into four quadrants representing four replications.

Table 1. Pesticides, buffering agents and surfactants, their active ingredients, rates/ac (along with the IRAC mode of action groups) and retail pricing for those applied in the grower standard diamondback moth control program. *Applied diamondback moth and aphid control

In the middle of each quadrant, one Suterra Wing Trap was set up with a Trécé Pherocon Diamondback Moth Lure. Lures were replaced once a month in early September and early October. Sticky liners of the traps were replaced every week to count the number of moths trapped. Traps were placed on Aug. 1, 12 and 24, Sept. 1, 11, 18 and 27, and Oct 6, and the moth counts were taken from respective traps on Aug. 8 and 20, Sept. 1, 11, 18 and 27, and Oct. 6 and 15. CheckMate DBM-F was applied at 3.1 fl oz/ac on Aug. 9 and Sept. 11. The number of larvae and their feeding damage on a scale of 0 to 4 (where 0=no damage, 1=light damage, 2=moderate damage, 3=high damage, 4=extensive/irrecoverable) were recorded from 25 random plants within each replication on Aug. 30 and Oct. 6 and 18. Data were subjected to analysis of variance using Statistix software and significant means were separated using Tukey’s HSD test. The retail value of various pesticides was also obtained to compare the cost of treatments.

When CheckMate DBM-F[(Z)-11-Hexadecenal (3) , (Z)-11-Hexadecen-1-yl Acetate (1)] was applied the first time on Aug. 9, Dibrom 8 Emulsive was replaced with Warrior II, the buffering agent Quest was not used, and the surfactant Dyne-Amic was replaced with Induce (dimethylpolysiloxane) to avoid potential compatibility issues. The impact of this substitution is expected to be negligible within the scope of this study. The retail cost of 3.1 fl oz CheckMate DBM-F is $45.60. The cost of lures and traps would be about $4 to $8 per acre for a six-month crop like Brussels sprouts.

 

Results and Discussion

Traps in replication 4 in both treatments on August 8 and replication 1 in the grower standard were missing, probably knocked down by a tractor. The day before CheckMate DBM-F was first applied, the mean number of adult DMB caught was 227 in the grower standard and 271 in the plots that would receive the pheromone application (Figure 1). There was a gradual decline in moth counts during the rest of the observation period in both treatments. However, the decline was higher in the plots that received CheckMate DBM-F. The number of moths per trap were about 19% higher in the pheromone-treated plots compared to the grower standard before the study but were nearly 98% lower by the end of the study (Figure 2). The reduction in moth populations from mating disruption was significant on September 18 (P =0.039) and October 15 (P = 0.006).

Figure 1. Mean number of diamondback moth adults found in the traps.
Figure 2. Reduction in moth populations by adding pheromone for mating disruption.

The mean number of larvae per 25 plants in a replication was zero on all observation dates except for 0.01 on Aug. 30 in the plots that received CheckMate. Four insecticide applications made by the time the study was initiated and the remaining six applications effectively suppressed larval populations.

Larval feeding damage ratings were consistently low (P < 0.0001) in the plants that did not receive CheckMate DBM-F (Figure 3). The damage was limited to the older leaves at the bottom of the plants and must have been from early feeding before the initiation of the study. The lack of larvae and the evidence of new feeding damage also confirm that the crop remained healthy and pest-free.

Figure 3. Feeding damage by diamondback moth larvae.

 

Yield and Cost Comparisons

Since frequent pesticide applications effectively suppressed larval populations and prevented their feeding damage, the effectiveness of mating disruption on larval populations or their damage could not be determined in this study. Moths found in the traps probably developed from the larvae in the field or could have been those that flew in from other areas.

However, lower moth populations in CheckMate DBM-F treatment demonstrated the overall influence of mating disruption and pest suppression.

It is common to make about 10 to 12 pesticide sprays during the six-month crop cycle of Brussels sprouts. The cost of each application varied from about $73 to $192 depending on the materials used with an average cost of about $128 per application in this study. The cost of two CheckMate DBM-F applications is $91. If diamondback moth populations could be reduced with mating disruption, it is estimated that two to three pesticide applications could be eliminated. That results in $164 to $292 of saving for the pesticide costs and additional savings in the application costs per acre by investing $91 in the mating disruption. Since DBM can develop resistance to several chemical and natural pesticides, eliminating some applications as a result of mating disruption also contributes to resistance management along with potential negative impact of pesticides on the environment. Compared to other mating disruption strategies, a sprayable formulation compatible with other agricultural inputs is easier and more cost-effective to use.

The grower’s yield data showed 762 cartons/acre from the grower standard block with pesticides alone and 814 cartons/acre from the block that received pesticide and pheromone applications. Although there seems to be a 7% yield difference, since data from individual plots could not be collected for statistical analysis, the impact of DBM mating disruption on yield improvement is inconclusive.

This study demonstrated that mating disruption with CheckMate DBM-F will significantly enhance the current IPM practices by reducing pest populations, contributing to insecticide resistance management, and reducing pest management costs. Additional studies with fewer pesticide applications that allow larvae to survive and cause some damage might further help to understand the role of mating disruption where pest populations are not managed as effectively as in this field.

Thanks to the PCA and grower for their research collaboration, Tamas Zold for his technical assistance in data collection, Ingrid Schumann for market research of pesticide pricing and Suterra for the financial support.

Feeding damage in cauliflower (photo by S.K. Dara.)

 

References

Dara, S. K. 2019. The new integrated pest management paradigm for the modern age. J. Int. Pest Manag. 10: 12.
Dara, S. K. 2020. Arthropod resistance to biopesticides. Organic Farmer 3 (4): 16-19.
Endersby, N. M., K. Viduka, S. W. Baxter, J. Saw, D. G. Heckel, and S. W. McKechnie. 2011. Widespread pyrethroid resistance in Australian diamondback moth, Plutella xylostella (L.), is related to multiple mutations in the para soidum channel gene. Bull. Entomol. Res. 101: 393.
Ferré, J., M. D., Real, J. Van Rie, S. Jansens, and M. Peferoen. 1991. Resistance to the Bacillus thuringiensis bioinsecticide in a field population of Plutella xylostella is due to a change in a midgut membrane receptor. Proc. Nat. Acad. Sci. 88: 5119-5123.
Leibee, G. L. and K. E. Savage. 1992. Evaluation of selected insecticides for control of diamondback moth and cabbage looper in cabbage in Central Florida with observations on insecticide resistance in the diamondback moth. Fla. Entomol. 75: 585-591.
Pu, X., Y. Yang, S. Wu, and Y. Wu. 2009. Characterisation of abamectin resistance in a field-evolved multiresistant population of Plutella xylostella. Pest Manag. Sci. 66: 371-378.
Zhao, J-Z., H. L. Collins, Y-X. Li, R.F.L. Mau, G. D. Thompson, M. Hertlein, J. T. Andaloro, R. Boykin, and A. M. Shelton. 2006. Monitoring of diamondback moth (Lepidoptera: Plutellidae) resistance to spinosad, indoxacarb, and emamectin benzoate. J. Econ. Entomol. 99: 176-181.

Life After Methyl Bromide in California Berries

By
Warren E. Clark
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Finding the right fumigation and herbicide program can help reduce long-term labor costs (photo courtesy L. Stoeckle.)

When methyl bromide was banned in 2005, California strawberry growers lost an effective tool in their crop care toolbox to control weeds, soilborne diseases, nematodes and symphylans. Special-use permits allowed them to continue using the fumigant through 2016, but growers feared final loss of the powerful soil fumigant might be the end of profitable production.

The California Strawberry Commission agreed, noting that elimination of methyl bromide fumigation brought forth several soilborne diseases for which there is no post-plant control. Of particular concern were Fusarium wilt (Fusarium oxysporum f.sp. fragariae), Verticillium wilt (Verticillium dahlia) and charcoal rot (Macrophomina phaseolina), according to the California Strawberry Commission.

But once again, necessity proved to be the “mother of invention,” said independent agronomist and PCA Lee Stoeckle, owner of Stoeckle Agricultural Consulting in Ventura, Calif.

Stoeckle has advised strawberry and caneberry growers for more than 30 years, and he noted that effective alternative fumigants have taken up the methyl bromide void and are now widely and successfully used. While California strawberry acreage has fallen, total production has actually increased thanks to innovation and application of new technology.

Stoeckle’s family-owned business provides recommendations on 3,000 acres of strawberries and 100 acres of blackberries in Santa Barbara County and San Luis Obispo County and 2,000 acres of strawberries in Ventura County. In addition, he consults on production of 800 acres of strawberry and 250 acres of raspberries in Baja Mexico. While he is primarily responsible for above-ground insect and disease control, he doesn’t write fumigation recommendations but instead advises based on what he learns about weeds and soil pathogens.

 A strawberry field at the beginning of the growing season in Oxnard, California.

 

A One-Two Punch

Top problem soilborne diseases, according to Stoeckle, are Fusarium, Macrophomina, Phytophthora, Anthracnose and Verticillium wilt. Top problem above-ground pests include two-spotted spider mites, Lygus bug, Botrytis fruit rot/gray mold and powdery mildew, depending on varietal susceptibility. Other pests necessary to watch include worms and aphids.

“The optimum disease and weed control program is a one-two punch,” Stoeckle says. “Hit it at the end of the season as a burndown and at the beginning before next planting via drip irrigation. The backbone of our control program is Pic-Chlor 60 EC (1,3-dichloropropene plus chloropicrin) at the max recommended rate (350 lbs/ac), going after the heavy hitting ‘big boys’ (soilborne diseases) when applied in August or September before planting.” He recommends Vapam or K-Pam to get the additional benefits to control these diseases (Fusarium, Verticillium, Macrophomina phacelia) and eliminate the inoculum reservoir in the crown.

Stoeckle says emulsified formulations of Telone® C-35 and chloropicrin can be applied with irrigation water through drip irrigation systems. Metam sodium is the active ingredient in Vapam® HL™ and metam potassium in the active ingredient in K-Pam® HL™. Both are AMVAC® soil fumigants, which give off methyl isothiocyanate (MITC) when combined with water via drip or otherwise. If drip fumigation is planned, good results have been obtained with a sequential application of chloropicrin or 1,3-dichloropropene plus chloropicrin, followed 7 days later with metam sodium or metam potassium.

Stoeckle considers fumigants essential and offers the math: “If we equate the value of every plant to be $2 each and you lose 10% of your plants, that’s 2,500 plants on a population of 25,000 per acre. Your choice is to fumigate or take a $5,000 loss. That’s an easy decision. The return on investment on fumigation is huge. We expect to see a 20% to 30% return on fumigant investment.”

He is quick to add there are other management decisions that help maximize production, noting, “I could go on and on about using the best plastic mulch or optimum fertilization practices.”

 

Best Practice Weed Control Reduces Production Costs

Fifth-generation Santa Barbara County grower Brett Ferini, owner of Rancho Laguna Farms, grows 400 acres of strawberries (300 fall plant, 100 summer plant) and 20 acres (adding another 20 for 40 total) of blackberries. Plus, he grows 35 acres of organic blackberries. He used Vapam on blackberries in February 2020.

“Vapam worked out really well in controlling nutgrass/nutsedge,” he said. “Pic-Clor 60 has no effect.”

For weed and disease control, “we hit everything at fall pre-plant with Pic-Clor 60 through drip tape and seven days after that we follow with Vapam,” he said. “It does a terrific job on nutsedge and other weeds, as well as the soilborne diseases including Verticillium, Phytophthora and, anecdotally, Macrophomina.”

Ferini says the operation plans to add a Vapam burndown at the end of the 2020 growing season

“We definitely [have] cut weeding by 60% on summer plant,” he says. “Labor is our highest cost. We saw $1,050 savings per acre vs. our normal costs of $1,800/acre. The big test will be on the fall plant when we get more rain. Our fall weeding cost normally runs $2,800 to $3,000 per acre. Using Pic-Clor 60 followed by Vapam treatment, I expect weeding cost savings of $1,000 to $1,500 per acre.”

Also seeing results with soil fumigants is Santa Barbara grower Josh Ford. He is COO of Ocean Breeze Ag Management LLC in Ventura, which grows 450 acres of strawberries, 50 acres of blackberries, and 25 acres of raspberries. Ford’s biggest soilborne pests are Macrophomina, Fusarium, and Phytophthora cactorum. Nutgrass is his most difficult weed to control.

“We’ve been using soil fumigants for many years,” he says. “We were using methyl bromide, but now we apply chloropicrin once a year and K-Pam once to twice a season. If nut grass is a bad problem, we will knock it down at the end of the season with K-Pam and also pre-plant K-Pam. Our ROI is good when you consider the increased cost of labor to manually remove nut grass.”

New Findings on Limb Dieback of Figs in California

By
Themis J. Michailides, David Morgan and Giorgio Gusella
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Dr. Themis Michailides sampling symptomatic fig trees. (photo courtesy T. Michailides.)

Back in 2004, and again in recent years, there were concerns by fig growers mainly in Madera and Merced counties about an excessive killing of major branches of their fig trees (Figure 1). Visits to some orchards back then and recently indicated that indeed they had a major problem. Initial close examinations of the dead branches showed symptoms which were similar to another disease: branch wilt of walnut.

Figure 1. Left, Fig tree affected by severe limb dieback; top right, still active canker; bottom right, inactive canker (branch is dead) (all photos courtesy G. Gusella.)

The bark of dead fig branches had cracks and one could easily remove large pieces of the bark, exposing the woody tissues underneath which were covered by a black powder. Rubbing this black powder with your finger could easily remove masses of it (Figure 2). The inner surface of the broken and removed bark pieces were also black due to these powder masses. A lot of trees had many dead major branches while others had one or two dead along with other branches bearing chlorotic and thin canopy, distinct from the green and dense canopy of healthy branches.

Figure 2. Top, Neoscytalidium dimitiatum, the cause of limb dieback producing spores (arthrospores) as the mycelia dried up and separate to small segments under the bark; bottom, easily rubbing off the spores under the bark (photo courtesy Beth Teviotdale.)

 

Pathogen Activity

To collect samples, we cut some of the symptomatic branches close to the interface of dead and alive-looking (green) tissues. We noticed that in a cross section, the dead woody tissues were delineated from the healthy tissues by a dark brown line while the living woody tissues were white (Figure 1).

Slices of these woody tissues from the branches were taken, isolations were made in the laboratory and a fungus known to be a pathogen of woody tissues was consistently recovered. The name of this pathogen is Neoscytalidium dimidiatum, which is a new taxonomic name of Hendersonula toruloidea fungus, which represents the pathogen first reported to cause the branch wilt disease of walnut.

Checking the literature, the same fungus under a different name (i.e. Nattrassia mangifera) was reported in 1945 on commercial figs in California as well as on Ficus religiosa and Ficus bengalensis, causing dieback and trunk cankers. In addition to walnut branch wilt, which is a common disease of walnuts grown in the San Joaquin Valley, the same fungus was reported in causing branch wilt and dieback of poplar, eucalyptus and mango. In other reports, we found out that this pathogen can cause killing of major branches of walnut, ash trees and grapefruit. More recently, it has been reported causing cankers and hull rot of almond. On fig shoots, the pathogen grows and infects injured bark (i.e. mechanical wounds, wounds by hail or sunburn), invades the woody tissues and kills the branch. When the branch is killed, it dries and usually the bark cracks, exposing large masses of black spores. These are not true spores but are small segments of mycelia that become black as the tissues dry up and break down into small pieces, producing a layer of black powder under the bark. The fungus also produces pycnidia that protrude through small cracks of the bark (See Figure 3). However, it is the spores produced in masses by the breaking mycelia called arthrospores. that can be spread readily by air and/or splashing rain and can cause infections of pruning wounds and other injuries of branches.

Figure 3. Left, pycnidia protruding through bark cracks; right, arthrospores of Neoscytalidium dimidiatum causing limb dieback of fig (photos courtesy T. Michailedes.)

 

Survey of Affected Areas

Before doing pathogenicity studies with the Neoscytalidium fungus, we wanted to make sure that this fungus was found frequently throughout the area where fig trees showed similar symptoms to the ones we initially observed in Madera County. Therefore, a survey of 16 fig orchards with branch dieback symptoms, representing all the major fig varieties (Black Mission, Calimyrna, Conadria and the male trees (Roeding and Stanford caprifig varieties)) was done in Fresno, Madera and Kern counties. Neoscytalidium was isolated in all of these orchards.

Limb and branch samples from the majority of these orchards had 60% to 100% Neoscytalidium, while three had 7% to 11%, and two 26% to 32%. In 12 of these orchards, a second pathogen, Phomopsis spp., was isolated along with Neoscytalidium in the first year of the survey. Phomopsis sinarencis has been reported in California causing an epidemic on Kadota figs back in 1935 and in other countries as an important fig canker pathogen. By the third year of this survey, less Phomopsis was isolated, and, very recently, almost none was isolated, probably because the very susceptible Kadota variety is rarely now planted in California. Phomopsis is known as a pathogen fungus associated with canker diseases in many other crops around the world, but more investigations are needed to figure out its role in fig limb dieback.

 

Differences in Susceptibility

To determine if there were any differences in susceptibility to the limb dieback pathogen, we inoculated six cultivars directly in the field. We found that three months after inoculations, the cultivars Kadota, Black Mission and Sierra developed twice as long canker size than the cultivars Brown Turkey, Calimyrna and Conadria (Figure 4). Growers also reported that they see the problem to be more severe in Black Mission than other cultivars. Inoculations of six cultivars showed that Neoscytalidium is a plant pathogen that likes high temperatures. For instance, it cannot grow below 50 degrees F; its optimum temperature for growth is 90 to 95 degrees F, and it can even grow at 104 degrees F to some extent. Therefore, this fungus likes hot summer temperatures and prefers to infect sunburned branches and pruning wounds.

Figure 4. Susceptibility of various fig cultivars to limb dieback pathogen Neoscytalidium dimidiatum.

In experiments, we inoculated shoots of fig of different ages, including current growth (green) shoots, one-year, two-year and three-year-old shoots, by wounding and inoculating with either a mycelial plug or a spore suspension. Interestingly, the three-year-old shoots developed almost three-fold larger cankers than the cankers on current and the one-year-old shoots. This suggests that larger cuts in the field during pruning seem to be more susceptible to infection than cuts made in current or one-year-old shoots. Also, inoculations done in May, June and July resulted in larger cankers than those done from August to November. ‘

When we compared infection on pruning wounds done in winter vs. those done in summer, pruning wounds in the summer developed almost threefold larger cankers than those done during winter months. Therefore, it is recommended that pruning of figs should be done in winter when pruning wounds seem to be less susceptible. Figs can be protected from infections of the branch wilt pathogen if shoots are painted with whitewash to protect them from sunburn. Applying Surround® on shoots also protected the shoots from infection, and this is recommended to become a routine practice by fig growers. Figure 5 shows results of inoculation experiments done following various treatments in the field and artificial inoculation with the pathogen.

Although wounding by only mallet or only sunburn resulted in larger canker than the non-inoculated, un-wounded/un-treated shoots, the shoots that were damaged by mallet wounding and sunburn at the same time resulted in the longest cankers. White wash or spraying with Surround protected the shoots even after wounding with mallets and inoculation (Figure 5 and 6). Therefore, pruning that exposes the shoots to sunburn and or any other type of wounding should be avoided, and spraying with Surround will help protect the fig shoots from the limb dieback pathogen.

The authors thank the California Fig Institute for funding this research and a number of fig growers who allowed us to sample their orchards.

Figure 5. Effect of stress factors (mallet wounding and sunburn) and treatment with white wash affecting the severity of limb dieback of fig.

 

Figure 6. Effect of Surround® spray on the severity of limb dieback of fig.

Plastic Mulches Reduce Spotted-Wing Drosophila Infestation in Fall-Bearing Raspberry

By
Hanna McIntosh, Christelle Guédot and Amaya Atucha
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Experimental plots of plastic mulches in fall-bearing raspberries in Wisconsin (photo by H. McIntosh.)

Spotted-wing drosophila (SWD), Drosophila suzukii, is an invasive vinegar fly and a major pest of soft-skinned fruit crops. The fly was first detected in the continental U.S. in 20081 and has quickly spread from its native range in Eastern Asia throughout the U.S. and into most major fruit-producing regions of the world2. For small-scale fruit growers, damage from this pest substantially reduces the yield of marketable fruit, making susceptible crops challenging to grow economically and sustainably3,4. For large-scale growers, the presence of SWD can lead to complete crop loss due to processors’ zero-tolerance policies for insect infestation5.

 

Biology

Vinegar flies typically lay their eggs in damaged or rotting fruit, but female SWD have a highly serrated ovipositor that allows them to saw through the skin of undamaged, ripening fruit6,7, which makes SWD an especially detrimental pest. Larvae emerge inside of and feed on the fruit, making it mushy and unmarketable. Recent research showed that around 80% of larvae drop from the fruit to pupate in the top layer of soil and that they are more likely to drop from the fruit if the fruit is overcrowded8,9. Larvae and pupae can also reach the ground when damaged fruit becomes mushy and falls to the ground.

SWD has a quick generation time, with many generations per year in most regions. The fly develops fastest in temperatures between 68 to 83 degrees F, but is unable to develop at temperatures above about 87 degrees F10,11. In temperate regions like the Upper Midwest or Pacific Northwest, SWD populations are highest during summer months. In hotter regions like California or Florida, fly populations are highest in the spring and fall and are much lower during their hot and dry summer months12.

SWD thrives in high humidity, developing fastest around 94% humidity13. Researchers found that females laid more eggs in the inner canopy of blackberry and blueberry plants, likely because the environment is more humid, cooler and darker14,15.

(Left) Spotted-wing drosophila adult female and male on a raspberry. Male flies are easy to identify by the large black spots on their wings. (Right) Spotted-wing drosophila females have a serrated ovipositor that allows them to lay eggs in undamaged, ripening fruit (photos courtesy Agri-Mag and Chris Thomas.)

Fruit crops that are the most susceptible to SWD include raspberries, blackberries, blueberries, strawberries, sweet and tart cherries, and some cultivars of wine grapes16-18. However, SWD can survive on alternative hosts like wild blackberries and apples, buckthorn and honeysuckle. It remains largely unknown how SWD survive in the winter and spring before fruit is available in the landscape and on farms, but one study found that SWD can develop on non-fruit hosts like mushrooms and bird manure19.

 

Traditional Management

Pest pressure from SWD is often very high due to the fly’s fast development time, optimal development conditions in the summer and high availability of host plants and food in the agroecosystem. Management relies heavily on chemical control in organic and conventional systems, which is costly to growers. In California, chemical controls for SWD cost around $470/acre for conventional and $1,210/acre for organic growers3.

Only a few insecticides approved for use in organic systems are effective at controlling SWD, limiting organic growers’ options for control20. Unfortunately, recent reports show evidence of insecticide resistance developing for some active ingredients in some regions, including spinosad (the main insecticide used to control SWD in organic systems)21,22.

Cultural practices can help reduce the fly’s population and are often used in tandem with chemical controls. Such practices include harvesting fruit promptly (every one to two days), frequent field sanitation, burial or composting of infested fruit and exclusion netting23,24. However, these methods are labor-intensive and expensive.

Since SWD is sensitive to temperature and humidity, cultural practices that modify the crop canopy microclimate have the potential to reduce infestation by deterring adults from laying eggs or disrupting larval development inside of fruit. Management strategies for SWD typically target adult flies in the canopy, but since the majority of SWD larvae fall to the ground before pupation, ground-based cultural management practices could also be important for reducing populations.

Spotted-wing drosophila life cycle (courtesy Jana Lee, USDA-ARS.)

Growers have used plastic mulches since the 1960s to modify the microclimate in fruit and vegetable agroecosystems. Plastic mulches are commonly used for weed control, promoting earlier ripening, improving fruit quality or color and increasing yield25,26. Different colors of plastic mulches have also been shown to successfully control insect pests including aphids, whiteflies, Asian citrus psyllid and Mexican bean beetles27-30.

 

Plastic Mulches for SWD

Based on the extensive body of literature reporting that plastic mulches can modify the crop microclimate, control some insect pests and provide other horticultural benefits, we tested the impact of three colors of plastic mulches on SWD adult and larval populations. Our study was conducted in 2019 and 2020 on a small fruit and vegetable farm in South Central Wisconsin in fall-bearing raspberries.

In this study, we tested black and white-on-black biodegradable plastic mulches (Organix Solutions AG film), metallic polyethylene mulch (Imaflex SHINE N’ RIPE) and a grower-standard control where grass filled in the space between the alleyway and the raspberry plants. We assessed the three mulches’ impact on SWD adult and larval populations in fall-bearing raspberry.

We laid the mulches by hand when the raspberry canes were just emerging from the soil in late April. We laid two mulch strips (25 feet long by 2.3 feet wide) along each side of the row, leaving a six-inch gap between the strips for the canes to grow. The edges of the mulches were secured with biodegradable sod stakes. All four treatments were randomly distributed in each of four rows of fall-bearing raspberries (cultivars “Polana” and “Caroline”), totaling 16 plots.

Starting when the first flies were detected in June, we measured the adult SWD populations passively using clear sticky cards placed in the fruiting zone, which were replaced weekly to estimate fly populations by week.

Larval infestation of fruit was evaluated by counting the number of larvae using the salt float method31. The evaluations were done two to four times per month starting in August.

Adult and larval populations were measured throughout the season until adult populations reached zero, usually in mid-October.

We also did a preliminary experiment to test whether plastic mulches could kill larvae that fell onto the mulch surface. We put lab-reared larvae into ‘corrals’ made from plastic sandwich containers and recorded their mortality and movement over three hours.

‘Corrals’ made from plastic sandwich containers used to test mortality of larvae on the mulch surface (photo by H. McIntosh.)

 

Population Reductions

In both years of our study, we found significantly lower SWD populations above all three plastic mulches compared to the control plots. Over the two-year period, the black and metallic mulches reduced the adult population of SWD by 51% and the white mulch reduced flies by 42% compared to the control.

Interestingly, the plastic mulches only reduced female fly populations and did not impact the number of male flies caught on the sticky cards. With fewer female flies in the canopy above the plastic mulches, it was unsurprising that we also found fewer larvae infesting the fruit in the mulched plots. Over the two-year study, the black mulch decreased the number of larvae in fruit by 72%, the metallic mulch by 61%, and the white mulch by 52% compared to the control.

Plastic mulches may be more effective than other types of mulches tested for managing SWD. In our study, we recorded the lowest adult fly populations and larval infestation of fruit above the black plastic mulch. A 2019 study tested black fabric weedmat as a cultural control for SWD in blueberry in several states and found no effect of the weedmat on SWD infestation of blueberries32. It is possible that some quality of the plastic mulch material (such as reflectivity or lack of permeability) makes it more deterrent to SWD than the weedmat.

In our preliminary experiment, larvae placed on the plastic mulches died quickly. Larvae on the black mulch died in less than one hour, and larvae on the white and metallic mulches died in less than three hours. We recorded high surface temperatures on the mulches, with all mulches heating up above 87 degrees F (SWD’s threshold for development) for two to four hours each day. On hot days, the black mulch got above 150 degrees F.

When placed on the mulch, we observed larvae struggling to crawl and visibly desiccating within minutes, making it unlikely that larvae could crawl off the mulch into the safety of the soil. We will collect more data in summer 2021 to confirm these promising results.

The results of our study provide evidence that black, white and metallic plastic mulches can reduce SWD adult and larval populations in fall-bearing raspberry in the Upper Midwest, showing promise for use of plastic mulches in sustainable pest management.

Combining plastic mulches with other cultural practices including short harvest intervals (every one to two days) and frequent field sanitation could have an additive effect on reducing SWD populations, potentially reducing the need for chemical controls in conventional and organic cropping systems.

 

Next Steps

Although the use of plastic mulches reduces SWD populations in the canopy of raspberry plants, the specific mechanisms causing this reduction are still unknown. We are still investigating how canopy light conditions, temperature and humidity are
influenced by the plastic mulches and whether these factors can explain the reduction in SWD populations we measured.

In the next two years of this project, we will conduct field experiments to determine whether the three plastic mulches we tested influence beneficial insects, including pollinators, and how the mulches impact soil health, raspberry plant growth, fruit quality and yield in Wisconsin’s climate.

Testing these mulches in other regions and fruit crops is warranted to determine if the reduction of SWD is maintained in different climates and other susceptible crops. Overall, plastic mulches are a promising new tool for more sustainable management of SWD in raspberry in the Upper Midwest.

 

References

1. Hauser, M. A historic account of the invasion of Drosophila suzukii (Matsumura) (Diptera: Drosophilidae) in the continental United States, with remarks on their identification. Pest Manag. Sci. 2011, 67, 1352–1357, doi:10.1002/ps.2265.
2. CABI Drosophila suzukii (spotted wing drosophila); 2016;
3. Farnsworth, D.; Hamby, K.A.; Bolda, M.; Goodhue, R.E.; Williams, J.C.; Zalom, F.G. Economic analysis of revenue losses and control costs associated with the spotted wing drosophila, Drosophila suzukii (Matsumura), in the California raspberry industry. Pest Manag. Sci. 2017, 73, 1083–1090, doi:10.1002/ps.4497.
4. DiGiacomo, G.; Hadrich, J.; Hutchison, W.D.; Peterson, H.; Rogers, M. Economic Impact of Spotted Wing Drosophila (Diptera: Drosophilidae) Yield Loss on Minnesota Raspberry Farms: A Grower Survey. J. Integr. Pest Manag. 2019, 10, doi:10.1093/jipm/pmz006.
5. Bruck, D.J.; Bolda, M.; Tanigoshi, L.; Klick, J.; Kleiber, J.; Defrancesco, J.; Gerdeman, B.; Spitler, H. Laboratory and field comparisons of insecticides to reduce infestation of Drosophila suzukii in berry crops. Pest Manag. Sci. 2011, 67, 1375–1385, doi:10.1002/ps.2242.
6. Kanzawa, T. Studies on Drosophila suzukii Mats. J. Plant Prot. 1939, 23, 66–70, 127–132, 183–191.
7. Walsh, D.B.; Bolda, M.P.; Goodhue, R.E.; Dreves, A.J.; Lee, J.; Bruck, D.J.; Walton, V.M.; O’Neal, S.D.; Zalom, F.G. Drosophila suzukii (Diptera: Drosophilidae): Invasive Pest of Ripening Soft Fruit Expanding its Geographic Range and Damage Potential. J. Integr. Pest Manag. 2011, 2, G1–G7, doi:10.1603/ipm10010.
8. Woltz, J.M.; Lee, J.C. Pupation behavior and larval and pupal biocontrol of Drosophila suzukii in the field. Biol. Control 2017, 110, 62–69, doi:10.1016/j.biocontrol.2017.04.007.
9. Bezerra Da Silva, C.S.; Park, K.R.; Blood, R.A.; Walton, V.M. Intraspecific Competition Affects the Pupation Behavior of Spotted-Wing Drosophila (Drosophila suzukii). Sci. Rep. 2019, 9, 7775, doi:10.1038/s41598-019-44248-6.
10. Ryan, G.D.; Emiljanowicz, L.; Wilkinson, F.; Kornya, M.; Newman, J.A. Thermal tolerances of the spotted-wing drosophila drosophila suzukii (Diptera: Drosophilidae). J. Econ. Entomol. 2016, 109, 746–752, doi:10.1093/jee/tow006.
11. Hamby, K.A.; E. Bellamy, D.; Chiu, J.C.; Lee, J.C.; Walton, V.M.; Wiman, N.G.; York, R.M.; Biondi, A. Biotic and abiotic factors impacting development, behavior, phenology, and reproductive biology of Drosophila suzukii. J. Pest Sci. (2004). 2016, 89, 605–619.
12. Wang, X.G.; Stewart, T.J.; Biondi, A.; Chavez, B.A.; Ingels, C.; Caprile, J.; Grant, J.A.; Walton, V.M.; Daane, K.M. Population dynamics and ecology of Drosophila suzukii in Central California. J. Pest Sci. (2004). 2016, 89, 701–712, doi:10.1007/s10340-016-0747-6.
13. Tochen, S.; Woltz, J.M.; Dalton, D.T.; Lee, J.C.; Wiman, N.G.; Walton, V.M. Humidity affects populations of Drosophila suzukii (Diptera: Drosophilidae) in blueberry. J. Appl. Entomol. 2016, 140, 47–57, doi:10.1111/jen.12247.
14. Diepenbrock, L.M.; Burrack, H.J. Variation of within-crop microhabitat use by Drosophila suzukii (Diptera: Drosophilidae) in blackberry. J. Appl. Entomol. 2017, 141, 1–7, doi:10.1111/jen.12335.
15. Evans, R.K.; Toews, M.D.; Sial, A.A. Diel periodicity of Drosophila suzukii (Diptera: Drosophilidae) under field conditions. PLoS One 2017, 12, doi:10.1371/journal.pone.0171718.
16. Lee, J.C.; Bruck, D.J.; Curry, H.; Edwards, D.; Haviland, D.R.; Van Steenwyk, R.A.; Yorgey, B.M. The susceptibility of small fruits and cherries to the spotted-wing drosophila, Drosophila suzukii. Pest Manag. Sci. 2011, 67, 1358–1367, doi:10.1002/ps.2225.
17. Kamiyama, M.T.; Guedot, C. Varietal and Developmental Susceptibility of Tart Cherry (Rosales: Rosaceae) to Drosophila suzukii (Diptera: Drosophilidae). J. Econ. Entomol. 2019, 112, 1789–1797, doi:10.1093/jee/toz102.
18. Pelton, E.; Gratton, C.; Guédot, C. Susceptibility of cold hardy grapes to Drosophila suzukii (Diptera: Drosophilidae). J. Appl. Entomol. 2017, 141, 644–652, doi:10.1111/jen.12384.
19. Stockton, D.G.; Brown, R.; Loeb, G.M. Not berry hungry? Discovering the hidden food sources of a small fruit specialist, Drosophila suzukii. Ecol. Entomol. 2019, een.12766, doi:10.1111/een.12766.
20. Sial, A.A.; Roubos, C.R.; Gautam, B.K.; Fanning, P.D.; Van Timmeren, S.; Spies, J.; Petran, A.; Rogers, M.A.; Liburd, O.E.; Little, B.A.; et al. Evaluation of organic insecticides for management of spotted-wing drosophila (Drosophila suzukii) in berry crops. J. Appl. Entomol. 2019, 143, 593–608, doi:10.1111/jen.12629.
21. Gress, B.E.; Zalom, F.G. Identification and risk assessment of spinosad resistance in a California population of Drosophila suzukii. Pest Manag. Sci. 2019, 75, 1270–1276, doi:10.1002/ps.5240.
22. Van Timmeren, S.; Mota-Sanchez, D.; Wise, J.C.; Isaacs, R. Baseline susceptibility of spotted wing Drosophila (Drosophila suzukii) to four key insecticide classes. Pest Manag. Sci. 2018, 74, 78–87, doi:10.1002/ps.4702.
23. Leach, H.; Van Timmeren, S.; Isaacs, R. Exclusion Netting Delays and Reduces Drosophila suzukii (Diptera: Drosophilidae) Infestation in Raspberries. J. Econ. Entomol. 2016, 109, 2151–2158, doi:10.1093/jee/tow157.
24. Leach, H.; Moses, J.; Hanson, E.; Fanning, P.; Isaacs, R. Rapid harvest schedules and fruit removal as non-chemical approaches for managing spotted wing Drosophila. J. Pest Sci. (2004). 2018, 91, 219–226, doi:10.1007/s10340-017-0873-9.
25. Tarara, J. Microclimate modification with plastic mulch. HortScience 2000, 35.
26. Kasirajan, S.; Ngouajio, M. Polyethylene and biodegradable mulches for agricultural applications: A review. Agron. Sustain. Dev. 2012, 32, 501–529, doi:10.1007/s13593-011-0068-3.
27. Greer, L.; Dole, J.M. Aluminum foil, aluminium-painted, plastic, and degradable mulches increase yields and decrease insect-vectored viral diseases of vegetables. Horttechnology 2003, 13, 276–284.
28. Croxton, S.D.; Stansly, P.A. Metalized polyethylene mulch to repel Asian citrus psyllid, slow spread of huanglongbing and improve growth of new citrus plantings. Pest Manag. Sci. 2014, 70, 318–323, doi:10.1002/ps.3566.
29. Nottingham, L.B.; Kuhar, T.P. Reflective Polyethylene Mulch Reduces Mexican Bean Beetle (Coleoptera: Coccinellidae) Densities and Damage in Snap Beans. J. Econ. Entomol. 2016, 109, 1785–1792, doi:10.1093/jee/tow144.
30. Nottingham, L.B.; Beers, E.H. Management of Pear Psylla (Hemiptera: Psyllidae) Using Reflective Plastic Mulch. J. Econ. Entomol. 2020, doi:10.1093/jee/toaa241.
31. Dreves, A.; Cave, A.; Lee, J. A Detailed Guide for Testing Fruit for the Presence of Spotted Wing Drosophila (SWD) Larvae. Oregon State Univ. Ext. Serv. 2014, EM 9096, 1–9.
32. Rendon, D.; Hamby, K.A.; Arsenault-Benoit, A.L.; Taylor, C.M.; Evans, R.K.; Roubos, C.R.; Sial, A.A.; Rogers, M.; Petran, A.; Van Timmeren, S.; et al. Mulching as a cultural control strategy for Drosophila suzukii in blueberry . Pest Manag. Sci. 2019, doi:10.1002/ps.5512.

A Review of Pythium Diseases in Row Crops

By
Steven Koike
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Large, established cauliflower plants can still become infected with Pythium (left), resulting in loss of roots, severe stunting and yield loss (all photos courtesy S. Koike.)

It is highly likely that growers, PCAs and other field professionals are familiar with the word “Pythium”. Pythium is the name of a soilborne, fungus-like organism that is notorious for primarily causing seedling diseases. Pythium is notable because many row crops are susceptible to it, the pathogen is very widely distributed and occurs in most cropped ground, and despite the use of IPM tools and strategies, Pythium problems can still show up in row crop production systems.

 

What is Pythium?

Pythium is a fungus-like organism. Previously considered to be a true fungus, molecular studies in recent years indicate that Pythium—as well as closely related organisms like Phytophthora and downy mildew—is more closely related to brown algae and diatoms. Formally, therefore, Pythium species are no longer part of the fungal taxonomic group but are classified in the kingdom Chromista, or Stramenopila. The Pythium genus contains over 200 species, most of which are not plant pathogens. There are Pythium species that are pathogens of animals (some of which can infect humans), and many species are saprophytes and only grow on dead and decaying organic material. Pythium species are mostly found in soil environments but are also present in aquatic habitats.

Plant pathogenic Pythium species are well equipped to cause problems on row crops. Most of these species form resilient, thick-walled sexual spores (oospores) that can withstand periods of unfavorable dry and warm conditions. These structures enable Pythium to persist in the soil for a long time. When favorable soil conditions are present, mostly in the form of abundant soil water, these Pythium organisms either produce hyphae that grow toward the roots or swimming spores (zoospores) that move through the soil water in search of susceptible plant tissues. Another feature that makes Pythium problematic for growers is the extremely fast growth rate of these organisms. Given suitable soil conditions, Pythium pathogens can rapidly grow from seed-to-seed, seedling-to-seedling and root-to-root.

The primary symptom of Pythium diseases is the dark discoloration and decay of roots, pictured here on lettuce.

 

 

Diverse Pythium Diseases

In contrast to many plant pathogens, Pythium causes several different types of problems on crops (Table 1). First, Pythium is a seed pathogen. Once placed in the ground, seed can be exposed to Pythium that is residing in the soil. If conditions are favorable for the pathogen, Pythium can invade and colonize the seed, causing it to rot before it can germinate. If the seed germinates, Pythium can cause a decay of the roots and shoots that just grew out of the seed. This early disease stage is often called damping-off. Damping-off is further divided into two phases. If the newly germinated seedling is infected so early and so severely that it dies before being able to break through the soil surface, this situation is called pre-emergent damping-off. However, post-emergent damping-off occurs if the diseased seedling is strong enough to emerge above the soil surface, only to succumb and collapse shortly afterwards. Collectively, seed decay, pre-emergent damping-off, and post-emergent damping-off can result in loss of plants very early in the production cycle, causing stand loss in the field.

Table 1. Categories of Pythium diseases of row crops

Healthy seedlings that escape death at the seed and newly germinated stages remain vulnerable to this pathogen; established seedlings can still be infected and become stunted and die due to diseased roots and crowns. Older, established plants have escaped the damping-off phase that kills seedlings but can be subject to infections that prune back the roots, leading to reduced plant vigor and yield. For example, Pythium can cause late infections in cauliflower and result in weakened roots and poorly yielding plants. This soilborne pathogen can even cause a foliar blight of leaves and shoots, though this type of disease is not very common. Bits of soil carrying Pythium can be splashed or moved up onto foliage and cause blights on crops such as spinach and bean. Finally, the fleshy parts of some vegetable crops are subject to Pythium pathogens. If in contact with infested soil, cucurbit fruits, sweet potato storage roots and potato tubers can develop a soft, watery rot that will result in a non-marketable commodity.

Of the hundreds of Pythium species worldwide, relatively few species infect row crops. These plant pathogens can be conveniently placed into two categories. One group consists of Pythium species that have a relatively narrow host range and infect only a few crops, with those few crops tending to mostly be within a particular plant family. Examples are Pythium mastophorum, which primarily infects celery and parsley (Apiaceae family), and Pythium uncinulatum, which reportedly only causes significant disease on lettuce (Table 2). The second group contains Pythium organisms that have very large host ranges. The two main species, P. aphanidermatum and P. ultimum, both infect scores of plants, including dozens of vegetable and row crops.

Table 2. Examples of Pythium pathogens with broad vs. narrow host ranges

Disease Development Development of Pythium diseases is straightforward. Initial inoculum is almost always linked with infested field soils and associated soil water. Pythium is a soilborne pathogen that resides in the soil primarily as dormant resting structures. Pythium inoculum is not seedborne or airborne. For Pythium to become active, grow, and produce those swimming zoospores, the soil must be wet for prolonged periods. Once susceptible seed, seedlings, and other plant parts are in close contact with Pythium inoculum, infection can take place and disease will be initiated. If wet soil conditions persist and temperatures are optimum for the pathogen, disease losses can be significant.

Pythium pathogens form thick-walled oospores that enable the pathogen to survive in soil for prolonged periods.

 

Diagnostic Considerations

Pythium is not the only soilborne pathogen that causes seedling damping-off and root rots of row crops. On spinach, damping-off and root rot can be caused by both Pythium and Fusarium; visually, one cannot distinguish between the symptoms caused by these two pathogens. Pythium and Phytophthora pathogens both cause dark, discolored roots of lettuce and cannot be differentiated in the field. Cauliflower transplants are susceptible to both Pythium and Rhizoctonia pathogens, both of which caused the roots to become discolored. Precise and accurate diagnosis of Pythium diseases will therefore require lab-based tests and assays.

When sufficient soil water is present, Pythium forms swimming spores that are released and search for host roots. Pictured here is a cluster of zoospores just prior to release.

 

Managing Pythium

Controlling diseases caused by Pythium requires the implementation of IPM practices.

Site selection: Choose to plant in fields that do not have a history of Pythium problems and have well-draining soils.

Crop rotation: If Pythium is an issue, avoid planting the same susceptible crop in the infested field. Rotate to crops that are not known to be susceptible to the Pythium species present at that location. However, remember that some Pythium species have very broad host ranges (Table 2).

Irrigation management: Because the Pythium pathogen is so strongly dependent on wet soil conditions, carefully schedule and limit irrigations to prevent overwatered, saturated soils.

Time of planting: In some cases, moving the planting date to a different time of year may help reduce losses to Pythium. For example, depending on the Pythium species of concern, planting the crop in the warmer, drier summer may be preferred to seeding the crop in the cooler, wetter spring.

Fungicides: Plant seed treated with a fungicide that is active against Pythium. Note that the fungicides used to control Rhizoctonia or Fusarium have no effect on Pythium. For some crops, applying fungicides to the emergent crop may provide additional protection. The repeated use of products having the same mode of action can result in Pythium isolates that are insensitive (=resistant) to those products; therefore, IPM strategies will require that thought be given to deploying different fungicides.

Resistant or tolerant cultivars: Unfortunately, there do not appear to be any row crop cultivars that have genetic resistance to Pythium.

Pythium plant pathogens can grow very rapidly. Pictured here are three-day-old cultures of Pythium, Phytophthora, Fusarium and Verticillium. The diameter of the petri dish is 85 mm.

Using N-Rich Reference Zones to Inform In-Season Nitrogen Fertilization Practices in California Small Grains

By
Michelle Leinfelder-Miles, Nick Clark, Taylor Nelsen, Tom Getts, Konrad Mathesius, Sarah Light, Giuliano Galdi and Mark Lundy
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Figure 1. The GreenSeeker held above a recently headed small grain and displaying the NDVI value. Values range from 0 to 1 (i.e. less-green to very green plants).

Over the last year, a team from UCCE has been working with California small grains growers on practices that can improve nitrogen (N) use efficiency. At demonstration sites, we have implemented practices that UC Grain Cropping Systems Specialist Mark Lundy has been investigating for several years, namely N-rich reference zones, a soil nitrate quick test, handheld reflectance devices and aerial imagery. We demonstrate how to use these tools to manage N fertilizers in small grain crops across variable soil and climatic conditions in the Sacramento Valley, Delta, San Joaquin Valley and Intermountain Region.

The demonstrations are funded by the CDFA Fertilizer Research and Education Program and a USDA-NRCS California Conservation Innovation Grant. Our goal is to help growers and consultants learn and implement these practices to guide N fertilization in small grains, thereby increasing crop productivity and N use efficiency while reducing potential for N loss to the environment.

 

What are “N-Rich Reference Zones”?

Reference zones are most useful to growers who can apply the majority of their seasonal N budget during or after the tillering stage of growth. Previous work has shown that N fertilizer applied during the season−between the tillering and heading stages of small grain development−results in higher yields, higher protein and increased fertilizer use efficiency compared to pre-plant applications. The reference zone is a relatively small area within the field where extra N fertilizer is added at the beginning of the season. This extra fertilizer ensures that the reference zone will not be N-limited from planting until an in-season fertilizer decision is made. When a grower is determining whether and how much N fertilizer to add in-season, measurements from both the reference zone and the broader field are compared to understand whether the broader field is sufficient in plant-available N.

 

Fertilizer N Rate and Field Variability

Fertilizer N rate and field variability are two important considerations when creating N-rich reference zones. The amount of N to apply in the N-rich zone will depend on several factors such as yield goal, protein goal and when the expected in-season fertilizer application will take place. There should be sufficient N applied to the reference zone at planting to ensure that the plants in the zone are not limited by N at the stages of growth when the in-season fertilizer is applied. Table 1 gives some examples of how much N fertilizer to apply to the N-rich zone for a range of potential yields.

Table 1. Approximate N fertilizer application rates suggested for use in N-rich reference zones based on a range of average yields and two stages of crop growth. Suggested rates ensure that crops within the reference zone are not N-limited when an in-season fertilizer application decision is being made at the crop stage indicated.

It is important to establish the N-rich zones in representative parts of the field. Areas of the field that are unique (i.e. low areas, high areas, gravel strips, etc.) should be avoided. It is also important that the zones capture field variability. If certain areas have distinct soil types or known patterns of yield or management differences, a grower should establish multiple zones to account for these sources of spatial variability if they represent large areas in the field. Soil maps (available from casoilresource.lawr.ucdavis.edu/soilweb-apps/) and historical aerial imagery can often help in identifying field patterns and good location(s) for reference zones.

 

How and When to Apply the N-Rich Zone Fertilizer

A grower can establish N-rich zones during the pre-plant fertilizer application. For example, a grower may apply 50 pounds N per acre across the field and then make another pass or two in the zone to apply an additional 50 to 100 pounds N per acre (depending on what the grower calculates is necessary, as described above.) This method might be most easily adopted by growers. We have observed, however, that if the fertilizer is placed too deep in the soil profile, the N may not be readily available to the seedling crop early in the season because it is below the root zone. Therefore, N-rich zones established by this method may not provide a reliable early-season point of comparison. Instead, we have found that broadcasting urea is the most effective way to establish N-rich zones. At our demonstration sites, we broadcasted urea after tillage or shortly after planting, but always ahead of a storm or irrigation event that could incorporate the fertilizer. Orienting the zones perpendicular to the rows or tractor passes also helps to capture field variability. When the zones are too narrow and run in the same direction as the field work, it can be hard to differentiate between a field pattern associated with equipment passes and a N effect, particularly early in the season.

 

Monitoring the Field

Once the crop begins to grow, the field should be monitored periodically to assess whether the crop is likely to respond to a N fertilizer application. A combination of the soil nitrate quick test (SNQT) and plant reflectance measurements taken from both the N-rich zones and the broader field can indicate when a top-dress fertilizer application may be beneficial. The soil nitrate quick test and plant reflectance measurements complement other important information like current crop growth stage, crop yield and protein goals, and local weather records to inform a site-specific N fertilizer recommendation.

The SNQT is a simple and low-cost test that provides a ballpark estimate of the soil nitrate-N concentration in the root zone. Nitrate is a highly plant-available form of N. Using the SNQT when N fertilizer decisions are being made will help to narrow a range of fertilizer rates appropriate for that field. More information on using the SNQT in small grains, including a sample protocol and demonstration video, is available at smallgrains.ucanr.edu/Nutrient_Management/snqt/. Over the past several years, UCCE agronomists have developed a strong relationship between the value measured using the SNQT and an estimate of fertilizer N equivalence.

Crop reflectance can be measured using a number of tools, including handheld devices, drones and satellite imagery. Common indices that result from measurements of canopy reflectance are normalized difference vegetation index (NDVI) and normalized difference red edge index (NDRE). These indices represent measurements of light reflected from the crop canopy at key wavelengths indicative of plant vigor. Relative differences in vigor among plants in the same field can be captured by comparing canopy reflectance measurements like NDVI and NDRE. We have been using handheld devices, drones and satellite imagery at our demonstration sites to compare crop reflectance values in the N-rich zones and the broader field.

One of the tools we are using is the GreenSeeker by Trimble Agriculture. This is a hand-held NDVI meter (See Figure 1) that emits light and detects how much is reflected from the crop canopy in the red and infrared wavelengths. The GreenSeeker’s canopy measurement indicates how well the plants are growing and covering the soil with greenness. This information about vigor is important early in the crop’s growth because it indicates the ability of plants to support grain production and yield potential.

We are obtaining similar information as from the GreenSeeker by measuring NDRE with a five-band multispectral camera (MicaSense RedEdge-MX) mounted on a drone (DJI Matrice M200 V2). NDRE is similar to NDVI but replaces the reflectance from the red wavelength with reflectance from the red edge wavelength. Because the drone is able to capture data from hundreds of feet above the ground, it allows us to measure a large area quickly and under conditions when entering the field is not possible. Figure 2 depicts side-by-side images from a field in Solano County where N-rich reference zones were implemented during the 2019-20 season.

Figure 2. A field in Solano County where three N-rich reference zones are visible at tillering using NDRE captured via drone (left), but not visible to the naked eye (right) (all photos courtesy M. Leinfelder-Miles.)

Another device we are using to monitor plant N is the atLEAF CHL by FT Green LLC, which is a chlorophyll meter that measures light absorbed by a single leaf (Figure 3). Like the GreenSeeker, it also emits and detects light. The atLEAF CHL, however, measures how much light passes through a single leaf instead of measuring reflected light. This information becomes increasingly valuable as an indicator of whether or not the crop has sufficient N as it begins heading out and filling grain.

Step-by-step instructions for using both the GreenSeeker and atLEAF CHL in small grains are available at ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=42903.

Since plant N is strongly related to plant greenness and chlorophyll content, measurements of NDVI, NDRE and leaf chlorophyll can serve as proxies for relative plant N status within a field. Many factors can affect absolute greenness or chlorophyll values, including variety, crop injury and environmental factors. Because of this, it is important to remember that the absolute values given by these devices are only meaningful when compared to a reference zone like the N-rich zone.

Figure 3. Small grain leaf inserted in the sampling area of the at LEAF showing a chlorophyll reading in the lower right corner of the display while the user’s back shades the device.

 

What do the Readings Mean?

Plant reflectance and transmittance measurements are best interpreted by expressing values measured in the broader field relative to the N-rich reference zones, according to the following equation:

Relative value= (Production area value)/(N-rich zone value)

The relative value is sometimes referred to as a Sufficiency Index (SI) and will usually result in a decimal value between 0 and 1. When the SI is below a certain threshold, it indicates that the production area is experiencing detectable N deficiency relative to the N-rich zone. Table 2 shows SI ranges for proximal and remotely-sensed data and the associated plant N status.

Table 2. Sufficiency Index (SI) values and associated plant N sufficiency status, calculated as the production area value divided by the N-rich zone value.

When it comes to deciding on N fertilization in California small grains, a N fertilizer response is almost certain when plant N status is “Highly Deficient”, very likely when the status is “Deficient” and uncertain when the status is “Sufficient”. The SNQT supplements the plant measurements with information about the current nitrate concentration in the root zone.

If a grower decides that a N fertilizer application is warranted based on the combination of plant and soil measurements, the next step is to figure how much N is necessary. This can be determined using a crop growth and N uptake model in conjunction with yield and protein goals. As part of our larger demonstration project, we will be releasing an online decision support tool in 2021 that integrates these components and provides customized predictions of crop response to in-season N fertilizer.

 

Summary

California farmers are under pressure to increase N use efficiency and reduce the potential for N loss to the environment. N-rich reference zones are a tool that can assist in these goals while considering and managing the risk of reduced yields. By implementing N-rich reference zones, using a suite of tools to monitor them during the season and comparing results to the broader field, a grower gets real-time knowledge to inform N fertilizer management in small grains. The information gained from implementing N-rich reference zones can help growers make fertilizer applications when increased yield and/or protein benefits are likely and avoid them when they are not. These improvements in N fertilizer decision-making can yield better economic and environmental outcomes in California small grain systems.

BMSB Targets Peach Crops

By
Cecilia Parsons
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Monitoring for BMSB in an orchard can be done with sticky panel traps with BMSB lures (photo by J. Rijal.)

Brown marmorated stink bug continues to spread in agricultural crops in California.

This large, invasive insect pest can cause significant damage to fruit crops, and peach orchards are one of this pest’s preferred feeding sites. First detected in the U.S. in the late 1990s, it has been causing crop losses in East Coast fruit orchards since 2010.

In his presentation for California Cling Peach Day, UCCE IPM advisor Jhalendra Rijal said BMSB is spreading slowly into the crop production areas of the northern San Joaquin Valley. As BMSB feed on 170 crop and non-crop host plants, the reproducing populations of BMSB have been established in residential areas of more than 16 California counties, with a majority of them being in the Central Valley. He emphasized the importance of identification of stink bug species when crop damage occurs.

Adult BMSB are about 0.75 inch in length, and larger than Consperse stink bug and red shouldered stink bug. BMSB can also be distinguished from other stink species by the two white bands on their antennae and legs.

In California, BMSB can have two generations per year. Adults will begin to emerge from overwintering sites in mid-March and may continue through May. Because of that, Rijal said the reproductive stages are staggered and both adults and nymphs can be found simultaneously in orchards. He also warned that populations can build quickly early in the season, especially when overwintering sites or non-crop hosts such as trees of heaven are present nearby.

BMSB feed on most of the plants out there with fruiting structures, Rijal said, but peach is one of the preferred hosts. BMSB can feed on all stages of the peach fruit development from early stage through ripening. Feeding causes both external and internal damage. Unlike almonds, early feeding does not cause fruit drop. Surface depression, gumming, necrotic lesions, cork-like lesions and whitish tissue beneath the surface are all signs of BMSB damage in peaches.

Damage is generally confined to the orchard edges. In a survey from 2017 to 2020, Rijal said that BMSB activity was cyclical. The severity of the damage to crops in 2020 was lower than previous years, but BMSB were found in more orchards

Monitoring for BMSB in an orchard can be done with sticky panel traps with BMSB lures. These panels are placed on a pole at the four-foot level from the ground in orchard edges. They attract both adults and nymphs. Beat trays and visual scouting can also confirm presence of this pest.

There are insecticides to reduce the BMSB population, mostly from the broad spectrum pyrethroid groups. These insecticides are often detrimental to natural enemies of other pests such as mites. Also, one spray is very likely not going to be effective if the population is present in the orchard throughout the season, similar to what has been seen on the east coast. The parasitic Samurai wasp is a specific natural enemy of BMSB and has been detected in many states including Washington, Oregon and in the Los Angeles area of California.

BMSB feeding on peach causes both external and internal damage (photo by J. Rijal.)

Biocontrol May be an Option for Grapevine Trunk Disease

By
Cecilia Parsons
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Fungal pathogens cause grapevine trunk diseases which can affect productivity and shorten the lifespan of a vineyard (photo by José R. Úrbez-Torres, Agriculture and Agri-Food Canada.)

Grape vine trunk diseases affect productivity and shorten life of a vineyard.

Akif Eskalen, UCCE Specialist and plant pathologist in the Department of Plant Pathology at UC Davis, in a presentation for Sustainable Winegrowing, explained his research into naturally occurring microorganisms for use as biocontrol against fungal pathogens that cause grape vine trunk disease.

Grapevine trunk diseases are prevalent in mature vineyards. The disease complex includes Eutypa, Esca, Botryospaeria and Phomopsis diebacks, but Eskalen said that at least 60 different fungal species have been identified in grapevines.

Delayed pruning and pruning wound protectants are two prevention routes identified in research, but a survey revealed that the majority of growers use neither preventative practice.

Antagonistic microorganisms already live in the plant tissue, Eskalen said, but they may become depleted. He and his research team are studying how to deliver the beneficial microorganisms back to plants, both in the nursery and in established vineyards.

Eskalen said there is evidence that these beneficial microorganisms not only increase the host plant’s defense mechanism, but also improve the health of the plant and potentially increase yield. Inside their host plant, the naturally occurring beneficial microbes secrete secondary metabolites that inhibit the growth of the harmful fungal organisms. Eskalen said they have identified several of the beneficial microbes and are now focusing on methods of delivery back into host plants.

Biological control can be an important tool in controlling grapevine trunk diseases, Eskalen said. The fungal pathogens that cause diseases each have a different mode to enter a plant and cause disease. No single fungicide can prevent all of those pathogens.

He also pointed out that naturally occurring microbes might be lost over time in vineyards where no other plants exist. Cover crops in vineyards foster more diverse microbe populations. Those populations will differ due to climate, soil type and other environmental factors. Eskalen said his research team is sampling vineyards in different areas of California to get a ‘big picture’ of beneficial microbe populations.

With this information, cultural practices can be adopted to encourage growth of the beneficial microorganisms. Increasing their levels in a plant, Eskalen said, will not only help with disease defense, but also improve overall health and yields.

Mass production and delivery of beneficial organisms is the goal of this research. Trials are introducing the microbes into cuttings in the nursery prior to grafting. In mature vineyards, vine injection and soil application are under study.

Citrus Breeding Efforts Aimed at HLB Resistance

By
Cecilia Parsons
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Citrus fruit from a hybrid HLB resistant tree. The fruits are 6×6 centimeters with 46% juice. According to Dr.Chandrika Ramadugu, most of the hybrid trees bear lime- or lemon-like fruit (photos by C. Ramadugu.)

UC Riverside scientists are working on breeding new varieties of citrus that will be resistant to citrus greening disease.

Citrus greening disease or Huanglongbing (HLB) is a bacterial disease that has killed citrus orchards worldwide. It has been detected in California citrus, but not in commercial citrus production. The disease is vectored by Asian citrus psyllid.

UCR researchers believe a sustainable solution to preserving the citrus industry is to develop varieties that carry natural resistance to HLB. The UCR research team, including Dr. Chandrika Ramadugu, has been awarded funding by the National Institute of Food and Agriculture to pursue breeding work. The research team includes collaborators from Texas A&M, University of Florida, Washington State University and USDA.

Resistant varieties of citrus have been identified, but the challenge is to use them to generate hybrids that will have the flavor consumers prefer along with resistance. The plan is to generate many hybrids and screen them for suitability for the citrus marketplace.

The Australian finger lime is one of six micro citrus varieties from Australia and is being used to create the hybrid varieties. It carries resistance to HLB, but few other attributes to fit the citrus market. The finger limes are about three inches long and roughly the size of an average person’s index finger, but fruit from juvenile trees can be less than one inch long.

The UCR team is currently studying the genetic makeup of the hybrids that have already been produced. Analyzing the new plants’ DNA will show if they carry enough disease resistance along with marketable qualities. Dr. Ramadugu said currently most of the hybrids have lime- or lemon-like fruits. The research team is still in the process of breeding other types of citrus.

One of the main challenges in this process is the length of time it takes before the hybrid citrus varieties bear fruit. With the help of UCR plant cell biology professor Sean Cutler, the team is hoping to accelerate the time it takes for the hybrid plants to bear fruit in the greenhouse. Clones of the best plants will be grown in Florida and Texas field trials.

Other approaches in the HLB fight at UCR include altering soil and root bacteria to improve plant immunity, and using an antibacterial peptide to clear HLB from an infected tree.

UCCE Looks at Whole Orchard Recycling in Walnuts

By
Cecilia Parsons
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Whole-orchard recycling site being prepared for planting new walnut orchard. Process involves pulling the old orchard, grinding the trees into woodchips, spreading the chips and incorporating them into the soil (photo by L. Milliron.)

Work has begun to determine if whole-orchard recycling (WOR) can be as successful in walnut orchards as it has been in almonds.

In a UCCE Virtual Walnut Series, UCCE orchard systems advisor Luke Milliron detailed WOR efforts in two walnut orchard trials.

Whole-orchard recycling involves tree removal and chipping, then spreading the wood chips over the orchard footprint and incorporating them into the soil. Burning restrictions and loss of cogeneration plants that would pay for wood chips spurred research in WOR over the past 10 years. UCCE farm advisor Brent Holtz has been studying WOR in almonds, and his research and field trials show both soil and tree benefits in replanted orchards.

Documented benefits include increased soil organic matter and carbon, increased soil nutrients and increased soil microbial diversity. There was no evidence of increased replant disease and no interference in pre-plant fumigation. There were also water use related improvements and increased orchard productivity.

Milliron shared that while his first trial, which begun in 2018, had some challenges with chip spreading, results of soil and leaf analysis were encouraging. Root lesion nematode levels in the soil were low, indicating successful fumigation. Leaf analysis showed no differences in nitrogen. Potassium and boron levels were slightly higher in trees grown on chipped ground. There were no growth differences between second leaf trees in chipped and non-chipped ground.

Due to the challenges of this trial, Milliron said the total tonnage of chips incorporated could not be determined.

A second WOR walnut trial in collaboration with Cliff Beumel at Agrimillora California is underway.

In this recent trial, Milliron said the dry chip tonnage came out to 91 tons per acre or 136 tons wet. In almond, chips are typically spread about two inches thick. In the walnut trial, the chips were spread at closer to three to four inches, and Milliron said they tried to put chips back at the same rate that trees were removed.

One of the “carrots” for growers is inclusion of WOR in CDFA’s Healthy Soils program designed to promote carbon sequestration and reduce greenhouse gas.

Some of the qualifications in the CDFA program include: trees must be at least ten years of age; orchards must be chipped and incorporated in place; chips must be evenly distributed throughout the orchard and incorporated into the soil to at least six inches in depth.

A list of whole orchard recycling providers can be found at orchardrecycling.ucdavis.edu/california-orchard-recycling-resources.

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