An Argentine ant perched on a citrus tree leaf (photo courtesy Mike Lewis, UC Riverside.)
The Argentine ant, or Linepithema humile, is a coffee bean-colored ant native to South America. Introduced in the early 1900s in California likely by exports, the ant has long been a problem for pest management in citrus orchards.
Mark Hoddle, UCCE Entomology Specialist at UC Riverside, spoke about issues surrounding Argentine ants, their mutualistic behavior with sap-sucking pests, monitoring tools, control options and field studies in a webinar presented by UC ANR.
“There are queens, males, workers and brood in underground colonies,” Hoddle said. “Their lifecycles are similar to butterflies, having egg, larval and pupal stages, and about 75 days to develop from egg to adult.”
The sheer number of Argentine ants that make up a colony, as well as their aggressive nature, makes them a daunting species to deal with. Each nest has multiple queens, which typically lay 20 to 30 eggs per day and have been found to lay up to 50 to 60 eggs in certain instances, according to Hoddle.
The mutualistic behavior of Argentine ants, however, is where the real issue lies in citrus orchards. The ants are able to co-exist with multiple sap-sucking, economically damaging pests in citrus such as mealybugs, aphids, whiteflies, soft scales and psyllids. Basically, ants living in orchards tend and feed off honey dew, a sugary excrement produced by these pests. In return, the ants protect their “livestock” from parasitoids and predators, the natural enemies that attack these pests and control their populations.
“They [ants] are specialized liquid feeders that feed on the sugary waste product excreted by phloem feeding pests,” Hoddle said. “Honey dew removal by Argentine ants actually protects these pest populations from essentially drowning in their own excrement, promoting population growth and driving ant infestation severity.”
Hoddle and his research team have been using a variety of monitoring tools to estimate ant densities in citrus orchards, an important way to determine control decisions. Among the monitoring tools used are sugar vials with 25% sucrose solution, counting ants moving past “landmarks”, and a new tool: infrared sensors attached to irrigation piping.
“Monitoring vials often overestimate ant populations,” Hoddle said. “Marking spots on a [citrus] tree and counting ant traffic over a specific amount of time can give good estimates to the densities and activity levels in trees.”
According to Hoddle, irrigation piping tends to be the easiest place to find Argentine ants in an orchard. The ants prefer moving in straight lines on the smooth surface of the piping which enables them to move more rapidly from nests to food sources and back again.
“Infrared sensors along irrigation piping collect physical data and send it to the cloud where the data are summarized and viewable with an app on a smart device,” Hoddle said. “The goal is to create a fully automated ant monitoring system for growers.”
Once the results can give an accurate idea of ant activity in an orchard, control options need to be decided. The most basic option is a physical barrier around the trunk of the tree. Composted organic mulch at the base of trees can deter ant walking speeds due to uneven terrain. Bait stations loaded with 25% sucrose water and ultra-low concentrations of insecticide (.0001%) can also be used.
In a field study completed by Hoddle and his research team, bait stations were used in six navel groves in Southern California where Argentine ants are most abundant. The results showed very quick diminished numbers of ants, with 50 percent reductions over the first month and 88 percent reductions over the next 11 months.
“The bait stations can work, but they need to be taken care of, monitored and replenished regularly,” Dr. Hoddle said. “Biodegradable hydrogel beads made out of seaweed, 25% sucrose water and .0001% insecticide proved effective as well. The ants feed off of these, return the toxin to the nest, and through communal food sharing the workers and queens are slowly poisoned.”
An unmanned vehicle pulls a tunnel-shaped frame with UV lights attached over a row of grapevines. The UV light kills powdery mildew at night to overcome powdery mildew evolutionary resistance to light damage (photo courtesy David Gadoury, Cornell.)
Ultraviolet (UV) light lamps attached to unmanned vehicles are being used to fight powdery mildew infections on research vineyards on the East Coast and may hold promise for West Coast vineyards as well.
Researchers at Cornell AgriTech in Geneva, NY, in collaboration with colleagues at Rensselaer Polytechnic Institute’s Lighting Research Center (RPI-LRC), the University of Florida, and the National Agricultural University of Norway (NMBU), have been working with a Norwegian manufacturer (SAGA Robotics) to develop the autonomous robots for commercial use.
Everywhere grapes are grown, powdery mildew poses a threat to the crop. This fungal disease is especially significant due to cost of control. According to the Robert Mondavi Institute Center for Wine Economics at UC Davis, powdery mildew management accounts for 74 percent of total pesticide applications by California grape growers and 17 percent of total pesticide use in California agriculture (by weight of active ingredient.)
David Gadoury, senior research associate in the Department of Plant Pathology at Cornell AgriTech, who is leading the project, said in a phone interview that interest in this unique control is high in California.
The UV lights are the same germicidal lamps used in hospitals; the lights penetrate plant canopies to reach and suppress certain pathogens. Research in using UV light to kill the powdery mildew pathogens is not new, Gadoury said, but it has accelerated with the discovery that it is effective at night.
He explained that powdery mildews have co-evolved with the plants they attack over millions of years and often develop resistance to chemical treatments. Their evolution has also given them an Achilles heel: adaptation to natural cycles of light and dark. UV light damages DNA of many organisms, but they evolved developed biochemical defenses against this damage, using a repair process that requires the blue light component of sunlight.
“What makes it possible for us to use UV to control these plant pathogens is we apply it at night,” Gadoury said. “At night, the pathogens don’t receive blue light and the DNA repair mechanism isn’t working.”
In field trials, the light arrays are carried within a tunnel-shaped frame that can be adapted to different trellis systems. The robotic factor can be a labor-saving feature, but the lamp array could also be pulled by tractor.
Gadoury said the UV treatment requires four hours of darkness after application for maximum effectiveness. At a speed of 5 mph, a towable or robotic array could travel over 20 miles during the available nighttime, even during the shortest nights of summer in most viticulture areas. Extensive trials have been completed on Florida and California strawberries. Grape trials for suppression of both powdery and downy mildew are underway in New York, Washington, and Oregon, with California trials planned for 2021.
The research group includes assistant professors Katie Gold and Yu Jiang at Cornell AgriTech, Natalia Peres at the University of Florida, Mark Rea at Rensselaer Polytechnic Institute’s Lighting Research Center, and Arne Stensvand at Norway’s Institute of Bioeconomy.
The research is supported by grants from the USDA-SCRI, USDA-OREI, the Research Council of Norway and the New York Farm Viability Institute. Support also came from lighting companies OSRAM and the Asahi Glass Co.
Drip germination in lettuce has resulted in fewer weeds than sprinkler irrigation (photo by Marni Katz.)
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.
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 large “other” 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.
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
Figure 1. Plant labels in tray of tomato seedlings prior to transplanting. The labels and tomato plants were transplanted together in the field.
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.
Spinach leaves are greatly deformed and discolored by Tobacco rattle virus (photo by S. Koike, TriCal Diagnostics.)
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.)
Almond Orchard Cover Crop study in Kern County, March 2019 (photo by S Shroder.)
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.)
Brennan, E. B. (2017). Can we grow organic or conventional vegetables sustainably without cover crops? HortTechnology, 27(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 Journal, 104(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 Research, 165, 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 Agriculture, 60(3), 146-153.
Figure 1. Romaine lettuce plant near maturity showing classic symptoms of outer leaf yellowing and necrosis. Symptoms may develop at any growth stage (all photos courtesy W.M. Wintermantel.)
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.
Choosing the right activator adjuvant can avoid phytotoxicity damage or losses from excess spreading and pesticide runoff from the target plant.
Agricultural spray adjuvants are materials added to the spray tank when loading the sprayer. They include products classified as activator adjuvants and marketed as wetters/spreaders, stickers, humectants, and/or penetrators. Activator adjuvants are marketed to improve the performance of pesticides and foliar fertilizers.
Activator adjuvants can have a place in tree (and vine) crop sprays, but matching the material to the job can be tricky. A bad match can lead to minor or major losses to the grower. Minor losses can result from excess spreading and pesticide runoff from the target plant. Phytotoxicity can cause major damage.
This article describes ingredients and functions of activator adjuvants commonly sprayed on tree and vine crops. Suggestions regarding activator adjuvant selection are offered. Growers must make their own activator adjuvant use decisions based on experience, particular needs, and risk tolerance.
Should You Use an Adjuvant?
Read and follow the specific instructions on the label. If the pesticide or foliar fertilizer label indicates the product should be used with certain types or brand of adjuvant(s), that’s what you need to use. For example, the Bravo Weather Stik® label cautions against using certain specific adjuvants and puts the responsibility in the PCA or grower court regarding adjuvant use. If the label includes phrases such as “use of an adjuvant may improve results” or “complete coverage is needed for best results” then you may want to look into selecting and using an appropriate activator adjuvant.
Before proceeding with use of an activator adjuvant, first look at your existing spray program. Are you already doing the best spray job you can? Good spray coverage begins with proper sprayer calibration and set up. Is your sprayer calibration dialed in for different stages of canopy development? Optimum sprayer set up—gallons of spray per acre, ground speed, fan output, and nozzle selection/arrangement—changes from dormant to bloom to early growing season to preharvest sprays. Adjusting your sprayer to best match orchard and vineyard conditions at each general stage in canopy development is the foundation of an effective, efficient spray program. An activator adjuvant will not make up for excessive tractor speed, poor nozzle arrangement and/or worn nozzles. Your money is best spent first dialing in your sprayer(s) for the whole season, before considering an extra material in the tank (that is not required on the label).
If you have your sprayer(s) dialed in for each orchard and stage of growth, now is the time to say “OK, I want to think about a little extra boost to my spray job.”
Which Activator Adjuvant to Choose?
First, know the properties of the pesticide you will use. Does it work on the plant surface or inside the plant? This is a key point in selecting adjuvants. Here is a quick review of the main classifications and characteristics of activator adjuvants as they currently appear in the field. Note: Certain products can provide more than one adjuvant property that can be beneficial in the field. For example, non-ionic surfactants can work as surfactants and penetrators, depending on use rate.
Wetters/spreaders: These materials contain surfactants that decrease the contact angle and increase the spreading of the spray droplet on the target. High rates of wetters/spreaders may also increase penetration of pesticide into the target tissue (leaves or fruit), potentially causing phytotoxicity. Excessive spreading of pesticide spray solution and runoff from the target may result when using a new or higher rate of spreader—especially when using silicon “super-spreaders”. Test new combinations of spreader material(s) and spray volume before regular use. Spray volume per acre or adjuvant use rate will probably have to be reduced if a labeled rate of adjuvant provides excessive spreading.
To check for excessive spreading, place a length of black plastic sheeting under several trees or vines in a row. Secure the plastic with spikes, wire staples, and/or weights. Spray the new adjuvant and pesticide combination using your current sprayer set up. Reenter the field right after spraying, wearing appropriate PPE, and evaluate coverage. If material is pooling at the lower portion of leaves and/or fruit, excessive spreading is occurring. Check to see if pooling is occurring only in a certain area(s) of the canopy or throughout the canopy. If more spray solution is landing on the black plastic tarp under the trees/vines than between them, then runoff is occurring. [Some ground deposit should be expected from standard airblast sprayer use.]
Compare the results of your adjuvant test with a similar application of your current pesticide/adjuvant combination on another portion of the row. If there is no pooling or runoff with the new adjuvant in the tank, you can use the adjuvant with confidence. A lack of pooling or run off with the new adjuvant also might mean that your old sprayer setup and tank mix didn’t deliver adequate coverage.
If the test with the new adjuvant showed pooling on leaves and/or runoff on the ground, you have several choices: 1) You can reduce spray volume per acre by replacing some or all nozzles with smaller nozzle sizes on the sprayer in an effort to reduce overspreading. If you saw overspreading on some portions of the canopy, but not others, reduce nozzle size only on the part of the spray boom that targets the over-sprayed part of the canopy. Recheck spray coverage if nozzling changes were made. 2) Reduce the adjuvant rate and recheck coverage/spreading. 3) You can just go back to your established program without the new adjuvant.
What’s the “best” course of action? That depends on your farming operation. Reducing spray volume per acre means more ground covered per full spray tank – a potential time and cost savings. If spraying is done during the heat of the day in hot, dry climate, spray water evaporation is a major issue and it may be best to keep the higher spray volume and reduce the spreader rate or eliminate it entirely. Checking coverage and overspreading allows you to make the best decision possible, avoid damage and, hopefully, save money. All farming operations are different. Make the choice that best fits your farm.
Stickers: These adjuvants can increase the retention time of the pesticide on the leaf and reduce rain wash off. They may limit movement of systemic pesticides into the plant, and are probably most beneficial when used with protectant materials (cover sprays). Do you overhead irrigate? Is there rain on the horizon? If you answer yes to either one of these questions, you may benefit from using a sticker.
Humectants: Under low humidity conditions humectants can help reduce spray droplet evaporation before and after deposition on the plant. This is especially valuable when small droplets and/or materials that must be absorbed into the plant (systemic pesticides, PGRs, nutrients, etc.) are used in the summer under high temperature and low relative humidity conditions.
Penetrators: Frequently used with herbicides, these products include oils (petroleum, vegetable, or modified vegetable oils) and non-ionic surfactants used at higher rates. In crop sprays, penetrators can be used to increase absorption of systemic pesticides (for example, oil with Agri-Mek) as well as translaminar materials. Penetrator adjuvants should be used with caution or avoided entirely with surface active pesticides such as cover sprays or else phyto may result. Finally, some penetrators can increase the rain-fastness of some pesticides.
What Adjuvant Material to Choose?
Use a product intended for crop spraying. Many activator adjuvants were developed and intended for use with herbicides. Products that are advertised for use with plant growth regulators should have a higher chance of crop safety compared with those that don’t. This is still no guarantee of a phyto-free application.
Ask for help from the adjuvant manufacturer’s sales rep if needed. How much do they know about the particular activator adjuvant in the spray mix you are planning?
Will the Adjuvant Work?
If you choose to use an adjuvant that is not specifically listed on the pesticide or foliar fertilizer label, jar test the planned spray solution first. Use the same spray water source. Include all leaf feeds, other adjuvants, and pesticide(s) that you plan to put in the spray tank. Do this before tank mixing these materials.
A lot of time and money rides on effective pesticide application. Do your homework before the spray tank is filled and you will be well on your way to solid results.
Mulches provide natural weed control in most strawberry growing regions.
Weed competition can have a serious impact on growth and productivity in commercial strawberry production. The diversity of climates among California’s growing regions requires different weed management options. The high-value of this crop, long-season and the fact that it is hand harvested twice weekly requires an effective weed control program.
Steve Fennimore, UC Cooperative Extension weed specialist, based in Salinas, said by far the most common tools used to suppress weeds in conventionally grown strawberry fields post planting are opaque mulches, pre-plant fumigation, hand weeding and herbicides.
Mulches are dark color films that restrict light and shade out weeds. In southern coastal areas, clear plastic is used to warm the ground and achieve an earlier harvest, but clear films can also promote weed growth. Fumigants and herbicides are used in conjunction with the clear film to control weeds.
In addition to pre-plant fumigation, placing clear film on the strawberry bed tops promote plant growth and opaque film on the sides of the beds is another weed control option. UC IPM Guidelines recommend securing opaque mulches to the soil prior to transplanting strawberry plants. Slits are made in the mulch at desired spacing. The smallest hole possible to insert plants will help minimize weed growth and seed deposition.
Of the common weeds in strawberry production, yellow nutsedge is one that cannot be controlled by using an opaque mulch as the shoots can puncture the mulch and grow through it. This is a perennial weed that grows from tubers capable of survival in the soil for three years.
Soil solarization is another technique for weed suppression in organic strawberry production. This strategy works where warm daytime temperatures are sustained, but is not effective in cooler coastal regions. This strategy, pre-plant, involves covering the soil with clear plastic and wetting the soil to field capacity. University of California research showed that temperatures of 108 to 131 degrees F could be reached in the top two inches of the soil. The plastic is left on four to six weeks. Solarization to kill weed seeds is most effective in the two 12 inches of soil. While solarization works for weed control, the heat generally does not penetrate deep enough to kill pathogens deeper in the soil.
Soil treatment with steam has also been effective in killing weeds, but lethal temperatures have to be reached in soil to kill weeds with propagules are present and it requires specialized equipment.
Anaerobic soil disinfestation can reduce the numbers of many annual weeds but it has limited efficacy on perennial weeds.
