The fall produce season is off to the races, and so is our diamondback moth (DBM) monitoring project. Although our area-wide trap network is still capturing zero adult DBM moths, AZDA inspectors are already detecting DBM larvae in incoming brassica transplants.

During the first month of inspections in August, AZDA reported 66 shipments of brassica transplants from six California counties. DBM was detected in 28 of those shipments—more than 40%.

These shipments enter Arizona under new phytosanitary requirements intended to reduce the movement of DBM, including inspection and certification within 24 hours of shipment. Understanding what an AZDA detection at arrival means for subsequent DBM risk in the field is therefore an important part of evaluating how well this prevention system is working.

Does a detection at arrival translate into DBM establishment and greater pest pressure after planting—or do some detections pose little subsequent risk?

With funding from the Yuma Center of Excellence for Desert Agriculture (YCEDA) Small Grants Program, we are following inspected transplant lots after planting and linking AZDA inspection findings with subsequent DBM observations and management in commercial fields.

Our goal is to determine which observations made during transplant inspections are useful indicators of subsequent field risk and use that information to develop a practical, risk-based inspection framework for AZDA, growers, and PCAs. Ultimately, better understanding the connection between detections at arrival and what happens in the field can help focus scouting and management where they are most needed—and provide evidence for improving transplant inspection and prevention efforts.

Additional Resources

AZDA Director’s Administrative Order 26-03: Exterior Quarantine for Diamondback Moth

A good stand and uniform plant population is a critical step in the crop production process. Crops that require expensive seeds and have tight production schedules require extra diligence in securing a good stand, which can be challenging. A review of the fundamental stages of seed germination and seedling development can be helpful in early season field evaluations and management.

Seeds are living organisms, and they need healthy embryonic tissue to germinate. Most plant species have food reserves to carry the seedling through germination and emergence. The embryo and food reserves are covered in a seed coat and the seeds are stimulated, or they “wake up”, in response to appropriate moisture, temperature, and light conditions that serve to initiate the germination process (Moore et al., 1998).

There are three distinct steps in the process of seed germination.

1. Imbibition. Water from the soil moves into the seed, the seed coat softens and swells.  
2. Interim or lag phase. In this part of germination, the internal seed physiology is activated, and cells begin to respire, and the seed begins to build proteins and metabolize its stores of food.
3. Radicle and root emergence. The cells in root tissue start to elongate and divide and this begins pushing the radicle, the embryonic root, out of the seed.

Several cool season plant species, including lettuce experience thermodormancy (Reynolds and Thompson, 1971), which is a state of dormancy induced by high temperatures that prevent germination. This is an adaptive trait that protects seeds from sprouting during hot conditions that can create plant stress and poor-quality growth (Gardner et al., 1985 and Maynard and Hochmuth, 2007).  

A range of optimum temperatures for a select group of common vegetable crops is shown in Table 1. Lettuce has an optimum temperature range for germination of 40-80°F with an optimum temperature of 75°F and a maximum temperature of 85°F.  

The soil temperature conditions commonly experienced in the low deserts in August, September, and October when thousands of acres of lettuce are being planted and established, commonly exceed these lettuce temperature ranges for germination.  

Table 1. Crop germination temperature ranges for a select set of vegetable crops. Source: Kemble and Musgrove, 2006. 

The temperature limits for lettuce have been modified to some extent by plant breeding and improvement programs that have produced lettuce varieties that can better accommodate hot conditions. However, there are limits to a plant’s genetic elasticity in this regard. Development of some genetic tolerance to heat is being accomplished in crops such as lettuce with modern crop improvement programs. But we must realistically recognize that we are not changing a cool-season plant to a warm-season plant.  

Several aspects of crop and soil management can help mitigate the extreme effects of heat on the soil surface. Perhaps the best and most common is the use of sprinkler irrigation systems to both moisten the soil and cool surface temperatures to a range that encourages lettuce seed germination and emergence (Figure 1). 

Figure 1. Sprinkler irrigation on a recently planted lettuce field, Yuma Valley, Arizona. 

The first step in germination is water entering the plant (imbibing) water from the soil. In dry lettuce seeds for example, this is primarily a physical process driven by the high-water potential gradient between the dry seed and the surrounding moist soil medium. For many seeds, including lettuce, a key entry point for water is the micropyle, which is a small pore or opening in the seed coat. In many species, water enters through the micropylar end, which is where the radicle (embryonic root) will later emerge.

The first thing to emerge from the seed is the primary root, called the radicle, which we often refer to as the “stinger” (Figure 2). The primary root serves to both anchor the plant to the ground and it also begins to absorb water.  

After enough water is absorbed into the seed, the shoot emerges. In dicot plants (most broadleaf plants), the shoot has three main parts: the cotyledons (seed leaves), the hypocotyl, the section of shoot below the cotyledons, and the epicotyl, the section of shoot above the cotyledons (DuPont, 2025).  

Figure 2. Germination stages for lettuce and other dicotyledonous crops. Source: Seeds and Seedling Biology, Penn. State University, 2025. 

These early stages of germination and emergence shown in Figure 2 are critical in establishing a healthy plant, a strong stand, and a good plant population. Root development includes an elongation of the radical and the formation of lateral and secondary roots. The epicotyl will be elongating simultaneously, proceeding towards soil emergence (Figure 3).  

Figure 3. Epigeal and hypogeal emergence. Source: Michael Knee, The Ohio State University. Further illustrations of emergent lettuce plants are shown in Figures 4 and 5.  

During the emergence process when a dicot plant is in the “hook” stage, it is a critical time when the hook is trying to push through the soil surface. Plant species with small seeds are commonly not very strong with this process and that can be exacerbated with any soil crusting. This is another reason sprinkler irrigation is often used to diminish soil crusting and facilitate germination and emergence in the low desert vegetable crop production areas. 

Figure 4. Lettuce cotyledons and first true leaves. Source: Harvest to Table. 

Figure 5. Stages of emergence. Source: Spider Farmer. 

Planting cool season crops, i.e., leafy green vegetables, in the lower Colorado River Valley in August, September, and October is a challenging operation. Seedling germination and early development are critical stages of growth that have lasting impact on the vigor and health of the crop.

References:

DuPont, T. Seed and Seedling Biology. 2025. Pennsylvania State University Cooperative Extension, ART 3208.

Gardner, Pearce, and Mitchell. 1985. Physiology of Crop Plants. The Iowa State University Press.

Kemble, J., and M. Musgrove. Soil Temperature Conditions for Vegetable Seed Germination. Alabama Cooperative Extension, 2006.

Knee, M. Development of Seed to Plant. 2024. The Ohio State University.

Maynard, D., and G. Hochmuth. Knott's Handbook for Vegetable Growers. Vol. 5. Hoboken, N.J.: John Wiley and Sons, 2007.

Moore R., W.D. Clark, and D.S. Vodopich. Botany. The McGraw-Hill Companies. 1998. ISBN: 0-697-38363-1

Galls observed on the stems of begonias submitted to the clinic 

Recently, we received a submission of begonias to the Yuma Plant Health Clinic that immediately presented a diagnostic challenge. At first glance, the symptoms seemed to point toward a familiar set of suspects. The plants had developed severe swelling of the stems, symptoms characteristic of physiological oedema or bacterial Agrobacterium-associated crown gall. But, as is often the case in plant diagnostics, appearances can be deceiving.

The begonias had been grown indoors in high-humidity terrariums and planter boxes. The symptoms were not associated with a particular begonia variety, nor were they consistently expressed across all plantings or cuttings. When the plants were removed from their planting substrate, however, another clue emerged: additional small galls were scattered throughout the root systems of those plants also showing the stem swellings. That observation changed the diagnostic picture considerably. 

Galls observed on the roots of begonias submitted to the clinic 

Dissection of both the swollen stem tissue and the roots revealed an abundance of eggs, juveniles, mature female nematodes, and giant cells, a classic signature of root-knot nematode (RKN) infection. Root-knot nematodes belong to the genus Meloidogyne, a group of microscopic roundworms that are among the most important plant-parasitic nematodes worldwide. Their typical symptom is familiar to many resulting in the formation of characteristic galls on roots.

These galls are not simply a physical wound created by the nematodes. Rather, RKNs actively manipulate their host. During infection, the nematodes secrete salivary compounds that alter plant growth and development, effectively hijacking normal cellular processes and redirecting them toward the formation of specialized feeding structures known as giant cells.

These giant cells become a nutrient-rich feeding site for the developing and mature nematodes. In doing so, the nematode establishes a sustained drain on the plant's resources. As infection progresses, the resulting disruption to the root system can interfere with nutrient acquisition and water uptake. Aboveground, this typically translates into stunted growth, declining foliage, wilting, and, in severe infestations, plant death. With more than 3,000 plant species reported as hosts to different root-knot nematode species worldwide, including many field and vegetable crops, the annual impact of these pathogens is enormous. Much research has been done on RKN in various crops, and reports of stem swellings occurring naturally is almost non-existent.

Yet the begonia case raises an especially interesting question: Why did a pathogen best known for producing root galls appear to be responsible for such dramatic swelling in the stems? This is where the environment becomes an essential part of the diagnosis.

The symptom is not the pathogen. It is a reaction of the plant to the presence of a pathogen. One of the most important principles in plant pathological diagnostics is that the same pathogen does not necessarily produce the same symptom under every set of environmental conditions. Likewise, similar symptoms can arise from entirely different causes.

Temperature, humidity, water availability, substrate conditions, light, nutrition, plant age, and even the way a plant is cultivated can all influence how a disease develops and how its symptoms are expressed. In other words, the symptom we see is the product of an interaction between the host, the pathogen, and the environment. Change any one of those components, and the resulting disease can change dramatically. This recent begonia case is a particularly useful reminder of this principle.

Root-knot nematodes are well known for their belowground symptoms, but reports of rare and isolated cases of infections on aerial plant tissues, including stems, leaves, and flowers, are reported in only one publicly available extension bulletin from 1985. These symptoms remain unusual and have rarely been observed unless aerial tissues are artificially inoculated, and they have only been documented in a limited number of hosts.

So, when confronted with the unusual stem symptoms in a begonia, it would be easy to focus exclusively on the appearance of the tissue and pursue diagnoses such as oedema or crown gall. These plants were being cultivated in enclosed, high-humidity conditions, where moisture levels around the plant were substantially different from those experienced by a plant growing in a more open environment. Such conditions can alter transpiration rates, water movement, tissue hydration, pathogen development, and the plant's own physiological responses. They also give rise to the wet, humid conditions necessary for oedema to develop or crown gall bacteria to thrive.

The small root galls were an important clue. Microscopic examination then provided the confirmation: eggs, juveniles, mature females, and the characteristic giant cells associated with root-knot nematode feeding. These signs of RKN were found in both the root and stem swellings. Understanding the environment in which the hosts were being grown, with its high moisture conditions, then provided supporting evidence that the nematodes had favorable environmental conditions to migrate into much higher foliar tissues than is typically seen in RKN infestations and produce an unusual symptom. Swollen galls on the stem told us that something was wrong, and if we relied on the most common culprits associated with this symptom alone, namely oedema or crown gall, we would have gotten the diagnosis wrong.

The begonia case is a fascinating example of how flexible disease expression can be. A pathogen with a reputation for being almost exclusively soilborne and producing galling symptoms on root tissues, under particular host and environmental circumstances, can be associated with symptoms in tissues where we do not ordinarily expect to find them.

Ultimately, environment is not simply a backdrop against which plant disease occurs. It is one of the determinants of how disease develops, how severe it becomes, and critically for anyone trying to ID the pathogenic cause, what it looks like.  

If you have any concerns regarding the health of your plants/crops please consider submitting samples to the Yuma Plant Health Clinic for diagnostic service or booking a field visit with me:

Christopher Detranaltes, Ph.D.

Cooperative Extension – Yuma County

Email: cdetranaltes@arizona.edu

Cell: 602-689-7328

6425 W 8th St Yuma, Arizona 85364 – Room 109 

It is with mixed emotions that I write to inform you that this will be my last University of Arizona Vegetable IPM Update. The reason – I am retiring. Thank you for your support over the years. It’s been an honor and a privilege working with you and serving this ag community.

Figure 1. Automated Thinning & Weeding Technologies Field Day.
(Photo credits: Rosa Bevington)

As lettuce operations are transitioning to drip irrigation rather than overhead sprinklers to maximize water efficiency in the lower Colorado River, farm management practices must adapt, especially weed control. Pre-emergence tools like Prefar 4-E (bensulide) and Kerb SC (propyzamide) remain vital for managing purslane, chenopodium species, annual bluegrass, and key broadleaves. However, both active ingredients depend on specific soil placement and water delivery to form an effective barrier against germinating weed seeds.

Historically, standard overhead sprinkler passes provided the uniform surface coverage and water volume needed to activate Prefar and wash Kerb SC into the target seed zone. As fields transition to surface or subsurface drip irrigation, moving away from overhead water creates a distinct field challenge: drip lines push moisture laterally and downward, often leaving the top 1 to 2 inches of soil too dry to activate surface-applied herbicides.

Matching the Tool to the Mechanism: Because these two herbicides interact with soil and water differently, switching to drip requires tailored incorporation strategies:

• Prefar 4-E (Mechanical Placement): Prefar binds tightly to soil particles and does not leach easily. Under drip, it demands precise mechanical incorporation into the top 1 to 2 inches immediately after application. Options include shallow rolling cultivators (Lillistons), rotary hoes, power tines, or integrating the spray pass directly into final bed-shaping operations before drip line installation.
• Kerb SC (Water Mobility & Timing): Kerb SC relies heavily on water movement to carry it into the active root/germination zone. Under drip, growers must balance mechanical soil mixing with strategic irrigation timing, such as running short, high-volume drip cycles to encourage capillary wicking to the bed surface, or using a brief initial sprinkler pass solely to activate the herbicide before switching to drip. 

Insect pests are constantly adapting to their environment. Over time, they have evolved ways to overcome control tactics. Repeated use of the same insecticides or tactics places strong selection pressure on pest populations. This allows the few individuals that can survive treatments to reproduce, leading to populations that are increasingly difficult to control, a process known as resistance.

Insects can develop resistance in several ways. Some break down insecticides more efficiently or develop changes at the target site that reduce product effectiveness. Others avoid exposure altogether by changing their feeding habits or movement patterns. In some cases, insects even develop thicker cuticles that reduce or slow down the absorption of insecticides.  

Production practices can influence the rate of resistance development. Large, uniform cropping systems and repeated use of the same control tools reduce opportunities for susceptible insects to persist, accelerating resistance.  

Figure 1: Chart illustrating how insecticide resistance develops when insecticides with the same mode

of action (MoA) are used repeatedly. 

What Should You Do to Mitigate Resistance in Insect Pests?

An integrated pest management (IPM) approach remains the best defense against resistance.  

Key practices include:

• Using crop varieties that are resistant to pest attack
• Rotating insecticides with different modes of action (MoA)
• Avoiding unnecessary insecticide applications  
• Preserving beneficial insects by selecting insecticides with no or minimal adverse effects on them
• Incorporating non-chemical management tools whenever possible  
• Maintaining refuges to sustain susceptible pest populations  
• Eliminating crop residues after harvest to remove food sources and breeding sites for pests.
  

No insecticide remains fully effective over the long-term if overused. Reducing selection pressure by avoiding consecutive applications of insecticides from the same IRAC group is critical. The more intensively a single tactic is used, the faster resistance will develop. Resistance can increase production costs, reduce marketable yield, and ultimately affect grower competitiveness. A diversified pest management approach helps preserve the effectiveness of available tools and supports long-term, sustainable pest control.

 

 

Additional Reading Materials

1. Calvin W., M. N. Keith, and B. McGrew. 2025. Guidelines for effective management of diamondback moth in brassica crops. University of Arizona Extension Publication. az2143. https://extension.arizona.edu/publication/guidelines-effective-management-diamondbackmoth-brassica-crops 
2. Calvin W. 2026. Effects of the Maximum Dose of Common Insecticides and Incipio on Diamondback Moth Populations Collected from Arizona and California Brassica Crops (Fall 2025– Winter/Spring 2026). https://acis.cals.arizona.edu/agriculturalipm/vegetables/vipm-archive/vipm-specialty-crops-view/effects-of-the-maximum-doseof-common-insecticides-and-incipio-on-diamondback-moth-populations-collected-fromarizona-and-california-brassica-crops-(fall-2025-winter-spring-2026) 

Yuma is heading into fall produce season after an unusually warm year. By July 31, 2026, the season had accumulated 553 heat units (86/55°F) more than the 2020–2025 average. Through mid-August, nighttime temperatures were the warmest for that period in the AZMet Yuma Valley record, with 15 of 17 nights never dropping below 80°F. So the simple question is: if we go into produce season with above-average days and above-average nights, what should we expect? So, the useful answer starts with a distinction. Warm days and warm nights are not the same problem. They affect different parts of the crop, on different timelines, and they call for different management responses.

Warm Days, Part One: A Water-Demand Problem You Can Calculate  Hot afternoons raise vapor pressure deficit and pull more water from the soil and crop. But the size of that effect is smaller than many people assume, and it is predictable. Averaged over 2020–2025, standardized reference evapotranspiration (ETo) during the establishment period was 1.69 inches per week in September, 1.26 inches in October, and 0.93 inches in November, a 45% decline across the window. Meanwhile, the spread between the warmest and coolest September in that record was only about one inch over the whole month. That comparison is the point: planting date moves water demand far more than how hot the year is. A warm September is not a reason to add a blanket percentage to every irrigation set. It is a reason to watch ETo and soil moisture closely during the two or three weeks when the crop has a limited root system and no canopy to protect the bed.

Warm Days, Part Two: A Germination Problem  
This is the issue that can cost stands, and it is easy to miss because it looks like a water problem. Lettuce germination can be inhibited above roughly 81°F, and the effect is stronger in darkness, which is exactly where the seed is. Prolonged exposure can push imbibed seed into secondary dormancy, at which point cooling the bed may no longer rescue it. Mean September soil temperature at 4 inches at the AZMet Yuma Valley station over the past six years was 85, 85, 86, 87, 89, and 93°F. Every year was above the threshold. In a warm year, the margin gets worse rather than better. More water alone does not fix this. Variety selection, seed priming, planting during the coolest part of the day, and sprinkler cooling of the seed zone are more targeted tools. Also, station soil temperature is a regional index, not the temperature in your bed. A bare, tilled, sunlit seedbed can run warmer. Put a thermometer in the bed at seeding depth and decide on that number.

Warm Soils and Surface Drying
Warm conditions also affect the seedbed and root zone. Faster surface drying can concentrate salts near the seed line, especially where moisture is not uniform. This matters during lettuce establishment because small seedlings have limited roots and are sensitive to both heat and salinity stress. The goal is not simply more water, but more uniform moisture in the right zone. Bed temperature, seed-zone moisture, and salt distribution should be checked together before and after planting.

Warm Nights: A Marketability and Quality Problem
Nights get less attention than hot afternoons, but this year they are the more distinctive signal. Lettuce spends energy overnight on maintenance respiration. When nights stay warm, more of the day’s photosynthate is burned rather than stored, and there is less recovery time after daytime stress. Later in the crop cycle, that can affect head development, firmness, uniformity, and tip burn risk. The cost lands on marketable yield. Lettuce is bought on quality, and stress that reduces uniformity or head structure can cost more than the water it appeared to save. The encouraging part is that nighttime minimum temperatures usually fall from about 74°F in September to about 62°F in October, so this pressure eases quickly with planting date.

Both Together: Pest Timing Can Move Up
Insect development is influenced by accumulated heat, so a season running 553°F-days ahead can also move pest timing ahead. That does not necessarily mean higher pest pressure; it means the timing may shift.

What to Expect, in Short

1. Water demand: modestly higher during warm periods, but still driven by ETo, crop stage, wind, solar radiation, humidity, and planting date. Track it; do not guess it.
2. Emergence: the highest-risk item. September seedbed temperatures can already sit near or above the lettuce germination threshold.
3. Salinity: faster surface drying can concentrate salts near the seed line. Aim for uniform seed-zone moisture, not simply more volume.
4. Quality: warm nights can affect recovery, head structure, firmness, uniformity, and tipburn risk, with pressure easing as nights cool in October.
5. Pests: accumulated heat can move insect development ahead of the calendar, so scouting should follow crop stage, degree-day conditions, and field observations.
   

Notice that four of those five pressures ease substantially between September and October. That is the practical thread running through all of this: in a warm year, planting date is one of the most powerful tools available, and the two weeks around establishment are where precision pays. A hot start does not call for guesswork, and it does not automatically call for more water or more inputs. Yuma growers have built their success by responding to heat with precision: better timing, closer measurement, stronger scouting, and careful root-zone management before produce season begins.

VegIPM Update Vol. 17, Num. 18

September 2, 2026

Results of trap catches below!!

Corn earworm: CEW numbers remained low at most locations over the last two weeks. About average for this time of the season. Expect to see numbers increase as we get into September.

Beet armyworm: BAW activity increased over the last two weeks. About average for this time of the year.

Cabbage looper: Cabbage looper moths remained very low over the last two weeks. Very few moths have been collected in the traps, usually numbers start picking up in September.

Diamondback moth: No diamondback moths have been collected in the traps since May 19th.  Based on the past six years of summer collection data, no DBM adults are collected in the traps in the summer months (Jun-Aug) until September.  

Whitefly: Adult activity remained steady across locations over the last two weeks; about average for this time of the year. Historically, whitefly numbers peak in July.  

Thrips: Adult thrips remained low over the last two weeks. About average for this time of the year. Based on previous years, thrips numbers remain low until Sept-Oct.  

Aphids: No aphids have been collected since April 21st. Historically, aphids are not found in the traps until November.