There is increased interest in the use of the nitrate quick test for managing fertilizer decisions in vegetable production. In this Blog, I will go over some details on obtaining a good representative sample in order to conduct the quick test analysis. 

Normally soil cores are taken down to 12 inches for lettuce and cole crops; however, for shallow rooted crops such as spinach and baby lettuce, soil cores to 6 inches are probably better because they better reflect the nitrate levels in the area of active roots. Do not include soil from the top 2 inches of soil as it may be high in nitrate, but too dry for the plants to access it. We have found that on some soil types (eg. clays, silty clays) it is important to angle the soil probe in the direction of the fertilizer bead or drip tape (in fertigated situations) (See Figures 1 & 2). The reason for this is that in these soils, sometimes the fertilizer does not move much with the irrigation water and by angling the probe, you get a more representative sample.  As a matter of habit, we angle the probe on all soil types to keep our sampling method uniform. 

Sample the field in a pattern that goes from one end of the field to the other, both sides of the field and through the middle – generally an “X” or “N” shaped pattern is fine.  In order to collect a good representative sample it is important to collect a sufficient number of samples from a representative area of the field. In general, 15 to 20 soil cores are sufficient. 

After collecting the sample, it is critical to homogenize the sample so that there is a uniform level of nitrate throughout. Some soils are easily homogenized, but sticky clays or even wet loams are too gummy to mix. In these situations, we do the “pinch” method by laying out the soil cores and pinching off small, uniform amounts from up and down each core. We then mix the pinches and use them for placing in the calcium chloride solution (see below).

Figures 1 & 2. Insert the probe in the seedline, but angle it to go beneath the bead of fertilizer or beneath the drip tape.

Here is information on conducting the nitrate quick test:

Equipment Needed

•  Merkquant® test strips. They can be purchased from Ben Meadows at 1-800-241-6401.  Order the 0 – 500 ppm nitrate strips (catalog No. 7JB-7830).  They come in tubes of 100 strips.
• 50 ml Centrifuge tubes. From VWR or Fisher Scientific. These companies are a bit difficult to deal with and want to sell larger batches of tubes. You can probably find companies on the web that will sell batches of 25 or so tubes. Here is one FYI: http://www.quasarinstruments.com/p-16378-50-ml-centrifuge-tubes.aspx
• Calcium chloride dihydrate. From Fisher Scientific. Fisher Chemical: C70-500. There are also companies on the web that sell aquarium or food grade (e.g. canning supply companies) calcium chloride. It is not “certified reagent grade”, but should be fine for conducting the test.
• 1 gallon of distilled water (add 5.6 grams of Calcium chloride to 1 gallon of distilled water to make up the 0.01 M Calcium Chloride solution)

 Procedure

1. Collect a composite soil sample as described above.
2. Fill a volumetrically marked tube or cylinder to the 30 ml level with 0.01 M Calcium Chloride (CaC1₂) solution.
3. Add soil to the tube until the liquid level rises to 40 ml; cap tightly and shake vigorously until soil is thoroughly dispersed.  Let sit until soil particles settle out.
4. When solution is reasonable clear, dip Merckquant nitrate test strip into the solution, shake off excess solution, and wait 60 seconds.  Compare color with the color chart provided.
5. To minimize variability inherent in soil sampling, run duplicate samples for each field soil evaluated. 

Interpretation

The test strips are calibrated in parts per million (PPM) NO₃.  Converting strip readings to (PPM) NO₃-N on a dry soil basis will require dividing by a correction factor based on soil texture and moisture: 

Strip reading (PPM NO3) ÷ correction factor = PPM NO₃-N in dry soil

Soil less than 10 PPM NO₃-N would be considered low; levels above 20 PPM NO₃-N have enough available N to meet immediate crop needs.

Yet another new race of downy mildew (Peronospora farinosa f. sp. spinaciae) on spinach has been identified in California’s Salinas Valley. The type, or original, strain was initially designated as UA2209 and was first detected in May 2009. Subsequently, it was found in an increasing number of locations throughout California in 2009 and 2010. This race breaks the resistance of several important cultivars. The race has been characterized on a set of differential cultivars and was designated as race Pfs 12 by the International Working Group on Peronospora (IWGP). The working group is located in the Netherlands and is administered by Plantum NL.

Race Pfs 12 poses a threat to the spinach industry because it is particularly well-adapted to most modern hybrids with resistance to race 1-11, which have been widely planted in the past few years. Race 12 is distinct from race 11 because of its virulence on the differentials Campania and Avenger. The appearance of a new race is not completely unexpected because hybrids with resistance to races 1-11 have been planted on a large scale. Similar developments have taken place when races Pfs 5 (1996), Pfs 6 (1998), Pfs 7 (1999), Pfs 8 and 10 (2004), and Pfs 11 (2009) were identified and named. The occurrence of Pfs 12 will create strong interest for Pfs 1-12 resistant spinach cultivars from both growers and breeders. 

The IWGP is a working group of Plantum NL consisting of spinach seed companies (Pop Vriend, Monsanto, Rijk Zwaan, Nunhems, Takii, Sakata, Bejo, Enza, Syngenta, Advanseed), Naktuinbouw, and the University of Arkansas. The efforts of the group are supported by research activities at the University of Arkansas and the University of California Cooperative Extension—Monterey County. The aim of the IWGP is to monitor and designate new races of downy mildew in spinach, and to promote a consistent and clear communication between the seed industry, researchers, and growers about all resistance-breaking races that are persistent enough to survive over several years, occur in a wide area, and cause a significant economic impact. 

IWGP is monitoring new races continuously by testing field isolates on a fixed, common host differential set of cultivars that contains the full range of available resistances. Researchers all over the world are invited to join the IWGP initiative and use the common host differential set to identify new isolates. For California, the Correll-Koike team will continue to receive and test spinach downy mildew samples for growers, pest control advisors, and seed companies. 

For more information on this subject you can contact Steven Koike (stkoike@ucdavis.edu), Jim Correll (jcorrell@uark.edu), Diederik Smilde (d.smilde@naktuinbouw.nl), or IWGP chairperson Jan de Visser (JandeVisser@popvriend.nl).



Downy mildew is the most damaging disease of spinach in California and causes yellow and tan leaf lesions.

To identify downy mildew races, a series of spinach cultivars is grown and inoculated; races are identified based on which cultivars become diseased. 

The seedcorn maggot (Delia platura) is a pest of many vegetable crops such as cabbage, broccoli, turnip, radish, onion, beet, spinach, pepper, potato, beans and peas.  Maggots usually feed on germinating seeds, roots or stems, resulting in reduction of seedling stands and contamination of the crop.  They also occasionally feed on head lettuce to make it unmarketable (the maggots damaging spring head lettuce were officially identified as seedcorn maggots.  For more information, please visit http://ucanr.org/blogs/SalinasValleyAgriculture/index.cfm?start=16, Spring Head Lettuce Crop Affected by Insect, Thursday April 29 2010).  The damage is especially severe during cool and wet winter or spring, and in fields with high organic matter.  The feeding damage often causes secondary infections by pathogen.  

The seedcorn maggot overwinters as pupa in soil.  The adult emerges in early spring and a female can lay an average of 270 eggs in the soil near plant stems.  The female prefers to lay eggs in fresh-tilled soil with high moisture and organic matter.  The eggs hatch in a few days and the maggots feed for 1 to 3 weeks on decaying organic matter or their host plants before pupation in soil.  

Prevention is the best management strategy for this pest.  Any cultural practice to speed up seed germination and plant growth will help to reduce crop loss.  Attach drag chains behind the planter during seeding can reduce egg laying in the seed row.  Fields with heavy manure or cover crop should be plowed at least 2 weeks before planting.  Fields with a history of seedcorn maggot problem may apply an insecticidal seed treatment at planting.  After damage is observed on the crop, rescue treatments are usually not effective.  Prompt resetting or replanting of the damaged crop may be necessary if stand loss is severe. 

Lygus bugs (Lygus species) are serious pest of strawberries in California even present at moderate densities.  Due to the emergence of pesticide resistance, it is essential to better time the few pesticides that are registered to control this pest.  The timing of pesticide applications is solely dependant upon a close monitoring of Lygus bug population dynamics and its developmental biology.  The sprays must be timed to kill the youngest immatures because the registered pesticides are not very effective against the adults. 

Lygus bugs feed on many host plant species.  In the Central Coast, they feed on strawberries and many flowering weed species and alternative crop hosts such as mustards, pepper weed, wild radish, vetch, alfalfa, and fava beans.  The adult bugs usually overwinter in these weed species and second-year strawberries.  They start to migrate to fall plantings in the spring, but only the adults can fly from one host to another.  In second-year strawberries, overwintered eggs are major sources of Lygus populations.  Our study showed that Lygus adults still lay eggs during December.

Monitoring to detect Lygus bugs on strawberries and the alternative hosts is the first step towards successful management of this pest.  Our monitoring program recently detected that the first generation of Lygus nymphs started hatching during the week from February 7 to February 11 in many second-year strawberry fields in the Central Coast region.  The cool weather and rain will slow down the Lygus emergence in these second-year fields but once the temperature starts to exceed 54 F for most of the day hours the emergence will again increase.  We suggest that growers check their second-year fields for the presence of small Lygus nymphs and apply treatments if the threshold is reached.

White mold disease, caused by the fungus Sclerotinia sclerotiorum, is causing damage to a number of vegetable crops in California and Arizona during the late 2010 and early 2011 months. On the coast of California, white mold is being found on crucifer crops such as broccoli and cauliflower. In the desert regions white mold is causing damage on broccoli, cauliflower, celery, lettuce, and other vegetables (for lettuce this disease is commonly called lettuce drop). White mold incidence on these crops appears to be greater than normally observed. See photos 1 through 6 below.

The first symptoms on most vegetable crop hosts are small, irregularly shaped, water-soaked areas on stems, leaves, pods, or flower heads. These infections quickly develop into soft, watery, pale brown to gray rots. Rotted areas can expand rapidly and affect a large portion of the plant. Diseased tissues eventually are covered with white mycelium, white mycelial mounds that are immature sclerotia, and finally mature, hard, black sclerotia. Mature sclerotia usually form after tissues are rotting and breaking down. Plants with infections on the main stems can completely collapse and fall over.

The black sclerotium is the survival stage of the fungus and can measure from ¼ to ½ inch long. Sclerotia are found in the soil and can directly infect plants if stems are in close proximity. However, these winter cases of white mold are due to ascospore infections. If sufficient soil moisture is present, shallowly buried sclerotia germinate and form small, tan mushroom-like structures called apothecia (photos 7 and 8). Ascospores (photos 8 and 9) are released from apothecia and carried by winds to the host plant. These ascospores are responsible for these winter infections and result in disease of the above-ground parts of plants. The relatively cool, moist weather found in most regions has allowed for the production of apothecia production and ascospore releases.

For ascospores to start colonizing plant tissues, nutrients and plant fluids from damaged tissues are usually needed. This is why white mold is very severe if ascospores land on compromised tissues such as lettuce leaves with tip burn, leaves and heads damaged by frost or other factors, stems with open wounds or exposed leaf traces (vascular tissue in the stem that is left exposed when a lower leaf falls off), and senescent leaves and stems.

Controlling white mold under these winter weather conditions is difficult. Protective fungicides provide some assistance and can be used effectively in lettuce. However, such fungicides need to be applied prior to ascospore flights and usually will require multiple sprays. Fungicides may not be warranted for crucifer crops.

Steve Koike thanks Jeff Rollins and Karen Chamusco for assistance with photographs for this article.

Photo 1: White mold (lettuce drop) on romaine lettuce.


Photo 2: White mold (lettuce drop) on romaine lettuce, showing white mycelium and two black sclerotia.


Photo 3: White mold on broccoli stems.


Photo 4: White mold on broccoli stem, showing white mycelium and one black sclerotium (center).


Photo 5: White mold on cauliflower head, showing white mycelium.


Photo 6:White mold on celery, showing numerous black sclerotia.


Photo 7: One sclerotium and several apothecia (spore producing structures) of Sclerotinia sclerotiorum.


Photo 8: Microscopic view of the spore-producing apothecium of Sclerotinia sclerotiorum. Note the lined-up ascospores (red) ready to be released. Photo used by permission (K. Chamusco).


Photo 9: Microscopic view of ascospores lined-up in a tube (called an ascus) and ready to be released. Photo used by permission (J. Rollins).