§      Watch out for low pH
§      Magnesium deficiency in geraniums
§      Leaf tip burn on Oriental lilies
§      Characterizing the diversity of domestic populations of Western flower thrips and their potential impact on floricultural crops

"The media pH is 6.0 and look at the damage." I've heard this comment way too often since we learned to recognize the symptoms of iron/manganese toxicity in geranium, marigold, Lisianthus and New Guinea impatiens. Symptoms of iron/manganese toxicity can develop three or more weeks after the pH drops below 5.8 so unless you're testing frequently you won't spot the problem until it's too late.

pH can drop very rapidly, in less than a week in geraniums, so you may need to test every three or four days in rapidly growing crops in four-inch or smaller pots.  You need to look at trends in pH  (either up or down), so graph your data using a spreadsheet program or on paper. It's a lot easier on the plants to make small adjustments to pH by switching fertilizers or rates rather than rescue them after the pH falls off a cliff. To give yourself a margin of safety, hold the pH of iron/manganese sensitive crops at 6.2 to 6.4.  The plants won't have any problems with iron uptake and you will have more time to catch falling pH and make corrections.

Magnesium deficiency in geraniums

Dean Krauskopf
Integrated Crop Management Agent
Southeast Michigan

I'm seeing geraniums with red lower leaf margins, which progress to interveinal chlorosis.  There could be several causes of these symptoms (which makes accurate diagnosis difficult when you only have symptoms to work with) including cold temperatures, holding the plants very dry for a long period, root problems and magnesium deficiency. The only way to confirm magnesium deficiency is by foliar or media analysis. But if you don't have magnesium in your water, don't have dolomitic lime in the media, aren't applying a fertilizer incorporating magnesium, and feeding predominately calcium and potassium, the problem probably is magnesium deficiency.  Even if you have magnesium in the media, high levels of calcium and potassium can interfere with magnesium uptake and the plant shows magnesium deficiency symptoms. 

Because of this interaction, you need to look at the balance between these three nutrients.  There should be adequate uptake of magnesium when the percent of total soluble salts (saturated paste media analysis technique) for magnesium is above 4% with calcium between 14 to 16% and potassium between 11 and 13%.  One application of Epsom salts at 8 to 16 ounces per 100 gallons will usually bring the nutrient balance back into line.  Don't over-apply magnesium because high levels can interfere with calcium and potassium uptake, although it would be rare.  Remember by the time you see symptoms, the plants have lost growth and quality that they won't regain.  Don't guess.  Media test.

Necrotic lesions on leaves of Oriental lilies (such as "Stargazer") is a common production problem and when severe can reduce the value of the crop (see photos). This physiological problem often occurs shortly before or at visible bud.  It is more common in double polyhouses than in glasshouses, and is more of a problem in late winter than during the spring or summer. The cause of the necrosis has been associated with a calcium deficiency, and is similar to what can cause marginal burn on poinsettia bracts. Calcium is considered an immobile nutrient in plants, meaning that deficiency symptoms occur on younger portions of the plant.  Uptake by the plant is passive meaning the plant has to transpire to absorb calcium into the roots.  In a humid greenhouse with low light conditions, water and calcium uptake by the plant is rather low.  If the temperature in the greenhouse is warm during cloudy weather, young leaves continue to expand with little calcium uptake (because transpiration is low).  If a sunny, warm day follows several days of cloudy weather, the young leaves are stressed by the high light and necrotic lesions develop, probably because cell walls are thin due to low levels of calcium.

There are a few strategies growers can employ to prevent leaf tip burn.  First, growers can use calcium nitrate as one of the nitrogen sources so that there are adequate calcium levels in the media.  However, this by itself will not prevent the problem.  Second, low humidity and airflow from horizontal airflow fans will promote transpiration (or water loss by the plants) under cloudy conditions, promoting uptake and distribution of calcium in the leaves.  This may or may not prevent the problem. Third, it is recommended to spray calcium chloride or calcium chelate (at about 400 ppm) on plants every three to four days, beginning at least one week before the first plants reach visible bud and continuing for about one week after all plants are at visible bud.  It is especially important to apply a spray just before a sunny day following several cloudy days.  These calcium sprays have proven effective in preventing the problem.  To apply 400 ppm using 77 to 80 percent calcium chloride, 5.6 grams (or 0.20 ounces) would need to be added for every gallon of water.  For a 100-gallon stock tank, that means you would have to add 560.7 grams (or 19.78 ounces).  Using these strategies can help prevent leaf necrosis that is common with Oriental lilies.

Abstract
Frankliniella occidentalis (Pergande), western flower thrips (WFT), obtained from native (N), laboratory (LC), or greenhouse (GH) environments in California (CA), Illinois (IL), Massachusetts (MA), Nevada (NV), or Texas (TX) were evaluated for feeding aggressiveness on Impatiens wallerana Hook.f. and for spinosad (Conserve®) resistance on Gerbera jamesonii Bol. ex. Adlam. In one experiment, insects from seven populations, CA-N2, CA-N3, CA-LC1, IL-LC1, TX-LC1, CA-GH1, and IL-GH1, were used to assess feeding aggressiveness or to initiate a laboratory colony. Feeding aggressiveness was assessed 0, 7, 14, and 21 weeks after collection using a digital image analysis system to determine the percent leaf area damaged by feeding. Damage varied the most at 0 weeks after collection and variation decreased until 21 weeks after collection. Declining damage was attributed to the standardization of fitness in the laboratory colonies or possibly to limited genetic diversity within the colonies reducing insect fitness over time. In a second experiment, nine populations, CA-N2, NV-N1, NV-N2, CA-GH1, IL-GH1, TX-GH1, IL-LC1, MA-LC1, and TX-LC1, reared for four months in the laboratory varied in percent survival when flowers inoculated with 25 WFT were sprayed with spinosad at label (0.81 mL×L^-1), half label (0.41 mL×L^-1), deionized water, or no spray. At the 0.41 mL×L^-1rate, CA-GH1 and IL-GH1 populations had the highest survival at 8.8 and 5.0 percent, respectively. At the 0.81 mL×L^-1 rate, 8.8 percent of recovered insects from IL-GH1 survived which was significantly more than any other colony. Feeding aggressiveness of WFT populations on impatiens leaves varies. Some resistance to spinosad (Conserve®) exists in greenhouse populations of WFT.

Introduction
Western flower thrips, Frankliniella occidentalis (Pergande), is a major pest of the floriculture industry worldwide. Western flower thrips (WFT) not only cosmetically damage crops valued for their appearance, but also vector plant viruses. The tendency of WFT to feed inside flower buds and unexpanded leaf tissue, resistance to insecticides¹, and government regulations to reduce chemical usage necessitates the use of alternative controls.

Host plant resistance to WFT varied among cultivars of chrysanthemum², impatiens³, roses⁴, and gladiolus⁵. Host plant resistance must be durable over environments and insect populations. The durability of resistance depends on screening crops with aggressive insect populations². Local environmental conditions, available host plants, and insecticide applications select for insects adapted to a particular set of conditions. Insect populations in different geographical regions may differ in genetics, morphology, and behavior. Of 10 internationally collected WFT populations, a population from the United States was the most damaging to chrysanthemums². More aggressive WFT populations may exist in the western U. S.

Population diversity changes over time, especially if a small sample is brought into a laboratory environment for rearing⁶. Laboratory colonies may become unrepresentative of sampled populations due to founder effect, genetic drift, food sources, and rearing environment⁷. Previous comparisons of WFT populations have standardized populations to laboratory conditions before evaluation²; however, variation present immediately after collection may be important to determine environmental impact on aggressiveness. For host plant resistance to be effective, colonies initiated with aggressive insects must maintain that aggressiveness and represent endemic populations.

WFT were obtained from native, greenhouse, and laboratory environments within the United States to determine 1) if WFT feeding aggressiveness on impatiens varied among populations reared for varying lengths of time in a laboratory and 2) if populations varied in resistance to spinosad, an insecticide used to manage WFT.

Materials and methods
Western flower thrips populations were obtained from laboratories, greenhouses, and native habitats in five states. Upon arrival at the University of Illinois, each population was sampled to determine initial feeding aggressiveness, to initiate a laboratory colony for subsequent experiments, and to verify species collected. Laboratory colonies were initiated with 50 to 100 insects from the initial samples collected by collaborators in five states. Each population was maintained in isolation in two laboratory rearing cages maintained at 26°C and a 16:8 L:D photoperiod for 21 weeks. Cut flowers, green bean pods, and dilute honey provided food and oviposition sites for the insects. Two experiments were conducted to determine initial feeding aggressiveness of each thrips population and to determine population resistance to spinosad (Conserve®).

Experiment 1: July 21 to December 6, 2001
Seven WFT populations, two native, two greenhouse, and three laboratory, were used to inoculate two impatiens F₁ hybrid cultivars. Single leaves on impatiens plants were inoculated with 2 WFT for 48 hours. To determine possible impact of laboratory rearing on feeding aggressiveness, leaves were inoculated and evaluated 0, 7, 14, 21 weeks after insects were collected (weeks after collection). The percentages of WFT alive, dead, or not recovered were determined along with the percentage of leaf area damaged and damage location. Data were analyzed as three-factor factorial in a RCB design with three blocks using mixed models

Experiment 2: November 3 to November 8, 2001
Nine WFT populations representing three native, three greenhouse, and three laboratory populations were used to inoculate cut gerbera daisies. Individual cut flower stems were inoculated with 25 WFT and maintained in isolation for 48 hours to allow insect establishment on the flower heads. After 48 hours, one of four spray treatments was applied; no spray, deionized water, or spinosad at 0.41 and 0.81 mL×L^-1. Flowers were dissected 72 hours after spray treatments and the percentage of WFT alive or dead was determined. Data were analyzed in a CR design with five replications using mixed models.

Results and discussion
In Experiment 1, survival rates of WFT varied by evaluation date (Figure 1a). Leaf cage failure resulted in a higher number of non-recovered WFT 0 weeks after collection than on any other evaluation date. Higher death rates were noted 0 and 14 weeks after collection, which coincided with sunny weather indicating that environment can negatively influence survival. WFT populations varied in feeding damage 0 weeks after collection (Figure 1b.) As populations were reared in the laboratory, variation in percent leaf area damaged caused by populations decreased. Standardizing population health and small founding numbers likely resulted in the observed decrease among populations. The reduced variation also indicates that environmental parameters influence WFT feeding aggressiveness.

In Experiment 2, WFT populations were screened for resistance to spinosad (Conseve®), an insecticide currently recommended to manage WFT in greenhouse floricultural crops. The percentage of insects recovered that were alive ranged from 37.0 to 72.8 for the no spray and deionized water controls (Table 1). WFT survival percentages greatly decreased when spinosad was applied at 0.41 or 0.81 mL×L^-1, half and label rate, respectively. Survival rates varied among populations with IL-GH1 exhibiting 8.8 % survival at both spinosad rates (Table 1). The CA-GH1 and IL-GH1 population survival percentages were similar when treated with 0.41 mL×L^-1 spinosad. When treated with insecticide, native and laboratory populations were effectively controlled (Table 1). Resistance to spinosad is developing in some greenhouse populations of WFT.

Conclusions
Domestic WFT populations vary in feeding aggressiveness based on impatiens leaf area damaged. This variation in feeding aggressiveness decreases as insect populations are standardized through laboratory rearing. As one might expect, environmental parameters influence insect aggressiveness and thereby impact feeding damage expressed on impatiens. The need to properly rotate chemicals listed to manage western flower thrips is highlighted by the identification of two greenhouse populations from two states showing increased levels of resistance to spinosad. Native and laboratory populations not exposed to spinosad had no resistance to this insecticide. Variation in WFT populations for feeding aggressiveness and insecticide resistance support the need to identify alternative control measures for WFT.

References
1)    Immaraju, J.A., T.D. Paine, J.A. Bethke, K.L. Robb, and J.P. Newman. 1992. Western flower thrips (Thysanoptera:Thripidae) resistance to insecticides in coastal California greenhouses. Journal of Economic Entomology 85:9-14.

2)    de Kogel, W.J., M. van der Hoek, M.T.A. Dik, F.R. van Dijken, and C. Mollema. 1998. Variation in performance of western flower thrips populations on a susceptible and a partially resistant chrysanthemum cultivar. Euphytica 103:181-186.

3)    Herrin, B.B. and D.F. Warnock, 2002. Resistance of impatiens germplasm to western flower thrips feeding damage. HortScience.37:802-804.

4)    Gaum, W.G., J.H. Giliomee, and K.L. Pringle. 1994. Resistance of some rose cultivars to the western flower thrips, Frankliniella occidentalis (Thysanoptera: Thripidae). Bulletin of Entomological Research 84:487-492.

5)    Zeier, P., and M.G. Wright. 1995. Thrips resistance in Gladiolus spp.: potential for IPM and breeding, pp. 411-416. In: B. L. Parker, M. Skinner, and T. Lewis (eds.). Thrips Biology and Management. Plenum Press, N.Y.

6)    Mason, L.J., D.P. Pashley, and S.J. Johnson. 1987. The laboratory as an altered habitat: phenotypic and genetic consequences of colonization. Florida Entomologist 70:49-58.

7)    Bartlett, A.C. 1985. Guidelines for genetic diversity in laboratory colony establishment and maintenance, pp. 7-17. In: P. Singh and R. F. Moore (eds.). Handbook of Insect Rearing. Elsevier, N.Y.

Acknowledgements
Funding for this project was provided in part by the University of Illinois Research Board and USDA CRIS Hatch Project ILLU-65-0308. The time, effort, and interest of the following collaborators who supplied thrips populations are greatly appreciated: Dr. Michael Brownbridge (University of Vermont), Ian Greene (Whitmire Micro-Gen Research Laboratories, Inc), Dr. Roy van Driesche (University of Massachusetts), Dr. Kevin Heinz (Texas A&M University), and Dr. Michael Parrella (University of California, Davis).