Source and Rearing of Aphid Species

Aphids were collected from citrus (Citrus sinensis Pers.), cucumbers (Cucumis sativus L.), scarlet milkweed (Asclepias curassavica L.), and cowpeas [Vigna unguiculata (L.)] in Alachua County, Florida between Apr and Jun 2005. Healthy adult females were hand transferred to clean host plants to obtain pure colonies. Aphid species were confirmed with identification keys by Blackman & Eastop (2000) and Halbert & Brown (1996) and confirmed by Keith S. Pike. Colonies of T. citricida, T. aurantii, and A. spiraecola were reared on potted grapefruit (Citrus × paradisi Macfad.), whereas A. gossypii and A. craccivora were reared on potted cucumber (Cucumis sativus L. var. Cucumber-Poinsett 76) and cowpea [Vigna unguiculata (L.) var. Cream 12 Peas] plants, respectively. Aphis nerii Boyer de Fonscolombe was reared on scarlet milkweed. Colonies were maintained in PVC cages (61 × 61 × 61 cm) covered with mesh sleeves (BioQuip Products, Gardena, CA) at 22-25°C, 60-70% RH under an 18:6 (L:D) photoperiod.

Rearing of L. oregmae

Lipolexis oregmae was reared on potted citrus trees infested with mixed stages of T. citricida (Walker & Hoy 2003). Seven citrus plants infested with approximately 500-700 aphids of mixed stages per plant were transferred to wood-framed mesh parasitoid cages (87 × 48 × 50 cm) twice a week (Walker & Hoy 2003). Parasitoid males and females were aspirated into a 50-mL conical centrifuge tube (USA Scientific, Ocala, FL) and fed honey and water for 6-12 h prior to release. At least 50 L. oregmae females and ∼20-30 males were released per cage. Each colony cage was provided with a honey strip and 2 water-soaked cotton pads at the time of parasitoid release and at the time of adult emergence of the next generation because adults require free water to drink and a high relative humidity. The colony was maintained at 25 ± 1°C, 60-70% RH under an 18:6 (L:D) photoperiod.

Host Plant Rearing

Grapefruit trees (35-40 cm tall) grown in 11.4-liter pots were pruned and soil in the pots was soaked with 250 mL of bleach (sodium hypochlorite 6.15% a.i., Ultra Clorox Germicidal Bleach, Clorox Professional Products Company, Oakland, CA) solution diluted (5 mL of bleach/100 mL water) to reduce fungal growth that could kill aphid mummies (Walker & Hoy 2003). After 20 min of treatment with bleach, pots were drenched with water. Plants were fertilized (Peter’s 20-20-20; United Industries, St Louis, MO) and sprayed with 0.26% (20 mL/7.57 L) of horticulture grade oil (Dyne-Amic, Setre Chemical Company, Memphis, TN) to reduce insect contamination. Plants were transferred to a greenhouse and kept there for approximately 2 weeks to allow them to produce tender new shoots (=flush) prior to being infested with aphids. Pittosporum tobira (Thunb.) W. T. Aiton plants purchased from Lowe’s (Gainesville, FL) were pruned, fertilized and sprayed with 0.26% of horticultural oil. Pruned plants were maintained in a mesh cage at 22-25°C, 60-70% RH under an 18:6 (L:D) photoperiod in the greenhouse until new growth was produced.

Cowpeas were raised from seeds in 3.8-L pots in a greenhouse. Once the plants produced 3 to 4 true leaves, 10 plants were transplanted to the center of another pot prior to being infested with aphids. Cucumber plants were raised from seeds in 3.8-L pots. Six seeds per pot were sown and thinned to 4 plants each when seeds emerged. Milkweed plants potted in 3.8-L pots were purchased (Lowe’s, Gainesville, FL). Cowpeas, cucumbers, and milkweeds were not treated with chlorox prior to being infested with aphids.

Assessment of L. oregmae Mummy Location When Reared on Different Aphids and Host Plants

Once the grapefruit trees had produced flush 1-2 cm in length, the soil surface was soaked for a second time with the bleach solution for 20 min and then flushed with water. The pots were then filled to the top rim with potting mix (Jungle Growth, Professional Grower Mix, Piedmont Pacific Inc., Statham, GA). The pot surface then was lined with plastic wrap (Borden Packaging & Industrial Products, North Andover, MA) and covered with 32 cm diameter filter paper (Whatman Ltd., U.K.) and taped to the sides of the pot with Duck tape (Henkel Consumer Adhesives, Avon, OH). The holes on the bottom of pots were sealed with tape. Trees were placed into the colonies of aphids for infestation. After 2 d, the plants were removed from the colonies and examined for a desired density of approximately 150-200 mixed stages of immature aphids per plant. Infested plants were placed into wood-framed mesh parasitoid cages in which the bottom was lined with a plastic sheet so that mummies could be seen, counted, and removed. The Pittosporum plants were treated similarly.

Cowpea, melon, and milkweed plants were allowed to establish for 3-4 d, the soil surface was soaked with the bleach solution for 20 min and then flooded with water. The pots were then treated as described for the grapefruit trees until being infested with aphids. After 2 d, plants were removed from the aphid colonies and examined for a desired density of approximately 150-200 mixed stages of immature aphids. Then infested plants were placed in PVC cages covered with mesh sleeves and the bottom of each cage was lined with a plastic sheet.

One female and 2 males of L. oregmae less than 24 h old were collected in a plastic vial and kept for 2 h to ensure mating. A honey strip was placed inside the vial and a water-soaked cotton pad was placed on the top of the vial covered with mesh to provide food and water for the parasitoids. The parasitoids were released after 2 h into the wood-framed mesh parasitoid cages (citrus and Pittosporum), or the PVC cages (milkweeds, cowpeas, cucumbers), which were monitored daily for the production of progeny (mummies). For all experiments, but one, 5 replicates were evaluated. In the case of cowpeas, 15 replicates were observed. In the case of grapefruit, milkweed, and Pittosporum, 1 plant per pot was used, whereas 10 plants per pot were used for cowpeas and 4 plants per pot for cucumber to yield the desired total of ca. 150-200 aphid hosts per cage. The number and location of the progeny mummies (on plant leaves, on stems, or on the bottom of the cage) was recorded 8, 10, and 12 d after parasitoids were released into the cage. The number of mummies produced on these different host aphids and plants was analyzed by ANOVA (SAS 1999) and means were separated by Fisher’s protected least significant difference (LSD) test, 5% level.

Aphid and Mummy Colors When Reared on Different Host Plants

The color of aphid mummies produced by L. oregmae on different host aphids and host plants was determined by matching mummy color to color chips in the Munsell book of color (Anonymous 1942). Five mummies were randomly selected from each host aphid and host plant combination. Individual mummies were matched to the color chips in daylight outdoors at noon under a 10× dissecting microscope following the instructions provided. For the Munsell charts, data were recorded as HV/C, where ‘H’ is hue name, ‘V’ is the value in the vertical column on the left of the chart, and ‘C’ is chroma in the horizontal line at the base of the chart. Similar procedures were followed to determine the color of wingless adult aphids from the colonies in order to compare aphid and mummy color. After obtaining the color values from the Munsell charts, the color designations were compared to the ISCC-NBS color name charts (Anonymous 1964) and the name of the colors observed was recorded.

Handling of L. oregmae Mummies for Adult Emergence

A single citrus shoot infested with brown citrus aphids was collected from cages of the laboratory colony of L. oregmae 4 d after release of adult parasitoids. The number of aphids per shoot was estimated and shoot length recorded for each of the 6 replicates. Each shoot was then placed in a plastic bag (60 × 30 cm) inflated with air along with 2 moist paper towels at 24 ± 1°C, 50-60% RH under an 18:6 (L:D) photoperiod. Mummies were removed from the bags after 6 and 10 d and counted; on each date, half were plated on 1.5% agar (non nutrient) plates (Difco Agar granulated, Becton, Becton, Dickinson & Company, Sparks, MD) and the rest were kept individually in gelatin capsules (No. 3, Eli Lilly and Company, Indianapolis, IN) within a 50-mL vial for 8 d to evaluate adult emergence. The number of mummies counted after 6 and 10 d were summed and the proportion of adults that emerged on both dates was calculated. The percentage parasitism of the aphids on the citrus shoots was calculated based on the number of mummies divided by the greater value (300) of the estimated number of aphids per shoot (range: 140-300) at the beginning of the experiment, and then average percentage parasitism was obtained for the 6 replicates.

Assessment of L. oregmae Mummy Location When Reared on Different Aphids and Host Plants

Host aphid and host plant did not affect the location of mummies of L. oregmae (Table 1). All mummies of L. oregmae produced in all host-aphid and host-plant combinations were found on the cage bottom or beneath the pots and sticking to the tape used for covering the holes on the pot bottoms. No mummies were found on the foliage and/or stems. Hill & Hoy (2003) obtained similar results with mummies of L. oregmae reared in brown citrus aphids on citrus and reported 98.6-99.2% of the mummies were on/in the soil surface. Thus, based on this sample of diverse aphids and host plants, it appears that L. oregmae only produces mummies off the host plant.

In these no-choice tests, when L. oregmae was reared in 5 different host aphids reared only on grapefruit, a significant difference (F = 98.29, df = 4, P < 0.0001) was observed in the mean number of mummies produced (Table 1). Lipolexis oregmae produced more mummies (mean = 188.2) in brown citrus aphid hosts followed by spirea aphid (144.0), black citrus aphid (132.8), melon aphid (58.2), and cowpea aphid (54.2) (Table 1). The higher rate of reproduction on brown citrus aphids on citrus is significant because this pest was the target of the classical biological control introduction.

When L. oregmae was reared in spirea aphid on grapefruit, a mean of 144.0 progeny were produced, which was significantly (F = 217.86, df = 1, P = 0.0001) higher than when L. oregmae was reared in spirea aphid on Pittosporum (51.0), suggesting that host plant may affect parasitism by L. oregmae. No significant difference (F = 0.09, df = 1, P = 0.7816) was observed in the mean number of mummies produced by L. oregmae when reared in melon aphids on grapefruit (58.2) or on cucumber (58.6). The mean number of mummies produced in cowpea aphids on grapefruit (54.2) was higher (F = 55.91, df = 1, P = 0.0017) than in cowpea aphids on cowpeas (0.7), which suggests that cowpeas affected oviposition or developmental success by the parasitoid. Despite the fact that we increased the number of replicates over a period of several months, L. oregmae mummy production remained low on cowpeas, indicating that this host plant and aphid combination would be unlikely to yield significant numbers of L. oregmae in a field survey.

Lipolexis oregmae produced an average of 48 mummies in the oleander aphid A. nerii (Boyer de Fonscolombe) on milkweed, which appears to be a new host record for this parasitoid (Table 1). This suggests that the host range of L. oregmae may be broader than previously recorded and that evaluation of nontarget effects in Florida should involve sampling diverse host plants and aphid species.

Because the number of aphid hosts provided (150-200) to the females was similar among the aphid-host plant combinations, these laboratory data indicate that L. oregmae performs better on brown citrus aphids on citrus, the target of the classical biological control program, than on other aphid hosts on citrus or on other host plants. In general, L. oregmae appears well adapted to aphids on citrus (Table 1).

Aphid and Mummy Colors When Reared on Different Host Plants

The color of the individual wingless adult aphids reared varied when reared on the same host plant under our laboratory conditions, but greater variability was observed when the aphids were reared on different host plants (Table 2). Brown citrus aphids, which probably develop only on citrus, were “Reddish yellow red” to “Yellow red” by the Munsell values and “Brownish black-Black” by the ISCC-NBS color chart, and black citrus aphids appeared similar to the brown citrus aphids when using the Munsell and ISCC-NBS color values (Table 2). Halbert & Brown (1996) described brown citrus aphids as shiny black and black citrus aphids as matte black in color. According to Blackman & Eastop (2000), the color of brown citrus aphids and black citrus aphids vary from very dark brown to black and reddish brown to brown black, respectively. Thus, different people may use different terminology to describe the same colors; for example, cowpea aphids reared on grapefruit are described in the Munsell Book of Color Values as “Greenish yellow to Yellow green yellow” while the ISCC-NBS color chart calls the same colors “Light olive gray to Olive black” (Table 2). However, by using either color chart, it is possible to provide consistent color designations, even though the color terminology used may not be consistent. These color evaluations were conducted between 1200 and 1400 h in sunlight and with a dissecting microscope, as recommended by the Munsell Book of Color Values, because color evaluations should be conducted with standardized light conditions.

The color of cowpea aphids also varied when reared on grapefruit and cowpeas (Table 2). Halbert & Brown (1996) and Blackman & Eastop (2000) described adult wingless cowpea aphids as shiny black in color. Melon aphids differed only slightly in color when reared on grapefruit and cucumber. Spirea aphids varied in color when reared on grapefruit or on Pittosporum. The oleander aphid could not be reared on citrus, but was a strong yellow when reared on milkweed (Table 2).

The color of mummies produced by L. oregmae when reared in different host aphids reared on the same or different host plants also varied (Table 3). Lipolexis oregmae, when reared in different host aphids but on the same host plant, produced mummies ranging from light gray yellowish brown to dark gray yellowish brown, medium brown to dark yellow brown, and grayish brown to dark grayish brown.

The color of mummies on same host aphid, but a different host plant, also varied (Table 3). In the case of oleander aphids reared on milkweed, the mummies produced were strikingly different from all other host aphid and plant combinations. The color of the mummies produced when L. oregmae was reared in spirea aphids on Pittosporum was reddish brown compared to grayish brown when mummies were produced in cowpea aphids, black citrus aphids, and brown citrus aphids.

These differences in the color of mummies produced by L. oregmae in different host aphids reared on the same or different host plants make it difficult to use color as a method for identifying L. oregmae mummies, especially if sampling among an array of host plant and aphid species. However, all mummies appeared similar in texture and size, even though they differed in color. Lipolexis oregmae mummies, at least when reared in these aphid hosts, are not shiny and appear somewhat ‘fluffy’ because silk strands protrude from the broken surface of the aphid exoskeleton. Typically, there are multiple breaks in the aphid exoskeleton surrounding the parasitoid with the parasitoid cocoon silk extruding. We have seen no evidence that L. oregmae glues the aphid exoskeleton to the substrate, unlike the behavior of many aphidiid species (Stary 1970).

Handling of L. oregmae Mummies for Adult Emergence

When L. oregmae was reared under laboratory conditions, the individual shoots had 140-300 brown citrus aphids per shoot, averaging 193.3 (SD = 54.3). The number of mummies produced per shoot ranged from 116 to 223, averaging 153 (38.6). Percentage parasitism ranged from 68.2 to 88.7, averaging 79.7% (7.5). A large proportion of the mummies recovered from the bags were found between the folds of the paper towels and were attached only very loosely to the rough paper by the silky strands protruding from the mummy. This behavior suggests that the parasitized aphids move to darker, more humid and secure areas before becoming mummified in the field.

Mummies (n = 202) of L. oregmae kept in gelatin capsules did not produce any adults. This was unexpected because many aphidiid mummies readily produce adults in gelatin capsules. We suspect that the failure to obtain adults was due to lower-than-optimal relative humidity conditions but other factors may influence emergence of adults from mummies. Walker & Hoy (2003) only achieved a mean percentage emergence (SE) of 44.5% (1.6) of L. oregmae adults from 100 second-instar brown citrus aphids reared on potted citrus plants enclosed in plastic cylinders under laboratory conditions, which suggests that mortality of L. oregmae mummies could have occurred due to fungal pathogens as well as to insufficiently high relative humidity on the surface of the soil.

For the 353 mummies that were placed on the 1.5% agar plates, the mean percentage emergence (SD) of adults was 95.7% (5.3). These mummies were 1-6 d old when placed on the agar and were held for 8 d. Most of the adults emerged within 4-6 d after being placed on the agar. Due to the high relative humidity in the petri dish, some fungal growth was observed growing on the mummies and on the nonnutrient agar after several days, but this apparently did not prevent emergence of the parasitoid adults.

The high rate of emergence of adults from mummies on the agar plates allowed us to ship L. oregmae mummies to Jamaica via express delivery service (in boxes that did not contain ice) during Oct and Dec 2005, where they were to be reared for a classical biological control program. D. Clarke-Harris (personal communication) kindly recorded the number of adults that emerged from these 2 shipments, which experienced unknown temperature conditions. In the first, 80.9% of the 110 mummies produced adults, while in the second shipment 109 adults were produced from 450 mummies (24.2%). Because adults of L. oregmae are short lived and difficult to ship, this agar plate method appears to be an effective method for shipping this species, as well as a useful method for obtaining adults for taxonomic confirmation.