The sugarcane aphid, Melanaphis sacchari (Zehntner) (Hemiptera: Aphididae), is a recent invasive pest of sorghum, Sorghum bicolor (L.) Moench (Poaceae), production in North America. Several species of grasses (Poaceae) have been recorded as hosts of M. sacchari (Singh et al. 2004). For example, M. sacchari is an economic pest of sorghum in Australia, Asia, Africa, and South America (Singh et al. 2004). Melanaphis sacchari was first reported in the United States from Hawaii in 1896 (Pemberton 1948), followed by reports from Belle Glade, FL, in 1977 from sugarcane, Saccharum officinarum L. (Mead 1978), and later in Louisiana in 1999 (White et al. 2001). Denmark (1988) documented sugarcane aphids feeding on sorghum in Florida, but did not report economic losses. The first significant infestation of M. sacchari on grain sorghum in the United States was reported along the Texas Gulf Coast and Louisiana (Villanueva et al. 2014). The insect was first detected in Georgia in August 2014 and had spread to South Carolina by mid-September of the same year (G.D.B. unpubl.). By the end of 2015, M. sacchari was reported from 17 states and Mexico feeding on grain sorghum, sorghum–sudan hybrids (S. bicolor × S. bicolor var. Sudanese), sweet sorghum, several millet varieties (Panicum miliaceum L.), and johnsongrass, Sorghum halepense (L.) Persoon (Bowling et al. 2015). Although M. sacchari can be found on field edges in corn, Zea mays (L.) (Poaceae), and cotton, Gossypium spp. (Malvaceae), these are nonreproductive hosts. The M. sacchari genotype specializing on sorghum likely resulted from transportation of the alates to North America by wind or through human activities (Bowling et al. 2016). This genotype, designated as MLL-F, belongs to one asexual clonal lineage that is believed to have originated either from Africa or Asia, the latter being the most probable (Bowling et al. 2016, Harris-Shultz et al. 2017, Nibouche et al. 2018).

Melanaphis sacchari is most commonly found on the abaxial side of sorghum leaves, where they establish and feed, avoiding direct sunlight. Like other types of aphids, M. sacchari is a phloem feeder (Singh et al. 2004). Plant damage occurs due to direct loss of plant nutrients, leaf chlorosis, and lost photosynthetic ability following development of secondary sooty mold that occur on honeydew excreted by the aphids. A loss of $62 to $432 per hectare has been reported from fields with a pest density of 250 aphids per leaf (Bowling et al. 2016). Yield loss is attributed to poor plant vigor and head emergence, in addition to inefficient harvest due to high aphid density and sticky honeydew (Villanueva et al. 2014). In the United States, M. sacchari propagates through asexual reproduction only. Overwintering adults and nymphs survive on remnant and ratooning sorghum as well as johnsongrass in the southern United States (Bowling et al. 2016).

A number of M. sacchari natural enemies are reported in studies from Texas. These beneficial insects include a number of aphidophagous predators and Aphelinus sp. nr. varipes (Hymenoptera: Aphelinidae) (Bowling et al. 2016, Colares et al. 2015a). Compared with a suite of generalist predators, coevolved parasitoid specialists often have the strongest impact in suppressing pest populations (Schmidt et al. 2003, Snyder and Ives 2003, Stiling and Cornelissen 2005). Therefore, identification and conservation of a key parasitoid attacking M. sacchari could lead to improved and effective biocontrol programs.

Because M. sacchari is a novel host for the aphid parasitoids in Georgia, any information about the associations between these species and the rates of parasitization is limited. In Kansas and Texas, Lysiphlebus testaceipes (Cresson) (Hymenoptera: Braconidae) was reported to attack M. sacchari in sorghum fields (Colares et al. 2015b, Knutson et al. 2016). Objectives of the study reported herein were to: (1) assess the fecundity of L. testaceipes parasitizing M. sacchari in the greenhouse, and (2) examine the temporal synchrony between L. testaceipes and M. sacchari in the field during 2017 and 2018.

Materials and Methods

Greenhouse study. A continuous supply of grain sorghum hybrid plants was maintained in the greenhouse to support the research. Three sorghum seeds (SS-800A, Southern States Coop. Inc., Richmond, VA) were sown per 11.4-liter plastic pots filled with Metro Mix 300 growing medium (Sun Gro Horticulture, Bellevue, WA) and fertilized bimonthly with 6.5 g of Osmocote 14-14-14 and 5.5 g of Micromax 90505 (The Scotts Co. LLC, Marysville, OH). This sorghum hybrid was selected due to its known susceptibility to M. sacchari. At the 2-leaf stage, two of the three seedlings were culled leaving one plant per pot. The potted sorghum plants were watered daily using automated sprinklers that provided 400 ml of water per pot.

A laboratory colony of M. sacchari was initiated from a natural infestation (approximately 300 adults) occurring on grain sorghum from Tift Co., GA, in October 2016. The colony was maintained in the greenhouse during the winter months on sorghum plants (starting with the 7-leaf stage) under a 13:11 (L:D) photoperiod and temperatures that fluctuated between 20 and 25°C and 60 ± 10% relative humidity (RH).

The greenhouse colony of M. sacchari was found to be parasitized during March 2017 (S.L. pers. obs.) (Fig. 1). The resulting aphid mummies were collected and held until the parasitoids emerged. Those adults were then identified as L. testaceipes using a taxonomic key (Pike et al. 1997) followed by a confirmation by N.C. Elliot (USDA-ARS, Stillwater, OK); voucher specimens have been deposited in the University of Georgia Collection of Arthropods in Athens. A subsequent parasitoid colony was maintained on live sorghum plants infested with M. sacchari inside a fine mesh cage (model BD2120, MegaView Science Co., Ltd., Taichung, Taiwan).

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Experimental exclusion cages were fabricated from polyethylene pipe insulation with wall thickness of 1 cm and internal diameter of 3 cm (Therma-Cel^®, Nomaco Insulation, Tarboro, NC). The pipe insulation was cut into 2-cm-thick rings using a band saw and then the opening on one side of the ring was covered with white organza fabric (Jo Ann Stores LLC, Hudson, OH) adhered with hot-melt glue. With the fabric-covered ends distal to the leaf surface, two sections of pipe insulation were tightly sandwiched over a sorghum leaf using hook-and-loop tape (Velcro USA Inc., Manchester, NH) that was attached to the pipe insulation using hot-melt glue. Small amounts of cotton lint were forced into any minute gaps between the cage and the leaf to prevent confounding emigration or immigration.

After the potted sorghum plants reached the 8-leaf stage of vegetative growth, the uppermost fully expanded leaf was selected for experiments. A single apterous adult was gently transferred from the aphid colony to the abaxial leaf surface using a fine bristle brush (number 0, Yangzhou Fuxin Artist Brushes Co., Ltd., Jiangsu, China), and the exclusion cage was fastened over the aphid to restrict the aphid from escaping (Fig. 2). The aphid was then allowed to feed and produce offspring. After 96 h, the leaf disk surrounded by the aphid cage with caged aphids was carefully cut from the plant and placed in an 8-cm-diameter polystyrene petri dish (P5856, Sigma-Aldrich, Milwaukee, WI) lined with dry filter paper (Fisherbrand^® Filter Paper P8-creped, Fisher Scientific, Pittsburgh, PA). Immediately thereafter, digital images of each leaf disk were acquired in order to accurately discern quantity and life stage of the individual aphids.

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Next, a single female L. testaceipes (24 h old) was introduced into the petri dish for 24 h, following the methods of Colinet et al. (2005). During this time the petri dishes were held in an environmental growth chamber maintained at 25 ± 1°C, 85 ± 1% RH, and on a 14:10 (L:D) photoperiod. Prior to the onset of each experiment, each freshly eclosed female L. testaceipes was held in a test tube provisioned with honey to generate parasitoids of known age and mating status (Lahiri et al. 2017). Two males per female parasitoid were present in each test tube for 4 h to ensure mating as Persad and Hoy (2003) observed L. testaceipes mating within 1 h of confinement. At the end of 24 h, the parasitoids in the petri dishes were removed using an aspirator and the leaf disk, along with aphids, was carefully transferred onto the organza fabric of the exclusion cage and the cage was attached to the abaxial side of a new leaf that was attached to a potted sorghum plant growing in the growth chamber. The aphids were held in the cage for the development of aphid mummies, which are tan in color. After 6 d, aphid mummies were collected with a fine brush and stored in a test tube held in the growth chamber until emergence. The percentages of aphid mummies and emerging parasitoids were scored relative to available M. sacchari hosts. A total of 10 replicates of this study were initiated on 23 March 2017 and 10 additional replicates were initiated on 12 April 2017. A total of 5 replicates had to be discarded due to parasitoids or predators gaining access to the exclusion cages during the experiment and confounding the results, leaving a total of 15 replicates for analyses.

Field survey. Field survey of M. sacchari and L. testaceipes occurrence in sentinel grain sorghum plots were performed in 2017 and 2018. In 2017, three irrigated plots (45.7 m × 16 rows with a 91.4-cm row spacing) were planted with a susceptible grain sorghum hybrid (SS-800A), located at the University of Georgia Lang-Rigdon Farm (N 31.5170, W –83.5483). Each of these three plots were planted on three different planting dates. Planting dates included 10 May 2017 (designated as May_Sorghum), 2 June 2017 (June_Sorghum), and 7 July 2017 (July_Sorghum). Seeds were not treated with any fungicides or insecticides and the fields remained unsprayed throughout the season. In 2018, a single planting date (6 June 2018) with a susceptible grain sorghum hybrid (DKS53-53, Bayer Corp, Whippany, NJ) was planted at the same location with a similar plot size. Each plot (45.7 m × 16 rows with a 91.4-cm row spacing) was observed for arrival of both M. sacchari and L. testaceipes. To facilitate this field sampling, 10 randomly selected sorghum plants per plot were sampled weekly where the ratio of tan mummies to healthy M. sacchari present on the bottom green leaf of each plant was recorded. During weekly sampling, the growth stage of the sorghum plants was assessed following the methods of Vanderlip and Reeves (1970).

Data analysis. Mean and standard error were calculated for responses, including M. sacchari progeny and L. testaceipes percent parasitization, male progeny proportion, and percent mummy hatch. Percent parasitization of M. sacchari was calculated as the number of mummies divided by the sum of apterous adults plus third and fourth instars. Alates were excluded from this calculation because they rarely formed mummies and, if they did, they very rarely produced adult parasitoids. Male progeny proportion of L. testaceipes was calculated as male progeny divided by total progeny. Lysiphlebus testaceipes percent emergence was calculated as the total L. testaceipes adults that emerged divided by total number of mummies formed in each replicate.

In the field study, pest and parasitoid abundance and percent parasitism were calculated for weekly observations in sentinel plots. Additionally, the presence and abundance of L. testaceipes and aphid mummies were recorded. Summary data were plotted to show temporal occurrence of the species in the field.

Results

Greenhouse study. Sugarcane aphids produced many offspring in the greenhouse. After 96 h, a mean of 93.3 ± 10.3 (n = 15) sugarcane aphid offspring (all life stages) developed in each exclusion cage. Fewer than 10 individuals per replicate completely molted to the adult life stage, as the majority of the individuals developed to 2nd and 3rd instars (Fig. 3). Lysiphlebus testaceipes parasitized M. sacchari at an average rate of 15.2 ± 3.3%. Adult parasitoids emerged from the majority of the mummies (88.7 ± 4.4%), with 38.6 ± 8.4% of the progeny being male.

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Field observation in sentinel plots. Although L. testaceipes were observed in the field, the percent parasitism was extremely low in both years. In 2017, M. sacchari alates initially appeared in the May_Sorghum plot on 30 May 2017, but L. testaceipes were never observed in that plot. In the June_Sorghum plot, M. sacchari alates were first observed on 13 June, while L. testaceipes were first observed 7 weeks later on 8 August (Fig. 4). The overall percent parasitism by L. testaceipes in this plot was found to be 4.5 × 10^–4 ± 1.5 × 10^–4%. In the July_ Sorghum plots, M. sacchari alates were first observed on 15 August, and L. testaceipes were first observed on 22 August. The overall percent parasitism by L. testaceipes in this plot was found to be 7.4 × 10^–4 ± 4.4 × 10^–4%. Due to overwhelming M. sacchari pressure, none of the plants in the May_Sorghum plot progressed beyond the boot stage. However, plants reached maturity and produced grain in the two later planting dates.

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In 2018, M. sacchari alates were first detected on 27 June, but L. testaceipes was not observed until 5 weeks later on 25 July. The overall mummy percentage in the June planted sentinel plot was recorded at 4.4 × 10^–5 ± 1.7 × 10^–5%.

Although it was not the target pest species in this study, Rhopalosiphum maidis (Fitch) (Hemiptera: Aphididae) was observed infesting the sorghum panicles and feeding on the flag leaf during the time that L. testaceipes started appearing in August 2017. Within 1 week of R. maidis infestation, L. testaceipes parasitized nearly 100% of R. maidis. One week after first observation, nearly all R. maidis were mummified, leading to a R. maidis population collapse (S.L. pers. obs.). To confirm the natural enemy that parasitized R. maidis, the mummies were collected from the field and held in a growth chamber until the adults emerged. Those adults were then keyed to L. testaceipes.

Discussion

The aphid parasitoid L. testaceipes was found to parasitize M. sacchari under both greenhouse and field conditions in Georgia. However, the rate of parasitization in the greenhouse assay was much greater than the very limited parasitism observed in the field. The fecundity of M. sacchari, under the present experimental conditions, was high (approximately 90 aphids in 4 d). The highest daily birth average of M. sacchari on artificial diet (30% sucrose + 0.5% yeast) during the first 4 d was reported as 28.9 ± 10.3 (Toledo-Hernandez et al. 2018). Access to a live host caged under optimal environmental conditions may explain the M. sacchari population explosion in the present study. Lysiphlebus testaceipes is capable of rapid host range expansion, as evident by its role as an exotic aphid parasitoid along the West Mediterranean and Atlantic coastal areas of southern France, Spain, and Portugal (Starý et al. 2004). Hosts of L. testaceipes in North America include R. maidis (Vinson and Scarborough 1991); bird-cherry oat aphid, Rhopalosiphum padi (L.); Russian wheat aphid, Diuraphis noxia (Kurdjumov) (Pike et al. 1997); brown citrus aphid, Toxoptera citricida Kirkaldy (Persad and Hoy 2003); and greenbug, Schizaphis graminum (Rondani) (Starks and Burton 1977). Therefore, the current host range of L. testaceipes in the United States already includes exotic invasive aphid species to which it has adapted to parasitize. However, rates of parasitism by L. testaceipes can vary widely by host aphid. For example, L. testaceipes parasitized 67% and 76% of S. graminum and 46% and 56% parasitism of Aphis gossypii Glover, under choice and no-choice test conditions in the laboratory (Rodrigues and Bueno 2001). Lysiphlebus testaceipes parasitized 86% of T. citricida in the field, despite the presence of red imported fire ants, Solenopsis invicta Buren (Hymenoptera: Formicidae). Interestingly, only 26% parasitization was observed in a laboratory study in the presence of Solenopsis invicta, due to preferential predation of parasitized aphids (Persad and Hoy 2003). Vinson and Scarborough (1991) documented that Solenopsis invicta selectively destroyed R. maidis mummies that were parasitized by L. testaceipes. After comparing performance of L. testaceipes on three aphid species, Aphis craccivora Koch, Aphis fabae Scopoli, and Aphis gossypii, at temperatures of 17, 22, and 27°C, A. fabae was found to be the ideal host at 20–22°C (Satar et al. 2018).

In this field study, L. testaceipes parasitized virtually every R. maidis present as soon as they appeared. Temporally, both R. maidis and L. testaceipes appeared in early August and L. testaceipes seemed to be spatially and temporally synchronized with R. maidis. Conversely, M. sacchari and L. testaceipes were also present during late July and throughout August, but the results were very different. One difference between the two aphids is that R. maidis and M. sacchari spatially occupy different zones of the plant. Feeding on the upper foliage, including the whorl and flag leaf, is typical of R. maidis infestation in corn and sorghum (Foott 1977, Li et al. 2008, Park and Obrycki 2004). Conversely, M. sacchari infestations start at the bottom of the plant and gradually move upward. A second difference is that M. sacchari infestations started much earlier in the year than R. maidis. Jackson et al. (1970) documented that L. testaceipes activity in the field in Oklahoma was not evident until August. In Georgia, the authors never observed L. testaceipes in the field until late July to August.

Although L. testaceipes does not seem to be a strong candidate for biological control of M. sacchari in Georgia at this time, there are other candidate species that need to be evaluated. Preliminary work in Georgia in 2017–2018 suggests that Aphelinus nigritus Howard is also present at multiple locations and could be an important parasitoid of M. sacchari (G.D.B. unpubl.). Additionally, there may be other native parasitoids that can adapt to M. sacchari over time.

Melanaphis sacchari is a highly fecund species, even compared with other aphids (Bayoumy et al. 2016). Future studies are required to elucidate new natural enemies or better adapted native species to serve as effective biological control agents in integrated pest management programs.