Pheropsophus aequinoctialis (Linné),
Frank, Howard, Erwin, Terry & Hemenway, Robert, 2009, Economically Beneficial Ground Beetles. The specialized predators Pheropsophus aequinoctialis (L.) and Stenaptinus jessoensis (Morawitz): Their laboratory behavior and descriptions of immature stages (Coleoptera: Carabidae: Brachininae), ZooKeys 14 (14), pp. 1-36: 18-34
treatment provided by
|Pheropsophus aequinoctialis (Linné)|
EGG ( Fig. 15View Figure 15). White. Rectangulate with moderately rounded apices. Surface polygonic, with numerous very close-spaced large perforations; micropore not obvious.
INSTAR I. Form. ( Fig. 16View Figure 16) Campodeiform planidium; head relatively small compared to prothorax, eyes absent. Frontale with three simple-tooth egg-bursters near base of head on frontale. Body setiferous dorsally, less so than in Stenaptinus (see above). Segment X (PY): sternite ( Figs. 16View Figure 16, 24View Figures 23-24. 23) medially with two widely spaced non-serrated recurved teeth and with seta PY7 normal. Urogomphi (cf. Fig. 16View Figure 16) absent.
Coloration. Mostly white color with creamy-colored head capsule and slightly rufescent mandibles darkened toward the tips.
Chaetotaxy. Head. ( Figs. 16View Figure 16, 17, 18View Figures 17-18. 17) Frontale ( Fig. 17View Figures 17-18. 17) with 9 “ancestral” setae (FR1 – FR9, FR10 and 11 missing), and one auxiliary seta each side, and 2 pores (FRd – FRe, a, c, and f missing) left side, right side devoid of pores in specimen illustrated. Parietale ( Figs. 16View Figure 16, 17View Figures 17-18. 17) with 18 setae (PA1 – PA18) and 8 pores (PAa – PAl; pores d, f, g, h absent) each side. Antenna ( Figs. 16View Figure 16, 17View Figures 17-18. 17): antennomere 1 with 5 “ancestral” pores (ANa – ANe) and one auxillary pore (unlabeled); antennomere 2 absent or fused with 3; antennomere 3 with 3 “ancestral” setae (AN1 – AN3), one auxillary seta, and 1 pore (ANf), plus a dome-shaped hyaline sensillum; antennomere 4 with 4 setae (AN4 – AN7) and 1 auxillary seta, no pores, and 2 small apical sensilla. Mandible ( Fig. 17View Figures 17-18. 17) falciform without setae and pores. Labium ( Fig. 18View Figures 17-18. 17): prementum with 1 seta (LA3) and 1 pore (LAa) each side; palpomere 1 with 1 seta and 3 pores, none of which correspond to the “ancestral” schema; palpomere 2 with 1 apical sensillum. Maxilla ( Fig. 18View Figures 17-18. 17): cardo without setae; stipes with 5 “ancestoral” setae ( MX 1 – MX 5), and 2 pores
(MXa – MXb), and no variable setae (gMX) on dorsal side; lacinia ( Fig. 18View Figures 17-18. 17) with 1 seta ( MX 10); galeomere 1 with 1 seta ( MX 7) and no pores; galeomere 2 with 2 minute dorsal setae, no pores; maxillary palpomeres without visible sensatory features.
Thorax. Prothorax: Notum ( Figs. 16View Figure 16, 19View Figures 19-22. 19) with 1 identifiable major “ancestral” seta (PR 9) and numerous auxiliary setae (not labeled), PR 1 absent, and no pores. Epimeron ( Fig. 20View Figures 19-22. 19) with 1 seta ( EP1), and no pores. Episternum and trochantin not defined. Prosternite ( Fig. 20View Figures 19-22. 19) with 1 seta “ancestral” (Pt1) and one auxiliary seta; gPS absent.
Mesothorax and metathorax: Notum ( Figs. 16View Figure 16, 19, 20View Figures 19-22. 19) with 1 identifiable major “ancestral” seta (PR 9) and numerous auxiliary setae (not labeled), PR 1 absent, and no pores. Mesepisternum ( Fig. 20View Figures 19-22. 19) with 2 setae (ES1, ES2) and no pores. Trochantin and epimeron not defined. Mesoprosterite ( Fig. 20View Figures 19-22. 19) with 3 setae (Pt1, Pt2, Pt3) each side; metaprosternite with 3 setae (Pt1, Pt2, Pt3). Metepisternum with 3 setae (ES1, ES2, ES4).
Abdomen. Figs. 16View Figure 16, 21View Figures 19-22. 19 -24View Figures 19-22. 19View Figures 23-24. 23. Tergite I ( Figs. 16View Figure 16, 21View Figures 19-22. 19) with possibly one “ancestral” seta (TE2) and numerous auxiliary setae (not labeled), and no pores. Tergites II – VIII as in
Tergite 1. Tergite IX, X and urogomphi ( Figs. 16View Figure 16, 23View Figures 23-24. 23), IX with 4 setae (UR8 – UR11) and no pores. Epipleurite IX ( Fig. 16View Figure 16) with 2 setae (EP1 – EP2) and no pores. Hypopleurite VII ( Fig. 16View Figure 16) with 2 setae (HY1 – HY2) and no pores. Segment VII sternite ( Fig. 24View Figures 23-24. 23) with 5 setae (ST1 – ST5) each side and no pores. Segment IX sternite ( Fig. 24View Figures 23-24. 23) with 3 setae (ST1 – ST3) each side and no pores. Segment X (PY) sternite ( Figs. 16View Figure 16, 24View Figures 23-24. 23) with 1 seta (ST1) each side, no pores. Medially with two wide-spaced nonserrated and recurved teeth ( Figs. 16View Figure 16, 24View Figures 23-24. 23).
Legs. ( Fig. 25View Figure 25) All legs stout, similar in proportions and setation; anterior leg slightly shorter than middle and posterior ones. Coxa with 7 setae (ancestral CO10 – CO17, with CO1-9 absent, and no pores. Trochanter with 5 setae (TR2 – TR5, and TR8) and one pore. Femur with 4 setae (FE2 – FE5) and no pores FEa and FEb. Tibia with 7 setae (TI1 – TI7) and no pores. Tarsus with 1 constant seta (TA1) and no pores. Claws simple, with no setae or tooth, symmetrical in shape and size.
INSTAR II. Form. (generally as in Fig. 35View Figure 35) Hypermetamorphic stage 2 instar.
Coloration. White; head capsule creamy-white with mouthparts slightly infuscated in part; mandibles piceous at tips.
Chaetotaxy. Head. ( Figs. 26-27View Figures 26-27. 26) Frontale ( Fig. 26View Figures 26-27. 26) with 7 “ancestral” setae (FR1 – FR7), and no pores. Parietale ( Figs. 26, 27View Figures 26-27. 26) with 12 setae (PA3, PA5– PA7, PA9, PA11 – PA13, PA15 and PA17) and no pores. Antenna ( Figs. 26View Figures 26-27. 26): antennomere 1 with one “ancestral” seta (AN1) and no pores. Dome-shaped hyaline sensillum absent. Mandible ( Fig. 26View Figures 26-27. 26) falciform without setae and pores. Labium ( Fig. 27View Figures 26-27. 26) without setae or pores. Maxilla ( Fig. 27View Figures 26-27. 26): cardo without setae; stipes with 3 “ancestral” setae ( MX 3 – MX 5), and no pores, nor variable setae (gMX) on dorsal side; lacinia ( Fig. 27View Figures 26-27. 26) without setae; galeomere without setae; palpomere 1 and 2 without setae, palpomere 3 with 2 minute apical setae, no pores.
Thorax. Prothorax: Figs. 28-29View Figure 28-33. 28. Notum ( Fig. 28View Figure 28-33. 28) with 12 major “ancestral” setae (PR 2 – PR 4, PR 6 – PR 14) and numerous auxiliary setae (not labeled), and no pores on each side. Epimeron, episternum, and trochantin not defined. Prosternite ( Fig. 29View Figure 28-33. 28) with a ring of auxiliary setae, gPS absent.
Mesothorax and metathorax: Figs. 28-29View Figure 28-33. 28. Mesonotum ( Fig. 28View Figure 28-33. 28) with 9 “ancestral” setae (ME1 – ME2, ME8 – ME14), and no pores on each side. Mesepisternum ( Fig. 28View Figure 28-33. 28) with 1 seta (PL1) and no pores. Trochantin and epimeron not defined. Sternum
( Fig. 29View Figure 28-33. 28) with 9 setae (unlabeled) in a median rosette. Metanotum ( Fig. 28View Figure 28-33. 28) with 5 “ancestral” setae (MT2, MT7 – MT9, MT12), and no pores on each side. Metepisternum ( Fig. 29View Figure 28-33. 28) with 5 setae (unlabeled) and no pores. Trochantin and epimeron not defined. Sternum ( Fig. 29View Figure 28-33. 28) with 14 setae (unlabeled) in a median rosette.
Abdomen. Figs. 30-34View Figure 28-33. 28View Figure 34. Tergite I ( Fig. 30View Figure 28-33. 28) with 11 “ancestral” setae (TE1 – TE11) and 5 auxiliary setae (not labeled), and no pores each side.Tergites II – VIII as in Tergite 1 with numerous auxiliary setae. Sternum with numerous setiferous rosettes. Tergite IX, X and urogomphi ( Fig. 32View Figure 28-33. 28), all with numerous setae in raised rosettes or on raised lobes, and no
pores. Epipleurites ( Fig. 33View Figure 28-33. 28) with numerous setae and no pores. Hypopleurite not defined. Segment VII sternite ( Fig. 33View Figure 28-33. 28) with numerous setae in rosettes and no pores. Segments VIII and IX with numerous setae (not in rosettes) each side and no pores. Segment X (PY) sternite ( Fig. 33View Figure 28-33. 28, 34) with numerous auxiliary setae in apical 2/3 rd, with no pores.
Legs. As in Fig. 35View Figure 35; reduced size and setation compared to instar I.
INSTAR III. Form. ( Fig. 35View Figure 35) Hypermetamorphic stage 3 instar.
Coloration. White; head capsule creamy-white with mouth parts slightly infuscated in part; mandibles piceous at tips.
Chaetotaxy. Head. ( Figs. 35View Figure 35 -37View Figure 35View Figures 36-37. 36) Frontale ( Fig. 36View Figures 36-37. 36) with 9 “ancestral” setae (FR1 – FR5, FR7 and FR9) and and no pores each side. Parietale ( Figs. 36, 37View Figures 36-37. 36) with 5 “ancestral” setae (PA4 – PA8) and 3 pores (PAa – Pac) each side. Antenna ( Figs. 36, 37View Figures 36-37. 36): antennomere 3 with 1 seta (unlabeled); antennomere 4 with 3 in an apical ring; the dome-shaped hyaline sensillum absent. Mandible ( Fig. 36View Figures 36-37. 36) falciform without setae and pores. Labium ( Fig. 37View Figures 36-37. 36): prementum with 1 seta (LA5) and no pores each side; labial palpomeres reduced and without vestiture. Maxilla ( Fig. 37View Figures 36-37. 36): cardo with 1ventral seta (ca1); stipes with 5 “ancestral” setae ( MX 1 – MX 5), and no pores and no variable setae (gMX) on dorsal side; lacinia and galeomere absent; maxillary palpomeres reduced with palpomere 1 unisetose (pa10) and without other visible sensory features.
Thorax. Prothorax: Notum ( Figs. 38 – 40View Figure 38View Figures 39-44. 39) with numerous long setae on front margin and numerous shorter setae posteriorly (not labeled), and no pores on each side. Parietal with 6 “ancestral” setae (PR 2 – PR 4, PR 6, PR 8, PR 14) and numerous auxiliary setae (not labeled), and no pores on each side. Epimeron ( Fig. 38View Figure 38) with 5 setae (EP3 – EP4, EP6, EP10 – EP11), and no pores on each side. Episternum and trochantin not defined. Prosternite ( Fig. 40View Figures 39-44. 39) with numerous medial auxiliary setae some in a rosette, gPS absent.
Mesothorax and metathorax: Notum ( Figs. 38View Figure 38, 39View Figures 39-44. 39) with 2 long setae medially and 3 shorter setae nearby (not labeled), and no pores on each side. Episternum ( Fig. 38View Figure 38) with numerous stout setae and no pores. Epimeron and trochantin not defined. Sternum with rosette of setae medially ( Fig. 39View Figures 39-44. 39).
Abdomen. Figs. 41 – 44View Figures 39-44. 39. Tergite I ( Fig. 41View Figures 39-44. 39) with 2 stout setae medially and numerous shorter auxiliary setae in patches (not labeled), and no pores each side. Tergites
II – VIII as in Tergite 1. Tergite IX with 2 stout setae each side and numerous shorter setae nearby. Tergite X (PY) with numerous short apical setae and no pores. Epipleurite IX ( Fig. 44View Figures 39-44. 39) with numerous long and stout setae on raised knob and no pores. Hypopleurite not defined. Segment VII and VIII sternites ( Fig. 44View Figures 39-44. 39) with numerous setae on raised knobs in rosettes each side and no pores. Segment IX sternite ( Fig. 44View Figures 39-44. 39) with subapical band of short setae each side and no pores. Segment X (PY) sternite ( Fig. 35View Figure 35, 44View Figures 39-44. 39) with numerous scattered short setae each side, no pores.
Legs. As in Fig. 35View Figure 35; reduced size and setation compared to instar I.
PUPA. Form. ( Fig. 45View Figure 45) Typical of carabid species. In addition, pygidium with fine short setae and dorsal surface with an array of small tubercules.
Notes on advancing an understanding of phylogenetic relationships. Much works still needs to be done in solving to infer the relationships between the Brachininae tribes Crepidogastrini and Brachinini and the subtribes Brachinina , Pheropsophina, and Masticina ( Erwin 1970). Here we have added larval traits that will, in part, add information toward a more robust phylogenetic analysis in the future. Immature stages of Crepidogastrini and Masticina are, as yet unknown, and we do not even know whether they are ectoparasitoids, or specialized predators. Likely, they are one or the other, but on what taxa? Larvae of Brachinus develop in 5 instars; they also have 6 eyespots (as in other carabids), whereas Pheropsophina larvae have 3 instars and at most a single eye-spot, usually none. First instar Brachinus have no egg burster and chew their way out of the egg ( Erwin 1967); Stenaptinus larvae possess a single-toothed egg burster; those of Pheropsopus have a triple-toothed egg burster. Pheropsophina larvae have pygidial hooks that aid them in attacking mole cricket egg clutches (Habu and Sadanga 1969), whereas Brachinus larvae do not. Larvae of Stenaptinus and Brachinus possess urogomphi whereas those of Pheropsophus do not.
Conclusion and discussion
Adult P. aequinoctialis and many S. jessoensis burrowed in sand-filled containers in the laboratory. Many S. jessoensis ̴ 91.4%) but few P. aequinoctialis ̴ 16.7%) were active on the surface in daylight. Th is supports field observations that P. aequinoctialis adults are active nocturnally. Very few P. aequinoctialis adults (1.2%) found brown paper towel on the sand surface to be as adequate a refuge as their burrows; perhaps more solid objects (as a result of photoperception or thigmoperception) would have been more acceptable as refuges. In contrast, more S. jessoensis adults (78.7%) sheltered under paper towel in daylight than sheltered in burrows, but most did not shelter at all. Th ere is a clear contrast between the mainly diurnal behavior of S. jessoensis and the mainly nocturnal behavior of P. aequinoctalis . Timing of daily activity will have an effect on ability to find food.
All diets presented to these adults were consumed, but we did not observe cannibalism by adults. Larvae of T. ni alone sustained adult S. jessoensis of unknown age for an average 12 months. Earlier authors showed that a broad diet of animal food is acceptable to them, and neither we nor previous authors tested acceptability of plant food alone. A diet of mealworm pupae, oatmeal and raisins was avidly fed upon by adult P. aequinoctialis , and sustained them well, so they will feed on plant food, supporting an observation of feeding upon palm fruits in nature ( Reichardt 1971). We did not attempt to produce an optimal diet for adults of either species. Adults of the two species produced many eggs on the diets provided. Females of both species oviposited abundantly on crumpled, moist, brown paper towel under highly artificial conditions. Chemicals produced by eggs or adults of mole crickets are not necessary to stimulate abundant oviposition. However, Weed and Frank (2005) found that more eggs were laid in tunnels excavated by mole crickets than in artificial tunnels, suggesting that perhaps allomones produced by adult mole crickets are detected by female P. aequinoctialis and influence placement of eggs.
Adult P. aequinoctalis oviposited in all months of the year; and adult S. jessoensis oviposited in most months of the year in the laboratory. Seasonality of oviposition in the field is mutable under laboratory conditions. We suspect that neonate larvae of both species suffer high mortality because they fail to detect suitable prey and thus die. Although P. aequinoctialis and S. jessoensis are highly fecund, Th iele (1977) gave examples of high fecundity among other carabids without such a specialized life cycle. Fertility of laboratory-produced P. aequinoctialis eggs varied for unknown reasons, at some times being very low. Th e evolutionary consideration is: Why are so many infertile eggs produced? Presence of two species of Wolbachia (Bacteria: Rickettsiae) in our 1992 P. aequinoctialis stock from Bolivia has been demonstrated. A suggested heat-treatment to eliminate the Wolbachia resulted in mortality of some adults, temporarily reduced oviposition, and failed to eliminate production of infertile eggs. Th e heat treatment may, of course, have killed some essential flora in the digestive system. Incubation of fertile eggs of S. jessoensis took 11.4 days and of P. aequinoctialis 13.5 days on average.
Most neonate larvae of S. jessoensis and P. aequinoctialis died when presented with mole cricket eggs in Petri dishes. Th ey wandered for days until they died, almost continually in motion. Neonate larvae were presented with alternative diets including A. domesticus and Gryllus sp. eggs, eggs and pieces of larvae of T. ni, intact pupae of T. molitor , and pieces of cucumber. All neonate larvae of both species died when presented with any diet other than mole cricket eggs although imbibition of fluid was observed from pieces of cucumber. However, a few began to feed on mole cricket eggs. Th ose eggs were of Scapteriscus abbreviatus , Sc. borellii , Sc. vicinus , and N. hexadactyla ; however, replication was inadequate to determine any differences in survival success between these mole cricket eggs diets. At least it can be stated that S. jessoensis can survive on mole cricket eggs other than those of Gryllotalpa , in contrast to unsupported claims by Habu and Sadanaga (1965, 1969). Once neonate larvae began to feed, their survival to the adult stage on the same diet was highly probable. Perhaps larvae will not begin to feed until they encounter enough eggs to complete their development ( Habu and Sadanaga 1965, 1969), but we have no data to support this claim. Faced with the impasse that neonate larvae would seldom develop on a diet of mole cricket eggs in a Petri dish, even when that dish was enclosed totally with aluminum foil to exclude light, we adopted a variant of the rearing method proposed for S. jessoensis by Habu and Sadanaga (1969). Using this method, an artificial mole cricket egg chamber is made in sand in a plastic vial, stocked with 30 mole cricket eggs, covered with sand, and a neonate larva is dropped onto the sand surface. The larva burrows down to enter the chamber and begins feeding on eggs. Th is resulted in high survival of larvae and pupae to the adult stage, and became our standard rearing method. Feeding and development seldom occurred in the more artificial conditions of a small Petri dish with mole cricket eggs piled onto a disc of paper towel, even in the dark, but we ran many feeding trials under those circumstances. Unfortunately, the method of an artificial egg chamber excluded frequent observation. Much later, we found by accident that the artificial egg chamber did not need covering with sand to exclude light. Th en, we conducted more feeding trials and confirmed that Acheta domesticus eggs, Tenebrio molitor pupae, and pieces of cucumber are not acceptable diets.Adult P. aequinoctialis are scavengers and generalist predators. Larvae, however, so far as determined, are specialist predators on mole cricket eggs. Th ey can develop under laboratory conditions on a diet containing only eggs of Scapteriscus abbreviatus , Sc. borellii , Sc. vicinus , or Neocurtilla hexadactyla but none survived using any other diet tried. Proof of restriction of the larval diet still is inadequate.
Each larva of the carabid genus Brachinus (Neobrachinus) feeds on only one water beetle pupa and is an ectoparasitoid ( Erwin 1967, 1979) replacing its host in a small mud chamber constructed by the water beetle larva. In Europe, Brachinus s. str. larvae feed on the pupal stage of the carabid genus Amara ( Saska and Honek 2004) . However, each larva of P. aequinoctialis and S. jessoensis requires tens of mole cricket eggs as food to complete its development. Such behavior is more aptly termed predation ( Van Driesche and Bellows 1996, p. 21 citing many earlier authors), so we consider Pheropsophus and Stenaptinus larvae to be specialist predators. At no time in our laboratory cultures did more than one P. aequinoctialis or S. jessoensis larva survive long on a single cache of eggs, so we believe they practice fratricide as has been noted in Brachinus ( Juliano 1984) . Erwin (1967) noted that fratricide did not occur in Brachinus pallidus , rather the first larva that began feeding became the “owner” of the pupa and the other larvae departed in search of another pupa. Th ese larvae, when offered a fresh pupa, developed to the adult stage.
Implications for biological control
The studies reported above were initiated because of a suspicion by T.L. Erwin that Pheropsophus spp. larvae might, as had been reported for Stenaptinus , develop only on a diet of mole cricket eggs. It was eggs of invasive species of the South American mole cricket genus Scapteriscus in the southern USA that were the target of our studies. These studies were initiated by J.H. Frank in the name of the University of Florida/ Institute of Food and Agricultural Sciences’ Mole Cricket Research Program ( Walker 1985; Frank and Walker 2006). Early mention of the studies was made by Hudson et al. (1987). Prey specificity of these beetle larvae was important because the native North America mole cricket Neocurtilla hexadactyla was not a target. Might the South American Pheropsophus be adapted to South American Scapteriscus mole crickets but the Old World Stenaptinus be adapted to the largely Old World genus Gryllotalpa ?
Habu and Sadanaga (1965) stated that S. jessoensis larvae feed only on eggs of G. africana Palisot de Beauvois. Th ey provided no evidence that they had experiment- ed with other diets. However, G. africana does not occur in Asia, and the species of mole cricket encountered by those authors may have been G. orientalis Burmeister ( Townsend 1983). We had no access to eggs of Gryllotalpa . By 1987, we had found that larvae of S. jessoensis and P. aequinoctialis would develop on a diet of eggs of Neocurtilla ( Gryllotalpinae ) or Scapteriscus ( Scapteriscinae ) mole crickets but, because of difficulties in getting neonate larvae to initiate feeding, we had conducted scores of failed trials with these and other diets.
Little that we had studied pointed to need for chemoperception. Adults laid eggs abundantly on paper towels. Neonate larvae may have used chemoperception to detect that mole cricket eggs are food, but there was no evidence that such detection occurred except in a pit in sand.
The Mole Cricket Research Program then concentrated on other biological control agents, which were successful, until its funding was ‘unearmarked’ in 1991 ( Frank and Parkman 1999). At this devastating event, to save expenses and because S. jessoensis clearly could not be a specialist of Scapteriscus mole crickets, cultures of both species were terminated.
A culture of P. aequinoctialis was reinitiated with stock from Bolivia in 1992. One reason was that an additional biological control agent that could be used in the vicinity of water bodies, on their banks in particular, could be beneficial in integrated pest management because application of chemical pesticides is prohibited from use in such habitats. A second reason is because of egg-guarding behavior by female Neocurtilla hexadactyla . Th ese excavate two side-by-side underground cells, one of which receives the eggs, the other serves as a resting site for the female, from which she emerges fom time to time to tend the eggs (J.H. Frank and R.C. Hemenway, obs.). In contrast, each Scapteriscus spp. female excavates only one cell and then, after oviposition of a clutch of eggs, leaves and blocks the entrance to the cell (J.H. Frank and R.C. Hemenway, obs.). It might therefore be possible for female N. hexadactyla to detect and kill intruding bombardier beetle larvae. If this can be demonstrated in the laboratory, it might justify release of P. aequinoctialis in Florida.
Research will not be complete until the subject of egg-guarding by N. hexadactyla females is adequately investigated. A major problem is that we have not devised a robust method for culturing N. hexadactyla . Survival of adults and nymphs was poor, perhaps because the diet we used was inadequate. We observed that females move their eggs when they are disturbed, which we believe to be a previously unreported facet of their presocial behavior.
Finally, some objection might be made to the release of a beetle whose adults are scavengers and generalist predators, even though this habit is shared with adults of many other insects, including adults of the ̴ 18 native species of Brachinus bombardier beetles in Florida. Still, population sizes must be limited by availability of mole cricket eggs, and we now have some idea of the quantity of food (≤ 2.3 large T. ni larvae per day) consumed by pairs of adult beetles.
Most of the work described here was supported by funds earmarked by the Florida Legislature for mole cricket research at the University of Florida in 1978-1991 (the UF/ IFAS Mole Cricket Research Program). Earmarking was withdrawn (but the program was not de-authorized) after three biological control agents had been released and established in Florida, but before research on them, let alone P. aequinoctialis , which has not been released, was complete. Thomas J. Walker (Entomology and Nematology Dept., Univ Florida), supplied Gryllus sp. eggs. Fred Adams, USDA-CMAVE laboratory (Gainesville, FL) supplied T. ni eggs and larvae. A. Jeyaprakash (Entomology & Nematology Dept., Univ. Florida) investigated and confirmed the presence of Wolbachia in P. aequinoctialis adults. Harold A. Denmark (Florida Dept. of Agriculture and Consumer Services, Division of Plant Industry, now retired) identified to generic level a paramegistid mite from P. aequinoctialis . Aquiles Silveira-Guido (Montevideo) was contracted to supply specimens and information. Karl Zinner (São Paulo) exchanged specimens of P. aequinoctialis for specimens of S. jessoensis . Herein is the background to a species of Rhabditis described as new by Smart and Nguyen (1994). Young Sohn provided the illustrations of the immature stages and Karolyn Darrow assembled the illustration plates; both individuals are members of the Department of Entomology at the Smithsonian Institution. Paul E. Skelley (Florida State Collection of Arthropods) provided the scanning electron micrographs of the eggs. We thank Robert McSorley and Frank Slansky, Jr. (Entomology & Nematology Dept., Univ Florida) for manuscript reviews.
No known copyright restrictions apply. See Agosti, D., Egloff, W., 2009. Taxonomic information exchange and copyright: the Plazi approach. BMC Research Notes 2009, 2:53 for further explanation.