Creophilus oculatus, (FABRICIUS)

Clarke, Dave J., 2011, Testing the phylogenetic utility of morphological character systems, with a revision of Creophilus Leach (Coleoptera: Staphylinidae), Zoological Journal of the Linnean Society (Zool. J. Linn. Soc.) 163 (3), pp. 723-812 : 789-794

10. CREOPHILUS OCULATUS (FABRICIUS)

( FIGS 1C, 3J, 4G , 5D , 7C , 9J, K, 32 , 33A, D, G, J, M, N, Q , 34 )

Staphylinus oculatus Fabricius, 1775: 265 . Type locality: ‘ nova Hollandia et Zelandia’ (here restricted to New Zealand, see below); Goeze, 1777: 724; Fabricius, 1781: 335 (not seen); Fabricius, 1787: 220; Gmelin, 1790: 2026 (not seen); Fabricius, 1793: 521; Olivier, 1795:(42): 11, pl. 2, fig. 19; Fabricius, 1801: 592; Turton, 1802: 510; Gravenhorst, 1806: 126; Olivier, 1808: pl.2, fig. 19 (not seen); Boisduval, 1835: 54, pl. 9, fig. 1; Erichson, 1839: 352; Dieffenbach, 1843: 273; White, 1846: 6; Erichson, 1859 (Fogg translation, not seen): 312; Murray, 1870: 57, 89; Wakefield, 1873: 299; Hutton, 1874: 160; Broun, 1876: 265, 271; von Kiesenwetter, 1877: 161; von Kiesenwetter & Kirsch, 1877: 154 (not seen); Broun, 1880: 107; Broun, 1882: 9; Hudson, 1892: 25, pl. 1, fig.5; Hutton, 1898: 155; Hutton, 1904: 183; Hudson, 1923: 362; 1934: 45, 180; Zimsen, 1964: 230; Radford, 1981: 185.

Creophilus oculatus ; Nordmann, 1837: 23; Motschulsky, 1858a: 49; Gemminger & Harold, 1868: 575; Fauvel, 1875: 56; Reitter, 1880: 166; Fauvel, 1885: 312; Olliff, 1887: 493; Alfken, 1903: 604 (not seen); Walker, 1904: 72, 121; Broun, 1909: 145; Broun, 1910: 292; Bernhauer & Schubert, 1914: 399; Brookes, 1925: 286; Lea, 1925: 229; Steel, 1949: 59, figs 3 and 6; Gourlay, 1950: 185, 202, figs 1 and 2 (pls. 21, 22); Brookes, 1951: 13, 29; Manson, 1960: 59; Eyles, 1961: 132; Coiffait, 1976: 218; Watt, 1980; Radford, 1981: 186; Andrews, 1986: 40; Andrews & Gibbs, 1989: 107, 113; Kuschel, 1990: 17, 26, 45; Klimaszewski & Watt, 1997: 116, figs 4 and 5, p.124, 57; Emberson, 1998: 35; Ward et al., 1999: 104; Herman, 2001a: 52 (senior primary homonym of Staphylinus oculatus O. Müller, 1776: 99 ); Herman, 2001b: 3325.

Emus oculatus ; Fauvel, 1877: 250; Fauvel, 1878a: 248; Fauvel, 1885: 311.

Creophilus erythrocephalus var. oculatus ; Lea, 1925: 229 (synonym of C. erythrocephalus ).

Type material: Staphylinus oculatus Fabricius. Lectotype (here designated). ♂, ‘Staphyl. oculatus| Fab. Entom.p. 265.4/ [red] LECTOTYPE | Staphylinus | oculatus Fabricius, 1775 | designated by| D. J. Clarke 2008’ (in BMNH). Specimen with antennomeres 4–11 and right metatarsomeres 3–5 missing, a smaller pin hole through base of pronotum, and prothorax glued to mesothorax. Wings are unfolded, intact. Paralectotypes (2). 1♂ [incorrectly as ♀ by Radford 1981], no data but with labels ‘[yellow] PARALECTOTYPE | Staphylinus | oculatus Fabricius, 1775 | designated by| D. J. Clarke 2008’ (in BMNH). Specimen missing left antenna, right antennomeres 3–11, and right metatarsus; glued to pin on ventral side. Wings are unfolded, intact. 1♀, no data but with labels ‘[yellow] PARALECTO- TYPE | Staphylinus | oculatus Fabricius, 1775 | designated by| D. J. Clarke 2008/ Creophilus oculatus ( Fabricius, 1775) det. D. J. Clarke 2010’ (in ZMUC). Specimen pinned in a unit tray with label pinned in tray: ‘ocula| tus.’ [Fabricius’s handwriting]. All three specimens are pinned through the right elytron with thick pins typical of the period ( Andrews & Gibbs, 1989).

Other material examined: 1443 specimens. See supporting information, Appendix S 1.

Diagnosis: With characters of the erythrocephalus - group; temporal regions and postgena orange-red, rest of integument and elytra uniformly pitchy-black ( Fig. 1C); apex of antennomere 11 produced into distinct ridge ( Fig. 32B ); macropterous ( Fig. 33N ); last protarsomere much longer than preceding two together ( Fig. 33J ); elytral discal series with three or four macrosetae; tergal chaetotaxic formula = 4-6-6-6- 4-6.

Description: Measurements ( N = 10♂, 10♀). Forebody length: ♂ 5.7–9.9 mm, ♀ 5.2–8.0 mm. See supporting Table S 5 for comparison of ranges of male and female ratios. Head. Head ( Figs 7C , 33A ) black between eyes, posteriorly including neck, and narrowly around mouthparts ventrally, temporal region and postgena orange-red, head thus appearing dorsally as ‘black with red spots behind eyes’; moderately to strongly trapezoidal, much wider posteriorly; HW/ HL = 1.38– 1.67; shining, without distinct microsculpture; eyes small to moderately large ( EYL / HL = 0.39–0.57), dorsolateral, lateral margins of head visible in dorsal view (not obscured by eye); HL 1/ HL 2 greater in females than males (♂ =1.21–2.00, ♀ = 2.00–2.50); antennae as in Figure 32B , antennomeres 1–6 black, 7–11 greyish-black, 11 slightly shorter than 9–10 together, with apex asymmetrically produced into distinct ridge ( Fig. 32B , arrow); mandibles as in Figure 32A , longer than head in large males, shorter than or subequal to head in females ( ML / HL ♂ = 0.90–1.37, ♀ = 0.86–1.00), right mandible with three teeth, T 3 largest, T 1 reduced in smaller specimens. Thorax and abdomen. Pronotum ( Figs 32F , 33D ) slightly transverse ( PW / PL = 1.04–1.24); moderately longer than elytral suture ( PL 1.23–1.50 ¥ ESL); with basolateral margins very slightly emarginate, hind angles indistinct; with sparse peripheral setae and short, sparse vestiture on anterolateral declivities; elytra uniformly black, disc moderately densely setose, humeral regions glabrous and shining, rugosely sculptured; wings fully developed ( Fig. 33N ), functional, clear black with distinct black spot in medial field between MP 3 and MP 4 veins; mesoventral intercoxal process broadly truncate to convex medially ( Fig. 33G ); metaventrite without posteromedial tumescence; protibiae always with outer row of long spines ( Fig. 33M ); last protarsomere much longer than preceding two together ( Fig. 33J ); abdomen shining and uniformly black; tergite VII with welldeveloped palisade fringe ( Fig. 1C). Male genitalia. Aedeagus as in Figure 32C and D ; median lobe with apex extended to short globular point ( Fig. 32I ); with basal and apical subtriangular membranous regions ventrally ( Figs 3J, 32C, I , stm); apicolateral sclerites fused to lateral edge of median orifice ( Figs 3J, 32 , as). Paramere shaft distinctly narrower near base than apically ( Fig. 32H ). Internal sac inverted as in Figure 32C and D , partially everted as in Figure 3J. Female internal genitalia. Internal female genitalia as in Figure 32G ; vaginal plate (vp) without median sclerotized strip, with paired lateral sclerites (pls), posterolateral areas membranous; vaginal fold (vf) strongly and finely rugosely sclerotized ( Fig. 32G , vf). Chaetotaxy. Elytral discal series with 3–4 macrosetae; metaventrital macroseta present; protrochanter with two macrosetae, mesotrochanteral macroseta present; tergal chaetotaxic formula = 4-6-6-6-4-6, medial macrosetae absent from tergite III, inner lateral macrosetae absent from tergite VII.

Comparison: Creophilus oculatus may be distinguished from all other Creophilus species (except rarely C. huttoni ) by the distinct red ‘spots’ behind the eyes and red postgena. It may be distinguished from C. huttoni , which has an identical colour pattern (though usually of different colours), by the pitchy black, generally shining cuticle, its fully developed and functional wings ( Fig. 33N, Q ), and the broader obtuse or blunt mesoventral intercoxal process.

Distribution ( Fig. 34 ): Many entomologists have considered C. oculatus to be introduced from Australia. Herman (2001b) recorded it from the Chatham Islands, New Zealand, Australia, New Guinea, and Society Islands ( French Polynesia). Coiffait (1976) recorded it from Tahiti (Arue) and only older literature records it from New Guinea ( Fabricius, 1781, 1801; Olivier, 1795); I have seen no specimens from either places. Gemminger & Harold (1868) recorded it from Brazil (an error according to Brookes, 1951). Brookes (1951) considered it adventive on the Auckland Islands, and it is possible that it did not naturally occur on the New Zealand subantarctic, Chatham, or Kermadec Islands previous to the whaling and sealing period of the 19 th century, when the species could easily have been transported to and between these islands via whaling ships. This is consistent with Gourlay’s (1950: 186) account and previous accounts of ship-mediated insect dispersal throughout the Tasman and South Pacific Region ( K. Wise, unpub. map in Lowe, 1973). An abundance of seabird or mammal carrion and other detritus on these islands could have facilitated establishment.

The type locality of C. oculatus was given by Fabricius as ‘ nova Hollandia et Zelandia’, the same as just one other of his beetle species (a dermestid). It has been historically recorded from (eastern) Australia by Fauvel (1877), but Olliff (1887) and Lea (1925) both doubted its presence there. In reference to one of few records with specific locality data (Queensland, Rockhampton), Steel (1949: 57) stated: ‘this is the only Australian specimen seen up to the present’. More recently, Andrews & Gibbs (1989: 107) noted that C. oculatus ‘may have been found in both New Zealand and Australia’ during Captain Cook’s H. M. S. Endeavour voyage and Kuschel (1990: 17) stated that it is ‘an Australian species, but was already in New Zealand at the time of Captain Cook’s visit in 1769’. Some ecological works also considered it adventive in New Zealand (e.g. Ward et al., 1999). Andrews & Gibbs (1989: 107) noted the likelihood that C. oculatus was actually collected aboard the Endeavour: ‘Any such dry or decayed animal materials [brought on board] are likely to have transported carrion beetles to the chart room table...’. It is therefore probably no coincidence that the carrion-associated dermestid ( Dermestes carnivorus F.) has the same type locality. Kuschel (1970: 191), Radford (1981: 158), and Andrews & Gibbs (1989: 110) detail many problems with the material collected on the 1769 Endeavour voyage (and later described by Fabricius), and most importantly they indicate where several New Zealand species had been erroneously described from Australia or South America, and vice versa. It now seems most likely that C. oculatus types were either collected on-board the H. M. S Endeavour or were victims of mixing of material, and that this species is indeed endemic to New Zealand or the New Zealand subregion. More recent literature lists C. oculatus as a New Zealand endemic ( Klimaszewski et al., 1996; Leschen et al., 2003).

Biology and ecology: Creophilus oculatus is the largest and one of the most common rove beetles in the New Zealand Region. It is widely recorded as a typical carrion associate, and Gourlay (1950) gives an account of maggot predation. When exposed to bright light, individuals more or less immediately attempt to fly, and when threatened discharge an unpleasantsmelling white substance from the abdominal defence glands (pers. observ.). Kuschel (1990) found it in open areas (pasture), and on beaches. It is also typically found in synanthropic situations and is commonly assumed to be a species primarily of open habitats (e.g. Watt, 1980; Kuschel, 1990), but evidently can be found in a wide range of habitats. Most specimens have been taken from carrion or carrion-baited pitfall traps, and many records are from forest habitat (c. 350 specimens). Other specimens were taken at light, in Malaise traps, window traps, under seaweed, and by many other collecting methods. Habitat: forest of all kinds (e.g. Agathis , Nothofagus , mixed podocarp/ broadleaf), swamp forest, scree and tussock land, penguin rookeries, pasture, coastal dunes, and beaches. Altitude: sea level to 2440 m.

Phenology: throughout the year, although few records from June–August. Larvae are known and will be described elsewhere. Pupae are unknown.

Remarks: Zimsen (1964) listed three syntypes for Staphylinus oculatus Fabricius , two in BMNH and one then in Kiel (now in ZMUC). Radford (1981: 186) listed a second label on the lectotype specimen (‘ BMNH Staphylinus oculatus ’) but this was not present in 2005 when I examined it. There is now ample evidence to correct the type locality to New Zealand, following Recommendation 76 A.2 ( ICZN, 1999). Under Article 76.1.1 ( ICZN, 1999) the type locality of C. oculatus is therefore corrected to ‘New Zealand’. Herman (2001b: 52) determined Staphylinus oculatus O. Müller, 1776: 99 (Type locality: Dania) to be a junior primary homonym of C. oculatus Fabricius (ex Staphylinus ), but regarded the Müller name as a nomen dubium and suggested a replacement name was not really needed.

Figures

Figure 4. Male copulatory pieces of selected species in the Creophilus-complex, dorsal view. In each figure the large paired dark structures are the sclerotized arms of the copulatory piece. A, C. variegatus; B, C. maxillosus maxillosus; C, C. m. villosus; D, C. galapagensis; E, C. flavipennis; F, C. incanus; G, C. oculatus; H, C. erythrocephalus; I, C. imitator; J, C. lanio; K, C. albertisi; L, Hadrotes crassus; M, Thinopinus pictus; N, Hadropinus fossor; O, Liusus hilleri. Abbreviations: iedg, apical inner edge of arms of copulatory piece; por, parallel oblique ridges of copulatory piece (see also Fig. 10); xbar, ‘cross-bar’ of appendix. Additional abbreviations as in Figure 3. Scale bar = 1 mm.

Figure 5. Female genitalia, details of morphology (all but E, ventral view). A, Creophilus m. villosus, internal genitalia; B, C. erythrocephalus, internal genitalia; C, C. huttoni, internal genitalia; D, C. oculatus, gonocoxites and styli (setae omitted from left side); E, Thinopinus pictus, abdominal segments VIII–X, dorsal (vestiture omitted); F, T. pictus, internal genitalia; G, Staphylinus caesareus, internal genitalia; H, Platydracus maculosus, internal genitalia; I, Hadrotes crassus, right gonocoxites and stylus (vestiture omitted) – arrow indicates membranous articulation; J, T. pictus, right gonocoxite. Abbreviations: bcx, bursa copulatrix; fl, flagellum (note that the complete structure is not shown); gc, gonocoxite; gcm, gonocoxal macroseta; gc1, gonocoxite I, gc2, gonocoxite II; mg, median groove; pls, paired lateral sclerites of vaginal plate; sc, sclerite of bursa copulatrix; sd, spermathecal duct; sp, spermatheca; ss, sclerotized strip of vaginal plate; sty, gonostyle; vf, vaginal fold; vp, vaginal plate. Scale bars = 1 mm.

Figure 7. Heads of some species of the Creophilus erythrocephalus species-group (antennal flagella omitted). A, C. erythrocephalus; B, C. lanio; C, C. oculatus; D, C. imitator; E, C. albertisi. A–D, large males; E, small female. Scale bars = 5 mm.

Figure 32. Creophilus oculatus, details of morphology (setae of mandibles, antenna, and pronotum omitted). A, mandibles, dorsal (bases omitted); B, right antenna, dorsal (only apical setae shown); C, aedeagus with details of inverted internal sac, ventral; D, same, right lateral (parameral setae omitted); E, ventral sclerite of internal sac, ventral (lower) and apical (upper); F, pronotum (female); G, female internal genitalia, ventral; H, paramere, dorsal; I, median lobe apex, ventral. Abbreviations: T1, T2, T3, mandibular teeth. Additional abbreviations as in Figure 3 (male) and Figure 5 (female). Scale bars: E = 0.1 mm; rest = 1 mm.

Figure 33. New Zealand Creophilus species, details of morphology (setae of heads, pronota, mesoventrites, and legs omitted). C. oculatus (A, D, G, J, M, N); C. huttoni (B, E, H, O); C. rekohuensis, holotype (C, F, I, K, L, P). A–C, head, dorsal; D–F, pronotum; G–I, mesoventrite; J, K, protarsus, dorsal; L, M, right protibia, anterior; N–P, left wing, dorsal; Q, scatter plot of wing length vs. pronotum length as proxy for body size (circles = C. oculatus, squares = C. huttoni, triangles = C. rekohuensis; black symbols = males, white symbols = females). Abbreviations: CuA, MP3, MP4, wing veins mentioned in text. Scale bars = 1 mm.

Figure 34. Distribution of all georeferenced specimens of C. oculatus (circles) on the New Zealand mainland (left) and Chatham Islands archipelago (right). This species is also known from several other offshore islands, including the subantarctic Auckland and Snares Islands in the south, and the Kermadec Islands in the north east. Rectangle at southern end of New Zealand indicates area shown in Figure 35.