Volume 7 Number

May 2007

The Taxonomic Report

OF THE INTERNATIONAL LEPIDOPTERA SURVEY

A Description of a New Subspecies of Lycaena phlaeas

(Lycaenidae: Lycaeninae) from Montana, United States,

With a Comparative Study of Old and New World Populations

Steve Kohler

125 Hillcrest Loop, Missoula, Montana 59803

United States of America

Abstract: The Palaearctic, Oriental and Ethiopian Region subspecies of Lycaena phlaeas are briefly discussed. A

more detailed account of the North American subspecies is presented, and a new subspecies, L. p. weberi, from the Sweet Grass

Hills, Montana is described. The possibility that the eastern United States subspecies hypophlaeas was introduced from the Old

World is discussed; however no conclusion can be reached with certainty. The relationship between Old World and New World

subspecies of L. phlaeas is discussed. Evidence presented supports the treatment of New World populations as subspecies of L.

phlaeas.

Additional key words: Polygonaceae, Rumex acetosella, R. acetosa, R. crispus, Oxyria digyna.

INTRODUCTION

Lycaena phlaeas (Linnaeus, 1761) is a widespread species with subspecies in Europe, North Africa,

Arabia, northern Asia, Japan, North America and tropical Africa. The nominate subspecies occurs in

northern Europe (Ackery et al ., 1995). Shields & Montgomery (1966) mentioned that European texts list

Polygonaceae (Rumex and Polygonum) as larval foodplants for L. phlaeas subspecies. Flight period is

April to November, in one to four generations, depending on local conditions; over-wintering is in the

larval stage (Tuzov, 2000). Bridges (1988) listed 19 subspecies in his catalogue, not including the North

American ones. Miller and Brown (1981) listed five subspecies for North America. Ford (1924)

attempted to cover the world-wide geographic races of L. phlaeas , but the emphasis was on the Old World

taxa. He only discussed two taxa from North America, hypophlaeas Boisduval and feildeni M’Lachlan.

Shields & Montgomery (1966) discussed the distribution and bionomics of L. phlaeas subspecies in North

America, as did Ferris (1974), with the description of a new subspecies, arctodon. Two more recent papers

also discussed taxa in L. phlaeas. Emmel et al. (1998) discussed hypophlaeas, with lectotype designation

and type locality restriction; and Emmel & Pratt (1998) gave a new name, alpestris , to the California

population. The Palaearctic, Oriental and Ethiopian Region subspecies will be briefly discussed below.

The North American subspecies will receive a more detailed accounting and a new subspecies will be

described. There has been speculation that the eastern United States populations were introduced from the

Old World by human agency. There has also been speculation by some authors that the North American

subspecies are not phlaeas , but constitute a different species. These theories will be discussed below.

PALAEARCTIC, ORIENTAL AND ETHIOPIAN SUBSPECIES

This section is presented to provide information pertinent to the discussion at the end of this paper

and in the hope that more light may be shed on the relationship between the Old World and New World

representatives of L. phlaeas. It is not intended to be an extensive and complete treatise on the Old World

subspecies of L. phlaeas and there may be unintended omissions.

Lycaena phlaeas phlaeas (Linnaeus, 1761). TL: Westermannia, central Sweden. The nominate

subspecies is widespread and common in Europe from south of the Arctic Circle to all of the larger

Mediterranean islands and island groups as well as NW Africa (Tolman & Lewington, 1997, plate. 21).

Typical specimens are shown in Figs. 1-16 of this paper. Probable synomyms of phlaeas are comedarum

(Grum-Grshimailo, 1890) (East Pamirs); oxiana (Grum-Grshimailo, 1890) (Bokhara, Pamirs); stygianus

(Butler, 1880) (West Pamirs, Baluchistan to Chitral and Ladak).

Material Studied : NORWAY: Skjeberg, Grimsoy, 28 July 1990, lc?; Els 20, Tune, Rakil, 6 June 1990, lc?, T. J. Olsen Coll.

ROMANIA: Hagieni Forest nr. Mangalia, 6 June 1984, 7c? c? 3 9 9, A. Popescu-Gorj Coll. GERMANY: R/M Hesse, Rhein

Main Air Base, 3 August 1971, lc? 1 9, R. L. Hardesty Coll. FRANCE: Aveyron: Naucelle Lespinassolle a Chateau d’ eau,

5200m, 12 July 1990, 2c?c?, 20 July 1990, 2c?c?, J. Moonen Coll.; Bretagne: Morbihan Arradon, 10 August 1987, lc?, J.

Moonen Coll.; Vaucluse: Luberon, 17 July 1983, 6c? c? 4 9 9. SPAIN: Barcelona, 4 March 1980, 1 c?; Sierra Nevada, 1300m, 20

June 1988, 1 9, J. Munoz Sariot Coll.; Madrid: Casa de Campo, 28 July 1984, 1 c?, A. Sanchez Conde Coll. ENGLAND: Essex,

Purfleet, 30 July 1924, 19, 6 August 1924, lc?, 10 August 1926, 1 9; Surrey, 27 July 1924, lc?. ITALY: Vergato, 3 May 1984,

3c? c?, D. Cappelli Coll.

Lycaena phlaeas abbottii (Holland, 1892). TL: “Eastern Africa”. It is found in northern Malawi,

Tanzania and Kenya (Ackery et al ., 1995), and was treated as a distinct species by Kielland (1990). It is

illustrated in D’ Abrera (1980, p. 525).

Lycaena phlaeas baralacha (Moore, 1884). TL: Baralacha Pass, 4875m, Ladak. It occurs in the outer

Himalayas (Kashmir-Kumaon) and Nepal (Shields, 1982).

Lycaena phlaeas coccineus (Ford, 1924). TL: Tian-Shan. Illustrated in Ford (1924, pi. LIV, figs. 3-4).

Lycaena phlaeas chinensis (C. Felder, 1862). Central China (Bridges, 1988). This subspecies is

illustrated in Ford (1924, pi. LIV, figs. 10-11) and Tuzov (2000, pi. 57, figs. 28-30).

Lycaena phlaeas daimio Seitz, 1908. TL: Japan (Bridges, 1988). Representative specimens are shown in

Figs. 25-28. Also illustrated in Ford (1924, pi. LIV, fig. 9) and Tuzov (2000, pi. 57, figs. 31-33).

Material Studied : JAPAN: Aomori: Kanagi, 16 August 1973, 2c?, A. Kitagawa Coll.; Hokkaido: Maruszppu, 7 June 1987, lc?,

Y. Yazaki Coll.; Hokkaido: Yudetsu, 27 August 1987, 1 9, Y. Yazaki Coll.; Kuroishi City, 28 July 1991, 1 d\ K. Dorbashi Coll.;

Miyazaki Omo, 2 June 1979, lc?, A. Kitagawa Coll.; Nagano: Hotaka, 8 June 1978, 1 9, A. Kitagawa Coll.; Saitama: Koma, 23

April 1977, 5c? c? 19,3 May 1978, 19, A. Kitagawa Coll.; Saitama: Dairokutenjin Iwatsuki, 21 April 1981, 2c? c? 29 9, S.

Ohshima Coll.; Tochig: Shiobara, 1 June 1978, 1 9, A. Kitagawa Coll.; Yamanashi: Shibiro, 5 July 1989, 1 9, K. Dorbashi Coll.

Lycaena phlaeas eleus (Fabricius, 1798). A representative male specimen is shown in Figs. 17-18.

Material Studied : MALTA: Buckett, June 1987, lc?, P. Samut, Collector.; Miseb, 26 May 1986, 2c?c?, P. Samut Coll.; San

Giljan Valley, 24 May 1986, 1 c?, P. Samut Coll.

Lycaena phlaeas ethiopica (Poulton, 1922). TL: Uganda: 6000’, in the extreme SW of Uganda; high

country near Lake Kivu and between it and the northern end of Tanganyika. Distribution includes alpine

areas in the Ruwenzori Mountains of the Kigezi District of south-western Uganda, adjoining areas of Zaire,

and NW Tanzania (Ackery et al., 1995). It is illustrated in D’Abrera (1980, p. 525).

2

Figs. 1-20. Old World Lycaena phlaeas ssp. Fig. 1. L. p. phlaeas, Hagieni Forest nr. Mangalia, Romania, 6 June 1984, A.

Popescu-Gorj Coll., d dorsal. Fig. 2. Same, ventral. Fig. 3. Same, 9 dorsal. Fig. 4. Same, ventral. Fig. 5. L. p. phlaeas,

Skjeberg, Grimsoy, Norway, 29 July 1990, T.J. Olsen Coll., d dorsal. Fig. 6. Same, ventral. Fig. 7. L. p. phlaeas, Naucelle

Lespinassolle a chateau d’eau, 500m, Aveyron, France, 20 July 1990, J. Moonen Coll., d dorsal. Fig. 8. Same, ventral. Fig. 9.

L. p. phlaeas, Luberon, Vaucluse, France, 17 July 1983, 9 dorsal. Fig. 10. Same, ventral. Fig. 11. L. p. phlaeas, Vergato, Italy,

3 May 1984, D. Cappelli Coll., d dorsal. Fig. 12. Same, ventral. Fig. 13. L. p. phlaeas, Barcelona, Spain, 4 March 1980, d

dorsal. Fig. 14. Same, ventral. Fig. 15. L. p. phlaeas. Sierra Nevada, 1300m, Spain, 20 June 1988, J. Munoz Sariot Coll., 9

dorsal. Fig. 16. Same, ventral. Fig. 17. L. p. eleus, Miseb, Malta, 26 May 1986, P. Samut Coll., d dorsal. Fig. 18. Same,

ventral. Fig. 19. L. p. lusitanicus, San Roque, Cadiz, Spain, 15 April, 1980, J.L. Torres Mendez Coll., d dorsal. Fig. 20. Same,

ventral. All figs, approximately 1.3X life size. Photos by Steve Kohler. _

3

shade of

Lycaena phlaeas flavens (Ford, 1924). TL: Lhasa, Tibet. The ventral hind wings are of an even

lemon-yellow, a unique feature. (Ford, 1924; Bridges 1988).

Lycaena phlaeas ganalica P. Gorbunov, 1995. TL: Kamchatka, Russia (Tuzov, 2000).

Lycaena phlaeas hibernica Goodson, 1948. TL: Ireland (Bridges, 1988).

Lycaena phlaeas hyperborea (Ford, 1924). Arctic Norway (Bridges, 1988). This subspecies is illustrated

in Ford (1924, pi. LIV, fig. 6).

Lycaena phlaeas japonica (Ford, 1924). TL: Japan (Bridges, 1988). This subspecies is illustrated in Ford

(1924, pi. LIV, figs. 2, 16). The type is in the Tring Zoological Museum.

Lycaena phlaeas kuriliphlaeas (Bryk, 1942). TL: Kurile Island (Bridges, 1988).

Lycaena phlaeas lusitanicus (Bryk, 1940). TL: Portugal (Bridges, 1988). Representative specimens are

shown in Figs. 19-22.

Material Studied : SPAIN: Cadiz: La Linea, 28 February 1980, Id*, J.L. Torres Mendez Coll.; Cadiz: San Roque, 16 January

1984, Id, 5 March 1982, 1 9, 15 April 1980, Id, 18 June 1983, Id, J.L Torres Mendez Coll.; La Coruna: Choren Mellid, 24

May 1986, 1 9, E.H. Fernandez Vidal Coll.

Lycaena phlaeas matsumuranus (Bryk, 1946). TL: Korea (Bridges, 1988). A representative male

specimen is shown in Figs. 23-24.

Material Studied : KOREA: Seoul, 30 April 1986, 1 d, 3 May 1986, Id.

Lycaena phlaeas phlaeoides (Staudinger, 1901). TL: Funchal, Madeira. Found only on Madeira (Tolman

& Lewington, 1997). The rich brown, somewhat mottled color and jagged whitish postmedian band of the

ventral hind wing on this subspecies are distinctive. It is illustrated in Ford (1924, pi. LIV, figs. 1, 8, 20)

and Tolman & Lewington (1997, pi. 21).

Lycaena phlaeas polaris Courvoisier, 1911. TL: Norwegian Lappland. Distribution is Arctic

Fennoscandia (Tolman & Lewingson, 1997). It is illustrated on their plate 21. This subspecies is

distinguished by the dove grey ventral hind wing ground color and the whitish postmedian band distally

bordering the postdiscal series of black spots.

Lycaena phlaeas pseudophlaeas (Lucas, 1866). TL: “Abyssinie”. It is found in the Highlands of Ethiopia

(Ackery et al ., 1995). It is illustrated in D’Abrera (1980, p. 525).

Lycaena phlaeas shima Gabriel, 1954. TL: Yemen: “Jebel Masnab, S. W. of Ma’bar, c. 8400 ft.” It is

found in the Highlands of south-western Arabia (Saudi Arabia and Yemen) according to Ackery et al.

(1995). It is illustrated in D’Abrera (1980, p. 525).

Lycaena phlaeas sibiricanus Kozhanchikov, 1936. TL: Siberia (Bridges, 1988).

Lycaena phlaeas timeus (Cramer, 1777). TL: North Western Himalaya (Bridges, 1988). The relationship

of comedarum, oxiana , and stygianus needs study.

Figs. 21-28. Old World Lycaena phlaeas ssp. Figs. 29-40. North American Lycaena phlaeas ssp. Fig. 21. L. p. lusitanicus, San

Roque, Cadiz, Spain, 5 March 1982, J.L. Torres Mendez Coll., 9 dorsal. Fig. 22. Same, ventral. Fig. 23. L. p. matsumuranus,

Seoul, Korea, 3 May 1986, d dorsal. Fig. 24. Same, ventral. Fig. 25. L. p. daimio, Koma, Saitama, Japan, 23 April 1977, A.

Kitagawa Coll., d dorsal. Fig. 26. Same, ventral. Fig. 27. Same, Iwatsuki, Dairakuteniin, Saitama, Japan, 21 April 1981, S.

Ohshima Coll., 9 dorsal. Fig. 28. Same, ventral. Fig. 29. L. p. arctodon, Beartooth Plateau, Carbon Co., Montana, U.S.A., 15

July 1989, S. Kohler Coll., d dorsal. Fig. 30. Same, ventral. Fig. 31. Same, 9 dorsal. Fig. 32. Same, ventral. Fig. 33. L.p.

arethusa, Hailstone Butte, Alberta, Canada, 24 July 1980, J. Johnstone Coll, d dorsal. Fig. 34. Same, ventral. Fig. 35. Same,

Plateau Mountain, Alberta, Canada, 26 July 1980, N.G. Kondla Coll., 9 dorsal. Fig. 36. Same, ventral. Fig. 37. L. p. weberi ,

Mount Royal, 6300-6900’, East Butte, Sweet Grass Hills, Liberty Co., Montana, U.S.A., 30 July 2004, S. Kohler Coll., holotype

d dorsal. Fig. 38. Same, ventral. Fig. 39. Same, 5 August 2003, allotype 9 dorsal. Fig. 40. Same, ventral. All figs.

approximately 1.3X life size. Photos by Steve Kohler. _

5

NORTH AMERICAN SUBSPECIES

Five subspecies of L. phlaeas occurring in North America are recognized. A sixth from the Sweet

Grass Hills of Montana is designated below. Each of the subspecies is discussed, and material examined

for this study is listed. Forewing length measurements (from the junction with the thorax to the wing apex)

are given in millimeters. Ferris (1974, p. 6) used a table to enumerate the differences among the named

subspecies, with the characters of dorsal forewing and ventral hind wing black spots; forewing black

borders; dorsal and ventral hind wing orange borders; and ground color of dorsal fore wing and ventral hind

wing. This table was also referred to by Emmel & Pratt (1998). The table of Ferris is reproduced with

revisions here as Table 1.

Lycaena phlaeas hypophlaeas (Boisduval, 1852). TL: “Nord de la Californie. II se retrouve dans tout le

nord des Etats-Unis”. It was restricted by Shields (1967) to northern California. It was further restricted to

vicinity of Boston, Massachusetts by Emmel et al. (1998). The lone syntype specimen is in the U.S.

National Museum (Emmel et al., 1998). The name americana Harris, 1862 is a junior synonym of

hypophlaeas Boisduval. This non arctic-alpine subspecies was known for many years by the name

americana ; however, the work of several authors has clarified the correct name. Shields & Montgomery

(1966) gave the English translation from Boisduval’s description of hypophlaeas in French, which was first

given by Wolley Dod (1907) as “North of California. It is found in all the northern United States”. Thus

they concluded the type locality is not “California” as listed by Klots (1951) and Comstock & Huntington

(1960), and said they did not know of a precise locality for hypophlaeas, nor where the type specimen(s)

were located. Shields (1967) then said that “north of California” should instead be translated “Northern

California”. He also said the probable type locality was “the Sierra Nevada Mountains, California”. Ferris

(1974) accepted the reasoning by Shields, and stated, “Boisduval’s type of hypophlaeas is in the collection

of the United States National Museum. The type was collected by J.M. Lorquin but does not bear exact

locality information”. Emmel et al. (1998) questioned the likelihood of hypophlaeas being from

California. They reasoned: (1) The lone syntype in the U. S. National Museum collection is typical of the

eastern United States phenotype and does not resemble any of the high-elevation California populations of

L. phlaeas ; and (2) Even if the type specimen was purported to represent an atypical variant of a California

population, it is extremely unlikely that Lorquin collected in any of the current arctic-alpine habitats of this

insect. Further, they reasoned that since Boisduval was aware of the presence of this insect in the eastern

United States, he undoubtedly already had material from that region and may have assumed that the species

occurred in northern California, without any Lorquin specimens to support this assumption. Thus, they

concluded that the name hypophlaeas was applicable to the L. phlaeas populations of the eastern U.S., but

not the ones from California. They designated the sole syntype as the lectotype and restricted the type

locality to the vicinity of Boston, Massachusetts, an area known to have populations with a phenotype

matching the hypophlaeas type, and an area which was, at the time, easily accessible to collectors

providing material to European lepidopterists. Thus the name americana Harris, 1862 becomes a junior

synonym of hypophlaeas Boisduval.

The subspecies hypophlaeas is widely distributed in eastern North America. Ferris (1974) gave its

range (as americana) as Nova Scotia and The Gaspe west through Canada to central Ontario and

Minnesota, south to Virginia and montane northern Georgia, Missouri and Kansas. One historical Cass

County, North Dakota record exists (Royer, 2003). It is generally rare or temporary on the Great Plains

westward. Marrone (2002) reported only three widely scattered South Dakota records. Layberry et al.

(1998) show a record for southern Manitoba. Hooper (1973) mentions one record near Regina,

Saskatchewan. Elrod (1906) said that C. A. Wiley found it not rare at Miles City, Montana. It has also

been taken in eastern Colorado near Colorado Springs (Ferris & Brown, 1981). Habitat where hypophlaeas

is most often found is disturbed areas, including vacant lots, weedy pastures, roadsides and lake shorelines.

6

Figs. 41-49. North American Lycaena phlaeas ssp. Fig. 41. L. p .hypophlaeas, Springdale, Sussex Co., New Jersey, U.S.A., 18

July 1978, W.B. Wright Coll., d dorsal. Fig. 42. Same, ventral. Fig. 43. Same, 9 dorsal. Fig. 44. Same, ventral. Fig. 45. L. p.

alpestris, N. Slope Mount Dana, 11500’, Mono Co., California, U.S.A., 6 August 1991, M. Grinnell Coll, d dorsal. Fig. 46.

Same, ventral. Fig. 47. Variation of some L. p. weberi paratypes from Mount Royal, East Butte, Sweet Grass Hills, Liberty Co.,

Montana, U.S.A. Left two columns dd, right two columns 9 9. Fig. 48. Variation of L. p. arctodon series from Beartooth

Plateau, Carbon Co., Montana, U.S.A. Left two columns d d, right two columns 9 9. Fig. 49. Variation of L. p. arethusa series

from Hailstone Butte and Plateau Mountain, Alberta, Canada. Left two columns d d, right column upper one d, two lower ones

9 9. Ligs. 41-46 approxmately 1.3X life size, figs. 47-49 approximately 2/3 life size. Photos by Steve Kohler. _

Klots (1951) reported the larval foodplants Rumex acetosella L. (Sheep Sorrel), R. acetosa L. and

R. crispus L. (Curled Dock). Opler & Krizek (1984) described the life history. The pale-green eggs are

laid singly on host leaves or stems. The young caterpillars chew holes in the underside of young host

leaves and later make longitudinal channels. Development takes about three weeks and pupation is under

leaves or rocks. Winter is spent as pupae. The caterpillars are covered with short hairs and are variably

colored rose-red to green. There is a red dorsal stripe on some caterpillars. The chrysalis is light brown,

tinged with pale yellow-green and spotted with black.

7

Allen (1997, pi. 33, p. 312) shows a photograph of the larva (as Americana). In the northern parts

of its range, hypophlaeas is bivoltine (June-early July and August-September) and probably has three

broods everywhere to the south (mid-April through May, mid-June through July and mid-August through

September). The ground color of the dorsal forewing of hypophlaeas is bright coppery red-orange, not

brassy or brassy-red like the arctic-alpine subspecies (Table 1). Typical adults are shown in Figs. 41-44.

Thirty-two males and 25 females were examined. Average fore wing length of males was 12.3 mm, with a

range of 11.5 to 14.5 mm. Average forewing length of females was 13.5 mm, with a range of 11.5 to 14.5

Material Studied : ILLINOIS: Palos Park, McMahon Woods, 23 May 1965, 8 c? c? 3 9 9, R. Arnold Coll.; IOWA: Polk Co.:

Des Moines, 850’, 9 July 1929, 1 ?, 21 July 1929, Id 1 9, 7 August 1932, 2d d, 21 August 1932, Id, 4 September 1927, Id,

O. E. Booth Coll.; MAINE: Penobscot Co.: Passadumkeag, 10 June 1976, Id 19, L.P. Grey Coll.; MARYLAND: Alegheny

Co.: 6 mi. E. Flinstone, 12 May 1983, 1 9, T.A. Greager Coll.; NEW JERSEY: Ocean Co.: Lakehurst, 11 May 1978, 2dd

69 9, W. B. Wright Coll.; Sussex Co.: Springdale, 16 July 1978, 2dd 19, 18 July 1978, 4dd 3 9 9, W.B. Wright Coll.;

PENNSYLVANIA: Elk Co.: 3 mi. W. Dent’s Run, 19 July 1983, 2d d, T.A. Greager Coll.; Indiana Co.: 1 mi. S. Clarksburg,

14 May 1977, 1 9, 2 mi. N. Shelocta, 5 July 1981, 19,5 June 1983, 1 d , 21 July 1983, Id, T.A. Greager Coll.; Westmoreland

Co.: 1.5 mi. W. Greensburgh, 14 May 1979, 1 9, 22 May 1981, 19,8 July 1977, Id, 2 August 1983, Id, T.A. Greager Coll.;

WEST VIRGINIA: Pendleton Co.: Franklin, 24 July 1978, Id, 16 August 1976, 2dd, J.E. Dewey Coll.; WISCONSIN:

Juneau Co.: Necedah Township, 5 June 1979, 29 9,30 July 1979, Id 1 9, T. Krai Coll.

Lycaena phlaeas feildeni (M’Lachlan, 1878). TL: Grant Land, Northwest Territories, according to Miller

& Brown (1981), who claimed the location of the type was unknown. However, Shields & Montgomery

(1966) referencing Tite (1957), stated, “M’Lachlan (1878) described L. p. feildeni from two males and one

female from ‘Lat. 81° 45’. The British Museum of Natural History contains these three specimens which

bear the label ‘Grinell Land west side of Smith Sound, Arctic America 78-83 Lat. (81-45) Capt. Feilden R.

N. 77-101’ ”. They also referenced Wolff (1964) stating that these were collected in 1875 or 1876. Ferris

(1974) placed the type locality of feildeni as “Ellesmere Island, Lat. 81° 45’N”. He showed the distribution

of this subspecies to be the Hayes Peninsula of western Greenland; Ellesmere Island, Banks Island, Baffin

Island, Simpson Peninsula, South Hampton Island and District of Keewatin, Northwest Territories, Canada.

He also stated that the insect is poorly represented in collections with the few extant specimens placed

primarily in the Canadian National Collection and the Natural History Museum (London). Layberry et al.

(1998) included the arctic coast of Yukon Territories and Alaska in the distribution of feildeni, while Ferris

(1974) considered these populations undescribed. The habitat of feildeni is tundra and the larval foodplant

is Oxyria digyna (L.) Hill (Mountain Sorrel) (Ferris, 1974; Layberry et al ., 1998). The subspecies feildeni

is illustrated in Shields & Montgomery (1966, figs. 1 and 2, pp. 232-233); in Ferris (1974, figs. 10-16, p.

12); in Layberry et al. (1998, pi. 10, fig. 1). There is one generation per year. The dull brassy color of the

dorsal forewing with smoky washed out aspect and the very small sometimes indistinct ventral hind wing

black spots characterize this subspecies (Table 1). No specimens were examined in this study.

Lycaena phlaeas arethusa (Wolley Dod, 1907). TL: “nr. Calgary, Alberta” in Miller & Brown (1981).

Restricted to the head of Fish Creek, Alberta by Kondla (1996). After giving reasons for restricting the

type locality, Kondla (1996) stated, “the locality was near Billings Lumber Mill as evidenced by label data

on additional paratypes in the Canadian National Collection, collected on 19 and 20 July 1903. In a brief

discussion about L. phlaeas , Wolley Dod (1904) stated, ‘About fifteen specimens of this were captured

near the spruce bush at the head of Fish Creek in southern Alberta’ ”. Kondla (pers. com., 2007) has

offered new information concerning the types and type locality of arethusa. The statement by Shields &

Montgomery (1966), “The holotype and allotype are in the U. S. National Museum and six paratypes are in

the Canadian National Collection” is not correct, nor is “HT in USNM” in Miller & Brown (1981). Wolley

Dod in the original description stated, “Described from five males and eight females. . . . Types, c? and ?

in U.S. National Museum, the rest co-types.” Wolley Dod did not designate one specimen as the name

bearing type and so all extant specimens in the type series are syntypes. Also, since one of the syntypes

came from the “south fork of Sheep Creek”, then the type locality should be amended to “the head of Fish

Creek and the south fork of Sheep Creek, Alberta”. Syntypes are in the U.S. National Museum and the

Canadian National Collection. Layberry et al. (1998) gave the distribution of arethusa as from the Rocky

Mountains of Alberta northward to Boreal Zone habitat in southern and central Yukon. A record by James

Scott from 1962 (pers. corresp., Scott, 1975)—Logan Pass, Glacier National Park, Flathead/Glacier

Counties, Montana is probably referable to this subspecies. Ferris (1974) in discussing the habitat and

larval foodplant of arethusa according to J.A. Legge, Jr. and C.D. Bird, stated that “on Plateau Mountain

south of Banff, Alberta, it flies in small grassy meadows at 8200’ in association with Oxyria digyna and

Rumex alpestris (Scop.)”. The flight period is typically the first two weeks in August. The dull, red-brassy

with smoky or dusky cast of the dorsal forewings in most males and the very small ventral hind wing black

spots characterize this subspecies (Table 1). Typical adults are shown in Figs. 33-36. A range of

phenotypic variation is shown in Fig. 49. For this study, seven males and two females were examined.

Average fore wing length of males was 13.1 mm, with a range of 12.5 to 14.0 mm. Average forewing

length of females was 14.5 mm, with a range of 14.0 to 15.0 mm.

Material Studied : CANADA: ALBERTA: Hailstone Butte, 24 July 1980, 2c? c?, J. Johnstone Coll.; Plateau Mountain, 26 July

1980, 3?c? 29 9,6 August 1979, Id', N.G. Kondla Coll.; Plateau Mountain, 8000’, 14 August 1973, 1 d\ L.P. Grey Coll.

Lycaena phlaeas arctodon Ferris, 1974. TL: E. side Beartooth Pass, Carbon Co., Montana. The holotype

is in the Allyn Museum of Entomology, now part of the McGuire Center for Lepidoptera & Biodiversity,

Gainsville, Florida. Ferris (1974) gave the distribution of arctodon as “the Beartooth Plateau on the Park

Co., Wyoming-Carbon Co., Montana border; the Teton Mountains, Teton Co., Wyoming; Yellowstone

National Park on Mt. Washburn; and from the Lemhi Range, Lemhi Co., Idaho”. He also referred

specimens from Sweet Grass Co., Montana to this subspecies, and tentatively assigned a single male

phlaeas from the Wallowa Mountains, Wallowa Co., Oregon to arctodon. Warren (2005) notes that the

original Oregon record was a single male from Matterhorn Mountain and that additional Oregon

populations have been found in similar habitats in other parts of the high Wallowas in Wallowa County

referencing Pyle (2002). Here the butterfly flies over rockslides and talus slopes above 7500’. Since the

description of arctodon by Ferris in 1974, it has been collected in the Wind River Mountains, Sublette Co.

and Fremont Co., Wyoming (Harry, 1981), and the Big Horn Mountains, Big Horn Co., Wyoming. New

localities in Carbon, Judith Basin, Silver Bow, Gallatin and Stillwater Counties, Montana have also been

documented (Fig. 56). Records also exist for the Delano Peak area, Beaver and Piute Counties, Utah (Clyde

Gillette, pers. com., 2007). Subspecies arctodon is found in lush moist alpine meadow habitat near or

above treeline where the presumed foodplant, Rumex acetosa is found. At the type locality the plants grow

in depressions in open meadows where some moisture remains from the spring snow melt. Harry (1981)

described the habitat in the Wind River Mountains, Fremont Co., Wyoming as quite different from

Beartooth Pass, “Here, the butterfly lives among the rocky slopes like that preferred by Erebia magdalena.

This type of habitat is typical of where Oxyria digyna exists”. Harry documented Mountain Sorrel as a

larval foodplant at this location on the Bear’s Ears Trail, collecting four larvae from it, and was able to rear

one to adult. Later at the same location, he obtained 38 ova from an adult female and reared them to pupa

on O. digyna from the Wasatch Mountains, Utah. The suspected foodplant, R. acetosa , at the Beartooth

Pass type locality has subsequently been verified by Clyde Gillette (pers. com., 2007). The subspecies

closest to arctodon in appearance is arethusa , but arctodon does not have the wide dark dorsal forewing

borders exhibited by arethusa nor the smoky cast of the fore wings of the males. The appearance of

arctodon is much brighter than arethusa, and the dorsal hind wing blue spots are also more prominent. The

black spots on the ventral hind wing of arctodon are more distinct than on arethusa (Table 1). Typical

adults are shown in Figs. 29-32. A range of phenotypic variation is shown in Fig. 48. Scott (1986), p. 387)

applied the subspecies name polaris Courvoisier, 1911 (TL: Norwegian Lappland) to all of the western

United States populations, including California, ignoring the name arctodon. This should not be followed,

as polaris represents Old World populations distributed in Arctic Fennoscandia that differ from any North

9

American populations in having extensive whitish spaces on the ventral hind wing distally from the

postdiscal series of black spots (Tolman & Lewington, 1997, pi. 21). For this study, 50 males and 29

females of arctodon were examined. Average forewing length of males was 12.8 mm, with a range of 11.0

to 14.5 mm. Average forewing length of females was 13.5 mm, with a range of 12.0 to 14.5 mm.

Material Studied : MONTANA: Carbon Co.: nr. Beartooth Pass, 1 August 1973, 4c?c? 5 9?, 1 August 1974, 4c?c? 2 9 9, S.

Kohler Coll.; Beartooth Plateau, 13 July 2006, 8c? c? 1 9, 14 July 2000, 3c? c? 29 9, 14 July 2006, 15c? c? 5 9 9, 15 July 1989,

7c?c? 39 9, 16 July 1985, lc?, 28 July 1976, 19, S. Kohler Coll.; Hellroaring Plateau, 31 July 1974, 19, S. Kohler Coll.;

Gallatin Co.: above Fairy Lake, Bridger Mountains, 11 August 1986, 1 9, S. Kohler Coll.; Silver Bow Co.: Table Mountain,

Highland Mountains, 24 July 1986, 1 9, S. Kohler Coll.; Stillwater Co.: Benbow Mine Rd., 17 mi. SW Fishtail, 9000’, 18 July

1989,4c? c? 1 9, B. Vogel Coll.; above Mystic Lake, 15 August 1986,4c? c? 6 9 9, S. Kohler Coll.

Lycaena phlaeas alpestris J. Emmel & Pratt, 1998. TL: north slope of Mt. Dana, 11,200-11,800’, Mono

Co., California. The holotype, allotype and nine paratypes are in the collection of the Natural History

Museum of Los Angeles County, California. For a long time this subspecies was known as hypophlaeas ,

but as pointed out by Emmel et al. (1998), the lectotype of Polyommatus hypophlaeas in the U.S. National

Museum does not resemble any California specimens and appears to be a typical example of L. phlaeas

populations of the northeastern United States (see discussion of hypophlaeas above). They restricted the

hypophlaeas lectotype to eastern U.S. populations of L. phlaeas and sunk a mericana Harris, the name those

populations were long known as, to a junior synonym. This left the California populations of L. phlaeas

without a name, which led to the description of alpestris by Emmel & Pratt (1998). The distribution of

alpestris given by them is “the higher elevations of the Sierra Nevada from Fresno County and Inyo

County on the south, north to Sonora Pass on the Tuolumne-Mono County line”. They reference Shields &

Montgomery (1966); D. Bauer & K. Davenport (pers. com.). Emmel and Pratt (1998) also recently

discovered a population in the White Mountains along the California-Nevada border. This subspecies flies

in a single brood from mid-July to mid-August, and the larval foodplant is O. digyna. Emmel & Pratt

(1998) referred to the table by Ferris (1974) in summarizing the distinguishing characters of alpestris. The

dorsal forewing ground color is a pale brassy red, often with a dusky aspect. The dorsal forewing spots are

prominent and well developed and the outer margin borders tend to be narrow. A typical adult male is

shown in Figs. 45-46. For this study, five males were examined. Average forewing length was 13.4 mm,

with a range of 11.5 to 14.5 mm.

Material Studied : CALIFORNIA: Mono Co.: N. Slope Mt. Dana, 11,500’, 6 August 1991, 5c? c?, M. Grinnell Coll.

Lycaena phlaeas weberi Kohler, new subspecies

During the winter of 2002-2003, Byron Weber of Missoula, Montana brought to my home a number

of pinned butterfly specimens that he had collected in the area of the Sweet Grass Hills in Toole and

Liberty Counties, north-central Montana. Looking through this material, I was very surprised to see two

male phlaeas specimens that Byron had collected on the East Butte, Sweet Grass Hills, Liberty County.

My experience with L. phlaeas in Montana prior to this had been in high elevation alpine habitats near or

above timberline. Needless to say, the two large, very dusky specimens from below 7000’ elevation

Canadian Zone habitat on a Prairie Island Range mountain grabbed my attention. Plans were made to

return to the area to obtain more specimens and study the population, which Byron and I did in August

2003. I made two additional trips to the area in 2004 and 2005 to accumulate an adequate study series.

Definition: Besides the larger size, the most striking characteristic of weberi dorsally is the extremely

dark, dusky appearance. In many males, the copper ground color of the forewing is almost completely

obscured by dark brown, which often obliterates the inner margin of the dark wing border. The dusky

brown is also present in many of the females, causing them to appear much more dark and dusky than any

arethusa females. The dark border of the fore wings of both males and females of weberi is very wide,

more so than any of the other subspecies (excepting possibly hypophlaeas ), as a percentage of total wing

10

length (Table 1). The pattern of dorsal fore wing blackish spots is also very distinct and heavy (Figs. 37-

40). Fig. 47 shows a range of phenotypic variation. Ventrally the ground color of the hind wing of weberi

is a darker shade of warm gray than arethusa. This color is continued on the ventral forewing border and

wing apex, where it is considerably darker than on arethusa, as well as the orange of the discal portion of

the ventral forewing being brighter and more intense than on arethusa. The black spots on the ventral hind

wing of weberi are as in arethusa, being very small with distal whitish edging present. However, further

distally from these whitish spaces there are spaces that are darker than the rest of the gray ground color of

the hind wing of weberi, forming an indistinct darker band and giving somewhat the impression of a two-

toned hind wing. There is also darker gray-black in the hind wing tornus area of weberi, being fairly

distinct and obvious, but only vaguely present on arethusa. The orange crenulate submarginal line on the

ventral hind wing is bright, narrow and distinct on weberi (Figs. 38, 40), but is narrower and sometimes

faint on arethusa (Fig. 34). The main differences between this new subspecies and the other named

subspecies from North America are outlined in Table 1. Of the North American subspecies, weberi is most

similar to arethusa, but larger. Forewing length of male arethusa studied averaged 13.1 mm, with a range

of 12.5 to 14.0 mm, while weberi males averaged 14.5 mm, with a range of 12.5 to 15.5 mm. Forewing

length of the male holotype is 15.0 mm. Fore wing length of female arethusa studied averaged 14.5 mm,

with a range of 14.0 to 15.0 mm, while weberi females averaged 15.2 mm, with a range of 13.5 to 16.0 mm.

Fore wing length of the allotype female is 15.5 mm.

Etymology: This subspecies is named for Byron Weber of Missoula, Montana, who discovered the

population at the type locality, and whose interest in the Sweet Grass Hills and energy expended in

exploring them are inspiring. Byron’s grandfather, Harry Demarest, came to the Sweet Grass Hills from

Nebraska around the turn of the 20 th century. He worked on ranches and hauled freight with a team of 12

horses and homesteaded in 1910 just north of the town of Whitlash. Today the ranch spreads from East

Butte to Middle Butte to the original homestead. As a child and young man, Byron spent his summers on

the ranch in the hay fields, but his favorite times were spent alone along the willows of Breed Creek and on

the native prairie, identifying wildflowers and birds and quietly observing the mammals. In 1995, he began

to seriously study the butterflies of the area and now has several drawers of pinned butterfly specimens.

Distribution and Phenology: To date, this subspecies is known only from the type locality (Fig. 56). It

flies in a single brood from late-July to mid-August. The adults are found in close association with Rumex

acetosa, which is the presumed larval foodplant. A preferred nectar source is Solidago multiradiata Ait.

(Goldenrod). In late July 2004, Byron Weber and I climbed the West Butte, Sweet Grass Hills, Toole

County, which is similar in elevation to the East Butte, but much more of the terrain is dominated by

rockslides. We did not find weberi, nor did we find the larval foodplant. There is some controversy about

whether R. acetosa is native to North America. Moss (1983) discussed two subspecies of R. acetosa L.,

ssp. acetosa —gardens and waste places, introduced; and ssp. alpestris (Scop.) Love—moist banks and

meadows to alpine elevations, native, more or less circumpolar, Alaska, Yukon to Wyoming. This was

confirmed by Lesica (2002) discussing R. acetosa in Glacier National Park, Montana,’’Uncommon in moist

meadows and talus slopes, upper montane to alpine; East, West. Our plants are ssp. alpestris (Scop.) Love.

Circumboreal south to OR, WY. A closely related ssp. is introduced from Europe and grown as a garden

herb”. Thompson & Kuijt (1976) reporting a study of the montane and subalpine plants of the Sweet

Grass Hills stated of R. acetosa, “Collected from only one area, on the moist north-facing slope of Mount

Royal where outcrops of Madison limestone have produced calcareous soils. Although this arctic-alpine

species, native to the American Arctic, has been naturalized from Eurasia in the eastern United States, it is

believed to occur as a relict in the Sweetgrass Hills rather than as a garden escapee, since it has been

reported in the vicinity of Montana only from alpine or subalpine areas in Glacier Park, the Bear Paw

Mountains, and the Beartooth Plateau”. They also pointed out that arctic-alpine disjunctions are often

correlated with calcareous substrates, and the close association of R. acetosa with soils derived from

limestone in East Butte suggests its persistence there as an arctic relict.

11

Types: Holotype male: MONTANA: Liberty County: Mount Royal, 6300-6900’, East Butte, Sweet

Grass Hills, 30 July 2004, S. Kohler Coll. (Figs. 39-40). Allotype female: MONTANA: Liberty Co.:

Mount Royal, 6300-6900’, East Butte, Sweet Grass Hills, 5 August 2003, S. Kohler Coll. (Figs. 41-42).

Paratypes (55 c? c? 33 9 9): MONTANA: Liberty Co.: nr. summit of East Butte, 6800’, Sweet Grass Hills,

15 August 1996, 2c? c?, B. Weber Coll.; Mount Royal, 6300-6900’, East Butte, Sweet Grass Hills, 5 August

2003, 6c? c? 13 9 9, 30 July 2004, 24c? c? 4 9 9, 28 July 2005, 28c? c? 8 9 9, S. Kohler Coll.; 5 August 2003,

lc? 7 9 9, B. Weber Coll.

Deposition of Types: The holotype male, allotype female, two male and two female paratypes will be

deposited in the Monte L. Bean Life Science Museum, Brigham Young University, Provo, Utah. Three

male and seven female paratypes are in the Weber collection, and the remaining paratypes are in the Kohler

collection.

Type Locality: MONTANA: Liberty County: north slope of Mount Royal, East Butte, Sweet Grass Hills,

from the summit (6914’) down slope (north) to the saddle (6300’) between Mount Royal and Mount

Brown. The upper part is forested with spruce ( Picea glauca x engelamnnii ), whitebark pine ( Pinus

albicaulis ), limber pine (P. flexilis ) and lodgepole pine (P. contorta ) (Thompson & Kuijt, 1976), and is

fairly steep and rocky in places, but the lower slope towards the saddle is more gentle and open and

supports open lush meadow (Figs. 51-53). The north-facing aspect of the slope allows the R. acetosa to

grow on the upper parts (Fig. 55). The Sweet Grass Hills in the northern part of Liberty County near the

Alberta-Montana border in north-central Montana are unique in that they are the highest isolated peaks in

the United States. Of volcanic origin, the Sweet Grass Hills are prominent landmarks, rising nearly 3000’

above the surrounding prairie with rolling hills extending to the north almost to the Alberta-Montana

border. They are visible for more than 50 miles and consist of three distinct butte complexes with scattered

grassy hills connecting them. The three buttes are West Butte (elevation 6983’, on left); Middle or Gold

Butte (elevation 6512’); and East Butte (elevation 6958’, on right) with two smaller features (on far right),

Grassy and Haystack Buttes (Fig. 50).

GENITALIC STUDY

Male genitalia of the following subspecies were dissected and subjected to microscopic

examination: L. p. phlaeas (Romania: Hagieni Forest near Mangalia, 6 June 1984, A. Popescu-Gorj Coll.,

1; France: Aveyron: Naucelle Lespinassolle a Chateau d’ eau, 520m, 12 July 1990, J. Moonen Coll., 1); L.

p. eleus (Malta: Miseb, 26 May 1986, P. Samut Coll., 1); L. p. lusitanicus (Spain: Cadiz, La Linea, 28

February 1980, J.L. Torres Mendez Coll., 1); L. p. daimio (Japan: Saitama, Koma, 23 April 1977, A.

Kitagawa Coll., 1); L. p. hypophlaeas (Pennsylvania: Indiana Co., 2 mi. N. Shelocta, 5 June 1983, T.A.

Greager Coll., 1); L. p. alpestris (California: Mono Co., N. slope Mt. Dana, 11500’, 6 August 1991, M.

Grinnell Coll., 1); L. p. weberi (Montana: Liberty Co., Mount Royal, 6300-6900’, East Butte, Sweet Grass

Hills, 30 July 2004, S. Kohler Coll., 1); L. p. arctodon (Montana: Carbon Co., Beartooth Plateau, 16 July

1989, S. Kohler Coll., 1); L. p. arethusa (Canada: Alberta, Plateau Mountain, 26 July 1980, N.G. Kondla

Coll., 1). After careful examination, I found no genitalic characters that were useful in separating these

taxa in this limited study. I concluded the male genitalia of these subspecies were virtually identical.

There have been few comparative studies of L. phlaeas subspecies in the literature. In Russia,

Gorbunov (2001) noted the apical part of the valve of chinensis (C. Felder) was wider than in subspecies

phlaeas and ganalica. Ford (1924) in reviewing Ethiopian populations, abbottii (Holland), ethiopica

(Poulton) and pseudophlaeas (Lucas) concluded from the genitalic descriptions of T.A. Chapman that “the

genitalia of these Ethiopian forms ... do not differ from those of H. phlaeas phlaeas save in a slight

diminution in size, most noticeable in the aedoeagus”, signs of geographical variation. Ford (1924) also

agreed with Chapman’s conclusions regarding the genitalia of “ hypophlaeas (Lapland and N. America) as

specifically identical with phlaeas .”

Figs. 50-55. Habitat of Lycaena phlaeas weberi at type locality. Fig. 50. Sweet Grass Hills, Montana, from the southwest,

looking north toward Canada; from left, West Butte, Middle Butte (Gold Butte), East Butte, Grassy Butte, Haystack Butte. Fig.

51. Near the summit of Mount Royal, East Butte. Fig. 52. From the summit of Mount Royal looking north to the saddle and

Mount Brown. Fig. 53. Looking back toward Mount Royal from the saddle. Fig. 54. L. p. weberi <S taking nectar from a

preferred source, Solidago multiradiata (Goldenrod). Fig. 55. Rumex acetosa, presumed larval foodplant of L. p. weberi.

Photos by Steve Kohler. _

14

Fig. 56. Distribution of Lycaena phlaeas ssp. in Montana.

• L. p. hypophlaeas

DISCUSSION

Opler & Krizek (1984) and Opler & Malikul (1992) suggested that the eastern United States

population of Lyaena phlaeas hypophlaeas was introduced from Europe in Colonial times, reasoning that it

is associated with waste places and introduced foodplants, Rumex acetosella (Sheep Sorrel) and

occasionally R. crispus (Curled Dock) and it resembled European material. Other authors (Ehrlich &

Ehrlich, 1961) have also put forth this theory. Layberry et al. (1998) pointed out, however, that unlike

European specimens, subspecies hypophlaeas (as Americana) has a pale gray rather than brown ventral

hind wing, with larger more sharply defined black spots. They also stated that in Europe second-generation

phlaeas tends to be duskier in color and have longer tails unlike subspecies hypophlaeas. I was not able to

find illustrations or specimens of Old World phlaeas that completely matched hypophlaeas in appearance.

Nominate phlaeas from Sweden is quite similar, as is subspecies polaris as figured by Tolman &

Lewington (1997, pi. 21), except that polaris has extensive whitish spaces distally from the postmedian

series of black spots on the ventral hind wing. Tuzov (2000) figures specimens on pi. 57, p. 337, from the

Chita Region and Altais, Russia under the name L. p. hypophlaeas , both spring and summer generations.

This is a very strange location for something conspecific with the eastern North American population.

There are no intermediate populations. Perhaps it represents convergence rather than conspecificity (per.

corresp., David Wright, 2007). From the dorsal aspect, these Russian specimens look similar to eastern

United States hypophlaeas , and the forewings are not dusky in the 2 nd generation, nor is there any evidence

of long hind wing tails. Ventrally, the hind wing ground color is gray, very similar to Nearctic

15

hypophlaeas , though the black spots are not quite as large or distinct as in U.S. hypophlaeas. There are Old

World populations of L. phlaeas that are similar-enough appearing to U.S. hypophlaeas, that this name has

been applied by some authors (Tuzov, 2000), as well as by Wolly Dod (1907), who stated that “In the

Staudinger Catalogue, Lapland, northern Scandinavia, Sajan-Geibel (Siberia), Amur and North America

are quoted as localities for ‘var. hypophlaeas ’, and some that I have bearing labels of some of those Old

World localities would pass anywhere as North American specimens, amongst which there is also an

occasional tendency to lose the spots, and so assimilate the typical European form”. Some of the driving

force for theories that eastern North American populations were introduced is the use of the name

hypophlaeas for Old World populations that has persisted through the years. Ford (1924) in discussing

hypophlaeas said, “Not only does it occur throughout the Nearctic Region, but it has an extended range in

Arctic Europe and Asia. There is a specimen from Siberia in the Hill Museum, Witley, and two from

Amurland in the Natural History Museum, South Kensington, while Staudinger also refers to specimens

from the later country. There can be little doubt that this form will ultimately be found distributed along

the north coast of Asiatic and European Russia, for it is known to occur in Lapland; there is a specimen

from this locality in the Tring Zoological Museum (Plate LIV, fig. 21), another in the Hill Museum,

together with one labeled ‘Norway’,which although it has no other data, must almost certainly have come

from the extreme north-east of that country”. Ford (1924, p. 739) then described subspecies hyperborea

from arctic Norway and Lapland, saying that it was not found in Siberia or North America. He stated that

specimens of hyperborea are far more frequent in collections than are Palaearctic examples of

hypophlaeas , and that some confusion exists in the literature dealing with the Far Northern races of

phlaeas. Although some individuals of some Old World populations of L. phlaeas are quite similar to

North American hypophlaeas , none match completely the description as translated from the French [from

Boisduval 1852] by Ford (1924), “Very near phlaeas , but smaller, with the spots more distinct, the wings

more rounded. The under side of secondaries of an ashy whiteness, with the fulvous marginal band well

marked”. I am of the opinion that the name hypophlaeas should not be used for any of the Old World

populations of L. phlaeas , and that currently there is no conclusive evidence that the North American

populations were introduced from Europe. Pratt & Wright (2002) presented an alternate hypothesis to an

introduction, positing that the eastern North American populations of hypophlaeas existed endemically in

the high elevations of the White Mountains in New England and expanded their range with the introduction

of Rumex acetosella. They said, “An expansion of this sort has been observed with alpine populations of L.

cupreus and L. editha. Both of these species have broadened their range with the introduction of Rumex

acetosella into western North America. Also high altitude California L. phlaeas from 12,000 feet elevation

can be experimentally reared on Rumex crispus at 800 feet elevation (and lower), suggesting that the

species has the ability to rapidly adapt to lowland conditions. Oxyria digyna is the primary host plant of

arctic-alpine L. phlaeas in North America. This plant occurs locally at high elevations on Mount

Washington in New Hampshire; the possible existence of high altitude L. phlaeas colonies there and

elsewhere in New England has not been studied”. If in the future, such colonies are discovered, it will

certainly be a valuable key in solving the introduction question. For the present, I am not able to answer

this question with certainty.

Do North American populations of Lycaena phlaeas represent a separate species? Evidence to

support a single widespread phlaeas species in the Old and New World is available in the literature. Maeki

& Remington (1960) showed the haploid chromosome number (n = 24) is the same for three subspecies of

L. phlaeas from the Palaearctic (Japan, Finland) and the Nearctic (United States). There is at least as much

adult phenotypic diversity among Old World subspecies as there is between the nominate phlaeas and New

World subspecies. Even the most phenotypically divergent Old World subspecies, phlaeoides , chinensis ,

matsumuranus, and daimio are still generally treated as phlaeas subspecies. Genitalic studies also suggest

conspecificity between Old and New World populations. Yet these facts may be inconclusive. Many

lycaenid complexes have multiple species with identical genitalia and chromosome numbers (pers. com.,

David Wright, 2007). Keilland (1990) in his recent treatment of the three east African taxa elevated

16

abbottii from a subspecies of phlaeas to full species. The conclusion reached in the present study is that

no real evidence exists to contradict the traditional placement of North American subspecies with Old

World phlaeas. Perhaps future molecular studies will shed more light on how many species are involved.

ACKNOWLEDGMENTS

Appreciation is given to David Dyer, Collections Manager, University of Montana Herbarium, Missoula for

access to the plant collection, and to Peter Lesica, Assistant Curator, for identification of the larval foodplants of

weberi and arctodon, as well as the foodplants for many other Montana butterflies. Appreciation is also given to

Clyde Gillette, Salt Lake City, Utah for information regarding the Utah, Wyoming and Montana L. phlaeas

populations. Harry Pavulaan, David Wright and Norbert Kondla assisted with the layout and review of this paper.

Special gratitude is given to Byron Weber, Missoula, Montana for freely sharing specimens of L. p. weberi , as

well as many other Montana butterflies, and information about the Sweet Grass Hills, and for being a pleasant

companion on hikes to the summits of the East and West Buttes of the Sweet Grass Hills.

REFERENCES

Ackery, P.R., C.R. Smith & R.I. Vane-Wright (editors). 1995. Carcasson’s African Butterflies An Annotated

Catalog of the Papilionoidea and Hesperioidea of the Afrotropical Region. Natural History Museum (London),

CSIRO Publications, Victoria, Australia, 803 pp.

Allen, T.J. 1997. The Butterflies of West Virginia and Their Caterpillars. University of Pittsburgh Press,

Pittsburgh, Pennsylvania., 388 pp.

Bridges, C.A. 1988. Catalogue of Lycaenidae & Riodinidae (Lepidoptera: Rhopalocera). Published by Author,

Urbana, Illinois, 782 pp.

Comstock, W.P. & E.I. Huntington. 1960. An annotated list of the Lycaenidae (Lepidoptera, Rhopalocera) of the

Western Hemisphere. J. New York Ent. Soc. 68:176-186.

D’Abrera, B. 1980. Butterflies of the Afrotropical Region. Lansdowne Editions, East Melbourne, Australia, 593 pp.

Ehrlich, P.R. & A.H. Ehrlich. 1961. How to Know the Butterflies. Wm. C. Brown Company Publishers, Dubuque,

Iowa, 262 pp.

Elrod, M.J. 1906. The Butterflies of Montana. Univ. of Montana Bull. No. 30, Biol. Ser. No. 10, 174 pp.

Emmel, J.F., T.C. Emmel & S.O. Mattoon. 1998. The types of California butterflies named by Jean Alphonse

Boisduval: designation of lectotypes and a neotype, and fixation of type localities, pp. 3-76, In Emmel, T.C. (editor).

Systematics of Western North American Butterflies. Mariposa Press, Gainsville, Florida, 878 pp.

Emmel, J.F. & G.F. Pratt. 1998. New subspecies of Lycaeninae from California and a type locality restriction for

Chrysophanus cupreus W.H. Edwards (Lepidoptera: Lycaenidae). pp. 661-680, In Emmel, T.C. (editor).

Systematics of Western North American Butterflies. Mariposa Press, Gainsville, Florida, 878 pp.

Ferris, C.D. 1974. Distribution of arctic-alpine Lycaena phlaeas L. (Lycaenidae) in North America with designation

of a new subspecies. Bulletin of the Allyn Museum (18):1-13.

Ferris, C.D. & F.M. Brown (editors). 1981. Butterflies of the Rocky Mountain States. Univ. of Oklahoma Press,

Norman, Oklahoma, 442 pp.

Ford, E.B. 1924. The geographical races of Heodesphlaeas , L. Trans, ent. Soc. Lond. 71(3/4):692-743.

1 ( 2 ): 5 ,

s, A.B. 1951. A Field Guide to the B

i, K. & C.L. R

me, G.M. 20C

Moss, E.H. 1983. Flora of Alberta (2 nd ed.). Univ. of Toronto Press

Opler, P.A. & G.O. Krizek. 1984. Butterflies East of the Great P

Hopkins Univ. Press,

Opler, P.A. & V. Ma

Pratt, G.F. & D.M. ^

. 1992. A Field Guide to E;

No. 1:

(INHELP) 1977,

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Volume 7, Number 2

12 June 2009

The Taxonomic Report

OF THE INTERNATIONAL LEPIDOPTERA SURVEY

Natural Life Histories of Alaska Colias (Lepidoptera: Pieridae)

Jack L. Harry

47 San Rafael Court, West Jordan, Utah, USA 84088

occasionally observed foraging in the area. The female was known by regular caribou hunters as being very good

about leaving people alone. The first day in camp in 2006 they came into camp to welcome me back. When they

were discovered in camp they were only 20 feet away. When the mother observed me she just took a quick curious

look and then began to wander away. Since she is so non-threatening it was probably good to have her in the area, as

she probably kept the males away while she had the cubs. She was not observed in 2007.

Climate and Weather

The weather information herein is mostly temperature data, but a few remarks are made about precipitation.

While in camp the daily high and low temperatures were recorded with my own thermometer. Naturally, some days

the exact high or low temperatures were missed, but the temperature that was obtained would be a good

representation for that day. Also at the campsite is a U.S.G.S. gauging station with a thermometer. Temperature data

was obtained for the years previous to the study, 1997 to 2004. Temperature data prior to 1997 and for the year 2000

was not available. Dan Bartu with the U.S.G.S. in Fairbanks obtained the data. Seven years of data is not sufficient

to establish an accurate daily average, but it does at least provide some idea of the normal temperature. Temperature

data was also obtained for May and June of 2005 and the last half of August when the camp was not occupied. The

U.S.G.S. thermometer was acting erratically in 2006 so temperature data for the last half of July and all of August

was not obtained. A synopsis of the temperatures is given in Table 1 (page 15).

The spring and summer of 2005 were very dry. It appeared obvious that conditions were dry upon arrival at the

campsite in June of 2005. Also, comments from Alyeska personnel indicated the spring had been dry. From June 25

to 29 of 2005 the weather was very warm (24-26°C) and sunny. Then on June 30 a north wind of 8-12 kts started

and continued until August 2. This wind caused July to be unusually cold. The first 20 days of July were 3.6°C

below average. The last ten days of July were nearly normal, then the first half of August was pleasant. Very little

rain occurred during July and August.

In 2006 the temperature from 21 May to 13 July was quite normal. When I arrived at the campsite on May 20 of

2006 the snow had just melted from the flat areas. There were still minor snowdrifts beside the pipeline, etc. There

was occasional snowfall until June 9. Thereafter there was much rain and limited sunshine until July 9. During the

first half of July the mountains to the south (Brooks Range) were almost constantly cloud covered and there was

In 2007 the first ten days of June were sunny and exceptionally warm. During that ten-day period the daily high

temperature averaged an incredible 10.1°C above average. After that there was a spell of three cool days then the

weather was warm for the rest of June.

By comparison, in 1971 there was a cold spell around the first of July that lasted for a couple of days. In 1991

there was no cold spell from June 21 to July 15, and there were only two non-collecting days during that period. The

primary flight and egg maturing time frame for Colias is June 20 through July. It is common that there is a cold spell

(even with snow) during this period but the spells normally last only two or three days then the temperature returns to

normal. The first 20 days of July 2005 were exceptionally cold which delayed egg development and caused many to

die.

Eggs and Plants

In 1991 and 1999 there was an extremely good butterfly flight (all species) in the research area and in the

Franklin Bluffs area. The Franklin Bluffs construction camp site was at Mile 378 Dalton Hwy, which is 37 miles (59

km) south of Deadhorse and on the coastal plain. From 2000 to 2003 the butterfly flight was moderate to poor. In

2005 and 2006 the butterfly flight at the study site was extremely poor. Except for Boloria frigga, Colias , and the

blues there were almost no butterflies and these taxa were very few. It is incredible that the butterfly population

could be so depauperate. In 2005 there were a few individuals flying as late as August 5 (including C. philodice and

C. gigantea ), this was probably because of the cool weather in July. In 2005 the flight at Franklin Bluffs was poor (C.

hecla was still flying on August 10) and in 2006 it was moderate, except that Colias boothii had an abundant flight.

At the research site in 2005 there was just barely enough Colias to be able to start the research project, except boothii

of which only one female was obtained. Because of the extremely warm first ten days of June in 2007 the flight of

most species was very early. However, the Colias flight was only a few days early. There was an extremely

abundant flight of all Satyrids and B. frigga. By June 24 there were only a few of these still flying and most of them

were worn. The abundance of other taxa was still low.

We always wonder why a particular year had a poor flight, yet another year an abundant flight. Sometimes it

seems obvious that weather had been a major factor, either positive or negative. Sometimes predators or parasites

2

legume feeding^o&w fly more abundantly along the pipeline than in the natural habitat. This may be because there

ix in wet areas but were n<

On May 23 and 24 of 2006 a few of the legume plants had already started to grow. As soon as there was growth

the larvae started eating. At two stations there were too many larvae so the new growth was entirely eaten, so the

larvae had to be moved to another plant. Some plants did not grow at all, so the larvae there had to be moved to a

growing plant. The Vaccinium and Salix are the last to start growing leaves. In 2006 the Salix did not start to grow

leaves until June 10, and many of the Vaccinium plants had buds starting to swell on June 5.

Larval Behavior

Colias larvae almost entirely eat the leaves of the hostplants. Rarely one was observed eating the buds or flowers

and even then there was very little eating of the buds or flowers. In general, the first through third instar larvae eat

the mesophyll of the leaf and leave the membrane and veins. This varies somewhat, especially with the third instars

which sometime eat the entire leaf. This also varies with the different plant species and the age of the growth. The

fresh growth in the spring is eaten entirely even by the first instars. The mature leaves of Salix and Vaccinium are the

most difficult to eat, so larvae smaller than fourth instar never eat the entire mature leaf.

The most common resting place for the first through third instar larvae is on the upper side of the leaf along the

mid-vein. However, this varies considerably in all species. Sometimes they rest on the underside of the leaf, along

the petiole, and occasionally along the stem. Fourth and fifth instar larvae that are eating Hedysarum, Vaccinium ,

and Salix stay on the plant and rest anywhere, while those eating Astragalus and Oxytropis like to rest on the ground.

These Colias larvae overwinter in various instars. The overwintering stages are discussed with each species. There

is no evidence anywhere in the world that any Colias overwinters as a pupa.

Very soon after the snow has melted the larvae break diapause and become visible while they are waiting for

their hostplant to start growing. Some plants started growing very soon while other took a couple of weeks. At a few

stations the plant never did grow, which required that the larvae be moved to a new plant. It was quite a nuisance to

have to find the first instar larvae so that they could be moved. It was rather amazing how long the first instar larvae

remained alive without eating. As time elapsed it became a concern that they would start to die. At three weeks

time, without any plant to eat, as many larvae as could be found were moved to a new plant. Every day or two the

old site would be visited and sometimes more larvae were found. At 27 days a few larvae were found still alive.

Mature fifth instar larvae leave the hostplant and wander about to find a place to pupate. This is usually near the

ground on a stem. Those confined under a sleeve would usually pupate on the netting near the ground.

In discussions concerning the elapsed time of development from egg to adult the followng terminology is used.

The first summer - this is the summer in which the egg was laid. The second summer is one year of elapsed time

from egg. The third summer is two years of elapsed time from egg.

Materials and Methods

Net sleeves were placed over the larval hostplants at numerous stations within the study sites. These sleeves

were to sequester the females for ovipositing on the larval hostplant and then left in place to protect the larvae. The

netting was green polyester netting that BioQuip Products sells for butterfly nets. The netting was supported by two

9-gauge wires. The wires were bent in U shape so that the sleeves would fit over them, and the ends were pushed

into the ground. The two wires were placed at 90° to each other to hold the sleeve in place. The bottom of the sleeve

was secured by rocks. The rocks had to be placed touching each other to provide a seal to keep predators out. Since

the sleeves (stations) are out in nature they are subject to being damaged or destroyed so several stations were

established for each species.

The stations were checked every 2 or 3 days and reset when they had been disturbed. When it became apparent

that the ova were about to hatch they would be monitored daily to find out when the ova hatched. Thereafter, the

contents would be checked occasionally to monitor the progress of the larvae. In 2005 the sleeves were removed on

August 30 for the winter and in 2006 the sleeves were removed on September 8. The sleeves were reset on May 24

in 2006 and on June 2 in 2007. Some of the data of the immature stages was obtained at home during the five years

prior to 2005. During this project some of the data was obtained in camp and some at home. The larvae that were

reared at home were put on potted plants or on cut stems that were placed in water. The potted plants were covered

with net sleeves; the cut stems were placed in a plastic container or a bucket with a netting cover. When the larvae

stopped eating and were preparing to molt they would be placed in a separate container (still on the leaf). First they

would be measured for length and then left to molt. After molting they would be returned to the hostplant and the

molted head capsule would be measured. It is rather difficult in the field to monitor the larvae to see when they are

getting ready to molt. However, some of the third instar, all of the fourth and fifth instar larvae of hecla, nastes, and

4

philodice were measured in the field. The first and second instar nastes and palaeno that were used for

measurements were reared in closed plastic containers in camp. Colias larvae that are reared in a closed container

eventually die (usually third instar), so these were sacrificed.

The larvae were left under the sleeves to pupate. Pupae were collected so that they could be measured. Then the

pupae were placed under a sleeve to pupate in natural conditions. Pupae not needed for measurements were put in a

communal sleeve near camp for easy monitoring.

Larvae that were reared in the lab were overwintered in the refrigerator. A large plastic container, with the lid

on, was used. Four small containers with water for humidity were set in the large container. The larvae were put in

small plastic containers that have numerous holes and these containers were put in the large container. No plant

material was put in the container with the larvae, since plant material would mold.

Morphological observations and measurements of the eggs, first instar larvae, and all the head capsules were

made with the aid of a stereomicroscope and a 0.1 mm scale. Length measurements of the second instar and larger

larvae and pupae were made with a mm scale and the aid of 3X reading glasses. Measurements of the head capsule

width are of the molted head capsule. Measurements of the length of the larvae were made at pre-molt stage. Length

of the fifth instar larvae is given for the mature larva during the most common resting position . Their length during

resting varies so this measurement is somewhat nebulous. N=10 for all measurements unless otherwise noted. Pupal

width is the width of the pupa from side to side. Pupal height is the back to the outside edge of the wing cases.

General Description of Colias Immature Stages

Eggs: Fig. 30. The eggs are typical of the Colias. The eggs are fusiform in shape with longitudinal ribs and small

transverse ridges. The top is rounded and contains the micropyle. Eggs are creamy white when oviposited and

become orange with a creamy tip within 4 or 5 days in the field (2 days in the lab). Prior to hatching the eggs exhibit

a black tip, the head of the larvae can be seen through the shell. Eggs hatch in 5 days at room temperature but vary in

nature as mentioned above. The size is given under each species.

Larvae: All Colias have five larval instars. The first instar larvae are similar and are described here rather than

repeating under each species. Any exceptions are given with the species.

First instar: Head is black with tiny white hairs. Body is green with dark green mid-dorsal stripe. On each side of

body there are 3 white hairs on each segment except the first segment which has 5 white hairs. The lateral line is

cream colored and there is a cream colored ring around the base of the hairs. Occasionally a few larvae have black

hairs on the head and a few on the body. There are many black dots on a yellow body under 20X magnification. The

spiracles are black, and the thoracic legs are dark green to black. There is a dark area above the anus on the majority

of the larvae but not all of them.

Second to fifth instar: These are similar so only the fifth instar is described. The size of each instar is given with

each species. The following characters are similar in all species. The head and body are green with many black

spots with give rise to black or white hairs. The spiracles are white with a black ring. Sometimes the spiracles exhibit

the color of the lower half of the lateral stipe. The dorsal stripe is dark green and the eyes are black. The other

features or variations of these features are described with each species.

Pupae: Fig. 31. The pupae of all species are similar so they are described here, exceptions are given with the

species. Head is green with front darker green, light green horizontal line in middle. Body is green, lighter green

posterior. The dorsal stripe is dark green. The sub-dorsal stripes are light green and faint. The lateral stripes are light

green or yellow and faintly expressed on wing cases. The spiracles are light green. The sub-lateral stripe is brown

on three segments immediately behind wings, there is a small black spot between lateral stripe and sub-lateral on two

segments immediately behind wings. The posterior end of the pupa is attached to a silk pad on the substrate and the

body is held loosely to the substrate with a silk girdle.

The larvae that died in the lab were diseased. They probably became diseased during the shipment to Salt Lake

City. Colias larvae become diseased easily when they have been in a closed container especially when there are

many in a container.

Colias gigantea inupiat

at north slope research site

Fig. 22

Eggs: Length 1.42 mm (range 1.22 to 1.56), width 0.55 mm (range 0.49 to 0.61).

Larvae:

First instar: Length 3.1 mm (range 2.7 to 3.2), head width 0.38 mm (range 0.36 to 0.40).

Second instar: Length 5.0 mm (range 4.5 to 5.5), head width 0.56 mm (range 0.50 to 0.65).

Third instar: Length 8.5 mm (range 7.1 to 10.5), head width 0.95 mm (range 0.78 to 1.25).

Fourth instar: Length 14.5 mm (range 13.8 to 15.2) head width 1.5 mm (range 1.46 to 1.65).

Fifth instar: Only black hairs have been observed except that sometimes the hairs are white on the lower half of

head and body. The sub-dorsal stripes are yellow and faint. The lateral stripe is white. Length 24mm, head width

2.4 mm (range 2.28 to 2.62).

Pupa: Length 19.7 mm (range 18.5 to 21.0), width 5.0 mm (range 4.5 to 5.5), height 6.3 mm (range 5.5 to 7.0).

Results

C. gigantea females have been observed ovipositing on Salix lanata L. (Salicaceae). They probably also use

other brushy willows. The larvae were reared on S. lanata at the research site. They were reared on Salix exigua

Nutt, in the lab.

C. gigantea was the last Colias to start flying in 2005. This was because none had started flying before the cold

spell started on June 30. From July 7 to Julyl5 five females were collected, which did not lay any eggs. Three

females were collected on July 21; these laid only four eggs. A few more females and some eggs were obtained in

the next few days. The results at the three stations that eventually had larvae are given next.

Station #36: On July 26 and 27 a few eggs were laid. On Aug 28 six larvae were observed. One of these, a

second instar, was still eating. In the spring of 2006 this station was submerged and no larvae survived.

Station #37: On July 24 about 12 eggs were laid, these eggs hatched on August 10. No larvae were observed

when the sleeve was removed on August 30, they were probably in hibernation in liter at the base of the plant. In the

spring of 2006 the station was slightly submerged, but on June 18 four larvae (first and second instar) were observed.

On July 4 four larvae (one third and 3 fourths were observed. These were left in the sleeve when the camp was

abandoned on July 13. That was unfortunate because on September 8 when the sleeve was removed, the plant,

ground, and sleeve were covered with dust (from nearby road construction) so no larvae were observed.

Station #38: On August 10 of 2005 there were 12 larvae at this station. On August 30 no larvae were observed

so they were in hibernation. On June 14 of 2006 three larvae (second instar) were observed. No larvae were

observed on Sept. 8.

During early June of 2007 one fourth instar at Station #37 and one fourth instar at Station #38 were found. Both

of these larvae became adults in late June.

On June 28 of 2007 one female was collected and about 70 eggs were obtained. These were reared in the lab

under constant light and one became an adult on Aug. 9. All of the other larvae diapaused as fourth instar. During

July of 2006 approximately 70 to 100 eggs were obtained in three different sleeves. In 2007 there were no larvae in

these three sleeves.

Discussion

This taxon had the poorest results of all of the Colias on the north slope. There were 16 known larvae which

resulted in only two larvae in 2007. Therefore all of the known larvae completed development the third summer.

Colias hecla glacialis

at north slope research site

Fig. 18

Eggs: Length 1.35 mm (range 1.30 to 1.44), width 0.55 mm (range 0.50 to 0.57).

Larvae:

First instar: Length 3.0 mm (range 2.8 to 3.4), head width 0.35 mm (range 0.31 to 0.38).

Second instar: Length 4.7 mm (range 4.3 to 5.0), head width 0.53mm (range 0.51 to 0.56).

Third instar: Length (N=6) 7.5 mm (range 7.2 to 8.0), head width (N=7) 0.83 mm (range 0.78 to 0.88).

Fourth instar: Length 13.7 mm (range 13.0 to 16.0), head width 1.33 mm (range 1.25 to 1.50).

Fifth instar: Only black hairs have been observed. The sub-dorsal stripes are yellow. There is a black patch on

the lateral side of sub-dorsal stripes on each segment. The lateral stripe is white. Length 24 mm, head width 2.3 mm

(range 2.18 to 2.48).

Pupae: There are a few faint dark streaks on wing cases. Length 18.3 mm (range 17.0 to 20.0), width 5.0 mm

(range 4.5 to 5.5), height 6.0 mm (range 5.2 to 6.2).

Results

Females have been observed ovipositing on Astragalus arcticus and Hedysarum mackenziei. Many more

observations have been on A. arcticus than on H mackenziei , so it appears that Hedysarum is used only occasionally.

The larvae were reared on arcticus and mackenziei at the research site and Astragalus cicer in the lab.

Only four females were collected in June of 2005. These females were put in two stations. Many eggs were

oviposited at both stations. During July it appeared that very few, if any, of the eggs hatched. From July 15 to

August 2 eight more stations were established for females to oviposit. Some of the females were taken at the

research site and some at Franklin Bluffs. The next spring approximately 45 post-diapause larvae were found among

all the stations. The plants at five of the stations did not grow. Larvae that could be found at these stations were

moved to another station.

On August 29 of 2005 five first instar and two second instar larvae were observed. The first instars were still

eating and the second instars were in hibernation. On June 12 of 2006 there were first, second, third, and fourth

instar larvae found. The fourth instar may have overwintered as third instar. As late as June 18 some first, second,

and third instar larvae were found at a station where the plant did not grow. It is amazing how long the first instar

can survive while waiting for some food. On June 24 the first pupae was attained, and on July 11 this pupa emerged.

All together there were 7 hecla (lm, 6f) that became adults in 2006. At one station on July 4 there were two fourth

instar and on July 7 they appeared to have diapaused. During June of 2007 only three larvae were found of the 25

that were known in July of 2006. All three of these larvae became adults in June of 2007.

During early July of 2006 three new stations were established with females for oviposition. There were no eggs

at one station but many eggs at the other two stations. On June 8 of 2007 there were two fourth instar and six fifth

instar larvae. All of these larvae became adults in June of 2007.

Discussion

Some larvae had definitely overwintered as first, second, and third instar. Most of the first instars would have

had plenty of time to attain second instar but apparently they wanted to remain first instars.

Of the larvae started in 2005 seven became adults in 2006 which means that they completed development in one

year. All of the larvae (only 3) in 2007 became adults in June. Of the many eggs that were started in 2006 only three

resulted in larvae. These three larvae became adults in 2007.

Colias nastes alias ka

at north slope research site

Figs. 21, 43

Egg: Length 1.30 mm (range 1.21 to 1.36), width 0.53 mm (range 0.50 to 0.57).

Larvae:

First instar: (N=6) Length 3.1 mm (range 2.7 to 3.4), head width 0.35 mm (range 0.33 to 0.38).

Second instar: Length 4.5 mm (range 4.0 to 4.8), head width 0.50 mm (range 0.49 to 0.53).

Third instar: Length 7.6 mm (range 6.7 to 9.0), head width 0.82 mm (range 0.75 to 0.90).

Fourth instar: Length 12.4 mm (range 9.5 to 13.8), head width 1.32 mm (range 1.12 to 1.44).

Fifth instar: The description is same as hecla. Length 23 mm, head width 2.3 mm (range 2.15 to 2.38).

Pupa: There are a few dark streaks on wing cases. Length 17.3 mm (range 16.0 to 18.0), width 5.0 mm (range 4.8 to

5.0), height 6.0 mm (range 5.8 to 6.5).

Results

Females have been observed ovipositing on Oxytropis borealis DC (Fabaceae). There has been no evidence that

females have been interested in ovipositing on any other species of plant. Larvae were reared on Oxytropis borealis

at the research site. In the lab first instar larvae would eat Thermopsis montana but did not grow and eventually died,

and they would not eat Astragalus cicer.

In June of 2005 two stations were established with females for ovipositing. One of these stations was a potted

plant. Approximately 30 ova were obtained on the potted plant and 40 to 50 ova at the other station (#15). None of

the eggs on the potted plant hatched and only 6 larvae were the most ever observed at #15. More results at this

station are discussed because the eggs were laid early and the larvae grew slowly. On August 28 six first instar

larvae were observed. On May 23 of 2006, when the sleeve was installed, two first instar larvae were observed and

they were apparently eating. On June 8 they were still first instar. On September 8 five third instar larvae were

observed. In June of 2007 nine pupae were taken from this station, so there was apparently no loss of larvae at this

station.

From July 15 to 25 of 2005 four more stations and one on August 2 were established for females to oviposit.

Many eggs were obtained and eventually over 130 larvae. On August 29 five first instar larvae were observed in

hibernation. On May 24 of 2006 one third instar larvae was observed which was already eating. On June 13 there

were first, second, third, and fourth instar larvae. The first pupa was attained on June 22 and the first adult on July 9.

All together there were 10 nastes (2m, 8f) that became adults in 2006.

During July of 2006 there were 134 larvae accounted for. During June of 2007 only 23 larvae were accounted for

and all of these became adults in June of 2007. In early July of 2006 three new stations were established for

ovipositing and many more ova were obtained. In the middle of June of 2007 55 larvae were accounted for. Only

four of them did not become an adult in 2007. One of these three stations had no larvae in 2007.

Discussion

A few of the individuals (10) that were started in 2005 completed development in one year. All of the rest (that

were found) completed development in two years. Nearly all of the individuals that were started in 2006 completed

development in one year.

Colins palaeno chippewa

at north slope research site

Figs. 19,31,44, 48, 49

Egg: Length 1.37 mm (range 1.25 to 1.53), width 0.49 mm (range 0.45 to 0.54).

Larvae:

First instar: Length 3.1 mm (range 3.0 to 3.3), head width 0.30 mm (range 0.28 to 0.33).

Second instar: Length 5.0 mm (range 4.1 to 5.5), head width 0.53 mm (range 0.46 to 0.59).

Third instar: (N=4) Length 6.9 mm (range 6.3 to 7.4), head width 0.85 mm (range 0.78 to 0.90).

Fourth instar: Length 12.4 mm (range 11.2 to 13.2). head width 1.22 mm (range 1.13 to 1.31).

Fifth instar: Only black hairs have been observed except that sometimes the hairs are white on lower half of

head and body. There are no dorsal or sub-dorsal stripes. The lateral stripes are yellow. Length 23 mm, head width

2.1mm (2.00 to 2.17).

Pupa: There are no sub-dorsal stripes and the sub-lateral stripes are faint. Length 17.5 mm (range 16.5 to 18.5),

width 4.7 mm (range 4.5 to 5.0), height 6.0 mm (range 5.5 to 7.0).

Results

Females have been observed ovipositing on Vaccinium uliginosum L. (Ericaceae). The larvae at the research site

were reared on V uliginosum. The larvae were reared on V. caespitosum Michx. in the lab.

From June 29 to July 21 of 2005 five stations were established for females to oviposit and many eggs were

obtained. At one station on August 28 there were some first and second instar larvae still eating. On June 17 of 2006

9

2005 and 2006.

h 1.25 mm (range 1.13 to 1.32), w

First instar: Length 3.5 mm (range 3.10 to 3.80), head width 0.35 mm (range 0.32 to 0.38).

Second instar: Length 5.1 mm (range 4.8 to 5.4), head width 0.55 mm (range 0.50 to 0.57).

Third instar: Length 8.2 mm (range 7.2 to 10.8), head width 0.88 mm (range 0.79 to 0.93).

Fourth instar: Length 15.9 mm (range 14.2 to 17.0), head width 1.58 mm (range 1.47 to 1.82).

Pupa: Length 20.4 mm (range 18.5 to 21.0), width 5.3 mm (r

£ 5.0 to 5.5), height 6.8 mm (range 6.5 to 7.0).

^ It was demonstrated that C. philodice completes development in one year.

>7 mm (range 1.32 to 1.45), w

t instar: Length 3.6 mm (range 3.50 to 3.80), head width 0.35 mm (range 0.32 mm to 0.39).

nd instar: Length 5.4 mm (range 5.2 to 5.8), head width 0.55 mm (range 0.47 to 0.60).

d instar: Length 8.9 mm (range 8.0 to 9.8), head width 0.90 mm (range 0.80 to 0.96).

h 16.1 mm (range 14.8 to 18.0), h<

Fifth instar: Some larvae have black hairs and some have white hairs. The sub-dorsal stripes are yellow and on

some larvae they are bordered by orange. A few larvae do not express sub-dorsal stripes. The lateral stripes are

white with red on ventral side. Length 33 mm, head width 2.6 mm (range 2.30 to 2.75).

Pupa: Length 22.4 mm (range 21.0 to 24.0), width 5.7 mm (range 5.0 to 6.0), height 7.0 mm (range 6.5 to 7.5).

Results

Females were observed ovipositing on Hedysarum mackenziei and Astragalus alpinus. Larvae were reared on

mackenziei at the research site and on H boreale, A. cicer, and Thermopsis montana in the lab. On June 17 of 2005

the station was established with two females for ovipositing. The researcher had to travel on to the north slope so it

was not known until August 30 if there were any larvae. On August 30 three second instar larvae in hibernation were

observed. At the end of September the sleeve was removed for the winter by a Federal Aviation Employee (FAA) at

the FSS. On May 15 of 2006 the sleeve was replaced. At this time the plant had not begun to grow. One larva was

observed which was waiting for food. The station was revisited on 15 July and three dead Christina adults were

Discussion

The larvae that were used for measurements were reared in the lab during 1999, 2000, and 2007. It was

demonstrated that C. Christina completes development in one year. Three individuals do not make a thorough

scientific study but they indicate that it is normal for C. Christina to complete development in one year.

Colias Christina X canadensis

at Northway

Figs. 14, 15

On June 10 of 1999 a female C. canadensis was collected at this location. Eggs were obtained from this female

and the larvae were reared to diapause by Ken Hansen. The post diapause larvae were reared by Ken Hansen, Jacque

Wolfe, and J. Harry. 16 adults were obtained and all of these adults display characters that demonstrate they are

hybrids of C. Christina and C. canadensis. The fifth instar larvae and adults are described.

Fifth instar: Like Christina with red along the stripes.

Adults, males: Upper surfaces ground color is orange with the orange completely covering the wing as in

canadensis. The dark border is wide as in C. Christina. Undersurfaces appear to be a combination of the two species.

Size varies from C. canadensis to C. Christina. The males that are the size of C. Christina are the largest all orange

Colias from Alaska, since the orange does not cover the entire surface of Christina.

Adults, female: Variable like C. canadensis and C. Christina but large like C. Christina. Undersurfaces appear to be

a combination of the two species.

Colias gigantea gigantea

at Northway

Figs. 39-41, 47

Egg: Length 1.44 mm (range 1.32 to 1.65), width 0.49 mm (range 0.46 to 0.53). The SEM pictures of the micropyle

demonstrate that different eggs of the same species have a different number of cells around the micropyle.

First instar: Length 3.6 mm (range 3.5 to 3.8), head width 0.37 mm (range 0.35 to 0.38).

Second instar: Length 5.9 mm (range 5.2 to 6.5), head width 0.59 mm (range 0.57 to 0.61).

Third instar: Length 9.3 mm (range 8.6 to 11.8), head width 0.98 mm (range 0.88 to 1.18).

Fourth instar: Length 16.5 mm (range 14.3 to 19.0), head width 1.46 mm (range 1.24 to 1.78).

Fifth instar: Only white hairs on head and body have been observed. The sub-dorsal stripes are yellow. The

lateral stripes are white with red on ventral side. Length 33 mm, head width 2.7 mm (range 2.30 to 2.84).

Pupa: Length 23.3 mm (range 22.0 to 25.0), width 6.0 mm (range 5.5 to 6.5), height 7.0 mm (range 6.0 to 8.0).

Results

Females were observed ovipositing on a Salix sp. Larvae were reared on Salix exigua in the lab.

12

Discussion

The natural life history was not determined in interior Alaska. The larvae that were used for measurements were

reared in 2003 and 2007.

Colins canadensis

at Northway

Fig. 26

Egg: Length 1.30 mm (range 1.20 to 1.40), width 0.51 mm (range 0.48 to 0.60).

Larvae:

First instar: Length 3.3 mm (range 2.9 to 3.5), head width 0.36 mm (range 0.32 to 0.37).

Second instar: Length (N=5) 5.4 mm (range 5.0 to 5.8), head width (N=6) 0.59 mm (range 0.55 to 0.61).

Third instar: Length 9.0 mm (range 8.3 to 9.7), head width 0.92 (range 0.80 to 1.00).

Fourth instar: Length 15.6 mm (range 14.5 to 17.3), head width 1.50 mm (range 1.38 to 1.65).

Fifth instar: Larvae are mostly without sub-dorsal stripes, only 4 of 52 exhibited a hint of sub-dorsal stripes.

Length 28 mm, head width 2.6 mm (range 2.25 to 2.85).

Pupa: Length 17.4 mm (range 16.5 to 19.0), width 5.25 mm (range 5.0 to 5.5), height 6.2 mm (range 5.5 to 6.5).

Results

Females have not been observed ovipositing at this location. The females readily oviposit on Lupinus arcticus S.

Wats (Fabaceae). but will not oviposit on Astragalus alpinus in captivity. Larvae were reared on ornamental lupine

(russel hybrid) and Hedysarum boreale in the lab. All larvae became mature fifth instar the first summer and then

diapaused. The larvae pupated the next spring without eating.

Discussion

The natural life history was not studied at this location. The adults start flying in early June which is 20-30 days

earlier than the other Colias.

Colias canadensis

at Mile 110 Dalton Hwy

Figs. 24, 25

This location is 0.9 km west of the Dalton Highway at Mile 110. The location is 9 km south of the Arctic Circle

at 66°29.36'N and 150°43.46'W at the elevation of 690 meters. This location is about 120 meters above treeline on a

mixed wet and dry hillside.

Results

Females have been observed ovipositing on Lupinus arcticus S. Wats. Larvae were reared on L. arcticus at the

research site, and in the lab on Hedysarum boreale and ornamental lupine (russel hybrid). On June 21 of 2005 five

stations were established with females for ovipositing. One female was put in each station. On August 14 four

second instar larvae were observed among the five stations. These were at base of the plants so they had already

diapaused even though the plants were still in good condition. On August 28 the sleeves were removed for the

winter. On May 20 of 2006 the sleeves were installed. At this time the plants had already started to grow with one

plant having grown 1.5 centimeters. On July 13 fourteen fifth instar larvae were observed. On September 8 when

the sleeves were removed sixteen fifth instar larvae were found. They all had left the plant and had hibernated on the

sleeve near the ground. Since they left the plants and tried to crawl away they did not intend to eat in the spring. If

they had been left there they would not have been found in the spring so they were taken to the lab. Five of these

larvae pupated the next spring without eating. The other larvae died during overwintering.

Discussion

The location at Mile 94 was a much better location than Mile 110. The Mile 94 location is 1.5 miles west of the

highway on a flat dry ridge. In 1999, 2001, and 2003 there was an abundant number of adults flying. In 2000 and

2002 there were only a few flying. It was apparent that C. canadensis has a two year life cycle at this location and

the research verifies this. The majority of adult males taken at this location are C. canadensis but a few C. boothii

13

have been collected. Some of the larvae are like C. canadensis at Northway, some are like C. boothii, and some are

intermediate. The two larvae in Figs. 24, 25 are from the same female. It appears that C. canadensis and C. boothi

commonly hybridize at this location or are conspecific. It was hoped to do further research at this location but the

area burned in 2004 and there were no adults flying in 2005. In June of 2001 females were sleeved on Lupinus plants

at Mile 94 Dalton Highway. When the researcher returned 11 days later the eggs were just beginning to hatch.

These larvae were reared on Hedysarum boreale and ornamental lupine in the lab.

The larvae that died during overwintering had become diseased. They probably became diseased while being

transported home.

Concluding Discussion

During the period of July 2005 to June 2006 seven C. hecla, ten C. nastes, one C. palaeno, and all (71) C.

philodice completed development. This proves that even during an unfavorable time span some individuals complete

development in one year. Then all of the remaining individuals completed development in 2007 (two years time

span). Unfortunately, there was considerable loss of larvae the second winter so the numbers were low that

completed development in two years.

Nearly all of the C. nastes and all of the C. palaeno (only one) that were started from eggs in 2006 completed

development in 2007. This demonstrates that during a favorable time span most individuals (except C. boothii) will

complete development in one year.

If a good supply of C. gigantea had been obtained it is probable that some individuals would have completed

development in one year.

There was a huge loss of larvae from Sept. 8 of 2006 to June 1 of 2007. The cause can only be speculated but all

species were affected. It would seem that the most likely cause would be the weather. Whatever the cause was it

sure didn’t inhibit the Satyrids from having an abundant flight in 2007.

The first winter larvae of all species overwinter in different instars (first, second, third). It is obvious that the

larvae do not progress at the same rate in nature. The second winter larvae of gigantea, hecla, nastes, and palaeno

overwintered as fully grown fourth instar. Anytime one of these become fifth instar they complete development

without diapausing.

During the second summer the larvae of boothi become fully grown fifth instar and then diapaused. C. boothi is

the only species on the north slope that diapauses as fifth instar. This is the reason that they fly 7-10 days earlier than

the other species. However, there are fresh boothi individuals that fly with hecla and nastes.

Colias canadensis and boothi are the only Colias in Alaska that diapause as fifth instar larvae.They both diapause

as mature fifth instar.

The larvae of some species of Colias vary in the expression of patterns and colors. In Utah some larvae of

philodice and eurytheme exhibit sub-dorsal stripes and some do not exhibit the

sub-dorsal stripes. In Alaska some larvae of Christina exhibit the sub-dorsal stripes and some do not. Also, some

Christina larvae have the orange along the sub-dorsal stripes and some do not. However, all observations indicate

that boothi is the only species in the hecla group that the larvae have the orange along the sub-dorsal stripes.

Quite often when stations were monitored there would be Arachnids on the sleeve or among the rocks at the base

of the sleeve. A few times a spider was observed in the process of eating a larvae that was resting on the sleeve.

Occasionally a dead larva which had been eaten was found stuck to the sleeve. Probably other larvae had been eaten

and the carcass fell to the ground. A few larvae were parasitized by Ichneumid wasps and Diptera. The parasites

must have stung the larvae while resting on the sleeve.

The larvae that were reared in the lab were under constant light. It is assumed that the constant light would have

no affect on the north slope larvae since they have constant light naturally. However one larvae of gigantea in 2007

went straight through to adults which was quite unexpected.

Many of the natural oviposition observations were made in years previous to the start of the study. All of the

plant species on which ovipositions have been observed were used in this research. All of the Colias species were

successfully reared to adults on all the corresponding plant species.

14

References

-sr.

: 6(3): 1-3.

Table 1. Daily high temperatui

Average

2005

*sr

2006

2007

*sr

May 11-20

2.8

May 20-31

6.1

9.1

3

11.5

103

-1.2

10.9

-0.6

21.6

10.1

June 11-20

15

16

1

16.7

1.7

18.1

1.1

June 21-30

17.1

18.6

1.5

17.4

0.3

19.1

2

July 1-10

15.9

11.6

-4.3

13

-2-9

July 11-20

18.4

15.4

-3 i

July 21-31

16.9

15.7

-0,8

17.4

0.5

Aug. 1-10

15.4

20.6

5.2

Aug. 11-20

12.1

15.6

3.5

Aug . 21-31

11.9

11.5

-0.4

17.8

5-Jun

29.2

15

Fig. 1 - C. boothii thula holotype Fig. 2 - C. boothii boothii lectotype

Mead River, Alaska, USNM Boothia Peninsula, Nunavut, USNM

Fig. 10 to 13 - female variation in C. boothii , Dalton Highway, north slope, Alaska

Figures 14 & 15 - C. canadensis X C. Christina hybrid male, Northway

Figure 14 - dorsal view Figure 15 - ventral view

16

17

18

19

20

Appendix 1

Notes on the natural life history of Papilio machaon in Alaska

Resume: The results of a brief study of the natural life history of Papilio machaon aliaska Scudder on the north

slope of Alaska in 2005 are annotated.

The site for this research on the north slope of Alaska is 92 miles (147 km) south of Deadhorse (Prudhoe Bay).

This is the same research site as that of the Colias research in this publication. The eggs and larvae of P. machaon

were found on Petasites hyperboreus Rydb. (coltsfoot) (Asteraceae) at Sagwon Hills which are 30 miles (48 km)

north of the camp site. The eggs and larvae were sleeved on hyperboreus near the camp. The materials and methods

are also the same as the Colias research.

On July 17 of 2005 eight eggs and one first instar larva were put in the first station. On July 18 six first and one

second instar larvae were put in the second station. On August 10 there was one fifth and four fourth instar larvae in

the first station. Also, on August 10 there were three fifth instar larvae in the second station. It is unknown what

happened to the missing larvae. On August 28 the sleeve was removed from the first station for the winter. At this

time there were three pupae which were attached to a dwarf birch stem near the ground. On August 28 in the second

station there was one pupa, one prepupa, and two fifth instar larvae. The pupa was attached to dwarf birch stem near

the ground. The prepupa was attached to the sleeve about 3 centimeters above the ground. The sleeve was reset to

another plant with the prepupa and the two larvae.

On May 24 of 2006 the pupae in the first station were missing. It is probable that they were eaten by a vole. At

the second station the one pupa not under the sleeve was found. A sleeve was put over this pupa and on June 19 a

female adult emerged. There was one pupa attached to the sleeve and 2 dead larvae carcasses. The pupa that was

attached to the sleeve died or may have already been dead on May 24. The larvae did not mature and pupate before

cold weather stopped them or the food plant became inedible.

The two larvae that did not pupate, died during the winter. Although results from two larvae do not constitute a

rigorous scientific study, they indicate that P. machaon larvae cannot survive the winter. The larvae and one pupa

were under a sleeve so they may not have had proper conditions. A short cold spell during July or August with mild

freezing temperatures and snow are not unusual so they must be able to survive these conditions. This demonstrates

that P. machaon larvae pupate the same season as the eggs are laid.

21

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22

Volume 7

February 2010

Number 3

The Taxonomic Report

OF THE INTERNATIONAL LEPIDOPTERA SURVEY

TIPS ON COLLECTING AND REARING IMMATURES

OF 375 BUTTERFLY AND SKIPPER TAXA

JACQUE WOLFE

459 East 2700 South Apt 16, Salt Lake City, UT 84115

JACK HARRY

47 San Rafael Court, West Jordan, UT 84088

TODD STOUT 1

1456 North General Drive, Salt Lake City, UT 84116

ABSTRACT: Rearing techniques are discussed for 375 different butterfly and skipper taxa from Utah and

beyond.

Additional keywords: ova, larvae, pupae, over wintering, obtaining and caring for immatures

INTRODUCTION

The authors of this paper, Jacque Wolfe, Jack Harry, and Todd Stout, with contributions from Dale Nielson

have over 100 years combined experience collecting and rearing butterflies. This publication includes natural and

lab host plants. We hope that this information will help you avoid some of the mistakes and losses we have

experienced. We also hope that this publication will encourage someone who has only collected adults to give

rearing a try.

For those new to rearing we encourage starting small. Not only can rearing provide perfect specimens but

also provide knowledge regarding the life histories of butterflies, which includes how to find caterpillars or how to

entice live females to lay eggs. The advantages justify the time and effort it requires.

Another advantage of rearing is that some species, like Papilio indra and Megathymus species, are difficult

to collect as adults. Therefor, rearing them can be much easier. For example, collecting larvae or netting a single

live female can result in obtaining a nice series of perfect specimens.

Remember not to be discouraged when you have setbacks. There is a learning curve involved with rearing

that this paper will help to accelerate. Good luck.

1 Staff Member, The International Lepidoptera Survey, Herndon, VA

GENERAL METHODS OF OBTAINING IMMATURES

For most species, it is both faster and easier to collect gravid females and confine them on their host plant

to obtain ova. For larger butterflies, such as Limenitis, Adelpha , and Papilio , we use a 12 to 24-inch cubicle cage

constructed with a frame made with thin strips of wood or aluminum and covered with nylon window screen (Fig.

7). With a screen lid and no bottom it can be placed on a flat surface. For Lycaenids a 6-inch cubicle cage works

well.

Another type of container that is used for larger females is a 5-gallon plastic bucket pail with a screen top.

Our favorite container for all but the largest butterflies is a 3-quart plastic container with a chiffon top. Place a 2-

inch thick piece of foam rubber that fits tightly in the bottom. A YA -inch diameter hole is made in the center so

that a EA-inch diameter by 2 1 / 2 -inch tall water container can be inserted. These containers, which are used to hold

pills, can be purchased at many pharmacies. The foam rubber keeps the host plant from falling, therefore creating a

very portable arrangement. It can ride on the dashboard or seat of your vehicle and sit in filtered sunshine while

you are out collecting. Place a small container that has a honeywater soaked sponge in it set on the foam rubber for

females to nectar (Figs. 1-2).

The proper amount of host plant placed in the container that houses a live female is critical. There needs to

be sufficient plant but the females must also have room to fly. Most butterflies will ovipsoit under artificial light.

Use a 60 or 100-watt bulb placed about 6 inches from the container. We use a 24-hour timer and have it set for 3

hours on and then 3 hours off during the day and then switched off overnight.

Using indoor light has several advantages. First, the females won't overheat, which is always a danger

when using direct sunshine that passes through a window. Second, you can get eggs on an overcast day. If you do

use direct sunlight take precautions to prevent overheating. Place the container in partial shade or filtered sunlight

through a window. Venetian blinds can be very helpful to filter sunlight. Having 3 or 4 females in a cage or

container is ideal as they keep each other active. Sometimes a solitary female will not be very active without other

stimuli. Sometimes females become active with a change of light from overcast to sunshine and vice versa.

For some groups of butterflies large numbers of ova or larvae may be found. With a few taxa, field

collecting of immatures is preferable to searching for elusive females. Actual methods will be discussed in each

genus or species account.

Searching for eggs or larvae in the field can be advantageous for several reasons. First, it can extend the

productivity of a day of collecting in the field. Not only can you can check host plants for immatures from sunrise

until the time of day when the adults begin to fly, but also from the time of day they stop flying until dusk. At the

same time, if overcast weather keeps the adults grounded, you can still spend a full day in the field looking for

immatures. Second, if you are unable to collect adult butterflies during one of their flight periods, you can always

look for immatures either before or after their main flight depending upon the species.

For species that diapause as larvae you can collect these in the early spring. This not only gives you a

wider timeframe for having immatures, but also extends your collecting season, which can be beneficial after a long

winter. Even in the dead of winter there are collecting opportunities. For example, you could have an enjoyable

and productive day looking for Limenitis hibemacula adjacent to river courses or Hypaurotis crysalus eggs near the

buds of oak trees until the new growth of spring makes them too hard to find.

Collecting post-diapause larvae means you should have some adults without having to overwinter your

larvae. A word of caution is that many species of checkerspot larvae, if smaller than 4 th instar, may feed for an

instar and then rediapause. There are many possible explanations for this. If the larvae are somehow overcrowded

or if the quality or quantity of hostplant is somehow restricted, these checkerspot larvae can diapause until later that

spring or for another year. Also exposing the larvae to extra warm temperatures can cause re-diapause.

GENERAL REARING

Lab rearing necessitates that larvae be protected from predators, such as spiders, earwigs, etc. The only

size requirement for containers and cages is that they must be large enough to hold sufficient host plant without

crowding the larvae. For rearing in larger numbers, common 3 or 5-gallon pails or 10-gallon terrariums (Fig. 14)

can be effective. When larvae are small, many can be placed in a container, but at most 15 to 20 larger larvae.

Overcrowding must be avoided.

As stated earlier, these containers or cages must have chiffon, fine mesh, or screen covering. Chiffon is

preferable as the porous top allows light and air into the container and allows moisture to escape. This is also

essential as it keeps caterpillar frass dry. Frass should be removed from the rearing environment frequently and

deteriorating host plant should be replaced regularly. Overexposing larvae to their own wet frass or deteriorating

plant can cause disease in many varieties of butterflies and should be avoided.

Place the food plant stems into a small bottle of water through the neck of the bottle. There must be no

spaces for the larvae to crawl or fall into the water through the neck of the bottle. Wrapping the stems with the

correct amount of plastic wrap is an effective way to seal the opening. Also, an effective method to create an

understory where the larvae can hide, rest or pupate is to place sections of white paper towel loosely around the

bottle.

As stated earlier, food plant cuttings deteriorate and lose nutritional value and need to be replaced

regularly. When larvae are small do not wait until all the plant has been consumed before changing it. The amount

of time that the hostplant remains useable depends on the plant species itself. Good rules of thumb to follow is to

replace host plant cuttings every 3 days in your cage or container and obtain a fresh supply of refrigerated cuttings

about every week or so. Again, the timing on these replacements needs to be fine tuned depending upon the plant

species.

Disinfection of any cage or container is also important. A mild bleach and water mixture applied for ten

minutes is effective. Lysol or any other 2-long-chain quat, sold in an aerosol can, can also be effective. Avoid

handling the larvae as much as possible, but when necessary, a small larva can be picked up with a camel's-hair

brush dipped in water. A toothpick dipped in water or teasing needle also works.

Most larvae can be transferred by laying the section of plant that they are attached to onto a new plant.

After the larvae crawl onto the fresh plant the old pieces can be removed. Larvae that are set to molt from one

instar to the next NEVER should be forced off their host plant which is why cutting them away from the host as

described above is always advisable.

Using potted hostplant is much less labor intensive and the plant is always fresh (Fig. 15). We recommend

VA to 2-gallon pots. They are small enough for easy handling and large enough for a good supply of plant. To

contain the larvae bend two 4-foot long 9-gauge wire lengths into a U-shape. Push the ends of the wire loop into

the soil with the loops at 90 degrees to each other. Place a mesh sleeve over the wire and secure to the pot with

twine or elastic cord. 5-gallon paint strainers from home improvement stores are ideal for this.

An effective method to keeping plants watered is to place the potted plant in a plastic pan with about 2

inches of water. If you keep plants outside replace water regularly as mosquitos can reproduce in standing water.

With this method the plant is watered from the bottom. Place your plants in the sun for 3 or 4 hours a day and then

back in the lighted lab before it gets dark. If left alone for 3-4 days, the plants will be okay inside.

A similar technique to potting plants is to place a mesh or chiffon sleeve around a live branch or branches

that contain larvae. Naturally, the plant must be in a secure area. Make sure that the netting is sealed tightly

around the branch. This technique is not recommended when rearing larvae that need to be exposed to 24-hour

photoperiod or reared in a different manner. All special handling will be listed in the individual accounts.

Pupae should be placed in a container with a paper towel or cardboard lining so that emerging adults can

climb to the mesh top to expand their wings. Pupae should be regularly misted with water until emergence or until

they are put in diapausing containers to be placed under an overwintering process.

After the freshly emerged adults have expanded their wings they should be put in an individual paper-lined

container and kept in subdued light or a dark closet for VA-2 hours. If they are not crowded, 3 or 4 that emerge at

the same time can be placed in the same container. They should then be put in the refrigerator for a minimum of 24

hours (5-6 days for Euphydryas). After being refrigerated they can be put in the freezer. Specimens should be

briefly thawed and placed in glassine envelopes and stored in an airtight container and kept in the freezer. They

will stay fresh enough to spread for a few months.

OVERWINTERING TREATMENT OF DIAPAUSING IMMATURES

It is necessary to have a refrigerator that is used exclusively for overwintering immatures. Put a

thermometer inside so the temperature can be monitored. Adjust the cold until the temperature is 30-32 degrees.

We use the largest plastic storage container that will fit in our large refrigerator. We place 2-4 small plastic

containers, containing water, into the large container, which provides 100 percent humidity in the large container.

The immatures are put in plastic containers that either have small holes poked in them or simply a chiffon top.

Some loosely wadded white paper towels can be put in a container for the larvae to cling to but it is not necessary.

The immatures should be kept in the winter conditions for at least 4 months but longer is okay. After the

immatures begin diapause, during the summer, we simply put them in the containers and store in the lab until fall.

They must be misted every other day or so until put in the refrigerator.

Post-winter treatment of immatures:

Ova: Overwintering ova should only be brought out of cold conditions and exposed to room temperature

when suitable hostplant is available. Specific strategies will be discussed in the species accounts.

Larvae: After overwintering, the larvae should be placed in the rearing container, on their host, just like

the one used to rear them to diapause. You can however, until diapause is broken, have many more larvae in a

container. Until the larvae begin to feed they must be sprayed 2-3 times each 24 hours to prevent dessication. As

is true with rearing pre-diapausal larvae, when plant condition starts to deteriorate it must be changed. Keep the

larvae under 24-hour photoperiod. This procedure may be modified for larvae that were reared to diapause using a

special method. This too will be covered in their account.

Mature larvae: Mature larvae finish all feeding prior to overwintering. Therefore, they need to be treated

exactly the same way pupae are treated both before and after they pupate.

Pupae: For the first 7 days, pupae should be kept moist. Soaking with a spray bottle quite often can do

this. For less attention, cover with a few layers of white paper towel and spray until soaked. This is necessary

during the transition time from being exposed to colder temperatures to the lab with a long photoperiod. After the

first week they can just be misted 2-3 times a day. Pupae that need different handling will be covered in the genus

or species accounts.

Adults: In spite of efforts, we have not as of yet, created a suitable overwintering technique for those

groups of butterflies, i.e; Polygonia, Nymphalis, Zerene, Phoebis, Eurema, etc. that overwinter or spend the dry

season as adults.

Please see http://raisingbutterflies. org for updated content from this article, which also includes text,

photos, and videos on raising butterflies. This site is collaborative and will invite others to share their best

practices as well.

ARRANGEMENT OF SPECIES/SUBSPECIES ACCOUNTS

Host(s): Larval food plants as documented in nature by one of the authors. Documentation is defined as finding

larvae or eggs on a species of plant and rearing them on that species of plant.

Lab host(s): Plants we have used in the lab to obtain ova or rear the larvae.

Other’s host(s): Ova or larvae were collected by persons other than the three authors but were reared by one of the

authors.

Remarks: Special handling or collecting methods, behavior, etc.

Parnassius smintheus savi

Host: Sedum lanceolatum

Remarks: Post-diapause larvae are easy to see as they feed out in the open. Larvae can be seen as one walks, so

many plants can be checked in a short time. This is preferable to getting eggs from a female because you have

adults one year sooner. This is one genus where wing-caught specimens may be more desirable than reared ones

because reared females do not have a sphragis. Females have been observed ovipositing on rocks adjacent to host

Sedum. Ova hibernate.

Parnassius eversmanni thor

Remarks: In nature, the females oviposit near the probable host, which is Corydalis pauciflora. In captivity the

females readily oviposit on material near the Corydalis plants.

Pavilio

Remarks: Papilio eggs perish if kept in an airtight container. They should be kept on white paper towel in a

plastic container with a chiffon top and misted occasionally. Larvae can be transferred to the host with a camel's-

hair brush or teasing needle as they hatch, or when hatching is near the eggs can be put on the host.

Papilio machaon group females normally oviposit readily in captivity. Papilio glaucus group females are rather

fussy about ovipositing in the lab. Females that were reared on Foeniculum vulgare, after mating, will oviposit on

F. vulgare.

Pavilio coloro

Host: Thamnosma montana

Lab hosts: Foeniculum vulgare, Ruta graveolens

Remarks: Adult abundance is greatly determined by sufficient and timely moisture. The best plants to find larvae

on are those growing on the shoulder of the road or other places where the road was bulldozed, stimulating fresh

growth (or germination) of the plants. Due to run-off, available moisture is much greater here, resulting in the new

growth on which the females love to oviposit. To be sure of getting both adult color forms, it is best to obtain ova

from captive females. Since Thamnosma plants do not have much food growing on them, the lab hosts are

preferable for rearing. In Utah, black adults (form clarki) seem to increase in abundance during the summer

months. They are never common in Utah.

EmM bairdii (including ssp. oregonim)

Hosts: Artemisia dracunculus, Cymopterus duchesnensis, Lomatium grayi grayi (In years with a population

explosion, we have found larvae on Lomatium junceum ).

Lab host: Foeniculum vulgare

Remarks: Population numbers can drastically fluctuate, producing large numbers in some years and very few in

other years. Checking plants that are growing near a watercourse can be very productive where population size

warrants. Where yellow ( brucei ) and black ( bairdii ) forms fly together, obtaining eggs from captive females is a

good way of obtaining numbers of both forms. P. bairdii is multiple-brooded so the pupae usually emerge in the

EmM brevicauda bretonemk

Lab hosts: Daucus carota, Lomatium dissectum, Foeniculum vulgare

Other’s host: Lingusticum scothicum

Remarks: Keep watch on pupae for a month or so, as some may break diapause. Most larvae reared on Lomatium

dissectum will die as large fifth-instar or prepupa.

Pavilio zelicaon (including form nitra)

Hosts: Lomatium dissectum, L. grayi grayi, Zizia aptera, Sphenosciadium capitellatum

Lab host: Foeniculum vulgare

Other’s host: Musineon tenuifolium

Remarks: The surest way to get both forms, in good numbers, is to get ova from captive females. Larvae can be

Pavilio machaon aliaska

Host: Petasites sagittatus

Pavilio machaon vikei

Host: Artemisia draucunculus

Lab host: Foeniculum vulgare

Remarks: With both machaon subspecies, some larvae can be found.

Pavilio polixenes asterius

Lab host: Foeniculum vulgare

Pavilio indra indra (includes western Nebraska ssp.)

Hosts: Lomatium graveolens, Cymopterus terebinthinus

Other’s host: Musineon tenuifolium

Remarks: P. indra colonies in Salt Lake and Davis counties use Lomatium graveolens. To the north in Cache

County, they use Cymopterus terebinthinus. Papilio indra in Cache County die when reared on local Lomatium

graveolens but do fine on Lomatium graveolens from Davis or Salt Lake County. This inconsistency is puzzling.

It may have to do with the soil where it is growing. We have found that L. graveolens and C. terebinthinus may be

used to rear many indra subspecies. We have had many losses on L. graveolans and none on C. terebinthinus. The

older stalks on the periphery of each Lomatium should be discarded when gathering plant to feed larvae. The stalks

in the center are fresher and last longer. Unlike most plants, Lomatium and Cympoterus spp. will stay fresh longer

if they are not put in a tub or vase of water. Put the cuttings in a plastic garbage bag and place in an ice chest.

Refrigerated plant is safe to use for 3 days but no longer. Eggs from a female are good of course but females of all

subspecies can be hard to locate at times. Eggs and larvae can be collected in good numbers. Plants that are

growing against rocks are the most productive. Eggs and small larvae will usually be found on the plant's

periphery. Larger larvae may be hiding in the center of the plant, or off the plant entirely.

6

Pavilio indra minori

Hosts: Lomatium junceum, L. eastwoodii, L. parryi, Cymopterus terebinthinus

Lab hosts: Lomatium graveolens

Remarks: Population numbers vary greatly due to parasitism and rainfall. Keep pupae under long photoperiod for

a month or more, watch closely, and mist daily. Some might emerge.

Pavilio indra fUT west desert segregate)

Host: Lomatium grayi var. depauperatum

Lab hosts: Lomatium graveolens, L. junceum, Cymopterus terebinthinus calcerea.

Pavilio indra calcicola

Hosts: Lomatium parryi, L. scabrum

Lab host: Lomatium junceum, Cymopterus terebinthinus

Remarks: A highly variable taxon.

Pavilio indra shastensis

Host: Cymopterus terebinthinus, Lomatium macrocarpum

Lab host: Lomatium graveolens

Pavilio indra versamus

Hosts: Taushiaparishi, T. arguta

Lab host: Lomatium graveolens

Pavilio indra fordi

Host: Cymopterus panamintensis

Lab host: Cymopterus terebinthinus

Remarks: Late instar larvae feed fine on Lomatium parryi but first-instars do not do as well on it.

Pavilio canadensis

Lab hosts: Populus fremontii, P. tremuloides

Pavilio rutulus

Hosts: Populus tremuloides, P. lombardii, Salix spp.

Lab host: Populus fremontii

Remarks: Immatures can be found.

Pavilio slaucus

Host: Prunus serotina

Lab host: Salix exigua

Pavilio multicaudatus

Hosts: Fraxinuspennsylvanica, Prunus virginiana

Remarks: Frequently large numbers of larvae can be collected. Try places of new construction where it has been

landscaped with many small host trees. Also, seek out isolated host plants with new growth. We have also found

larvae in numbers in Evanston, Green River, Rock Springs, and Rawlins, Wyoming, where cultivated ash grows in

corporate areas of these cities.

Pavilio eurymedon

Hosts: Ceanothus velutinus, Prunus virginiana

Lab host: Prunus serotina

Remarks: On several occasions we have had a small number of pupae remain in diapause until the second spring.

EMM cresphontes

Host: Citrus sspp.

Remarks: Larvae can be found in good numbers on isolated plants. Always seek out new growth especially

suckers coming out from near the trunk of the mature tree. Larvae do not feed on older leaves.

Pavilio xuthus

Host: Citrus sspp.

Battus vhilenor vhilenor

Host: Aristolochia watsoni

Lab hosts: Aristolochia tomentosa, A. druior, A.fimhriata

Remarks: Aristolochia watsoni is a very prostrate plant. It would usually go uncut if the area were mowed. Many

larvae can be collected in the late afternoon, however. As soon as the sunlight hits at a sharp angle many larvae

crawl up on tall plants. They can be easily seen from quite a distance. By carefully searching the understory below

the larvae that you see you will usually find its host. More often than not there will be more larvae on it. You can

collect until it is too dark to see or for a while later if you have a good lantern.

Battus vhilenor hirsuta

Host: Aristolochia californica

Remarks: Pupae can be found in the winter, often in good numbers. The host is a large vine usually growing

around a tree or shrub. Pupae can be found on the vine or nearby trees and posts. The pupae on the trees or posts

are usually no more than 3 feet above the ground. They are easy to see when there is no foliage to block your view.

In season, larvae should be easy to find.

PIERIDAE:

Remarks: Monitor hatching ova or separate ova, as hatching larvae will often eat other ova.

Remarks: For large numbers of immatures of all Colias, it is best to get ova from females. The larvae of Colias

eurytheme, Colias alexandra , and Colias philodice will not diapause if reared with a 24 hour photoperiod. It is best

to use a 24 hour photoperiod for all species of Colias. Constant light accelerates larval development and, with

some individuals or species, prevents diapause. Any taxon that might not diapause (that we are aware of) will be

mentioned in the species account. Colias are very hard to rear as they are highly prone to disease. The easiest and

best way to rear all Colias is to use potted plants.

Colias larvae that always diapause will not leave drying plant in search of fresh food, but will just diapause on the

plant. Potted plants stay fresh and the larvae will grow to their maximum diapausing instar. If only cut plant is

available the containers must be designed so they can be placed in the sun for a few hours a day and then put in the

lighted lab before sundown. Ultraviolet rays greatly reduce the chance of disease. Once a day carefully remove the

sleeve and check the plant for freshness and add water to the water bottle. When the plant freshness starts to

decline transfer each larvae to the leaf of a fresh plant.

Colias eurytheme

Hosts: Medicago sativa, Lupinus argenteus, Astragalus lentiginosus

Lab host: Thermopsis montana

Colias philodice eriphvle

Hosts: Medicago sativa, Astragalus cicer, A. lentiginosus, Lupinus argenteus

Lab host: Thermopsis montana

Colias interior

Lab hosts: Vaccinium caespitosum, V angustifolium, Vaccinium sp.

Remarks: Females fly in open timber. When shadows are long few females are flying.

Colias velidne skinneri

Lab host: Vaccinimum myrtilloides

Remarks: In timbered areas, females do not fly much when sunlight comes at too low of an angle.

Colias 2 in ante a sis an tea

Lab host: Salix exigua, Salix sp.

Remarks: An occasional larva will not diapause when reared with 24-hour photoperiod. These have always been

females.

Colias sisantea inuviat

Host: Salix lanata

Lab host: Salix exigua

Colias sisantea h arroweri

Host: Salix spp.

Lab host: Salix exigua, Salix sp.

Colias scudden

Hosts: Vaccinium caespitosum, V myrtilloides, Salixplanifolia

Lab host: Salix exigua

Colias boothi thula

Hosts: Hedysarum mackenziei, Astragalus arcticus

Colias canadensis

Host: Lupinus arcticus

Lab hosts: Hedysarum boreale, Lupinus (russell hybrids)

Colias hecla glacialis

Hosts: Astragalus arcticus, Hedysarum mackenziei

Lab Host: Astragalus cicer

Colias nastes nastes

Remarks: Oviposition observed on Astragalus arcticus

Colias nastes aliaska

Host: Oxytropis borealis

Colias nastes streckeri

Lab hosts: Astragalus cicer, Astragalus sp.

Colias valaeno chivvewa

Host: Vaccinium uliginosum

Lab host: Vaccinium caespitosum

Colias mossi

Remarks: Oviposition observed on Astragalus uniflores.

9

Pieris protodice

Hosts: Brassica nigra, Sisymbrium officinale, Stanley a pinnata, Cleome serrulata, Cardaria dr aba, Sisymbrium

altissimum

Lab host: Cardaria dr aba

Pieris beckerii

Hosts: Stanley a pinnata, Brassica nigra, Cleome serrulata, Descurainia pinnata, Sisymbrium altissimum

Lab host: I satis tinctoria

Remarks: Occasionally larvae can be found in large numbers. Last instar larvae provide conspicuous feeding

damage as they strip the inflorescense of the host.

Pieris sisvmbrii

Hosts: Arabis microphylla, A. sparsiflora, A. perennans, Isatis tinctoria, Descurainia pinnata, Cardaria dr aba

Lab hosts: Sisymbrium altissimum

Pieris sisvmbrii nisravenosa

Hosts: Streptanthus cordatus, Descurainia pinnata, Arabis holboelli

Lab hosts: Arabis perennans, Cardaria dr aba, Isatis tinctoria, Sisymbrium altissimum

Anthocharis Imceolata lanceolate

Lab hosts: Arabis holboelli, A. sparsiflora

Other’s host: Streptanthus tortuosus

Anthocharis cethura vima

Hosts: Streptanthella longirostris, Descuraniapinnata, Caulanthus lasiophyllum var. utahensis, Sisymbrium irio

Lab host: Arabis perennans

Remarks: We have had mixed results using Sisymbrium irio as a lab host even though females occasionally

oviposit on it in nature. Pupae can diapause for up to 11 years in the lab and still produce healthy adults.

Anthocharis sara sara

Host: Dentaria californica

Lab hosts: Arabis sparsiflora, Isatis tinctoria

Anthocharis sara nseudothoosa

Hosts: Descurainia pinnata, Arabis perennans (Fig. 4)

Lab hosts: Arabis glabra, Isatis tinctoria

Anthocharis thoosa thoosa

Hosts: Arabis perennans, Arabis holbellii, Descurainia pinnata, Streptanthella longirostris, Isatis tinctoria

Lab hosts: Any species of Arabis will serve as a lab host. Larvae eventually die on Streptanthus cordatus and

Brassica nigra.

Anthocharis thoosa Colorado

Hosts: Descurainia pinnata, Arabis spp.

Lab hosts: Any species of Arabis will serve as a lab host.

Anthocharis thoosa inghami

Hosts: Arabis perennans, Descurainia pinnata, Streptanthella longirostris,

Lab hosts: Any species of Arabis will serve as a lab host.

Anthocharis iulia iulia

Hosts: Arabis glabra, A. holboelli, A. perennans, Descurania pinnata, Streptanthella longirostris

Lab hosts: Isatis tinctoria

Anthocharis iulia browninsi

Hosts: Arabis glabra , A. perennans, A. sparsiflora var. subvillosa, Arabidopsis thaliana, Descurainiapinnata

Lab hosts: Arabis microphylla, Isatis tinctoria, Streptanthella longirostris. (Note: Any species of Arabis will

serve as a lab host. Larvae accept but perish on Sisymbrium altissimum, Cardaria draba, and Chorispora tenella.

Larvae refuse Capsella bursa-pastoris and die.

Anthocharis julia Stella

Hosts: Arabis perennans, Descurainia pinnata

Lab hosts: Isatis tinctoria

Remarks: Females from the Anthocharis julia Stella TL near Marlette Peak, Carson City, Nevada, do not oviposit

on the inflorescense as do many other taxa within the Anthocharis sara complex. They oviposit on the center stalk

towards the middle. This likely happens because deer or other animals consume inflorescenses.

Anthocharis julia sulfuris

Hosts: Descurainia pinnata, Arabis drummondii, Arabis sp.

Lab host: Isatis tinctoria

Remarks: Females of Anthocharis julia sulfuris from Boise County, Idaho oviposit near the inflorescense of

Descurainia pinnata whereas females of Anthocharis thoosa thoosa oviposit more towards the middle to the upper

two thirds of the plant.

Anthocharis iulia flora

Lab hosts: Arabis glabra, Isatis tinctoria

Anthocharis iulia alaskensis

Lab host: Arabis glabra

Remarks: Pupae develop to produce adults approximately 6 days earlier than do Colorado Anthocharis julia julia

under identical lab conditions.

Anthocharis midea annickae

Lab host: Arabis holboelli, A. sparsiflora, A. perennans

Other’s host: Arabis glabra

Euchloe hvantis lotta

Hosts: Sisymbrium altissimum, Descurainia pinnata, Stanleya pinnata, Streptanthus cordatus, Caulanthus

lasiophyllum utahensis

Lab host: Isatis tinctoria

Remarks: Can occasionally find larvae in good numbers. Females oviposit towards the inflorescense of the host.

Adults fly usually after Pieris sisymbri nigravenosa and Anthocharis thoosa thoosa have reached their peak flight.

Euchloe hvantis hvantis

Lab hosts: Arabis holboelli, Arabidopsis spp.

Other’s host: Streptanthus tortuosus

Remarks: Larvae accept but die on Cardaria draba in the lab.

Euchloe ausonides coloradensis

Hosts: Arabis sparsiflora, A. perennans, Isatis tinctoria, Descurainia pinnata, Cardaria draba

Lab hosts: Arabis holboelli, Arabidopsis spp.

Other’s host: Streptanthus tortuosus

12

Euchloe olympia

Hosts: Arabis glabra, Boechera fendleri, Descurainiapinnata,

Lab hosts: Arabis sp., I satis tinctoria

Remarks: Larvae are similar to Euchloe ausonides coloradensis at early instars but look darker at later instars.

Like other Euchloini larvae prefer fruits and flowers.

Ascia monuste monuste

Ascia monuste vhileta

Hosts: Capparis sp.

Lab host: Tropaeolum majus

Other’s host: Lepidium virginicum

Remarks: Length of photoperiod does not appear to affect the darkness of adults.

Neovhasia menavia menavia

Hosts: Pinus edulis, P. ponderosa

Remarks: Overwinters as ova. Larvae camouflage well against its host. Before years of heavy flights, post¬

diapause late instar larvae can be found if portions of tree are mist sprayed with water because the larvae jerk back

and forth violently. This suggests that the mist may be perceived as some sort of predator making the larvae very

conspicuous.

Daunus vlexivvus

Host: Asclepias speciosa (Fig. 3)

Remarks: Females seem to prefer plants that are growing among scattered trees or shrubs.

Daunus uilimms thersiimsus

Hosts: Sarcostemma cynanchoides, Asclepias speciosa, A. erosa

Agraulis vanillae incamata

Host: Passiflora sp.

SATYRIDAE:

Remarks: Overall, female satyrids will oviposit liberally in the lab. If you feel comfortable dealing with potted

grasses and sedges rearing many multivoltine taxa of satyrids is not too difficult. The problem arises with some

univoltine taxa that are difficult to force through to adult under lab conditions. Diapause for many satyrids is not

the rigid concept it can be for many other species of butterflies, except skippers. For example, some species of

satyrids, i.e., Coenonympha tullia brenda , will feed through to 4 th instar and then slow down its feeding and growth

rate to the point that larvae will not grow nor progress to the next instar. It is not always clear as to when these

larvae should be placed into diapause or attempted to be forced through to adults under a 24-hour light scheme.

Megisto cymela, Erebia magdalena, and Neominois ridingsi are other examples of satyrids which feed extremely

slow under lab conditions. Their feeding rate seems to slow down even further as they approach mature fifth-

instars, suggesting that some Neominois may overwinter in that stage.

Coenonympha tullia amvelos

Coenonympha tullia brenda

Coenonympha tullia erynsii

Lab Host: Poa pratensis

Remarks: Obtain ova from live females. Females prefer to oviposit on dead blades. Larvae will feed on many

species of potted grass. It is not generally difficult to push larvae of multivoltine populations of the Coenonympha

tullia complex through to adult under lab conditions. However, univoltine populations either diapause at 3 rd or 4 th

instar, or slow down their feeding rate as to continue feeding without growing or molting to the next instar.

13

Cercvonis spp.

Remarks: Use a similar rearing strategy for Cercyonis pegala, C. sthenele, C. meadi and C. oetus. Set up live

females in a small cage. Females seem to prefer to oviposit on dead blades of any bunch grass, including Poa

pratensis. In the lab ova postpone hatching for roughly 17 days and unfed 1 st instar larvae diapause. This is oddly

similar strategy as compared to Speyeria spp. Larvae can be forced out of diapause by placing them on fresh grass

blades and placing them under 24 hours of light. Once larvae start feeding it is advisable to then rear them through

the fall/winter months on any convenient species of potted grass.

Cercvonis meadi mexicana

Lab hosts: Bouteloua gracilis, Poa pratensis, Most species of grasses will serve as a lab host.

Mesisto cvmela

Lab host: Bromus inermis

Remarks: Both Megisto cymela and Megisto rubricata larvae have taken a long time to rear through to adults

under lab conditions (4-5 months) either due to the usage of problematic grasses or artificial photoperiod problems.

Mesisto rubricata chenevorum

Lab host: Bromus inermis

Remarks: Like many satyrids, larvae will accept many species of grasses in the lab. Mature larvae will aestivate

and not pupate until exposed to treatments of mist spraying, simulating summer monsoons in the desert southwest.

Neonvmvha areolata areolata

Lab host: Poa pratensis

Remarks: Females will oviposit on grasses as well as sedges. Larvae will accept grasses in the lab but generally

do better on sedges.

Satyr odes avvalachia avvalachia

Hosts: Carexstricta

Lab host: Cyperus esculentus

Remarks: Females will oviposit on sedges. Larvae will accept many varieties of sedges in the lab. Finding larvae

in the field can be difficult.

Erebia mazdatena magdalena

Lab host: Poa pratensis

Remarks: In the field females oviposit haphazardly around rocks and talus near its native grasses. Under lab

conditions, females do not necessarily even need grasses or a lot of sunlight to oviposit. Young instar larvae are

extremely wary and should be raised on potted grasses surrounded by nylon netting or chiffon to avoid escape (Fig.

15). Young instars that wander off into water have proven to be drown-resistant. Larvae take many months in the

lab to feed from first to fifth-instar. Larvae likely diapause as mature 5 th instar; but this is not proven.

Neominois riding si dionvsius

Lab host: Pseudoroegneria spicata

Remarks: Caterpillars grow very slowly in the lab and likely diapause as 5 th instar.

Neominois wvominso

Host: Pseudoroegneria spicata

Remarks: Unfed 1 st instars diapause; however, larvae will feed on fresh grasses if provided. Larvae grow very

slowly in the lab.

14

Asterocamva cbjton texana

Asterocamva clvton clvton

Asterocamva clvton flora

Host: Celtis reticulata

Lab host: Celtis occidentalis

Remarks: Larvae and pupae can be found.

Asterocamva celtis celtis

Asterocamva celtis montis

Asterocamva celtis antonia

Host: Celtis reticulata

Lab host: Celtis occidentalis

Remarks: Larvae can be found. New growth on isolated plants may be best.

Asterocamva leilia

Host: Celtis pallida

Remarks: Larvae can be difficult to find. The best strategy is to look for new growth on isolated plants. 1 st instar

larvae refuse Celtis reticulata in the lab

Nymvhalis antiova

Hosts: Salix exigua, Celtis reticulata, Ulmuspumila

Remarks: Occasionally larvae can be collected in large numbers.

Nymvhalis catifomica

Hosts: Ceanothus velutinus, C. martinii

Remarks: Large numbers of larvae can be found, sometimes in varying sizes in close proximity. They can

defoliate their host.

Nymvhalis milberti

Host: Urticadioica

Remarks: Larvae are easily collected in large numbers. Larvae are gregarious at early instars and then tend to

scatter somewhat at later instars and make nests similar to Vanessa atalanta.

Vanessa atalanta

Host: Urtica dioica

Remarks: Occasionally larvae can be found in good numbers. Larvae create nests.

Vanessa cardui

Hosts: Cirsium undulatum, Carduus nutans, Lupinus argenteus, L. sericeus, Helianthus annuus

Remarks: Larvae can be found in good numbers. It can be a common urban dweller.

Vanessa annabella

Hosts: Malva neglecta, Sida hederacea

Remarks: Larvae and ova can be collected in good numbers on isolated plants. Late summer, until a hard freeze,

is the most productive time. Host Malva neglecta is somewhat cold weather resistant. Normally adults overwinter

but larvae at any instar can survive the winter as well. Instead of diapausing larvae can feed at a very slow rate

during the winter when temperatures are above freezing.

Vanessa virginiensis

Host: Anaphalis margaritacea, Gnaphalium palustre

Remarks: Can find ova and larvae in good numbers.

Precis coenia

Hosts: Plantago major, P. lanceolata, Antirrhinum majus

Remarks: Larvae can be found on host in agricultural areas. Adult phenotypes can vary in the lab when larvae are

subjected to variable photoperiod and temperature.

Precis nigrosuflkm

Lab host: Antirrhinum majus

Other’s host: Mimulus sp.

Remarks: Larvae can be found.

Anartia iatrophue guantanamo

Host: Lantana sp.

Lab host: Plantago major

Polygonia spp.

Remarks: Some Polygonia species have two seasonal forms. Larvae of these reared with 24-hour photoperiod

produce the non-hibernating form. Those reared with 8-hour photoperiod produce the hibernating form.

Polygonia faunus arcticus

PolXgonia fuuiuis cenverag

Host: Salix scouleriana

Lab host: Salix sp.

Remarks: Larvae can be found.

Polygonia satyrus (coastal and inland forms)

Host: Urticadioica

Remarks: Larvae make a conspicuous nest. See http://utahbutterflies.ning.com/video/satvr-comma-and-milberts

for video tutorial showing the differences between larval nests of Polygonia satyrus vs. Nymphalis milberti.

Polggonia gracilis . zephmis

Hosts: Ribes montigenum, R. viscosissimum, R. cereum

Lab host: Ribes oxyacanthoides

Remarks: Last instar larvae tend to feed and strip leaves towards the ends of branches on shrubs. Larval strip

pattern is noticeable and last instar larvae can be found if timing is correct. Larvae that feed on Ribes montigenum

can camouflage themselves well.

Polygonia oreas oreas

Polygonia oreas threatfuli

Lab hosts: Ribes oxyacanthoides, Ribes ^p.

Other’s host: Ribes divaricatum

Remarks: Females will often hover next to the host. They prefer using scattered bushes interspersed in an open

clump with other shrubs or small trees.

Polygonia interrogationis

Lab host: Ulmus pumila

Adelvha eulalia

Hosts: Quercus turbinella, Q. gambelii

Lab hosts: Quercus robur, Q. alba

Remarks: Can find an occasional larva. Most females are reluctant to lay in captivity. In nature females have been

observed ovipositing on leaves that are more concealed within the body of the bush, making the finding of

immatures more difficult. Similar to Limenitis , 1 st instar larvae construct a perch extending the vein of the leaf.

16

Limenitis Ionium lorguM

Limenitis lorguini burrisonii

Limenitis lorguini vallidafacies

Hosts: Salix spp., Populus angustifolia, Amelanchier alnifolia

Lab hosts: For all Limenitis, Populus fremonti, Salix exigua

Remarks: Most Limenitis that are reared with a 24-hour photoperiod will not diapause. Freshly molted 2 nd instars

are especially sensitive to monitoring photoperiod. 1 st instars do not. In habitats where the hosts are scattered or

confined ova and small larvae can be collected. During the late fall or winter when most of the leaves have fallen it

is not too difficult to spot hibemacula. As is true with other Limenitis species, females oviposit well in a cage with

high humidity and exposed to filtered sunlight. Immatures of all Limenitis can be found.

Limenitis lorguini x Limenits weidemeveri

Hosts: Prunus virginiana, Salix sp.

Remarks: A very occasional hibemacula can be found along a willow-lined stream. Less than a hundred feet up

on the dry hillside, short, scattered chokecherries were growing. Nearly every Prunus plant had a hibemaculum

and sometimes as many as six. This is a typical scenario in the dry southwest.

Limenitis weidemeveri weidemeveri

Limenitis weidemeveri oberfoelli

Limenitis weidemeveri latifascia

Hosts: Amelanchier alnifolia, Populus tremuloides, P. angustifolia, Salix sp., Prunus virginiana

Limenitis archivvus archivvus

Limenitis archinvus obsoleta

Limenitis urchippus lahonUmi

Limenitis arcUimms jloridensis

Hosts: Salix exigua , S. laevigata, Populus fremontii

Remarks: Females prefer to lay on branches that are hanging over water

Limenitis archivvus obsoleta x Limenitis astvanax arizonensis

Host: Salix sp.

Lab host: Populus fremontii

Limenitis astvanax astvanax

Limenitis astvanax arizonensis

Hosts: Prunus serotina, P. virginiana, Salix sp., Populus sp.

Limenitis arthemis rubrofasciata

Host: Salix sp.

Lab hosts: Populus balsamifera, P. tremuloides

Euvhvdrvas phaeton phaeton

Euph y dry as phaeton qwM

Hosts: Chelone glabra

Lab hosts: Castilleja chromosa, Penstemon cyananthus

Remarks: Post diapause larvae have been found on Castilleja coccinea and Virburnum recognitum.

17

Euphydryas gilletti

Hosts: Lonicera involucrata, Veronica wormskjoldi

Remarks: This taxon has a 2-year life cycle; therefore, the larvae overwinter for two winters. Afer the first winter

the second diapause can be avoided by rearing with a 24-hour photoperiod. Egg clusters and small larvae are easy

to locate. Females like scattered plants near a stream or on a dry hillside above it. Post-diapause larvae can be

found in good numbers in leaf litter under the host or on branches in the plants interior.

Euvhvdryas anicia anicia

Euvhvdryas anicia maria

Euvhvdryas anicia alena

Euvhvdryas anicia bernadetta

Euvhvdryas anicia windi

Euvhvdryas anicia macvi

Euvhvdryas anicia veazieae

Euvhvdryas anicia wheeled

Euvhvdryas anicia hermosa

Hosts: Penstemon gloriosus, P. utahensis, P. cyananthus, P. palmeri, Penstemon sp., Castilleja chromosa,

Lonicera involucrata, Symphoricarpus oreophilus

Lab hosts: All Euphydryas that use Castilleja or Penstemon will switch from one to the other.

Other’s host: Bessaya wyomingensis

Remarks: Egg clusters and prediapause larvae can be found. Prediapause larvae feed gregariously in silk nests

and disperse to plants up to 500 feet away before diapause and can be found as post-diapause larvae on these plants

the next year. An individual female will normally lay her eggs in a relatively small area. Euphydryas break

diapause early and can be found as soon as their host has useable growth. Even if no larvae are visible on a plant,

search all the understory and debris nearby. Post-diapause larvae that feed on shrubs are not sensitive to dry and

warm conditions and may be taken directly to the lab for rearing. Post-diapause larvae like to rest on dead foliage.

Post-diapause larvae can rediapause if overcrowded. However, larvae that have re-diapaused can break diapause

yet again the same year if exposed to another cold treatement of one month.

Euvhvdryas colon nevadensis

Euvhvdryas colon sverryi

Hosts: Symphoricarpus oreophilus

Lab hosts: Penstemon cyananthus, Symphoricarpus sp.

Remarks: Post diapausal larvae can be found in good numbers. They are easiest to see in early spring when the

host has just leafed out. Penstemon and Castilleja grows very common among the Symphoricarpus but no larvae

were found on them.

Euvhvdryas chalcedona chalcedona

Euvhvdryas chalcedona olancha

Euvhvdryas chalcedona klotsi

Euvhvdryas chalcedona macslashani

Hosts: many Penstemon spp., Keckiella antirrhinoides

Lab hosts: Penstemon cyananthus, Penstemon palmeri

Euph v dry as editha lehrnmi

Eiwh (inas editha colonia

Euvhvdryas editha baroni

Hosts: Castilleja chromosa

Lab hosts: Penstemon cyananthus, Collinsia sp.

Other’s host: Penstemon sp., Plantago lanceolata

Phyciodes mvlitta mylitta

Hosts: Cirsium undulatum, C. vulgare

Remarks: Occasionally larvae can be found in good numbers. When reared with a 24-hour photoperiod most

Phyciodes will not diapause. Females will sometimes oviposit on young Cirsium vulgare basal rosettes in the fall

where the larvae overwinter under the leaves.

Phyciodes orseis orseis

Phyciodes orseis ssp. (CA)

Lab host: Cirsium undulatum

Other’s host: Cirsium cymosum

Remarks: At least a few larvae will not diapause when reared with a 24-hour photoperiod. In some populations,

most of the larvae will not diapause. Immatures can be found with regularity. Females like to use plants that are

growing against a rock, tree or windfall.

Phyciodes vulchella vulchella

Phyciodes vulchella shoshone

Phyciodes vulchella Camillus

Phyciodes vulchella ssp. (CA)

Host: Asterfoliaceus

Lab Hosts: Aster chilensis. Aster sp.

Remarks: Larvae can be found.

Phyciodes vallida pallida

Phyciodes vallida barnesi

Lab host: Cirsium undulatum

Phyciodes cocyta cocyta

Phyciodes cocyta selenis

Lab hosts: Aster foliaceus, A. chilensis, Aster sp.

Phyciodes texana texana

Lab hosts: Dicliptera resupinata, B eloper one guttata

Remarks: Immatures have been found.

Poladryas arachne arachne

Hosts: Penstemon utahensis, P. humilis

Lab host: Penstemon cyananthus

Remarks: Occasionally post-diapause larvae can be found. Post-diapause larvae refuse P. cyananthus but larvae

obtained from ova thrive on it. Most larvae reared from ova will not diapause when reared with 24-hour

photoperiod.

Thessalia leanira leanira

Thessalia leanira wrishti

Thessalia leanira alma

Thessalia leanira elesans

Thessalia leanira oresonensis

Hosts: Castilleja chromosa, C. sulphurea

Lab host: Castilleja spp.

Other’s host: Castilleja affinis

19

Remarks: Depending on sufficient moisture, large numbers of post-diapause larvae can be collected. In areas

where Euphydryas are sympatric, leanira will be found after most Euphydryas have finished. The reason for this is

that post-diapause larvae will molt before resuming feeding in the spring, whereas Euphydryas anicia complex

larvae will feed immediately, placing Euphydryas larvae ahead of Thessalia larvae by approximately 1-2 instars.

Most larvae will be found in the understory and surrounding vegetation. Larvae can be found as much as 3 feet

from the host. Larvae of elegans will go through to adult if reared with a 24-hour photoperiod. The other

subspecies diapause. Larvae of wrighti that have re-diapaused will resume feeding 1 month later if separated and

placed on fresh host after a cold treatement.

Thessalia theona thekla

Hosts: Castilleja lanata, C. laxa

Lab host: Castilleja chromosa

Remarks: Post-diapause larvae can be found in good numbers. Larvae of T. fulvia and T. cyneas may be found in

the same area. The larvae lack bright coloration.

Thessalia fulvia fulvia

Thessalia fulvia coronado

Thessalia fulvia ssp. (mostly AZ)

Lab host: Castilleja chromosa

Other’s hosts: Castilleja laxa, C. lanata, Castilleja sp.

Remarks: Immatures can be found in good numbers.

Thessalia cyneas

Lab host: Castilleja chromosa

Other’s host: Castilleja laxa

Remarks: Post-diapause larvae can be found in limited numbers.

Thessalia chinatiensis

Host: Leucophylum minus

Remarks: Larvae can be found in numbers

Chlosvne ianais

Host: Anisacanthus wrighti

Remarks: The host is a low spreading shrub that grows in dense colonies. Often larvae of all instars can be found

as well as pupae. When the plants are bathed in sunshine the larvae are hiding in shady interior branches or the

understory. Larvae are up on the plant feeding in early morning and evening.

Chlosvne californica

Host: Viguiera deltoides

Remarks: When moisture has been sufficient larvae can be found in good numbers.

Chlosvne lacinia form adiustrix

Chlosvne lacinia form crocale

Host: Helianthus annuus

Remarks: Larvae of all sizes can be found. Early spring larvae usually produce form adjustrix adults. Larvae

collected mid-summer will produce both forms.

Chlosvne sorsone sorsone

Lab host: Helianthus annuus

20

Chlosvne harrisi ha nisi

Chlosvne harrisi lissetti

Chlosvne harrisi albimontana

Lab host: Aster engelmannii

Other’s host: Aster umbellatus

Remarks: Have obtained ova and reared larvae on A. engelmanni. C. harrisi liggetti post-diapause larvae can be

found in good numbers on Aster umbellatus near Spruce Knob, West Virginia.

Chlosvne nvcteis nvcteis

Chlosvne nvcteis drusius

Lab host: Helianthus annuus

Other’s host: Helianthus annuus

Chlosvne valla valla

Chlosvne valla eremita

Chlosvne valla flavula

Chlosvne valla ssp. (3-CA)

Chlosvne valla ssp. (MT)

Host: Aster engelmannii

Lab Host: For all subspecies: Aster englemannii

Other’s hosts: Aster sp.

Remarks: When reared with a 24-hour photoperiod only an occasional larva will not diapause. Rearing post¬

diapause larvae with constant light is best. With a natural photoperiod about 50 percent of the larvae will

rediapause. With 24 hours of light, 30 percent or less will rediapause.

Chlosvne hofjmam segregate

Lab host: Aster engelmannii

Other’s host: Aster brickellioides

Remarks: With 24-hour photoperiod only an occasional larvae will rediapause.

Chlosvne sabbi sabbi

Lab hosts: Aster chilensis, Corethrogyne filginifolia

Remarks: Four females laid 5 egg clusters (one was very small) on Aster engelmannii as no natural host was

available. Upon hatching the larvae fed for about 30 days before eventually dying. Females would not lay on A.

chilensis but larvae thrived on it. When reared with 24 hour photoperiod an occasional larva will not diapause.

Chlosvne sterove sterove

Chlosvne sterove dorothvi

Chlosvne sterove acastus

Hosts: Chrysothamnus viscidiflorus, C. greenei, Machaeranthera canescens, Pyrrocoma radiatus

Lab host: Aster engelmannii

Remarks: Ova and larvae can be found in good numbers including an occasional pupa. Prediapause larvae of C,

sterope acastus have always diapaused when reared under constant light. Of approximately 300 C. sterope

dorothyi larvae reared from collected ova and 1 st instar larvae, 92 went through to adults. 17 larvae of C. sterope

sterope went through to adults (all females) and 87 larvae diapaused.

Chlosvne neumoeueni

Host: Machaeranthera tortifolia

Remarks: As with all desert species abundance is determined by sufficient timely moisture. Larvae can be

collected in good numbers. They can be seen at a distance but a stealthy approach is advised. The slightest

vibration of the host plant causes them to “jump ship” and drop to the understory.

21

Chlosvne damoetas ssp. (UU WY)

Chlospie damoetas ssp. (MT)

Lab host: Aster engelmanni

Remarks: These taxa are associated with Solidago multiradiata and Erigeron leiomeris the likely larval host

plants. Females from all locations have laid egg clusters and the larvae have been reared on Aster engelmanni.

Obtained ova from the UT and WY populations in 2001 and reared the 162 larvae to diapause. The majority

diapaused as large 2 nd instars, 14 molted to 4 th instars, which fed very briefly and diapaused. The remainder

diapaused as 3 rd instars. In 2002, raised post-diapause larvae under 24-hour photoperiod where all but two larvae

grew an instar and rediapaused. The two that went through to adult emerged as females. In nature, I think all

would have diapaused. 2003 was a repeat of 2002. Two more female adults emerged and the rest diapaused as

either large 3 rd or 4 th instars. In 2004 two males and one female emerged. All of the larvae diapaused as 4 th instars.

Sometime in the fall of 2004 all of the diapausing larvae died. In 2006 we obtained 200-300 eggs from the

distinctive Montana subspecies and 13 larvae went through to adults; 5 males and 7 females. 31 diapaused as large

4 th instars and the rest as large 3 rd instars. In 2007 fifteen went through to adults. About 30 percent of the larvae

diapaused as 4 th instars; the rest as 3 rd instars. In 2008 only a few larvae were alive when spring arrived, all died

without breaking diapause.

Mona SPP-

Remarks: There is an alien species of Viola that is toxic to Boloria and Speyeria larvae. This Viola is frequently

grown in gardens.

Boloria kriemhild

Host: Viola sp.

Remarks: Larvae diapause at mid-instars when reared in the lab.

Boloria frissa sasata

Lab host: Salix exigua, Salix sp., Viola sp.

Remarks: If reared with 24-hour photoperiod a good percentage will not diapause. The females oviposited on

Salix sp. Larvae do well on Salix or Viola.

Boloria eunomia ursadentis

Host: Viola adunca

Speyeria diana

Lab Host: Viola sp.

Other’s host: Violapapilionacea

Remarks: Speyeria diana can be reared on potted Viola tricolor but some do not do well on it. Unfed 1 st instars

will diapause. If you have Viola available you can break the larval diapause. Place the newly hatched larvae in a

petri dish with new growth Viola leaf. Use 24-hour photoperiod and make sure the Viola is always fresh. This

method gives you adults several months sooner and avoids the danger of your diapausing larvae dying. Speyeria

females will oviposit well if placed in a paper grocery bag with a couple of Viola leaves. Fold the top shut and hold

it closed with clothes pins or large paper clips.

Speyeria cvbele charlotti

Speyeria cybele leto

Speyeria cybele letona

Speyeria cybele ssp. (MT)

Lab hosts: Viola adunca, V. tricolor

Other’s host: Viola papilionacea, Viola sp.

Speyeria nokonm apacheana

Lab host: Viola sp.

22

Speveria idalia

Lab host: Viola sp.

Other’s host: Violapapilionacea, V pedatifida

Speveria mormonia luski

Lab host: Viola sp.

Other’s host: Viola adunca, Viola sp.

Speveria atlantis nausicaa

Lab Host: Viola sp.

Other’s host: Viola sp.

Speveria edwardsi

Lab host: Viola sp.

Other’s host: Viola nuttalli

Eiwtoieta daudia

Lab host: Viola sp.

Other’s host: Passiflora sp.

Apodemia mormo virsulti

Apodemia moi

”mo deserti

Apodemia moi

*mo cvthera

Apodemia moi

”mo meiicana

Apodemia moi

ymo durvi

Apodemia mormo nigrescens

Hosts: Eriogonum inflation, E. corymbosum, E. fasciculatum, E. brevicaule, E. umbellatum, E. wrightii, Krameria

glandulosa

Remarks: Larvae can be collected in good numbers. Larva usually live in a nest of leaves silked together. Larva

using Eriogonum inflatum chew a hole into the inflated pod when it is green. They hide and eventually pupate in

the dried pod. If the hole in the pod is silked over you know it is occupied. The larger the larvae you collect the

better because they grow very slowly.

Apodemia palmeri

Hosts: Prosopis glandulosa, Prosopispubescens

Remarks: When rearing in the lab, small larvae are reluctant to crawl from unusable plant if its cutting is laid on

fresh plant. The larvae should be transferred with a camel's-hair brush.

LYCAENIDAE:

Remarks: When possible remove prepupae from rearing containers. Larvae are highly cannibalistic and will often

eat prepupae or pupae.

Theclini:

Remarks: Almost all of the Theclini can best be reared by obtaining ova from females. The only reared Theclini

that were obtained by other methods are Habrodais grunus, Hypaurotis crysalus citima, Satyrium californicum,

Satyrium tetra, Sandia mcfarlandi, Incisalia fotis, Callophrys comstocki, and Strymon bazochi. These will be

discussed with each species. Habrodais grunus and Hypaurotis crysalus are notorious for refusing to oviposit in

captivity. To rear, they must be obtained as ova or larvae. Both species overwinter as ova. All Satyrium

overwinter as ova. All Callophrys, Incisalia , and Mitoura overwinter as pupae.

23

Habrodah uninus

Host: Quercus chrysolepis

Remarks: Larvae can be collected in good numbers. Larvae can be found by inspecting the foliage or by placing a

tarp under the branches and beating them with your net handle. If most of the larvae are small, beating will be

more profitable. Some larvae may be Satyrium auretorum. There will also be a good supply of spiders, earwigs,

moth larvae, etc.

Hyyaurotis crysalus citima

Host: Quercus gambelii

Remarks: During the winter and spring following a good flight, eggs can be found in good numbers. Eggs are

deposited at the base of next year’s leaf buds. Cuttings of host do not stay fresh very long.

Harkenclenus thus immaculosis

Lab host: Prunus virginiana

Remarks: To obtain ova confine females on cuttings that have cracks and imperfections in the stems. Few leaves

are needed. Also lay some twigs at the base of the cuttings to construct an understory.

Satyrium californica

Hosts: Purshia tridentata, Cercocarpus montanus, Amelanchier utahensis, Ceanothus velutinus, Prunus

virginiana

Remarks: All attempts to obtain ova from females in captivity have been unsuccessful. Oviposition has been

observed in nature (although this is very time consuming) and the ova were taken. Larvae are usually difficult to

find. However, in one good colony we were able to find a good supply of larvae by looking for attendant ants.

Racemes of P. virginiana or C. velutinus are the best to use for rearing. We have found pupae in good numbers on

two occasions.

Satyrium svlvinus vutnami

Satyrium sylvinus mesapallidum

Host: Salix exigua

Remarks: Females lay best on cuttings with cracks, holes and other imperfections. If each cutting has twig stubs,

that helps also. Females like to tuck all their eggs in such places. Very few leaves are needed.

Satyrium saevium

Hosts: Ceanothus velutinus, Ceanothus sp.

Remarks: Small larvae prefer flowers. Larvae were obtained by beating the hostplant branches adjacent to the

flowers, and allowing them to fall into a net or other container.

Satyrium behri behri

Hosts: Purshia tridentata, P. mexicana, Cercocarpus montanus, C. ledifolius

Satyrium fulisinosum semiluna

Hosts: Lupinus sericeus, L. argenteus

Remarks: Ants, in some colonies, hollow out areas under most of the plants where S . fuliginosum flies and the

larvae hide in these holes, tended by the ants. Often there are 3 or 4 larvae in a hole. The ants not only protect

them from 6 and 8-legged predators but also from each other. They are very cannibalistic and must be reared

separately. If you check the same holes a month earlier, you will find Plebejus icarioides larvae. Occasionally you

find a late finisher when the fuliginosum larvae are quite small Females oviposit in the ground at the base of the

Lupine plants. To obtain ova from females the base of the plant must have some soil surrounding it. It probably

also helps to have a few twigs and leaves on the soil.

Satyrium auretorum

Host: Quercus chrysolepis

24

Satyrium tetra

Host: Cercocarpus betuloides

Remarks: S. tetra larvae were obtained by the beating method. Many were parasitzed.

Satyrium livarovs alivarovs

Lab host: Primus virginiana

Satyrium calanus sodarti

Host: Quercus gambeli

Atlides halesus estesi

Hosts: Phoradendron juniperinum, P. californicum

Remarks: Host will last for days in the lab with the stem of the mistletoe hostplant in water. If only raising a few

larvae, using a closed container works when you replace host plant and container daily.

Sandia mcfarlandi

Host: Nolina erumpens

Remarks: Larvae can be found in good numbers, and can be found in the blossoms and resting at the base of the

plant. Also, ova have been obtained from females. Four different color morphs of larvae were found on the same

Incisalia eryyhon eryyhon

Incisalia eryyhon vuryurascens

Lab host: Pinus monophylla

Other’s hosts: Pinus contorta, P. monophylla

Remarks: Larvae like the new growth of spring. Females prefer to oviposit on new growth and refuse to oviposit

on older growth.

Incisalia ausustinus

Hosts: Ceanothus velutinus, Purshia tridentata, Arctostaphylos uva-ursi, Prunus virginiana

Remarks: Oviposition observed on Cuscuta sp. Larvae have been collected from Ceanothus velutinus using the

beating method.

Incisalia mossii

Host: Sedum lanceolatum

Incisalia fotis fotis

Host: Purshia mexicana

Remarks: Larvae can be found in good numbers. Since larvae can be found, no attempt has been made to obtain

ova from females. Adults will emerge soon after pupae are put into a warm room in winter or spring. Some have

even emerged in an unheated garage (or even in a refrigerator) in February.

Mitoura svinetorum

Hosts: Arceuthobium americanum, A. divericatum

Mitoura iohnsoni

Lab host: Arceuthobium americanum

Other’s host: Arceuthobium campylopodum

Mitoura siru siva

Mitoura siva chalcosiva

Hosts: Juniperus osteosperma, J. scopulorum

25

Mitoura loki

Lab host: Juniperus californica

Mitoura loki thornei

Lab host: Cupressus forbesii

Callovhrys sheridani neoverylexa

Hosts: Eriogonum heracleoides, E. racemosum, E. corymbosum, E. umbellatum, E. brevicaule

Remarks: Females will oviposit in a squat tub closed container with host lying at the bottom (Fig. 8). Females

respond well to artificial light that is utilized intermittently, turn the light on and off every 20 minutes or so.

Callovhrys affinis affinis

Hosts: Eriogonum umbellatum, E. elatum, E. racemosum, E. heraclioides

Remarks: Larvae can be found.

Callovhrys avama avama

Hosts: Ceanothus fendleri, Eriogonum racemosum, E. alatum

Callovhrys comstock cornstocki

Hosts: Eriogonum corymbosum, E. hermannii, E. fasiculatum

Remarks: Larvae can be found in good numbers on the dorsal side of host leaves. Ova have been obtained from

females, but most of our rearing was done by finding larvae.

Callovhrys dumetorum

Host: Lotus scoparius, Eriogonum fasiculatum

Strymon melinus franki

Hosts: Eriogonum racemosum, E. kearnyi, E. alatum, E. heraclioides, E. umbellatum, E. corymbosum, Astragalus

utahensis, Malva neglecta, Malvella leprosa, Hibiscus mocheutos, Althea rosea, Medicago sativa, Hedysarum

boreale, Lupinus argenteus, Sidalcea oregana, Iliamna rivularis

Remarks: Larvae can be very abundant. If you are collecting other Lycaenid larvae and you find one with a

double row of dashes down the back, you have a melinus. They have many color morphs. Larvae are very

cannibalistic so must be reared separately.

Strymon bazochii

Host: Hyptispectinata

Lab host: Lantana camara

Remarks: Ova and larvae have been found in good numbers. Therefore, no attempt has been made to obtain ova

from females.

Erora laeta

Lab host: Salix exigua

Other’s host: Corylus cornuta

Erora quaderna

Lab host: Salix exigua

Lycaena spp.

Remarks: In the Rocky Mountains, all Lycaena overwinter as eggs except L. cupreus and L. phlaeas. To obtain

ova of the species that use Rumex, Oxyria, and Polygonum (except cupreus and phlaeas). It is necessary to have

soil surrounding the base of the plant because females oviposit in the soil. A few twigs and leaves on the soil help.

L. cupreus and L. phlaeas females oviposit on the leaves of the hostplant.

26

Lycaena gorgon

Host: Eriogonum nudum

Remarks: Larvae and ova can be collected in good numbers. Large larvae can be seen at a distance, eating the

very top of the flower stem.

Lycaena mariposa penroseae

Lab host: Vaccinium caespitosum, Vaccinium sp.

Lycaena vhlcieas arctodon

Host: Oxyria digyna

Lycaena vhlaeas weberi

Lab hosts: Rumex acetosella, Oxyria digyna (reared by Dale Nielson), R. acetosa alpestris

Remarks: Obtained ova on Rumex acetosa alpestris from the colony and Oxyria digyna. Larvae refused Rumex

acetosella hybrid that Lycaena cupreus and Lycaena editha do well on. Larvae accepted but perished soon

thereafter on Rumex crispus. Reared under natural photoperiod but in a warm lab many larvae pupated and

emerged. Some of the ova diapaused. As day length lessened; pupae and half-grown larvae diapaused.

Lycaena ghlaeas feUdeni

Lab hosts: Oxyria digyna, Rumex acetosa

Lycaena xanthoides xanthoides

Lycaena xanthoides dione

Lab host: Rumex crispus

Lycaena rubidus sirius

Host: Rumex crispus

Remarks: Larvae can be found in large numbers. If the plant is growing in hard-packed soil with no understory

there will be no larvae. Larvae are found in loose understory, plant debris or gravel. Larvae may be 4 or 5 inches

below the surface and 7 or 8 inches away. Once 92 larvae were found under one small plant.

Lycaena hyllus

Host: Rumex maritimus

Lab host: Rumex crispus

Remarks: Larvae can be found in large numbers. Look in the understory as with prior species. Females will also

oviposit on plants growing in water. On these, the larvae will be on the Rumex.

Lycaena helloides helloides

Lycaena helloides mesaloceras

Lab hosts: Rumex crispus, R. paucifolius

Remarks: Obtained ova of megaloceras on Potentilla concinna. Larvae were offered two other species of

Potentilla besides concinna, and they refused all.

Lycaena heteronea heteronea

Lycaena heteronea northi

Hosts: Eriogonum umbellatum, E. racemosum, E. corymbosum

Lab host: Eriogonum heraclioides

Lycaena nivalis browni

Lab hosts: Rumex crispus, Rumex sp., Polygnium douglasii

Remarks: Some larvae eat Rumex but only a few lived. Most females will not oviposit on Rumex. An occasional

female will lay but usually only a few eggs. Females oviposit well on Polygonum douglasii and larvae flourish on

it. It is best to pot thick clumps of douglasii. It takes several individual plants to rear a single larva.

27

Lycaena arota schellbachi

Host: Ribes leptanthum

Lab host: Ribes spp.

Remarks: As soon as it is warm and sunny in the morning the females are active. Look for yellow composites

near the host.

Lycaena editha montanci

Lab host: Rumex acetosella hybrid

Remarks: Larvae will feed on other Rumex spp. besides their known hosts but most will die in the late instars.

Lycaena cuvreus cuvreus

Lab hosts: Rumex acetosella hybrid, Rumexpaucifolius

Remarks: Most larvae diapause at 3 rd instar; but some will go through to adult in the lab. Most females are

reluctant to oviposit in captivity.

Lycaena cuvreus sjiowi

Host: Oxyriadigyna

Remarks: Larvae can be found but normally only a few. Females oviposit readily on leaves of digyna in a closed

container.

Brevhidium exilis

Hosts: Salsola iberica, Atriplex canescens, Portulaca oleracea

Remarks: Put cardboard or a dark tarp under plant and beat with net handle. Larvae and pupae can be collected in

good numbers using this technique.

Asriades sp. (MT)

Other’s host: Douglasia montana

Remarks: Post-diapause larvae can be found. Females will oviposit well in the lab.

Lamvides boeticus

Lab host: Pisum sativum

Remarks: Reared on fresh snow peas purchased at the supermarket.

Hernia runs ceraunus gyas

Host: Melilotus officinalis

Lab host: Pisum sativum

Remarks: Female oviposited on Melilotus officinalis but larvae accepted fresh snow peas purchased at the

supermarket.

Celastrina ladon echo

Host: Eriogonum racemosum, E. wrightii, Ceanothus velutinus

Remarks: Larvae and ova can be found. Larvae found on Eriogonum racemosum in Sevier County, Utah, were

oddly not tended by ants.

Celastrina ladon lucia

Host: Ledum palustre

28

Everes amvntiila

Host: Vida americana

Lab host: Lathyrus lanszwertii, L. latifolius, L. odorata

Remarks: Young larvae burrow into flower buds and remain there for as long as the bud will support them.

Larvae turn brown before diapausing or pupating. Some larvae diapaused and some did not, when reared with 24-

hour photoperiod.

Glauconsvche Ivsdamus Ivsdamus

Glaucovsvche Ivsdamus oro

Hosts: Astragalus sp., Lupinus sericeus, L. argenteus, Hedysarum boreale, EL. mackenziei, Eriogonum wrightii

Remarks: Larvae can be collected in good numbers. Look for ants tending larvae on inflorescens.

Glaucovsvche viasus daunia

Hosts: Lupinus sericeus, L. argenteus

Remarks: Where lygdamus and piasus fly together, lygdamus larvae are usually in mid to late-instar when piasus

is found as eggs or small larvae. The majority of the larvae will usually be lygdamus. Females are reluctant to

oviposit in captivity.

Plebeius acmon acmon

Plebeius acmon lutzi

Hosts: Eriogonum umbellatum, E. kearneyi, E. racemosum, E. wrightii

Remarks: Have found larvae usually when looking for Callophrys or Euphilotes larvae. Females oviposit well on

E. umbellatum. Most lutzi larvae that feed on leaves in the lab do not diapause (John Emmel, pers. comm.).

Plebejus melissa melissa

Host: Medicago sativa

Plebeius icarioides ardea

Host: Lupinus sericeus

Remarks: Find larvae in ant holes beneath the plant. Mid-instar larvae usually diapause in nature but larvae can

be forced through to adult if ova are obtained from live females in the lab.

Plebeius shasta minnehaha

Host: Astragalus sp.

Remarks: Larvae can be found in fair numbers.

Philotes sonorensis

Host: Dudleya cymosa

Remarks: Larvae can be found.

Euvhilotes svaldinsi

Host: Eriogonum racemosum

Remarks: Larvae can be found in good numbers. Examine food plant with a magnifying glass when you get to the

lab. Often you will find small larvae. Rear separately. 1 st and 2 nd instar larvae, if reared under 24-hour

photoperiod, will pupate and emerge the same year. The pupae must have the long photoperiod also. All of these

methods apply to all Euphilotes we have reared. Diapausing Euphilotes pupae must be kept outside and monitored

until emergence time (this coincides with the blooming of their hostplant). Without natural photoperiod and

temperature they will not emerge.

29

Euphilotes vallescens vallescens

Euvhilotes vallescens ricei

Euph ilotes vallescens arenamontana

Host: Eriogonum kearneyi

Remarks: Larvae can be found in good numbers.

Euvhilotes enoptes ancilla

Hosts: Eriogonum heraclioides, E. umhellatum

Remarks: Females tend to oviposit only on the open flower petals of Eriogonum heraclioides. Finding larger

larvae on the hostplant can be difficult because they camouflage or hide themselves quite easily on the flowers or

seedpods of the hostplant.

Megathyminae:

Remarks: The "Megs" included in this list (and possibly all Megs) can be reared with good results. Megathymus

use only small to medium size plants and the larva usually kill the plant. To rear Megathymus Yucca plants were

planted in the yard. Over a period of several to 25 years the crop of Yucca herrimaniae spread by rhizomes to

constitute a sizable and dense patch of Yucca plants. The plants become too dense to use and must occasionally be

thinned by cutting the stems at ground level, also plants that become large must be cut off at ground level. Yucca

angustissima plants spread only sparingly by rhizomes, however, individual plants would continue to grow and

develop a large root. These plants also must be cut off at ground level when they become large. After being cut

off, the root would grow two new stems, each one on opposite sides of the root. When there is a large root the

larva does not kill the plant and after a few years the plant is usable again.

Agathymus use Agave plants of any size in Utah. Agave plants were transplanted to the yard in the spring when

they were going to be used. Most of the plants did not live permanently but would survive until the larvae pupated.

When transplanted, the plants must be cleaned by removing the dead leaves and washing out the remaining good

leaves. The dead leaves and rubbish harbor unwanted critters. It is easy to obtain ova from Agathymus females but

very few of the newly hatched larvae become established in a leaf. It was quite successful to harvest first and

second-instar larvae in nature and transplant to plants in the yard. Plants or group of plants must be covered with

screen to keep the parasites and predators away from the larvae. Even with a screen cover an occasional larva

would get parasitized or spiderized.

Megathymus ova are glued to a leaf and are quite easy to find. Agathymus ova are dropped in or near a plant and

are nearly impossible to find.

Mesathvmus vuccae

Hosts: Yucca angustissima, Y. bacata, Y. harrimaniae

Remarks: Larvae build a silk tent in the center of the plant. This tent can easily be found and many M. yuccae

pupae can usually be found. The pupae can be harvested shortly before flight time. Larvae finish eating long

before flight time but diapause until shortly before flight time to pupate. If larvae are found within a couple months

of flight time they can be kept in their silk tube and will eventually pupate or a paper tube can be used.

Mex at hymns streckeri

Hosts: Yucca harrimaniae, Y. angustissima, Y. glauca

Remarks: Larvae build their tent in the ground near the plant. The top of the tent is at ground level so it is very

difficult to find the pupae. This taxon is best obtained by rearing. Many ova can be found easily after the females

have been flying a few days.

Asathvmus neumoeseni

Host: Agave scabra

Remarks: Pupae can be found by spotting the trap door in the Agave leaf or by spotting the frass.

30

Agathvmus marine

Host: Agave lechugilla

Remarks: Pupae can be found by spotting the trap door in the Agave leaf or by spotting the frass.

Asathvmus alliae

Host: Agave utahensis

Remarks: Larvae make their trap door on the under side of the leaf It is possible to find pupae in the field but

searching can be a long arduous process with limited results. To get a good series it is necessary to rear this taxon.

Pyrginae, Hesperinae:

Remarks: Larvae of most hesperine and pyrgine (spread-wing skippers) construct nests. These skipper larvae

differ from most moth larvae in that they exit in order to launch frass whereas most moths that make nests do not.

In other words, finding larval nests on plants with frass means you have found a moth nest not a skipper nest.

Females of most hesperine skippers will oviposit on and larvae will usually accept a far greater variety of grasses

than is normally utilized in nature making lab rearing not too difficult. Rearing of some hesperine skippers from

humid areas of the Midwest and Eastern U.S. requires the use of humidity (mist spraying) both for unhatched ova

and for unfed first-instars in order to stimulate hatching and feeding.

Similar to Megathyminae, many (but not all) pyrgine and hesperine skippers pupate in the same nest they fed as

larvae but seal up the entrance of the nest with silk. This extra layer of white silk is very noticeable in hesperine

skippers. This increases the possibility of finding pupae in the wild.

Larvae of the Hesperine genera Hesperia, Polites, Pseudocopaeodes, Hylephila, and Atalopedes make nests down

at the bases of bunch grasses and are nearly impossible to locate. Larvae of Amblyscirtes, Atrytonopsis, Lerodea,

Panoquina, Ochlodes, Poanes, Piruna, Thymelicus, Wallengrenia, Copaeodes and Oarisima feed on taller, wide-

blade species of grasses higher where they can be much more conspicuous. Some "skipperling" hesperine larvae

make smaller nests or none at all.

Finding larvae of many pyrgine skippers is not too difficult when their population numbers are significantly high

and hostplants are not too prevalent.

Eparcvreus clarus

Hosts: Glycyrrhiza mendota, G. lepidota, Robiniapseudoacacia, Lotus crassifolius

Remarks: Larvae construct a nest among the leaves of the hostplant and are easily found in any instar. This is

especially true when the host is Glycyrrhiza mendota. They are easiest to rear by finding 5 th instar larvae so that

they don’t have to be fed very long. When the pupae are put into a warm room in the spring to emerge they must be

moist for some time (to adjust to the warmth) or they will dessicate. This is easily accomplished by placing them on

wet paper towels and covered by wet paper towels. Use lab host Robinia pseudoacacia over natural host

Glycyrrhiza lepidota in an open terrarium setup. The problem with using cuttings of Glycyrrhiza lepidota is that

cuttings will wilt in an open terrarium. The only way to keep cuttings erect is to increase humidity in the setup

similar to a closed terrarium setup recommended for Limenitis. However, this increased humidity in the closed

terrarium does not allow frass to dry and larvae can die as pupae. Robinia pseudoacacia is a much drier host.

Larvae use this host naturally in the Eastern U.S. An open terrarium setup is fine with this host and larvae stay

healthy. For multivoltine populations such as Epargyreus clarus californica keeping pupae humid is not necessary.

PohL'onus leo sarianv

Host: Pisidium piscipula

Remarks: Larval nests can be found on new growth of host. Larvae will pupate in nest.

31

Phocides oiamalion okeechobee

Host: Rhizophora mangle

Remarks: Larval nests can be found on host. Larvae will pupate in nest and pupae can be occasionally found. It

is not advisable to use cuttings of the host, as larvae do not do well on cuttings. Try and obtain R. mangle (red

mangrove) starts from a nursery and rear on potted plants. Use saline solution for potted plants.

Thorybes mimics mludes

Host: Melilotus officinalis

Lab Hosts: Trifolium repens, Medicago saliva

Remarks: Mature larvae hibernate. Larvae make nests. Females have oviposited on Astragalus cicer in nature

but first-instar larvae refused to feed.

Thorybes mexicana nevada

Lab host: Medicago sativa, Melilotus officinalis

Remarks: Mature larvae hibernate. For some reason pre-diapausal 5 th instars can easily die during the wintering

process.

Thorybes diversus

Lab host: Trifolium repens

Remarks: Mature larvae hibernate.

Erynnis icelus

Hosts: Populus tremuloides, Salix sp.

Remarks: Mature larvae hibernate. Pupae do not hibernate. Occasionally, lab reared larvae in the fall will pupate

and will emerge a few weeks later. This also applies to other species of Erynnis.

Erynnis brizo bursessi

Hosts: Quercus gambellii, Q. turbinella

Lab host: Quercus alba

Remarks: Seek out seedlings to find nests in the late summer or early fall. Mature larvae hibernate. Females

oviposit only on very young tender new growth of leaves. Ova are white, usually turning orange 24 hours later.

Erynnis telemachus

Host: Quercus gambellii

Lab host: Quercus alba

Remarks: Finding late instar larvae in late summer can be a challenge even during larger flights. Finding isolated

seedlings growing along roads or fresh cuts can improve chances of finding immatures. Larvae take 3-4 months to

mature to last instar in the lab.

Erynnis meridianus

Host: Quercus turbinella

Lab hosts: Quercus alba, Quercus gambellii

Remarks: Double-brooded in southwest Utah. The key to finding immatures is to seek out ovae on very fresh

succulent new growth of the host during the summer brood in mid to late August. New growth is usually hard to

find on the host during this time of the year, therefore, when you do find it you usually will find an ova. Timing is

important because finding smaller instar larvae on the same new growth is usually much less productive as young

instars can be quickly consumed by predation.

Erynnis vacuvius lilius

Host: Ceanothus velutinus

Remarks: Larvae can be found.

32

Ermnis persius fredericki

Host: Lupinus argenteus

Remarks: Larvae can be found.

Erynnis afranius

Host: Hedysarum boreale

Lab host: Lupinus argenteus

Remarks: There are up to 3 broods at low elevations. Late instar larvae are not too difficult to locate. Poor-

quality natural host in late summer produces smaller larvae and hence smaller adults for the first flight in the

spring. Second brood adults are almost as large as Erynnis telemachus.

Heliovetes ericetorum

Host: Sphaeralcea sp.

Lab hosts: Malva neglecta, Sida hederacea, Althaea rosea

Remarks: Larvae of all instars overwinter.

Pyrsus scrivtura

Hosts: Sphaeralcea ambigua, Sida hederacea

Lab host: Althaea rosea

Remarks: Larval nest is somewhat distinctive to Pygus communis. In the same habitat; larvae that are found on

short hosts in dry areas are P. scriptura. P. communis larvae prefer healthier plants in wetter areas. Pupae

overwinter.

Pyrsus communis

Hosts: Sphaeralcea ambigua. Sida hederacea, Malva neglecta,

Remarks: Females will oviposit in a small cage. Finding larvae in disturbed or suburban areas is not too difficult

where the adults fly. Larvae change color from green to brown when overwintering.

Pyrsus ruralis ruralis

Hosts: Potentilla glandulosa, Fragaria vesca

Lab Host: Potentilla fruticosa

Remarks: Females will oviposit fairly well in captivity on the hostplant. Finding immatures in the wild can be

very difficult. Larvae create nests but do not pupate in their nest. Pupae overwinter.

EXISUI centaunme loki

Host: Potentilla diversifolia

Lab Hosts: Potentilla glandulosa, Fragaria vesca.

Remarks: Larvae accept Fragaria vesca in the lab but attempts to get pupae to emerge the following spring have

failed. Current overwintering techniques that work for Pyrgus ruralis ruralis have not worked for Pyrgus

centaureae loki.

Svstasea zamva

Host: Abutilon abutiloides

Lab host: Alcea rosea

Remarks: Conspicuous larval nests are not too hard to find.

Pholisora catillus

Hosts: Chenopodium album, Amaranthus retroflexus

Remarks: Larvae can be found.

33

Hesperopsis libya libya

Host: Atriplex canescens

Remarks: Larvae can be found in areas where the host is not overly abundant. Look on plants along the periphery

of the population. Larvae create nests and will pupate in nests.

Hesveroysis libya confertiblanca

Host: Atriplex confertifolia

Lab Host: Atriplex canescens

Remarks: Look on isolated plants. Sometimes multiple larvae can be found on one plant. 3 rd through 4 th instar

larvae will semi-aestivate during the hot summer months and will not usually pupate until mid to late-July.

Therefore, it is advisable to sleeve larvae on local Atriplex canescens then harvest the late instars or pupae later.

Hesveroysis alvheus oricus

Host: Atriplex canescens

Remarks: Larvae can be found. Like Erynnis, Thorybes, and Amblyscirtes mature last instar larvae overwinter

and pupate the following spring.

Hylepbila phvlens pin-lens

Host: Cynodon dactylon

Lab host: Poapratensis

Remarks: Females will oviposit fairly well on Bermuda grass. Under lab conditions the amount of time between

the 1 st instar larvae and pupae is roughly 9 weeks, which is much quicker than many species of Hesperia and

Polites.

Atalovedes campestris camnestris

Host: Cynodon dactylon

Lab host: Poa pratensis

Atalovedes campestris huron

Lab host: Distichlis spicata

Remarks: Females will oviposit on Distichlis spicata. Larvae are generalists and will accept many species of

grass in the lab.

Polites vibex

Lab host: Poa pratensis

Remarks: Larvae feed quickly from hatchling 1 st instar to pupa in about 9 weeks.

Polites origenes rhena

Lab host: Bromus inermis.

Remarks: Raising this skipper can be difficult. Exposing larvae to 24 hours of light does not guarantee that they

will go through and pupate the same year. Many larvae slow down their metabolism at late instars neither

completely diapausing nor growing. Some overwintered larvae resume feeding at a normal rate, pupate and

emerge. Whereas others continue feeding at a very slow rate and eventually die.

Polites themistocles

Lab hosts: Poa pratensis, Phalaris arundinacea

Remarks: Females oviposit somewhat sparingly on Kentucky Blue Grass. Larvae will accept Phalaris

arundinacea in the lab and will go through to adults the same year.

Polites sabuleti sabuleti

Host: Poa pratensis

Remarks: Last instar larvae can burrow at the base of the plant to make a nest. Rearing larvae on potted grass is

advisable.

34

Polites sabuleti chusca

Host: Distichlis spicata

Remarks: Females will not oviposit on Poapratensis in the lab.

Polites veckius

Lab host: Poa pratensis , Phalaris arundinacea

Polites sonora utahensis

Lab host: Sedges

Remarks: Females oviposit very sparingly on grasses, if at all.

Hesperia iuba

Host: Distichlis spicata

Lab Host: Phalaris arundinacea

Remarks: As is the case with many Hesperia , larvae grow at a fairly slow rate, about 2-3 months from ova to

adult.

Hesperia pahaska nr. martini

Host: Bouteloua gracilis

Lab Host: Phalaris arundinacea

Remarks: As is the case with many Hesperia , larvae grow at a fairly slow rate, about 2-3 months from ova to

adult. Rear larvae under 24 hours of light and provide fresh host to avoid diapause.

Hesperia uncas lasus

Host: Bouteloua gracilis

Lab Host: Phalaris arundinacea

Remarks: Females will oviposit on Distichlis spicata in the lab. As is the case with many Hesperia , larvae grow

at a fairly slow rate, about 2-3 months from ova to adult. Rear larvae under 24 hours of light and provide fresh host

to avoid diapause.

Hesperia nevada nevada

Lab Host: Phalaris arundinacea

Remarks: Females prefer not to oviposit on Poa pratensis. Other natural bunchgrasses should be better.

Pseudocopaeodes eunus eunus

Host: Distichlis spicata

Remarks: To get a series, try and get as many females as is possible. Compared to other hesperine skippers, ova

of eunus are very large and females are limited to how many ova they can oviposit.

Poanes taxiles

Host: Bromus inermis

Lab host: Phalaris arundinacea

Remarks: Larvae attempt diapause (or a feeding slowdown) at 4 th to 6 th instar. If larvae are given fresh host

regularly they will feed through and produce adults the same year.

Poanes zabulon

Lab hosts: Bromus inermis, Phalaris arundinacea.

Remarks: Unlike Poanes taxiles, Poanes zabulon larvae only have five instars. Throughout much of its range, P.

zabulon is multivoltine. Therefore, if larvae are given fresh host regularly they will feed through to pupae and

produce adults the same year.

35

Pomes hobomok

Lab hosts: Bromus inermis, Phalaris arundinacea.

Remarks: Larvae have 6 instars and seem to insist on diapausing at the 4 th instar.

Ochlodes vuma vuma

Host: Phragmites australis

Remarks: Larvae and pupae can be found. Ova have been found on Sorghum halepense. See

http://utahbutterflies.ning.com/video/finding-yuma-skipper-larval for a video tutorial on how to find larval nests on

Phragmites.

Ochlodes svlvanoides nava

Hosts: Phalaris arundinacea, Bromus inermis

Remarks: Larvae can also be found. Larval nests are conspicuous. Last instar larvae diapause for roughly 2-6

weeks before finally pupating, producing a late summer flight. Unfed 1 st instars hibernate by silking the tips of the

Amblvscirtes vialis

Host: Bromus inermis

Lab Host: Phalaris arundinacea

Remarks: Females will oviposit in the lab. Mature last instar larvae hibernate. In the lab some mature last instar

larvae wifi pupate, producing adults soon thereafter.

Amblvscirtes fimbriata

Host: Phalaris arundinacea

Remarks: Larvae can be found. Mature last instar larvae hibernate. However, in the lab many last instars wifi

pupate and produce adults soon thereafter.

Amblvscirtes eos

Host: Sorghum halepense

Lab Host: Phalaris arundinacea

Remarks: Larvae can be found.

Lerema accius

Host: Sorghum halepense

Lab Host: Phalaris arundinacea

Remarks: Larval nests can be found.

Lerodia eufala

Host: Sorghum halepense

Lab Host: Phalaris arundinacea

Remarks: Larval nests can be found. Larval nests, coloration, and feeding patterns are oddly similar to

Thymelicus lineola.

Wallensrenia eseremet

Host: Panicum sp.

Lab Host: Phalaris arundinacea

Remarks: Females will oviposit on a variety of grasses. Ova seem to need humidity to hatch.

Oarisima sarita

Host: Bromus inermis

Lab Host: Phalaris arundinacea

Remarks: Larvae can be found. Using potted grasses is advisable for rearing this taxon.

36

Thvmelicus lineola

Host: Phalaris arundinacea

Remarks: It is not too hard to find larval nests in the field. However, late instars use nests to a lesser degree or

not at all. Larvae pupate on grass blades and are not too difficult to locate. This skipper overwinters as ova.

Ancvloxvvha numitor

Remarks: It’s not too hard to find larval nests in semi-wet habitats. Larvae pupate in grass blades in such a tight

fashion as to making their removal quite problematic. It is advisable to leave the nest intact for emergence

purposes.

Ancvloxvvha arene

Lab Host: Phalaris arundinacea, Bromus inermis

Remarks: Populations are very local in wetland areas. Larvae pupate in grass blades in such a tight fashion as to

making their removal quite problematic. It is advisable to leave the nest intact for emergence purposes.

Covaeodes aurantiacus

Host: Cynodon dactylon

Lab Host: Phalaris arundinacea

Remarks: To get a series, try and get as many females as is possible. Compared to other skipperlings ova are very

large and females are limited to how many ova they can oviposit. Larvae show interesting similarities to some

Satyrids.

Covaeodes minima

Lab Host: Phalaris arundinacea

Remarks: Females will oviposit in the lab.

Piruna virus

Host: Bromus inermis

Lab Host: Phalaris arundinacea

Remarks: Females seem to be fussy about ovipositing in the lab in numbers. Larvae feed through rapidly to adult

in the lab even though they diapause in nature.

37

Fig 1-2. A. julia browningi and P. eurymedon females nectar on honey water. Fig 3. D. plexippus female oviposits on

Asclepias speciosa. Fig 4. A. sara female oviposits on Arabis perennans (Dennis Walker). Fig 5. Ovum of A. julia

browningi on Arabis sp. Fig 6. Ovum of M. siva chalcosiva on Juniperus osteosperma. Fig 7. Female A. eulalia set up

in a screen cage with cuttings of Quercus gambellii. Fig 8. Female C. sheridani neoperplexa set up to ovipsosit in a

squat tub with Eriogonum racemosum. Figs 9-12. Last instar caterpillars of C. affinis affinis , P. bairdi, P. indra minori,

A. cethura pima, and T. leanira wrighti. Fig 13. L. lorquini burrisoni 3 rd instar caterpillar crawls out of its hibemaculum.

Fig 14. Open terrarium setup for Colias meadi larvae under 24 hours of light (Nicky Davis). Fig 15. Potted grass setup

for E. magdalena larvae. Fig 16. P. eurymedon pupa.

38

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39

Volume 7

31 March 2010

Number 4

The Taxonomic Report

OF THE INTERNATIONAL LEPIDOPTERA SURVEY

Xfy '

Observations on Anthocharis julia browningi and Anthocharis thoosa thoosa

Including Tension Zones near Nephi, Juab County, Utah

TODD STOUT 1

1456 North General Drive, Salt Lake City, UT 84116

ABSTRACT: Two visibly distinct taxa of the sara orangetip fly sympatrically and synchronically in a zone

northeast of Nephi, Juab County, Utah. This paper discusses how these two taxa interact based upon six character

sets—adult phenotype, adult male behavior, distribution, habitat, larval coloration, and pupal shape.

Introduction

There currently are different schools of thought regarding how many species there are in the

Anthocharis sara complex (sara orangetip.) The purpose of this report is to discuss a small piece of that

puzzle by sharing observations in the distribution, habitat, behavior, adults, and immature stages of

Anthocharis julia browningi and Anthocharis thoosa thoosa (Fig. 1.) More importantly, this report will

also provide a historical account of tension zones between these two taxa in Northern Utah as well as

present a closer in-depth study carried out during the spring of 2009 near Nephi, Juab County.

Because of my research in progress on much of the Anthocharis sara complex, browningi is

currently treated as a subspecies of the southern rocky mountain orangetip {Anthocharis julia) based upon

larval and pupal characters as well as adult blend zones amongst this and other races of A. julia. At the

same time, the southwestern orangetip {Anthocharis thoosa) is treated as a distinct species consistent with

Opler (1999), because of adult, larval, and distributional differences withH. sara.

t'TH

tjg.

pH

jT V

w

fei ni

fe*.

Fig 1. Adult reared series of A. julia browningi (left) and A. thoosa thoosa (right) from Northern Utah.

1 Staff Member, The International Lepidoptera Survey, Herndon, VA

Anthocharis julia browningi in Utah

General:

The Type Locality of Anthocharis julia browningi is City Creek Canyon, Salt Lake County, Utah;

Skinner, 1905. Topotypical A. j. browningi is phenotypically very similar to other Wasatch Front

populations of this butterfly. In both sexes, the dark markings are reduced or washed out as compared to A.

julia Stella and A. julia julia. The background color in males is off white with a slight yellowish tint that

can be more pronounced in some individuals in Cache County populations. The background color in

females is a light yellow. The discal cell bar is narrow, offset, and frequently disconnected with the bottom

black apical border. This subspecies has been recognized as being a race of A. sara. However, consistent

larval differences between this and other races of A. sara and A. thoosa coupled with consistent larval

similarities with ssp. julia, sulfuris, Stella, flora, and alaskensis, place browningi as a subspecies of julia.

Utah Distribution and Habitat:

In Utah, browningi flies throughout most of the Wasatch Range from Box Elder to Juab County as

well as the Bear River Range found in the extreme northeastern corner of the state. Intermediates between

browningi and nominotypical julia have been taken by Alan Myrup in the Uinta Mountains. A. j.

browningi also flies in the San Pitch Mountains of Juab County and in Carbon County in Price Canyon.

Bionomics :

Host plants for browningi include many rock cresses including Arabis sparsiflora var. subvillosa,

Arabis perennans, Arabis microphylla, Arabis drummondii, and Descurainia pinnata (tansy mustard.)

Oviposition has also been observed on Arabidopsis thaliana. Larvae will readily accept Isatis tinctoria

(dyars woad) in the lab. With the exception of dyars woad, most of these larval host plants can be found

either between rocks {Arabis spp.) or taking refuge under trees {Descurainiapinnata.)

The ova are white turning orange after 24 hours; hatching in about 4-5 days. First instar larvae are

cannibalistic and will consume other pierid ova if it finds them. The young first instar larva is light colored

with a dark head. Second and third instar larvae of browningi are greenish. The fifth instar larva is light

green and is pictured below (Fig. 2) and darkens as it approaches pupation. Hibernation is as pupa.

2

Anthocharis thoosa thoosa in Utah

General:

The Type Locality of Anthocharis thoosa thoosa is Mokiak Pass, Mojave County, Arizona;

Scudder, 1878. The amount of dorsal yellow coloration in females seems to be greater with southern Utah

populations as compared to northern ones. This butterfly is at home in great basin piny on-juniper habitat

where males patrol up and down ravines or dry washes occasionally leaving the gully to investigate nearby

Juniperus osteosperma trees in search of females. Females can be found flying in association with Juniper

trees either resting or ovipositing on one of its larval host plants, Descurainia pinnata , which take refuge

under or near the same Juniper trees. Females also oviposit on rock cresses from the genus Arabis.

Utah Distribution and Habitat:

The general distribution of thoosa in Utah ironically forms a general U shape surrounding and

circumventing the Wasatch Range and the distribution of browningi. In the northwestern part of the state,

A. thoosa flies in Utah’s Great Basin and West Desert Ranges south to Utah’s Dixie located in the

southwestern corner of the state. Populations then extend east towards the four corners region of the state

and then north again towards the East and West Tavaputs Plateau. Eastern Utah populations are designated

as Anthocharis thoosa Colorado. See Scott and Fisher (2008).

Current studies of adults, larvae, and pupae of topotypical Colorado from McElmo Creek,

Montezuma County, Colorado, may place it as a junior synonym to nominotypical thoosa. To date, my

furthest documented northeastern population of thoosa is Slaughter Canyon, near Sunnyside, Carbon

County, Utah. As you head north, populations of this butterfly from Eastern Utah are replaced by

Anthocharis julia in the Uinta Range in Summit and Daggett counties in the NE section of the state.

Bionomics:

Host plants for A. t. thoosa include Descurainia pinnata, Arabis perennans, Arabis holboelli , and

other species of Arabis. The young first instar larva is light orange colored with a dark head. Second and

third instar larvae of A. thoosa are greenish. The fifth instar larva is green and is pictured below (Fig 3).

Hibernation is as pupa. Because the butterfly flies in more or less xeric habitat, most pupae will bypass

emergence after one year and will emerge during the second, third, or fourth year of diapause.

3

Differences in Immatures

Larvae:

Because last instar caterpillars of many species of the pierid tribe Euchloini change color as they

advance through their last instar, it is imperative that any larval coloration comparisons between taxa be

made from the same timeframe after a fourth instar larva molts to fifth instar. For my larval comparison

studies, I compare larvae that have been fifth instars for 54 - 60 hours because this interval demonstrates

the most visible, consistent differences to the naked eye.

Fifth instar browningi larvae have a broader white lateral stripe and the ground color is lighter green

as compared to thoosa (Fig. 4.) At the same time, to the naked eye, the transitional color change from the

white lateral strip to the green base color of a browningi larva is much more subtle as compared to thoosa

larvae. A. t. thoosa larvae show larger green pinacula surrounding the setae or tubercle giving the larva the

appearance of being a much darker green to the naked eye. The pinacula of browningi fifth instar larvae

also enlarge and darken as the larva progresses towards pupation; but do so 12 to 24 hours later.

These consistent differences between last instar caterpillars of these two taxa are also applicable

when comparing other subspecies of A. thoosa to A. julia. (This is not just a consistent phenomenon in

comparing thoosa to browningi.) Interestingly, coloration of fifth instar larvae of topotypical A. sara sara

and topotypical A. sara pseudothoosa are consistently a much darker green above the white lateral stripe

than those of A. julia or A. thoosa; irrespective of the darkening of the pinacula surrounding the setae.

Pupae:

The shape of browningi pupae are consistently different to those of thoosa even though color

differences, whether green or tan are not reliably consistent. The main difference is that the browningi

pupal cone bends back whereas those of thoosa are erect and upright (Fig. 4.) These pupal differences are

only consistent on a population or subspecific level; but, not necessarily a specific level. For example,

pupae of A. julia sulfuris and A. julia flora tend to have pupal cones that bend back and are longer than

those of A. julia julia, A. julia browningi, and A. julia Stella. (Pupal cones of A. julia alaskensis are erect;

similar to A. thoosa.) At the same time, pupal cones of A. thoosa inghami are similar in shape to those of

A. thoosa thoosa; but, the cone is slightly longer and sometimes slightly tilted back.

Cross section Comparison of Fifth Instar Larvae Comparison of Pupae

A. julia browninsi A . thoosa thoosa

Fig. 4. Comparison of the mid segments of same-age fifth instar larvae (left) and pupae (right) of A. t. thoosa

and A. j. browningi. Oddly enough, color larval differences appear more noticeable to the naked eye or when

you squint your eyes—which is how I noticed their differences in the first place.

4

Differences in Adult Characters

There are a few contact or tension zones of different taxa within the Anthocharis sara complex

including but not limited to A. sara and A. julia Stella near Fresno Dome, Madera County, California (Ken

Davenport, personal communication), A. sara sara and A. julia Stella in Northern California—see Geiger

and Shapiro (1986), A. sara Siskiyou segregate and A. julia nr. Stella at Klamath River Canyon, Klamath

County, Oregon (Andy Warren, personal communication), as well as A. julia and A. thoosa Colorado near

the Four Corners Region—see Scott and Fisher (2008). Scott also references several other contact zones in

California, Colorado, and New Mexico.

The best opportunity to visibly note possible intermediates between two sympatric taxa was to

select those that visibly looked the most distinct from each other. A. julia browningi and A. thoosa thoosa

seemed to fit these criteria. (See table and Figure 5 below.)

Fig 5. Dorsal and ventral examples of an Anthocharis julia browningi male (left), Anthocharis

thoosa thoosa male (middle) and hybrid male (right) from 2009 study area in Juab County, Utah.

Previous Northern Utah Contact Zones

Willow Creek; 3.0 miles ESE of Mona, Juab County

My personal interest in discovering contact zones of A. j. browningi and A. t. thoosa was piqued on

7 May 1997, when Bob Hardbarger discovered both taxa flying together at Willow Creek; 3.0 miles east of

Mona, Juab County, Utah. Three days later, Bob, Steve Sommerfeld and I returned to Willow Creek and

confirmed Bob’s finding when I vouchered several Anthocharis males—some of those were thoosa and the

rest were browningi. (I do not have exact numbers as those specimens were later sent to Paul Opler for

research and study.) However, we did not locate any apparent intermediates or hybrids on that day.

Subsequent trips to Willow Creek in 2006, 2007, and 2008 were somewhat frustrating as the

dominant taxon was A. j. browningi. All males and females collected during this timeframe were

browningi. However, I also collected and reared several eggs and caterpillars to adult—all browningi ;

except two were thoosa. I obtained no intermediates from reared material.

Deep Creek Canyon; 3.6 miles South of Levan, Juab County

On 27 May 1999, Steve Spomer, Jim Reiser, and I found mostly A. t. thoosa and a few browningi

flying together at Deep Creek Canyon, Juab County. Of the males I collected that day, one appeared to

have intermediate characters between thoosa and browningi (Fig. 6.) The rest were parentals. We also

found a population further up the canyon where patrolling males were browningi.

Fig 6. Male of an apparent hybrid of Anthocharis thoosa thoosa x Anthocharis julia browningi taken

on 27 May 1999 at Deep Creek Canyon, Juab County, Utah.

Gardner Creek; 2.7 miles NNE of Nephi, Juab County

Because A. j. browningi was the dominant taxon at Willow Creek in 2007, I decided to investigate

Gardner Canyon—which was typical “ thoosa ” habitat located 2.5 miles south of Willow Creek and 2.7

miles NNE of Nephi. On 14 Apr 2007, because it was a relatively dry year for that region, I only collected

two thoosa males, 1 thoosa female, six ova on Descurainia pinnata (which were reared to adult and turned

out to be thoosa) and one apparent hybrid (Fig 7); but did not observe or collect any parental browningi.

In a tension zone, the key to finding thoosa and browningi flying sympatrically may have a lot to do

with habitat. The mouth of this canyon, even though officially part of the Wasatch Mountains, was typical

Great Basin habitat ubiquitous with Artemisia tridentata , Juniperus osteosperma and Purshia mexicana.

6

Fig 7. Male of a possible hybrid of Anthocharis thoosa thoosa x Anthocharis julia browningi taken

on 14 Apr 2007; Gardner Canyon; 2.7 miles NNE of Nephi, Juab County, Utah.

Rock Canyon; 2 miles East of Provo, Utah County

On 17 April 2004, after a Utah Bug Club meeting held at the Monte L. Bean Life Science Museum

at BYU, I took students on a field trip and collected several males of A. j. browningi as well as 2 males of

A. t. thoosa flying in what was previously considered to be a “browningi- only” population in Rock Canyon,

just east of BYU in Provo, Utah County, Utah. These two thoosa males were examined and appeared to be

pure parental thoosa without any visible evidence of gene exchange with browningi.

2009 Study Area

During the winter of 2009, because I located two study areas within 2.5 miles of each other where

either one taxa or the other dominated in northeastern Juab County, I decided to take a closer look at other

canyons, draws, and/or gullies between these two areas where I might find both taxa flying sympatrically

and synchronically.

I decided to study three small accessible canyons—Birch Creek, Little Birch Creek, and an

unnamed draw several hundred feet south of Little Birch Creek. These study areas (Fig. 11) were almost

nestled between Willow Creek to the north (where browningi dominates) and Gardner Creek to the south

(where thoosa dominates.)

Birch Creek; 3.7 miles SSE Mona; 4.0 miles North of Nephi, Juab County

On 20-21 Apr 2009,1 visited Birch Creek and found several A. t. thoosa males patrolling along the

base of the canyon flying in typical Juniperus osteosperma habitat at an elevation of 5400 feet (Fig. 11.)

As I hiked about 1,000 feet to the east up Birch Creek to an elevation of 5585 feet, I noted the subtle

difference in habitat change where oaks and maples replaced Juniper trees. It was here where I collected

three patrolling browningi males and one apparent hybrid male.

About fifty feet away from where the males were flying, I collected a female (Fig. 8.) My initial

impression was that she was a hybrid based upon her proximity to browningi males, smaller discal cell bar

than typical thoosa , faded dark markings (which turned out to be the result of age), and weak flight.

However, of the 25 eggs she laid; 16 were reared to pupae where larvae and pupae conformed to thoosa ;

without showing any noticeable browningi characters.

7

Fig 8. Three A. j. browningi males (left); one hybrid male and one faded/worn female A. thoosa

thoosa (right) collected at Birch Creek on 20 Apr 2009.

Little Birch Creek; 4.2 miles SSE Mona; 3.6 miles North of Nephi, Juab County

Although I was unable to locate A. t. thoosa and A. j. browningi flying in the exact same spot at

Birch Creek, I was able to collect 4 male browningi, 9 male thoosa, and 4 apparent male hybrids patrolling

sympatrically and synchronically at Little Birch Creek at an elevation of 5400 feet (Fig. 9.) This might

have been the case because this ravine was so narrow that it didn’t really separate browningi from thoosa

habitat.

It was interesting to note the behavioral differences in the patrolling males at Little Birch Creek. A.

j. browningi males tended to fly much slower than A. t. thoosa males. Also, thoosa males left the ravine

from time to time to investigate nearby Juniper trees in search of females; before scampering back to the

ravine to patrol. Oddly enough, it seemed to me that the behavior of the hybrid males was either slow and

deliberate {browningi) or fast and scampering {thoosa.)

Approximately 250 feet below the ravine, I collected two female thoosa flying amongst the Juniper

trees which laid eggs in the lab on native host Descurainia pinnata. Both the larvae and pupae of those

immatures were analyzed as thoosa without any visible evidence of browningi immature traits.

I had hoped to find ova on several plants of Arabis perennans growing in the outcroppings near the

ravine where the males of both species were flying; but didn’t find any pierid immatures on them;

including those of Pontia sisymbri nigravenosa.

MW

MW

MW

Ml

M-

MW

MW

MW

Fig 9. I collected a total of four browningi , nine thoosa ,

and four apparent hybrid males on 20-21 April, 2009 at

Little Birch Creek, Juab County, Utah. (If you have an

electronic copy of this file, you can examine these

specimens more closely by zooming in.)

MW

MW

MW

MW

MW

MW

W

MW

I didn’t spend as much time in the draw located roughly 0.1 miles south of Little Birch Creek; but, I was

able to collect 1 male ^4. thoosa thoosa and 1 mal eA.julia browningi flying there (Fig 10.)

A. jutia browningi d

A. thoosa thoosa d

T,;

Fig. 10. Male browningi and thoosa collected at the unnamed draw located 0.1 miles south of Little Birch Creek

9

Review

During my 2009 study of contact zones of Anthocharis julia browningi and Anthocharis thoosa thoosa

between Mona and Nephi, Juab County, Utah, I noted that of the 23 collected males, 43.5 percent were

parental A. t. thoosa , 34.8 percent were parental A.j. browningi, and 21.7 percent showed apparent

intermediate traits. I also collected three females who were parental thoosa where all offspring and

emerged adults showed notable thoosa characters; although at press time, these have not yet been spread.

The fact that the percentage of hybrids in this study area is roughly 22 percent with the rest showing

parental traits coupled with a narrow overlap in distribution, consistent differences in habitat preference,

male adult flying behavior, larval coloration characters and pupal shape characters, suggest that subspecies

thoosa and browningi belong to different species.

Also, outside of noticeable adult characters, there may be other factors that might show a higher percentage

of gene exchange using better available technologies such as electrophoresis (Geiger & Shapiro, 1986), or

nuclear dna studies.

10

Acknowledgments

Special thanks are given to Jon Pelham, Dr. Andy Warren, Paul Opler, and Harry Pavulaan for reviewing this

paper and to Norbert Kondla for his help with plates. Also, because of information shared with James A.

Scott in Papilio #18, I did not reference Scott who was referencing me regarding larval studies; but have

referenced original material from James A. Scott. Acknowledgment also goes to COL. Clyde F. Gillette,

who has proven records of either browningi or thoosa in every county in Utah.

Bibliography

Davenport, Kenneth E. 2004. The Yosemite butterflies. Taxonomic Report of the International Lepidoptera

Survey 5(1): 1-75, figs. {29 Dec 2004}

Davenport, Kenneth E., Norbert G. Kondla, Charles Grisham and C. Howard Grisham. 2007. The Yosemite

butterflies: color plates. Taxonomic Report of the International Lepidoptera Survey 5(2): 1-83, color figs.

{15 Mar 2004}

Geiger, H. & A. M. Shapiro. 1986. Electrophoretic evidence for speciation within the nominal species

Anthocharis sara Lucas (Pieridae). The Journal of Research on the Lepidoptera 25(1): 15-24.

Opler, P.A. 1999. A Field Guide to Western Butterflies. Houghton Mifflin Co., Boston. 540 pp., 44 pis.

Pelham, Jonathan P. 2008. A catalogue of the butterflies of the United States and Canada: with a complete

bibliography of the descriptive and systematic literature. Journal of Research on the Lepidoptera 40: xiv +

658pp.

Scott, James A. and Michael S. Fisher. 2008. Anthocharis “sara” group, especially in Colorado and vicinity

(with new research from Todd Stout) (one ssp. coauthored by Norbert G. Kondla). Pp. 1-14, pi. 2, in: J.

Scott (Ed.), Geographic variation and new taxa of Western North American butterflies, especially from

Colorado. Papilio (New Series) 18 {3 Dec 2008}

Warren, Andrew D. 2005. Butterflies of Oregon: Their taxonomy, distribution, and biology. Fort Collins;

Contributions of the C.P. Gillette Museum of Arthropod Diversity, Colorado State University: 405 pp., 3

maps {15 Mar 2005}

11

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12

Volume 7 Number 5

20 May 2010

The Taxonomic Report

OF THE INTERNATIONAL LEPIDOPTERA SURVEY

The Nomenclatural Status of Ten Names

in the Genus Pieris (Lepidoptera: Pieridae)

>0 - 17 Avenue SE, Calgary, Alberta T2A 0W6 Canada eg

clatural status of ten names in the genus Pieris are r.

■a Verity, 1911 is an in

teand location of pi

:dunnoughii Remington, 1954 is the correct and original^spelling; Miller and Brown (1981) provideThe incorrect subsequent

Hing mcdunnoughi. The name passosi Warren, 1968 is an available species-group name. A lectotype for passosi is

nenclature (ICZN) 1999; henceforth referee

le is not “law”, but is the ca

TABLE OF CONTENTS

The topics addressed in this paper, with their page numbers, are:

1. PSEUDOBRYONIAE VERITY, 1908.2

2. ADALWINDA FRUHSTORFER, 1909.4

3. PSEUDOBRYONIAE BARNES AND McDUNNOUGH, 1916.5

4. ARCTICA VERITY, 1911.7

5. arctica Barnes and McDunnough, 1916.7

6. pseudonapi Verity.8

7. MA CDUNNOUGHII REMINGTON, 1954.9

8. passosi Warren, 1968.11

9. PALLIDISSIMA BARNES & MCDUNNOUGH, 1916.13

10. ANGELIKA ElTSCHBERGER, 1983.13

11. PUBLICATION DATE OF ElTSCHBERGER’S BOOK SYSTEMA TISCHE UNTERSUCHUNGENAM PlERIS NAPI-

BRYONIAE-KOMPLEX (S. L.) (LEPIDOPTERA, PlERIDAE) .14

12. Summary of Conclusions.16

ANALYSIS OF THE NAMES

1. PSEUDOBR YONIAE VERITY, 1908

Kudma (1983) catalogued (p. 60) this name as:

pseudobryoniae (fm) - Pieris napifrigidapseudobryoniae Verity, 1908 - 010:

146- [U.S.A.]: Alaska: Nulato; Scandinavia: [N. Norway]: Finmark.

where the “(fm)” indicates that Kudma considered the taxon to be an infrasubspecific form. We agree with Kudma’s

conclusion but the nomenclatural saga of the word “pseudobryoniae” does not end there, as detailed below.

Verity described Pieris napi var. frigida form pseudobryoniae on page 146 [publication date 31 Jan 1908 '] of Verity

(1905-1911), with the phrase

“certains exemplaires se rapprochent cependant de bryoniae par leurs nervures larges et diffuses et meritent

le nom de pseudobryoniae (fig. 36 et 37).”

This can be translated to English as

“Certain examples nevertheless compare themselves to bryoniae by their wide and diffuse veins and merit

the name of pseudobryoniae (fig. 36 and 37).”

The species level taxonomy of “ Pieris napi ” and the variety ‘ frigida ” are clearly stated by Verity, and he indicates

that the name pseudobryoniae applies only to certain individual examples of var. frigida. The name pseudobryoniae

Verity, 1908 is therefore infrasubspecific, and is not available under the Code, Article 10.2:

“10.2. Availability of infrasubspecific names. An infrasubspecific name is not available [Art. 45.5] from

its original publication, unless it was published before 1961 for a “variety” or “form” and is deemed to be

available under Art. 45.6.4.1. If an author uses a name, previously published at infrasubspecific rank, in a

way which makes it available for a species or subspecies, that author thereby establishes it as a new name i

and it takes his or her authorship [Art. 45.5.1] (see also Articles 23.3.4 and 50.3.1)”

Several other Articles are cited in Article 10.2. Article 45.5 includes the statement that “A fourth name published as

an addition to a trinomen automatically denotes an infrasubspecific entity”. Article 45.6.4.1 is only applicable if the

name is deemed by the Code to not be infrasubspecific under Article 45.6.4, which is not the case for pseudobryoniae

Verity, 1908. The application of Articles 45.5.1, 23.3.4 and 50.3.1, which deal with the consequences of a later

author using an infrasubspecific name at the subspecies or species level, are not relevant to the availability of

1 Publication dates for parts of Verity (1905-1911) are provided by Kudma (1983), which is in part based on Verity (1914).

2

pseudobryoniae Verity, 1908, as discussed in the next section. Therefore, the name pseudobryoniae Verity, 1908 is

an infrasubspecific name not available for taxonomic use under the Code.

Verity’s figures of two specimens of pseudobryoniae , with the associated figure captions were published on 30 Apr

1909 as Plate XXXII Figures 36, 37. Neither specimen of pseudobryoniae Verity, 1908 was identified as the “type”

in the original description. However, Verity (1905-1911) starts with an Index to all taxa in the work; the Index has

the publication date of 31 Oct 1911. The explanatory heading on page XIII of the Index is shown in Figure 1, and the

entry on page XXVIII for the species Pieris napi, subspecies frigida, race arctica, form pseudobryoniae is shown in

Figure 2. The explanatory heading for the Index (Fig. 1) states that an asterisk, as in “XXXII, 37*”, specifies that

figure 37 is of the “type” specimen. Therefore, this index entry is the designation, by Verity in 1911, of the specimen

illustrated in his Plate XXXII Figure 37 (Fig. 4, 5) as the “type” of form pseudobryoniae.

INDEX SYSTEMATIQUE ET TABLEAU SYNOPTIQUE

DE LA VARIATION ET DE LA DISTRIBUTION GEOGRAPHIQUE

GENRE

sous-espece

forme (morpha)

aberration

aberration abortive ou strictement pathologique

Figure 1. Heading to the Index (Verity 1905-1911).

Figure 2. Index entry for pseudobryonieae (Verity 1905-1911).

[the phrase “[XLVII, 16 *-17” pertains to the previous line of the legend, taxon arctica]

Verity’s index + text + illustrations that were published on 31 October 1911 are all part of the same work, therefore

the index is part of the original descriptions of the new taxa that were named at that time. Use of the asterisk to

indicate "type" is “holotype” designation for the new taxa published 31 October 1911. However, this particular

“type” was designated in 1911, three years after the 1908 description of the taxon. Hence it would be the designation

of a “lectotype” by Verity, if the Code applied to an infrasubspecific name (which it does not). This “lectotype” is a

specimen from “Finmark, Scandinavie”, which is the type locality. According to Kudma (1983), “Finmark”

corresponds to northern Norway. However, this “lectotype” designation by Verity was actually without meaning

because it was preceded by a “lectotype” designation by Fruhstorfer (1909), as discussed below. In any case the

“lectotype” designation is moot, because pseudobryoniae Verity, 1908 is an infrasubspecific name and hence is

unavailable under the Code; hence terms such as “lectotype” are not actually applicable.

3

Figure 3. Plate VI, Figures 42bb and

43b of Wright (1905).

Figure 4. Figures 36 and 37 from Plate XXXII (Verity, 1905-1911).

30.

31.

32.

33.

34.

35.

36.

37.

P. napi, L. var. frigida, Scudd. S (Kamtchatka) [coll. Deckert]

Id. S (He d’Yesso, Japon) [coll, de Joannis]

Id. S (Norvege sept.) [coll. Stephanelli]

Id. $ (Nulato, Alaska) [coll, de Joannis]

Id. S Revers (Kamtchatka) [coll. Obth.]

Id. $ (Kamtchatka) [coll. Deckert]

Id. $ (Nulato Alaska) [coll, de Joannis]

Id. $ (Finmark, Scandinavie) [coll. Obth.]

146

146

146

146

146

146

146

146

Figure 5. Legends for Plate XXXII, Figures 30 to 37 (Verity, 1905-1911).

2. ADALWINDA FRUHSTORFER, 1909

Frahstorfer (1909), after seeing Plate 32 of Verity (1905-1911) that was published on 30 Apr 1909 (see above),

determined that Verity’s two illustrated specimens represent two separate taxa:

"Unter dem Namen pseudobryoniae vereigt Verity pag. 146 und t. 32 f. 36 und 37 zwei heterogene

Rassen aus Alaska (Type) und Finnmarken. Letztere ist viel groBer und steht naturlich der alpinen

bryoniae O., namentlich der f. obsoleta Rober viel naher als der nordamerikanischen Polarform. Fur

unsere nordische Rasse, charakterisiert durch seisslichere Grandfarbe und im distalen Teil der

Vorderflugel viel dunklere braune Flecke fuhre den Namen adalwinda ein."

This can be translated to English as:

“Under the name pseudobryoniae Verity, page 146 and Plate 32 Figures 36 and 37, is combined two

heterogeneous races from Alaska (Type) and Finland. The latter is much larger and stands naturally

much closer to the alpine bryoniae O., namely / obsoleta Rober, than to the North American polar

form. For our northern race, characterized through whitish ground colour and in the distal part of the

forewings much darker brown spots, is given the name adalwinda

Frahstorfer (1909) designates the specimen illustrated in Verity’s Figure 36 (from Nulato, Alaska) as the “type” of

pseudobryoniae Verity, 1908. This “type” is the first “lectotype” of pseudobryoniae , with publication date priority

over the “lectotype” designation of Verity (1911), which was discussed above. This is of no importance, given that

pseudobryoniae Verity, 1908 is an unavailable infrasubspecific name.

Frahstorfer (1909) called pseudobryoniae Verity a “race” from Alaska, and he clearly considers pseudobryoniae

Verity to have the same taxonomic rank as what he names as race adalwinda. A “race” named at that time is

equivalent to “subspecies” under Article 45.6 of the Code unless there is clear evidence to the contrary. There are

two pieces of information that provide such contrary evidence. First, in the remainder of his paper Frahstorfer (1909)

proceeds to describe four new subspecies in the genus Pieris, using the headings “Pieris napi leovigilda nov.

subspec.”, “ Pieris napi nesis nov. subspec.”, “ Pieris rapae micipsa nov. subspec.” and “ Pieris rapae lysicles nov.

4

subspec.” He therefore clearly and consistently used the genus-species-subspecies trinomial concept, with explicit

use of the term “subspecies” in contrast to his previous use of “race”. Second, Fruhstorfer references race

pseudobryoniae Verity in the same taxonomic context as when he names race adalwinda, and hence the taxonomic

placement of pseudobryoniae and adalwinda is that used by Verity - as a quadrinomial inlfasubspecific taxon. These

two lines of evidence demonstrate that Fruhstorfer deliberately and consistently uses the term subspecies in a

trinomial name, and used “race” for pseudobryoniae and adalwinda as a quadrinomial inlfasubspecific taxon.

The names pseudobryoniae and adalwinda Fruhstorfer, 1909 are therefore not available as species-group names

because they are inlfasubspecific names. We leave it to other taxonomists to decide whether to apply to the

International Commission on Zoological Nomenclature to take action to conserve the name adalwinda Fruhstorfer,

1909 in the interest of nomenclatural stability, as recommended by Kudma (1986).

3. PSEUDOBRYONIAE BARNES AND McDlJNNOUGH, 1916

Barnes and McDunnough (1916) reviewed the North American Pieris, and stated:

“In the extreme north [of North America] three distinct forms are separable; in the inland Arctic region

(Barren Plains) we have the form arctica Verity with strongly blackish marked veins on the underside in

both sexes and on the upper side in the $; there is however no suffusion of black and the markings are clear

cut; we figure a $ and $ from Chatanika, Alaska (Figs. 6, 7). Along the Alaskan coast we meet with the

form pseudobryoniae Verity which is what has been considered until recently to be bryoniae Ochs., a race

now restricted to the Alps of Europe; Wright’s figures (Butt. W. Coast PI. VI, Fig. 43b and 42bb) are typical

of the variation of the $. On the numerous islands of the Behring Sea and Alaskan coast the form hulda

Edw., is found in which the secondaries on the under side are almost totally suffused with greenish in the S

sex, leaving only dashes of yellowish ground color; the $’s are usually less suffused and on the upper side

are intermediate between arctica and pseudobryoniae ; we figure a S underside and $ upperside (Figs. 8,

9)”.

They do not illustrate pseudobryoniae , however in the figure captions they treat arctica and hulda as subspecies

names. Furthermore they state that “along the Alaskan coast” Wright’s figures “are typical of the variation [of

pseudobryoniae\\ They clearly considered pseudobryoniae to be a geographic subspecies that occurs along at least

part of the Alaskan coast, at the same taxonomic level as subspecies arctica and subspecies hulda. In this paragraph

they are using the word “form” with the generalized meaning of “phenotype”, not “form” in the taxonomic sense.

This is relevant in relation to Code Article 45.5.1:

“45.5.1 A name that has infrasubspecific rank under the provisions of this Article cannot be made 1

I available from its original publication by any subsequent action (such as "elevation in rank") except by a |

ruling of the Commission. When a subsequent author applies the same word to a species or subspecies in

a manner that makes it an available name [Arts. 11-18], even if he or she attributes authorship of the name

to the author of its publication as an infrasubspecific name, that subsequent author thereby establishes a

new name with its o wn auth orship and date.”

The first sentence means that an infrasubspecific name can only be elevated in rank through a ruling of the

Commission.

(a) therefore pseudobryoniae Verity cannot be “elevated in rank” by Barnes and McDunnough (1916); and

(b) the action taken by Barnes and McDunnough (1916) is the establishment of a new name, not the elevation of

Verity’s name.

The second sentence says that:

(a) the same word that was used for the inffasubspecific name can be used by a new author, to establish a new

(b) the establishment of the new name must conform to Articles 11-18; and

(c) it is irrelevant whether the new author thinks he is using someone else’s species-group name.

The name pseudobryoniae Barnes and McDunnough, 1916 meets the requirements of Articles 11-18, of which

Articles 13-18 are not relevant. All the provisions of Article 11 are met. For Article 12, Barnes and McDunnough

(1916) do not provide even the slightest hint of a description or definition. However, Barnes and McDunnough

5

provide a clear indication by bibliographic reference to specific illustrations in a specific publication by Wright

(Article 12.2.7); therefore the specimens represented by those illustrations, reproduced in Figure 3, are syntypes of

pseudobryoniae Barnes and McDunnough, 1916 and the name is available through that indication. The relevant parts

of Article 12 are:

Article 12. Names published before 1931.

12.1. Requirements. To be available, every new name published before 1931 must satisfy the

provisions of Article 11 and must be accompanied by a description or a definition of the taxon that it

denotes, or by an indication.

12.2. Indications. For the purposes of this Article the word "indication" denotes only the

following:

12.2.1. a bibliographic reference to a previously published description or definition even if the

description or definition is contained in a work published before 1758, or that is not

consistently binominal, or that has been suppressed by the Commission (unless the

Commission has ruled that the work is to be treated as not having been published [Art. 8.7])

12.2.7. the proposal of a new genus-group name or of a new species-group name in association

with an illustration of the taxon being named, or with a bibliographic reference to such an

illustration, even if the illustration is contained in a work published before 1758, or in one that is

not consistently binominal, or in one that has been suppressed by the Commission (unless the

Commission has ruled that the work is to be treated as not having been published [Art. 8.7])

The reference to “ pseudobryoniae Verity” in the above quotation is not a clear indication in itself. However, the first

line of the Barnes and McDunnough’s (1916) treatment of Pieris napi states: “Verity has lately (Rhop. Pal. Vol. I)

dealt at considerable length with the various races and forms of this species; we offer the following remarks as to the

arrangement of our North American races as it is probable that Verity’s work is inaccessible to the majority of

American entomologists”. Hence, Barnes and McDunnough gave an adequate bibliographic reference for

pseudobryoniae and Article 12.2.1 is applicable. Therefore the two specimens on which the name pseudobryoniae

Verity was based (Figure 4) are also syntypes of pseudobryoniae Barnes and McDunnough, 1916 and the name is

also available through that indication.

Barnes and McDunnough (1916) also imply that they examined other specimens that they considered to be

pseudobryoniae , if so, those specimens are also syntypes (Article 72.4.1). The location of these syntypes, if they are

identifiable, is unknown to us.

Therefore pseudobryoniae Barnes and McDunnough, 1916 is an available name, and the syntypes are the two

specimens illustrated by Verity (1905-1911), plus the two specimens illustrated by Wright (1905), plus any other

specimens (identity and location unknown) that Barnes and McDunnough examined and considered to be

pseudobryoniae (Article 72.4.1).

The type series of pseudobryoniae of Barnes and McDunnough likely contains more than one taxon (Eitschberger

1983), therefore a lectotype needs to be designated to provide both taxonomic clarity and foster nomenclatural

stability. We therefore designate the specimen illustrated in Plate 32 Figure 36 of Verity (1905-1911), reproduced

here in Figure 4, as the lectotype of Pieris napi pseudobryoniae Barnes and McDunnough, 1916, with the type

locality being Nulato, Alaska. This is consistent with historical, although irregular, use of the name pseudobryoniae ,

with various authors attributed to it.

In our opinion, Pieris marginalis browni Eitschberger, 1983 (Type Locality: Seward Peninsula, Alaska) is a junior

subjective synonym of Pieris napi pseudobryoniae Barnes and McDunnough, 1916. Eitschberger (1983, p. 349)

recognized that Plate 32 Fig. 36 represented his taxon browni , but did not recognize the availability of the name

pseudobryoniae Barnes and McDunnough, 1916.

After Barnes and McDunnough (1916), other authors also used the word pseudobryoniae as a subspecies-level name

and, through indication to Verity (1905-1911), met the Code requirements for to make the name available with their

new authorship and date (e.g. dos Passos 1965). These later uses of the name are not available names, because the

author of a name is the person who first publishes it (Article 50); in this case Barnes and McDunnough (1916).

6

4. arctica Verity, 1911

Kudma (1983) catalogued (p. 60) this name as:

Ida arctica Verity, 1911 - 020:334 - Scandinavia: [N.

• & Kautz (1939): Pieris arctica [nec Verity], species.

on page 334

31 Oct 1911] of Verity (1905-

d’eloigner une fo

lie des Alpes. Mes figures 32 et 33 (PI. XXII [sic - actually XXXII] auxquelles j’ajoute deux

s types dc ,- y (PI. LXVII, fig. 16 et 17) donneront une idee exacte de cette race et la

.arison du 3 (fig- 32) avec le <? alpin (fig. 25) Pen distinguee bien.”

This can be translated to English as

it

by the name of arctica in

distinguished from the one of the Alps. My figures 32 and 33 (PI. XXII [sic - actually XXXII]), to

which I add two other typical ones (PI. LXVII, fig. 16 and 17), will give an exact idea of this race

and its comparison (Fig. 32) with the alpine one (fig. 25) will distinguish it well.”

is Verity’s Plate LXVII and its legends for figures 16 and 17 (Fig. 6

P. napi var. arctica Verity, 1911. In isolation this arctica name is

it is

on page 334, and Plate LXVII were al

The Index and the text on page 334

The above evidence demonstrates that the name arctica Verity, 1911 is infrasubspecific and therefore is not available

under the Code (Article 10.2).

16. P. napi, L. var. arctica. Verity $ (Saltdalen, Norvege sept.) [e coll. Murray]

| 17. Id. $ (Laponie) [e coll. Leech]

Figure 6. Plate LXVII legends for arctica (Verity 1905-1911)

5. arctica Barnes and McDunnough, 1916

The name arctica Bames and McDunnough, 1916 meets the requirements of Articles 11-18, of which Articles 13-18

are not relevant. All the relevant provisions of Article 11 are met. For Article 12, Bames and McDunnough (1916)

provide a definition and figures of two syntypes from Chatanika, Alaska. However, they also provide an indication to

Verity’s original description, including the figures, through the same mechanism discussed above in Section 3 for

pseudobryoniae. Therefore the specimens represented by the Verity’s illustrations for his arctica are also syntypes of

arctica Barnes and McDunnough, 1916.

Bames and McDunnough (1916) also imply that they examined other specimens that they considered to be arctica ; if

so, those specimens are also syntypes (Article 72.4.1). Their identity and present location is unknown.

Therefore arctica Bames and McDunnough, 1916 is an available name, and the syntypes are the four specimens

illustrated by Verity (1905-1911), plus the two specimens illustrated by Bames and McDunnough (1916), plus any

other specimens (identity currently unknown) that Bames and McDunnough examined and considered to be arctica

(Article 72.4.1).

The locations from which the six known syntypes of arctica Bames and McDunnough originate range from

Scandinavia to Alaska, and more than one taxon is likely represented; hence designation of a lectotype is required for

taxonomic clarity and to stabilize the nomenclature for on-going revisions of the genus Pier is. Bames and

McDunnough (1916) considered the name arctica to be represented by the “types” of the unavailable name arctica

Verity. Therefore, we designate the lectotype of arctica Bames and McDunnough, 1916 to be the specimen

represented by Plate XXXII Figure 32 of Verity (1905-1911). This is the specimen designated as the “holotype” of

arctica Verity and shown above (Fig. 4), with the type locality being “Norvege sept.”, which is “Scandinavia: N.

Norway” according to Kudma (1983).

It could be argued that the “holotype” of arctica Verity is automatically the holotype of arctica Bames and

McDunnough. This would certainly be the case if arctica Verity was an available name being replaced by another

name due to homonymy. However, the circumstance of a name being available through indication of the description

of an infrasubspecific name is not addressed by the Code. Hence, designation of a lectotype that is the same as the

putative holotype under an alternative interpretation of the Code (with which we disagree) achieves the objective of

nomenclatural stability.

The name arctica Bames and McDunnough, 1916 is the available name for the European populations to which the

unavailable name adalwinda Fmhstorfer, 1909 is presently applied.

After Bames and McDunnough (1916), other authors also used the word arctica as a subspecies-level name and,

through indication to Verity (1905-1911), met the Code requirements for to make the name available with their new

authorship and date (e.g. dos Passos 1965). These later uses of the name are not available names, because the author

of a name is the person who first publishes it (Article 50); in this case Bames and McDunnough (1916).

6. pseudonapi Verity

Kudma (1983, p. 60) catalogued this name as:

pseudonapi (ra) - Pieris melete melete pseudonapi Verity, 1911 - 010:330 - Japan:

Yezo [= Hokkaido]: Ichikiri.

where the “(ra)” indicated that Kudma believed that Verity had described the taxon with the rank of “race”, and that

the taxon is infrasubspecific and the name is not available (the reasoning behind this conclusion is doubtful in the

context of the Code, but does not need to be discussed here). This conclusion suggested that the name pseudonapi

Bames and McDunnough, 1916 had been incorrectly determined to be a homonym by Remington (1954), who

replaced it with the name macdunnoughii. However, Kudma’s conclusion was incorrect because the text on page 330

of Verity (1905-1911) was not the actual original description of pseudonapi.

Verity’s Plate LIX and the associated figure legend (Figures 7 and 8 below) were published on 31 Jan 1911. The

plate and figure legend together are a valid original description (Code Articles 12.1 and 12.2.7), and that description

has date priority over the 31 Oct 1911 text description on page 330 of Verity. The legend for Plate LIX Figures 13-17

used the term “var.” (variety) to indicate the rank of pseudonapi , which the Code specifically states must be

considered to be equivalent to subspecies rank (Article 45.6.4), in the absence of clear evidence to the contrary - as

in this case. The name pseudonapi. Verity, 1909 is therefore available, and the specimens represented by Verity’s

Plate LIX Figures 13-17 are the syntypes.

The name pseudonapi Barnes & McDunnough, 1916 is therefore a primary homonym of pseudonapi , Verity, 1911,

validly replaced by macdunnoughii Remington, 1954.

Figure 7. Copy of the figures of the syntypes of P. melete var. pseudonapi (Verity, 1905-1911, Plate LIX)

Figure numbers re-typed for clarity. Figure legends shown above (Figure 7).

13. P

melete, M6n. var. pseudonapi, Verity cf (lekikishiri, Yesso, Japon)

14.

Id.

d Revers (lekikishiri, Yesso, Japon)

15.

Id.

$ (lekikishiri, Yesso, Japon)

16.

Id.

? (lekikishiri, Yesso, Japon)

17.

Id.

$ Revers (Ichikiskiri, Yesso, Japon)

Figure 8. Copy of the figure captions for P. melete var. pseudonapi from Verity’s Plate LIX

7. MACDUNNOUGHII REMINGTON, 1954

The action of Remington (1954) in replacing the North American name pseudonapi McDunnough, 1916 was correct,

because of homonymy, although he cited the wrong date and location (1911, p. 330) for the original description of

pseudonapi Verity (1 Jan 1911, Plate LIX Figures 13-17; see above). Regardless of the publication date, pseudonapi

McDunnough, 1916 is a primary homonym of the available name pseudonapi Verity, 1911, and hence a new name

such as macdunnoughii was required to replace it.

The spelling macdunnoughii , used by dos Passos (1964), was exactly the same as appeared in the original description

by Remington (1954). It therefore could not be an “unjustified emendation” (Code Article 33.2.3) as stated by Miller

and Brown (1981) in the note for their checklist entry “ mcdunnoughi ”: “[Note] 259. Unjustifiably emended to

“macdunnoughi ” [sz'c] by dos Passos, Mem. Lepid. Soc. (1): 40 (1964).”

Miller and Brown (1981) use the spelling mcdunnoughi ; which has two spelling changes from the original

description - a change of “mac” to “me” and a change of double “z'z” to single “z”. The relevant sections of the Code

33.2. Emendations. Any demonstrably intentional change in the original spelling of a name other

than a mandatory change is an "emendation", except as provided in Article 33.4.33.2.1. A change in

the original spelling of a name is only to be interpreted as "demonstrably intentional" when in the

work itself, or in an author's (or publisher's) corrigenda, there is an explicit statement of intention, or

when both the original and the changed spelling are cited and the latter is adopted in place of the

former, or when two or more names in the same work are treated in a similar way.

>ued that, in parallel to the Code example, the statement in the paper that the taxon was n

of “Me” to “ mac ” is part of the latinization

of the

“21. Personal names bearing prefixes should be treated as follows in forming zoological names:

(a) The prefixes "Mac", "Me", or "M" should be spelled "mac" and united, as in maccooki (McCook),

macoyi (M'Coy).” _

Incorrect latinization is specifically stated to be not considered an inadvertent error, and in this case the spelling

change was clearly correct latinization by the standard of the day. As a point of interest, the same requirement is still

8. PASSOSI B. Warren, 1968

Figure 9. Syntype of Pieris passosi

Scale larger than life-size; forewing span = 42:

9. PALLIDISSIMA BARNES & McDUNNOUGH, 1916

Bames and McDunnough (1916) described the taxon pallidissima with the words:

“In Utah we meet with a second generation (July, August) which is extremely pale, being practically

immaculate in both sexes on both sides; the underside is tinged with pale yellow on secondaries and apex of

primaries and the $ on the upperside of primaries shows faint traces of upper black spot; it is a further

development of castoria apparently differing from both this form and pallida in the reduction of the black

spots in the ?; we propose the name PALLIDISSIMA for the race and figure the type $ and $ from Provo,

Utah (Figs. 4, 5, 10).”

The captions for the figures of pallidissima are clearly in trinomial form, with pallidissima treated as a subspecies

(Plate VI, Figs. 4, 5, 10), as shown in Figure 10.

Fig. 3. Pieris napi pseudonapi B. & McD. Paratype, 9 Silverton, Colo.

Fig, 4. Pieris napi pallidissima B. & McD. Type, S Provo, Utah.

Fig. 5. Pieris napi pallidissima B . & McD . Type, 9 Provo, Utah.

Fig. 6. Pieris napi arctica Verity. $ Chatanika, Alaska.

F'ig. 7. Pieris napi arctica Verity. 9 Chatanika, Alaska.

Fig. 8. Pieris napi hulda Edw. 9 Pribilof Is., Alaska.

Fig. 9. Pieris napi hulda Edzv. underside Pribilof Is., Alaska.

Fig. 10. Pieris napi pallidissima B. & McD. S, underside Provo, Utah.

Figure 10. captions for the figures of Pieris napi pallidissima

The phrasing of the text could be misinterpreted, without close examination, to indicate that pallidissima was named

as the summer form of the Utah populations; this was the interpretation of Remington (1954). However, the use of

the word “race”, combined with the format of the captions of the figures, makes it clear that pallidissima was named

as a geographically defined group of populations - a subspecies - that is characterized by the appearance of the

summer generation.

Miller and Brown (1981) and Pelham (2008) asserted that there is a holotype of pallidissima. However this is

impossible because (1) no holotype was explicitly designated in the original description, and (2) the name is based on

more than one specimen and hence holotype designation by monotypy does not apply (Article 73.1). The specimens

in the type series therefore are all syntypes.

If the taxonomic decision is made that the Utah and Colorado populations of Pieris marginalis are the same taxon, as

suggested by authors such as Remington (1954) and Warren (1968), then macdunnoughii Remington, 1954 may

become a subjective synonym of pallidissima Bames & McDunnough, 1916. Given the taxonomic uncertainty

related to Pieris populations in western North America; we consider it essential to have a clear and objective standard

of reference for the name pallidissima. We therefore designate the specimen illustrated in Plate VI Figure 4 of Bames

and McDunnough (1916) to be the lectotype of the name pallidissima.

10. ANGELIKA ElTSCHBERGER, 1983

The name Pieris angelika was first proposed by Eitschberger (1981). When translated to English, the original text

“4) Pieris angelika angelika n. spec.

This species so far has gone under the unjustified name of Pieris napi pseudobryoniae auct. (not VERITY,

1908) .... The populations from Alaska and Northwestern Canada I hereby call Pieris angelika angelika n.

spec, after the name of my wife, who not only suffers Entomology, but is rather actively involved in

promoting and supporting my work. Of this species, there are so far, from diverse localities in the above

named regions nearly 200 males and females in the coll. EITSCHBERGER-STEINIGER. Further material

is at hand from various private and museum collections. Exact analysis and description of this species

follows in the earlier mentioned revision. But in order to already determine the species at this time, a few

black and white photographs shall be shown here. These specimens, as well as all other taxa of the napi-

13

r (1983) are, in our

glacial period was likely spread through much of the ice-free area that extended from the

eastern Siberia (= Beringia), with a land connection where the Bering Strait is now present. T

TL: Keno (el. 4600 feet), Yukon,

b TL S } cl ^ t ! m j Steri Klt ^ cllhcrgcr '

11. PUBLICATION DATE OF ElTSCHBERGER’ S BOOK SYSTEMA TISCHE UNTERSUCHUNGENAM

PlERIS NAPI-BRYONIAE-KOMPLEX (S. L.) (LEPIDOPTERA, PlERIDAE)

14

(ICZN), Article 21 and 22.

The 1964 edition of the Code, which was in force when Reissenger published his opinion, states:

Article 9. What does not constitute publication. - None of the following acts constitutes publication

within the meaning of the Code:

(6) mere deposit of a document in a libra ry. _

Therefore, under the 1964 Code, if only the one copy had been deposited in the library, it would not have constituted

Definition:

date of publication, n. Of a work (and of a contained name and nomenclatural act): the date on which

copies of the work become available by purchase or free distribution. If the actual date is not known, the

date to be adopted is regulated by the provisions of Article 21,2-7. _

The criteria of ‘delivery and shipping’ and ‘general public accessibility’ used by Reissinger (1986) are not Code

criteria for establishing date of publication. The Code only requires that copies ‘become available’, and it has been

21.2. Date specified. The date of publication specified in a work is to be adopted as correct in the absence

of evidence to the contrary.

21.3. Date incompletely specified. If the day of publication is not specified in a work, the earliest day on

which the work is demonstrated to be in existence as a published work is to be adopted as the date of

publication, but in the absence of such evidence the date to be adopted is

21.3.1. the last day of the month, when month and year, but not day, are specified or demonstrated, or

_21.3.2. the last day of the year when only the year is specified or demonstrated._

We are not aware of any other taxonomic publication regarding the genus Pieris in December 1983; therefore further

refinement of the date is unnecessary, even if possible.

15

(1983):

corrected from 1911 to 1909.

4. The name arctica Verity, 1911 is an unavailable infrasubspecific name.

The name macdunnoughii Remington, 1954 is the correct and original spelling

(1954);^the spelling macdunnougM by dos Passos^(l%4) ^correct and is not an

Eitschberger, 1983.

16

ik were distributed during December 1983, demonstrating the Code requirement of

ts of the Code for a>

(1985),

(1985), and Ferris (1989).

i he § may disagree with sonw^of the Conclusions), Philip Tubbs Cf the

Articles regarding hybrids, Kenelm Philip for providing color illustration;

excellent review comments, and Ulf Eitschberger for clarifying the dates of pr

y„* i

r, Ulf 1981. Die m

Arten aus der Pieris napi-bryoniae-Gmppe (lej

le group (Lep., Pieridae)]. Atalanta 11(5):366-71.

(Lep., Pieridae) [The

by Roger

lex (s.l.)” (I

■, L.D., a

;)byU

. 1981. A c

zu Kudmas & Geigers "A Critical Review

- Lepid. Pieridae). Atalanta (Markleuthen)

17

Shapiro, A.M. 1985. Book review. Journal of the Lepidopterists’ Society 38(4):324-27.

Verity, Ruggero. 1905-1911. Rhopalocera palaearctica, Iconographia et description des papillons diumes de la region

palearctique par Roger Verity (Papilionidae et Pieridae). Florence, publ. by author. 368 pp., 72 pis.

Verity, Ruggero. 1914. Dates of publication of "Rhopalocera palaearctica, Iconographia et description des papillons

diumes de la region palearctique par Roger Verity (Papilionidae et Pieridae)." Novitates Zoologicae 21:426.

Warren, B.C.S. 1968. On the Nearctic species of the bryoniae- and oleracea -groups of the genus Pier is.

Entomolgist’s Record and Journal of Variation 80:60-66.

Wright, W.G. 1905. The butterflies of the west coast of the United States. San Bernardino, CA: W.G. Wright. 257 +

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19

Volume 7 Number 6

15 June 2014

The Taxonomic Report

OF THE INTERNATIONAL LEPIDOPTERA SURVEY

Larval host plants of Enodia anthedon, Satyrodes appalachia and

S. eurydice in Vermont, USA

David J. Hoag

173 West Shore Rd.

Grand Isle, Vermont 05458

Abstract: Field observation and captive rearing was used to clarify larval host plant use in Enodia anthedon,

Satyrodes appalachia and S. eurydice in Vermont, USA. In nature S. appalachia larvae were found on Carex

lacustris, C. lupulina , C. gracillima and C. tuckermanii. E. anthedon in nature was found to oviposit on grass and on

Carex lacustris. Larvae of E. anthedon were found on C. lacustris and C. lupulina. In captivity, all but two E.

anthedon larvae chose Carex over grass. Both E. appalachia and E. anthedon larvae thrived on a diet of C. lacustris

and C. lupulina in captivity. Early instar captive larvae refused C. sparganioides but late instar captive larvae

accepted said plant. Grass may be an acceptable alternate host for Vermont S. eurydice. Miscellaneous observations

on early instars are reported.

Additional key words: Lepidoptera, Nymphalidae, Satyridae, life cycle, habitat

INTRODUCTION

Various grass species and Carex sedges are listed respectively as host plants for Enodia anthedon and Satyrodes

appalachia by many authors, including Howe (1975). Scott (1986) wrote: [ anthedon ] “Hostplants grasses” and

[<appalachia ] “Larvae refused grasses in the lab.” Handfield (1999) lists eight grasses for E. anthedon, and Carex for

S. appalachia leeuwi. Nielsen (1999) and Douglas & Douglas (2005) similarly list various species of grasses for

anthedon vs. sedges for appalachia. O'Donnell, et al. (2007) list grass ( Dactylis glomerata ) for anthedon, and sedges

(incl. Carex stricta) for appalachia. Allen (1997) wrote: “Grasses are the primary host for [anthedon]” and “Third

instar larvae from the last brood overwinter in a rolled grass blade tied together with silk.” Allen describes appalachia

as overwintering in a similar way, but adds, “In West Virginia the Appalachian Brown also uses grasses as hosts.”

METHODS

I searched for ovipositing females and larvae in natural habitats during 2007, 2008, 2009 and 2010. I searched the

range of habitats used by E. anthedon and S. appalachia in the Grand Isle area. Larvae were raised, in containers, in

locations receiving partial sunlight - somewhat imitating wooded habitat. A screened narrow porch on the west side of

the house provided partial late afternoon sunlight; the duration of which was limited by trees to the west. Indoor

locations were used when high winds were thrashing the sedges and when close monitoring of larvae was desired (also

in cold months!): windows provided partial direct morning and afternoon light, with only a distant 60w bulb extending

light hours throughout each evening.

Figures 1-3. Host - Carex sedges. Fig. 1. 31-May-2009, small sixth-instar E. anthedon larva eating container-grown

sedge. Fig. 2. 30-Jun-2009: Typical habitat at the swamp, a 1.75 acre (0.71 hectare) vernal pool. Fig. 3. E. anthedon

male at swamp.

Figures 4-12. Life cycle of E. anthedon. Fig. 4. E. anthedon larvae hatching. Fig. 5. Recently hatched E. anthedon

larvae after first meals of sedge. Fig. 6. First-instar larvae: E. anthedon (one) & S. appalachia (two), 19-Jul-2009.

Fig. 7. Second-instar E. anthedon. Fig. 8. Third-instar E. anthedon. Fig. 9. Fourth-instars: E. anthedon (left) & S.

appalachia (right), on grass, 26-Aug-2009. Fig. 10. Sixth-instar E. anthedon , after molt from fifth-instar as collected

on Carex sedge, 4-Jun-2009. Fig. 11. Late sixth-instar E. anthedon. Fig. 12. E. anthedon female, ex-pupa 30-Jun-

2009.

2

Figures 13-15. S. appalachia. Fig. 13. Female (54.5mm), ex-ovum Jul-2008; ex-pupa 12-Jun-2009. Fig. 14. Various-

aged S. appalachia larvae and molted head capsules illustrating the often prominent dark line extending from the base

Figures 16-19. 5. appalachia. Fig. 16a. Male, ex-pupa 26-Aug-2008; from ovum laid 30-Jun or 01-Jul-2008. Fig.

16b. Female, netted at swamp 02-Sep-08. Fig. 16c. Male, netted at lake site 03-Sep-08. Fig. 16d. Male, from swamp

05-Sep-08. Fig. 17. Newly h;

RESULTS AND DISCUSSION

An E. anthedon was observed ovipositing on grass, the expected host, in the woodland near the swamp on July 27,

2008. However, in the same swamp, on Sept. 25, I collected three small anthedon larvae from Carex sedges in two

locations. These three larvae were stored over winter in my refrigerator, along with two dozen appalachia larvae. Due

to mold encroachment, some larvae were prematurely transferred to container sedge in late January, too early for

extended larval survival - and in late-February, when newly released appalachia larvae shortly resumed robust

growth. However, the two surviving anthedon larvae soon died - only one eating but one meal on Mar. 4, 2009.

Room temperature was 14°C (57°F).

Subsequent searches of the woodland swamp, May 30 to June 14, 2009, revealed ten post-diapause sedge-eating

anthedon larvae. Another was found on June 4, 2009, in a lakeside wet woodland 10k (5.4 miles) away. Two larvae

were monitored at the swamp; the other nine were collected, the smallest of which died; it appeared to be deformed, or

possibly was injured when collected. Another larva escaped from its container. Although two of the larvae had been

feeding on a fine-leaf grass, they were situated in sparse vegetation where they had initially eaten sedge. The two

undoubtedly would have returned to sedge - the available grass was insufficient to provide many meals. When

collected and offered a choice of sedge or the same grass species, the two larvae chose sedge. These seven surviving

container-bound larvae successfully matured on the sedge diet; the first two emerged from their pupae on June 25, the

same day that the first flight of anthedon (n = 4) was observed. The two uncollected larvae also pupated successfully

on their host sedges.

From early Sept, through Oct. 5, 2009, twelve pre-diapause anthedon larvae were found eating sedge at the swamp -

some of these larvae were found again in May of 2010.

The two woodland study sites (44.75000N, -073.30556W and 44.69391N, -073.34210W) for E. anthedon were

chosen for the presence of colonies of S. appalachia with which the anthedon associated. The nearby drier site, a

former pasture reverting to shrubs, was at 44.69548N, -073.34010W. Observed flight numbers in 2009 for anthedon

were low, perhaps related to frequent rainy weather; the flight seemed to be delayed a week or two. However, the one-

day maximum of 120-140 appalachia was higher at the lake site than numbers observed in previous years - in 2010,

an early first flight peaked at 250. The lake site is dominated by young ash ( Fraxinus pennsylvanica ) and elm ( Ulmus

americana) with a dense carpet of sedge. The woodland swamp is a 1.75 acre (0.71 hectare) vernal pool shaded by

red maples ( Acer rubrum), and surrounded by other mature trees and patchy forest-floor vegetation.

An interesting observation was made: At the dry upland site, up to 15 male anthedon clustered together on shrubbery,

instead of engaging in territorial aerial pursuit as observed in the two wet habitats. A few years previously, I had

observed similar communal behavior at the lake site. These Vermont sites fall within the “contact zone“ of E. a.

anthedon and E. a. borealis - taxa showing behavioral differences (Grkovich & Pavulaan, 2003).

Captive Rearing

In 2009,1 collected five female E. anthedon , and a few female S. appalachia to obtain ova. Two of the anthedon were

collected from the woodland swamp, two from the lakeside woodland (one was ovipositing on sedge), and the fifth

from a nearby dry upland habitat. I later noted no discernible differences among the anthedon larvae.

Of >65 E. anthedon and S. appalachia eggs, most were deposited on sedge (it being the dominant plant in each

container), one egg on grass, one on twine, and several inside the plastic cap of a support stake and on the sides of the

Rubbermaid® containers.

Larvae were free to chose between sedge and grass - all but two of the anthedon larvae opting for sedge. By late-

Aug., three anthedon larvae were feeding on grass - two having moved from sedge onto grass, while one of the

previous two had abandoned the grass. On Sept. 8, four were on grass, but some of the anthedon were already idle, in

diapause, as were most of the appalachia larvae sharing the containers. Two of the S. appalachia larvae were also

feeding on grass. One quit its grass diet for sedge in mid-Aug; the second appalachia remained on grass, molting to a

normal straw-colored 4th instar in late-Aug. All appalachia (n = 45-50) entered diapause as 4th instar larvae by Sept.

14, while six anthedon were still feeding. Larvae of both species ate at any hour of the day, the daytime feeding being

especially noticeable with the appalachia. Conversely, Cech and Tudor (2005) wrote, “[< appalachia ] Caterpillars feed

at night, hiding near hostplant base by day.”

Whereas most authors state that larval diapause occurs with the third or fourth instar, Layberry, et al. (1998) wrote,

“[anthedon] larvae overwinter in the first instar.” Watching and measuring each anthedon larva, I attempted to

confirm that every larva molted from third to fourth instar prior to diapause, and am confident that all larvae did. By

Sept. 14, the six ex-ova anthedon larvae still eating sedge (five) and grass (one) had reached their maximum instar

length: 18.5-19mm. The two smallest of the five larvae collected from the swamp molted to fourth instar on Sept. 13

and 17. The lack of frost in September ensured that all larvae had time to enter the fourth instar. In most years, the

surrounding Lake Champlain buffers Grand Isle from early frost. The average first frosts normally arrive in mid-Oct.

In all years, none of the diapausing larvae, anthedon or appalachia , made any attempt to create a shelter of rolled

leaves as reported by Allen (1997), although in 2008 some appalachia did hide under dried deciduous leaves, rather

than stay on sedge stems and leaves. I did notice that occasionally a cut section of refrigerated sedge leaf, while

drying, by chance would partially curl around the attached larva - the larva’s silk track having no influence on the

direction of the curl. It was also noted that anthedon (and appalachia) larval length shrinks during diapause.

When appalachia larvae resumed growth in springtime, they required two more molts prior to pupation. In 2008, the

one container-raised appalachia larva that skipped diapause, apparently did skip one late larval instar; I saw no

evidence of the missing (and necessarily short-timed) molt. In 2009, one exceptional appalachia larva unexpectedly

molted to a large, boldly-striped SEVENTH-instar, but the ensuing pupa was slightly deformed and eclosion failed..

From my limited observations of May-June anthedon larvae, I noted only two molts occurring post-diapause - a total

of six larval instars (final max. length 41-43.5mm) - the same sequence as the local appalachia. Again in early 2010,

the same two post-diapause molts were observed for anthedon larvae which had survived fourth-instar diapause in my

refrigerator, and which successfully eclosed as adults, after feeding on sedge. Additionally, in 2010, a warm year in

which appalachia and anthedon had early (mid-August) second flights, I obtained some ova from a first flight

anthedon female, as well as from some appalachia. Three of nine surviving anthedon larvae skipped diapause,

eclosing in late August and early September.

Although the majority (50) of my first-brood appalachia larvae also skipped diapause in 2010, twenty-two entered

diapause at fourth-instar as I had anticipated. However, from a mated pair of the second-flight appalachia , six of the

seven ex-ova larvae entered diapause earlier, at third-instar; only one underwent one more molt.

Both species were again raised on C. lacustris and/or lupulina, with access to a very limited amount of grass in one of

the four containers of sedge. Larvae of both species were observed eating sedge and grass, with no apparent

preference. Also present in two of the containers were C. sparganioides (Muhl. ex Willd) and an unidentified common

wide-leaf sedge. Both of the latter sedges were refused by early-instar larvae, thus unlikely to be utilized in the wild -

although both, especially the sparganioides, were accepted by late-instar larvae.

Phenotypically, my anthedon larvae match published descriptions. However, larvae of this Vermont population of

appalachia leeuwi differ slightly from descriptions - the black stripe on the Vermont leeuwi head-capsule horns

typically extends beyond the base of the horns to the eyes (the stripe may vary in intensity, but is often very bold).

Carde, et al. (1970) and other authors indicate that for appalachia, unlike S. eurydice, the dark stripes stop at the base

of the horns.

CONCLUSION

Although sedges are not currently listed as host plants for E. anthedon. Car ex species are accepted by, and may be the

preferred host of, this northern New England population of anthedon. Oviposition by anthedon on C. lacustris, and

on a grass species, was observed, and larvae were found feeding on C. lacustris and C. lupulina. E. anthedon larvae

raised from ova thrived on a diet of C. lacustris and C. lupulina. Host sedges for larvae of S. appalachia in Grand

Isle include C. gracillima, C. lacustris, C. lupulina and C. tuckermanii.

ACKNOWLEDGMENTS

I thank Alex Grkovich, Norbert Kondla, Harry Pavulaan and David Wright for their encouragement and assistance in

writing this report.

REFERENCES

ALLEN, T. J. 1997. The Butterflies of West Virginia and Their Caterpillars. University of Pittsburgh Press,

Pittsburgh, PA. 388 pp.

CARDE, R. T„ A. M. SHAPIRO, & H. K. CLENCH. 1970. Sibling Species in the Eurydice Group of Lethe

(Lepidoptera: Satyridae). Psyche 77(1): 70-103.

CECH, R. & G. TUDOR. 2005. Butterflies of the East Coast - An Observer's Guide. Princeton University Press,

Princeton, N.J. 345 pp.

DOUGLAS, M. M. & J. M. DOUGLAS. 2005. Butterflies of the Great Lakes Region. University of Michigan

Press, Ann Arbor, MI. 345 pp.

GRKOVICH, A. & H. PAVULAAN. 2003. The Case for Taxonomic Recognition of the Taxon Enodia anthedon

borealis A. H. Clark (Satyridae). Taxonomic Report of The International Lepidoptera Survey 4(5): 1-15.

HANDFIELD, L. 1999. Le Guide des Papillons du Quebec. Vol. I. Broquet, Boucherville, Quebec. 982 pp.

HOWE, W. H. 1975. The Butterflies of North America. Doubleday & Co., Inc. Garden City, N.Y. 633 pp.

LAYBERRY, R. A., P. W. HALL, & J. D. LAFONTAINE. 1998. The Butterflies of Canada. University of Toronto

Press, Toronto, Ontario. 280 pp.

NIELSEN, M. C. 1999. Michigan Butterflies and Skippers - A Field Guide and Reference. Michigan State

University Extension, East Lansing, MI. 248 pp.

O'DONNELL, J. E., L. F. GALL, & D. L. WAGNER. 2007. The Connecticut Butterfly Atlas. State Geological and

Natural History Survey. Bulletin No. 118. 376 pp.

SCOTT, J. A. 1986. The Butterflies of North America - A Natural History and Field Guide. Stanford University

Press, Stanford, CA. 583 pp.

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Volume 7 Number 7

16 June 2014

A case of sympatric Celastrina ladon (Cramer),

Celastrina lucia (W. Kirby) and Celastrina neglecta (Edwards)

(Lycaenidae: Polyommatinae) in Northern Virginia,

with additional records of C. lucia in Virginia

Harry Pavulaan 1

P.O. Box 1124, Herndon, VA 20172

ABSTRACT. A case of fully sympatric Celastrina ladon , C. lucia and early spring brood C. neglecta is documented

at a site in northern Virginia. Observations indicate that all three species occupy the same habitat, fly during the same flight

period and utilize the same hostplant with no evidence of hybridization. C. ladon and C. lucia are obligate univoltines while C.

neglecta is multivoltine. A later flight (second brood) of Celastrina neglecta at the same site utilizes eriophyid mite-induced

leaf galls on the very same host tree species. Additional Virginia records of C. lucia are documented.

Additional key words: sympatry, androconia, Prunus serotina, leaf galls, Comusflorida

INTRODUCTION

Celastrina systematics has long remained in flux. Following the separate descriptions of C. ladon

(Cramer, 1780), C. lucia (W. Kirby, 1837) and C. neglecta (W. H. Edwards, 1862), the notion of how

many species of Celastrina inhabit North America has fluctuated from author to author and has been a

source of endless debate and confusion. Disagreement among checklist authors continues to the present

day. For example, Pelham (2008) lists nine Celastrina species north of Mexico, thus adopting species-

rank for the three species recorded in this study. On the other hand, the North American Butterfly

Association's most recent checklist as of this writing (NABA, 2001), recognizes only three Celastrina

species, relegating neglecta to subspecies status under C. ladon , and does not recognize lucia at any rank.

Until the 1990's, most authors traditionally treated neglecta as a summer form of C. ladon (e.g. Iftner et

al., 1992). Since that time, the majority of newly published regional-level guides have treated C. neglecta

at full-species rank. Wright and Pavulaan (1999) identified a unique primary dorsal wing scale character

that differentiates C. ladon from C. neglecta and all other North American Celastrina (except C. nigra) as

a full-species taxon (Figs. 1 & 2). This character breeds true within C. ladon from annual generation to

generation without variation. It is also expressed in lab-produced "false summer generation" adults and

never appears in lab-reared C. neglecta. This unique character is used as a convenient method to

differentiate C. ladon from both C. lucia and C. neglecta at a northern Virginia site in the present study.

The name Celastrina lucia has been applied to a broad grouping of phenotypically similar, though

apparently distinct, Celastrina populations spanning the northern portion of the North American continent

and extending southward in the Appalachian and Rocky Mountain regions. The taxonomic standing of

these various populations is under current review and will likely be revised to include two or more sibling

species once it can be determined precisely which population Kirby described as lucia. The Appalachian

lucia population reported here is tentatively retained as a member of the lucia species-group until further

1 Staff Member, The International Lepidoptera Survey, Herndon, VA.

research resolves the type locality issue and clarifies the relationship of continental populations currently

treated as lucia. All references given here to the name lucia are tentative and follow the line of reasoning

given above. [As an alternative, lucia in Virginia may be referable to as " lucia Auct." until its taxonomic

standing can be resolved.]

Fig. 1: Scanning electron micrograph (SEM 640X) of dorsal forewing of C. neglecta showing androconia

between blue scales. Specimen taken September 17, 1987, Harleysville, Montgomery Co., PA. Fig. 2:

(SEM 640X) of dorsal forewing of C. ladon showing long overlapping scales and lack of androconia.

Specimen taken April 23, 1992, Green Ridge State Forest, Allegany Co., MD. Photos by David M.

Wright.

DISCUSSION

A distinctive hilltop site known as Old Knob located near Gore, VA, Frederick County, was

discovered in June, 2005. This hill (elevation 1,300 ft.) is not impressive by regional standards, but the

summit area contains an exceptional array of butterfly species. During a multi-year study of the local

colony of Papilio ( Heraclides ) cresphontes and summit stand of Ptelea trifoliata (Wafer Ash, Hop Tree),

an unexpected assembly of sympatric Celastrina species {ladon, lucia , neglecta) was uncovered.

A substantial colony of Celastrina neglecta was first observed on the summit of Old Knob on June

6, 2006. It was noted that Prunus serotina (Black Cherry) comprised a significant portion of the forest

understory. Many of these trees were infested with leaf galls formed by eriophyid mites, attributed to

Phytoptus cerasicrumena. Several second-brood adult C. neglecta females were observed ovipositing on

the leaf galls and many eggs were located, but not collected. Eriophyid mite leaf galls are the primary

larval host of C. serotina throughout much of the northeastern United States (Pavulaan & Wright, 2005).

Oviposition by C. neglecta on P. serotina leaf galls was previously observed in the area surrounding Big

Meadows Recreational Area of Shenandoah National Park in Page County, VA in 1985. C. neglecta has

been observed to utilize hosts from a broad range of plant families (Pavulaan and Wright, 2005), thus this

behavior at Old Knob was not deemed unusual. At both the Old Knob and Big Meadows sites, mound¬

building ants were very common (Fig. 27); the ants built huge mounds and defended their turf against

intruders rather aggressively. The ants at Big Meadows were of an unidentified type of large stinging ant,

while the ants at Gore did not sting, but inflicted painful bites in large numbers as experienced by the

author. Throughout the study the ants vigilantly guarded and protected larvae against predators in

exchange for larval honeydew secretions. The presence of ant colonies likely contributed to the longevity

of the associated Celastrina colonies in this region.

On April 23, 2007 adults of both the Celastina lucia (Figs. 3, 9, 15) and C. ladon (Figs. 4, 10, 16)

populations were collected on the summit of Old Knob. No early spring brood individuals of C. neglecta

were recorded on Old Knob in 2007. The C. lucia males were easily distinguished from C. ladon males

by dissimilarity of their forewing scale characters (Figs. 1 and 2). Furthermore, it was immediately

surmised that C. lucia and C. ladon resided in sympatry at this northern Virginia site without evidence of

interbreeding and were capable of retaining their separate species-level identities. It was further observed

that both species were found in close proximity to a potential host tree Prunus serotina and associated ant

mounds. No attempt was made to obtain ova on this date. Further exploration demonstrated C. ladon was

also found in considerable numbers on the south slope of the hill below the summit along Knob Road at

approximately 700-860 ft. elevation. Despite the relatively short distance from the summit, no C. lucia

individuals were found below the summit along Knob Road. This suggests that C. lucia may have a

stronger inter-dependency on mound-building ants on the summit than C. ladon.

Subsequent early-spring surveys on Old Knob on April 11, 2008 and April 18, 2009 revealed the

same pattern, with only C. ladon and C. lucia evident at the summit and only C. ladon along the roadway

at lower elevations. In 2008, several ova were obtained from an unidentified female confined with

Prunus serotina flower buds. The larvae were first reared on flower buds, but were later switched to

eriophyid mite galls. These produced a “false (lab-induced) second generation” of C. ladon adults (Figs.

7, 13, 19) with all male specimens displaying the unique dorsal scale character of ladon forewings. It was

subsequently observed that both C. ladon and C. lucia utilize Prunus serotina flower buds and eriophyid

mite galls at the summit site.

On April 6, 2010 at the summit, several adults of what appeared to be spring form of C. neglecta

(Figs. 5, 11, 17) were found flying with C. ladon and C. lucia , and also in close association with Prunus

serotina. Several females of the three distinct phenotypes were confined with Prunus serotina buds in

separate containers and many ova were obtained. Unfortunately, the eggs of only one female successfully

emerged, while the rest failed to emerge. It is unclear why the remainder did not emerge; mold or fungal

infection is suspected or the females may not have mated. The surviving larvae from the single female

yielded typical summer form C. neglecta adults (males distinguished by typical neglecta dorsal forewing

scale structure) (Figs. 8, 14, 20), which emerged over the period from May 20 to 29. An additional trip

was made April 14, 2010, but no females were found. A trip to Old Knob on May 27, 2010 detected a

second (summer) brood of C. neglecta was flying. This confirmed a spring flight (April) and subsequent

late May flight of C. neglecta occurred at Old Knob. Both broods apparently utilized Prunus serotina.

Even later summer broods of C. neglecta also occurred at Old Knob. These have not been studied other

than the collection of specimens to confirm the presence of additional broods of C. neglecta.

On May 5, 2011, a worn female of an unidentifiable phenotype was collected on Old Knob and

confined on developing flower buds of Prunus serotina. Several eggs were obtained and larvae were

reared to maturity on both flower buds and leaf mite galls. Many larvae perished during the course of

rearing due to cannibalism. Two “false second generation” adults of C. lucia (Figs. 6, 12, 18) eclosed on

June 6, 2011. The male displayed a dorsal forewing scale structure identical to spring brood individuals.

The fact that three different Celastrina populations occupy the same ecological niche in Virginia

without apparent hybridization or intergradation is strong evidence of reproductive barriers maintaining

species-level distinctness. It is surmised that a small isolated C. lucia population on the summit of Old

Knob would have been obliterated long ago by natural hybridization, if this were to have occurred. A

fourth species, C. neglectamajor , was also recorded on Old Knob, but not at the study site. Two

individuals were collected on the lower portion of Knob Road on May 12, 2009. Due to the close

proximity of the neglectamajor colony, there is a high likelihood that stray individuals briefly traveled

into the study site. The host plant ( Cimicifuga racemosa ) of C. neglectamajor (Pavulaan & Wright, 2001)

was not present on the summit.

3

COMPARISON OF CELASTRINA TAXA

A comparison of the three taxa including the summer form of C. neglecta is presented here. All

three sympatric species display a similar phenotype to the naked eye during their early spring flight

periods. Only C. ladon males are easy to distinguish from C. lucia and spring form C. neglecta.

Celastrina ladon. Adults (Figs. 4, 10) at the study location are typically of the spotted ventral

hindwing phenotype (Fig. 16), showing no tendency to develop darkened ventral hindwing margins or

dark ventral hindwing discal patches, which are more frequent in the northern portions of the species’

range. The spotted form is referred to as form " violacea " (W.H. Edwards, 1866), which is technically a

species-level junior synonym of the name ladon. One individual of the dark-margined form [form

"marginata" of authors] was collected here (Fig. 23). Males of C. ladon are easily distinguished from all

other blue Celastrina species by their unique male wing scale structure (Fig. 2). Adults of C. ladon tend

to be slightly more violet-blue in color than either C. lucia or C. neglecta , thus the name violacea.

C. ladon is univoltine throughout its range. In lab rearing, an artificial summer phenotype can be

produced (Figs. 7, 13, 19), which bears the unique ladon male dorsal wing scale structure and has only a

superficial resemblance to the natural summer form of C. neglecta (which does not bear the unique male

dorsal wing scale structure). No adults resembling the lab-reared false summer form of C. ladon have ever

been found at the study site.

Celastrina lucia. Adults (Figs. 3, 9) at the study location are generally of the spotted ventral

hindwing phenotype (Fig. 15), with some individuals displaying darkened ventral hindwing margins

[described as form marginata (W. H. Edwards, 1883)] or rarely the dark ventral hindwing discal patch

(form "lucia"), which is characteristic of C. lucia in northern latitudes. The ventrally-spotted form has

been incorrectly referred to as form " violacea " of authors, however the name violacea technically applies

only to the spotted form of C. ladon. Two interesting individuals are figured from here, one is form

"marginata " (Fig. 24) and the other is form " lucia " (Fig. 25). Males of C. lucia are easily distinguished

from males of C. ladon which bear the unique wing scale structure (Fig. 2). Adults of C. lucia tend to be

noticeably more metallic blue in color than C. ladon when fresh, but have a peculiar tendency to become

more violet when flight-worn with age. Males of C. lucia can be distinguished from males of spring form

C. neglecta by the lack of very distinct white wing veins along the leading forward edge of the dorsal side

of the forewing, which are characteristic of spring form C. neglecta. Also individuals of spring form C.

neglecta bear clear hindwing fringes, while in C. lucia they are either darkened or checkered black and

white. Females are generally difficult to distinguish, as both C. lucia and C. neglecta females tend to be

very similar in appearance, both being noticeably lighter blue than females of C. ladon. In spread series,

C. lucia females from the Appalachian region are noticeably smaller than C. neglecta females and have

narrower black outer margins on the dorsal forewing.

C. lucia is univotine throughout its range, and is thus presumed to be univoltine at the study site.

In the lab, an artificial summer phenotype can be produced (Figs. 6, 12, 18), which bears no resemblance

to the natural summer form of C. neglecta. No naturally occurring individuals resembling the lab-reared

false summer form have ever been found at the study site.

Celastrina neglecta spring form. Adults (Figs. 5, 11) at the study site are typically of the

distinctly-spotted ventral hindwing phenotype (Fig. 17), showing no tendency to darkened ventral

hindwing margins or dark ventral hindwing discal patches. The ventrally-spotted form has traditionally

been referred to as form "violacea" by authors, but the name violacea technically applies only to the

ventrally-spotted form of C. ladon. Males of C. neglecta spring form are easily distinguished from males

of C. ladon which have the unique forewing scale structure (Fig. 2). However, in all other respects, they

are extremely similar to C. ladon and very difficult to distinguish by ventral markings alone. Adults of

both sexes of C. neglecta spring form tend to be bluer in color than C. ladon , but similar in color to C.

lucia. Males of C. neglecta spring form can often be distinguished from sympatric males of C. lucia by

the presence of distinct white veins along the leading edge of the dorsal forewing. This character state is

useful when examining fresh individuals, but is not always reliable as veins become subdued with age.

Also C. neglecta spring form individuals have clear white fringes on the hindwing edge. In C. lucia these

fringes tend to be darkened or checkered and in C. ladon they also tend to be darkened, but may appear

light in some individuals. Females are generally difficult to distinguish from C. lucia, but in general they

are larger and have broader black outer margins on the dorsal forewing.

Celastrina neglecta summer form. The summer form of neglecta is uniquely different from the

three spring phenotypes, in that the venter is very white, and dark markings are reduced to mere dashes

and dots (Fig. 20). On the dorsum, the males and females both display characteristic distribution of white

insuffusion on the hindwings, arranged in rays between the wing veins (Figs. 8, 14). This phenotype has

been produced in reared offspring of spring form females throughout the range of neglecta.

The summer form of C. neglecta was recorded during surveys conducted on the following dates:

June 6, 2006 and May 27, 2010 (associated with mite galls on both dates); June 7 & 23, July 1, and

August 29 in 2010.

ADDITIONAL NOTES ON CELASTRINA LUCIA IN VIRGINIA

In addition to the Old Knob study site, C. lucia has been confirmed from six additional sites in

Virginia, totalling four counties (Fig. 28). It is interesting to note that all of these populations are closely-

associated with Prunus serotina with the exception of the Great North Mountain population:

Great North Mountain, near Havfield, Frederick County, VA. A visit to the top of this ridge on

April 21, 2014 found a colony of C. lucia flying in an area of remnant Pitch Pine/Scrub Oak barren at

2300 ft. elevation. Most of the forest canopy now consists of various Oak species with an understory of

Scrub Oak ( Quercus ilicifolia ) and Mountain Laurel ( Kalmia latifolia). Interestingly, unlike the other

Virginia C. lucia sites, Prunus serotina was not observed to be a primary component of the ridgetop

forest. Rather Blueberries ( Vaccinium sp.) were very common everywhere especially along the roadside

edges and are suspected to be the host. Five males were collected and an additional 15 individuals were

observed in the same area, presumably all C. lucia. During a return trip on April 24, 2014, six males were

collected and an additional 12 individuals were observed. A final trip on May 3, 2014, following several

days of damp rainy weather, only found one worn male and three fresh females were found. One of the

females was form “lucia” with the distinctive ventral hindwing patch (Fig. 26).

Cacapon Mountain, north of Cross Junction. Frederick County, VA. A visit to the top of this

ridge, literally within a few hundred feet of the very northern border of Virginia on May 3, 2014 produced

a single male specimen of C. lucia. C. lucia occurs more commonly northward along the top of this same

ridge on the West Virginia side of the border.

Lake Thompson area, lower east slope of Blue Mountain. George Thompson Wildlife

Management Area, north of Markham. Fauquier County, VA. Few C. lucia adults have been documented

at this location among thousands of spring-flying Celastrina individuals that were either collected,

examined in-hand (net/release) or observed (resting only) at very close range. All C. lucia specimens have

been collected toward the end of their flight period with most adults being faded from age, thus leading

one to suspect they have flown in from some distance, likely from somewhere on Blue Mountain or along

the Blue Ridge. All specimens were collected along woodland trails at the fairly low elevations of 970-

6

1200 ft. Interestingly, no C. lucia specimens have ever been collected on top of Blue Mountain. These

specimens are based on adult phenotype (Fig. 21) and are a close match to C. lucia found to the north in

central Pennsylvania or on the Allegheny Plateau in West Virginia. Prunus serotina is present in the

woodlands here and is the only host tree on which ovipositions were observed. Blueberries ( Vaccinium

sp.) are uncommon in the dense mixed Appalachian woodland of Blue Mountain; thus they are likely not

utilized. Collection dates: April 10, 1999; April 29, 2000; April 13, 2002; April 27 & May 3, 2003; April

17, 2004; April 11, 2005; April 13 & 14, 2006; May 17, 2008; April 17, 2009; April 11 & 23, 2010.

Tanners Ridge (part of the Blue Ridge), near Big Meadows Recreational Area, Shenandoah

National Park, Page County, VA. A single female specimen of form "marginata" was collected at this

location [under permit], just off Skyline Drive at an elevation of 3387 ft. In general, Celastrina are

noticeably scarce along the top of the Blue Ridge, except in late May and early June when C.

neglectamajor flies. Prunus serotina is common on the crest of the Blue Ridge and several sightings of

unidentified Celastrina adults around P. serotina suggest this might be the primary host for C. lucia or

other Celastrina species on the Blue Ridge. Blueberries ( Vaccinium sp.) are also very common in open

places at higher elevations, but apparently not utilized by Celastrina. Over several years (1985-2008), I

have carefully scanned forest clearings, roadside edges and a power line cut in the forest around the Big

Meadow, but have seen no Celastrina associated with Vaccinium. Collection date: May 5, 2001.

Tanners Ridge (part of the Blue Ridge), along Route 682, near Stanley, Page County, VA. Early

in this study I explored a location on the west slope of the Blue Ridge just outside the National Park,

where one could collect fairly high in elevation (up to 2884 ft.). Several female Celastrina were collected

along this road in May, 1987, which for several years I kept in a papered series of C. ladon from the

location. Only upon examination of these specimens, which were mounted for this report, did I discover

three specimens with features distinctly those of C. lucia , i.e. very light blue dorsal color with narrow

black dorsal forewing margins (Fig. 22). Collection dates: May 14 & 16, 1987.

Shenandoah Mountain, west of Briery Branch, Rockingham County, VA. Several individuals

were collected along the upper portion of State Road 924 (at elevations of 2062 ft.-3467 ft.) and also

along Forest Road 85 (approximately 3845 ft.) going north along the ridge top, which delineates the

Virginia-West Virginia border. A female captured on April 29, 2001 was confined with Prunus serotina

flower buds in a rearing container; eggs laid in confinement subsequently produced several "false summer

brood" adults on June 1, 2001. These individuals resembled lab-produced false brood adults from C. lucia

populations in West Virginia, Pennsylvania and New Jersey. Collection dates: April 29, 2001; May 4 &

6, 2006.

HISTORICAL RECORD

In The Butterflies of Virginia, under Cyaniris argiolus pseudargiolus , Clark & Clark (1951)

wrote: "We have taken the form lucia only in western Frederick County in Virginia." It is presumed that

the authors were referring to the ventral hindwing dark-patched form. However, since the patched form

has been recorded in all Celastrina species in the eastern region of the United States, it is not known

whether the authors collected C. lucia , C. ladon or C. neglecta. Assuming they did have a specimen of C.

lucia in hand, that would be the first record of C. lucia in Virginia. The present study would corroborate

the older Frederick County record.

A NOTE ON HOSTPLANT ACCEPTANCE BY CELASTRINA NEGLECT A

As an extension of this study, an effort was attempted to corroborate previous host intolerance

findings, specifically that spring Celastrina neglecta females do not oviposit on and neonate C. neglecta

larvae do not utilize Cornus florida (Flowering Dogwood), a common C. ladon host. On April 21, 2014,

a survey of the Old Knob study site found no C. lucia, but two females of C. ladon and one female of C.

neglecta were captured. These females were separately confined in containers with cuttings of C. florida

flower buds. The two C. ladon females readily oviposited on C. florida within the first day, whereas the

C. neglecta female refused to oviposit on the same plant under identical conditions. On the third day of

confinement, cuttings of Viburnum prunifolium (a documented C. neglecta host in this region) were added

to the container containing the female C. neglecta. She immediately oviposited approximately 50 eggs on

V. prunifolium within 24 hours, while still ignoring C. florida. Subsequently, individual flower buds

containing C. neglecta eggs were removed and strategically placed onto cuttings of C. florida flower buds

so that newly-hatched larvae would have the direct choice of feeding on Cornus florida. Newly hatched

larvae were also transferred from the V. prunifolium buds to C. florida , thus leaving them no choice but to

feed on C. florida. By May 3, 2014, most of the C. neglecta larvae had hatched and ignored the C. florida,

subsequently starving and leaving only shriveled corpses on the container sides. A few remaining larvae

attempted to feed on C. florida , not on the flower buds but rather boring into the base of the underside of

the white bracts or into the basal portion of the flower buds. By May 8, 2014, all C. neglecta larvae had

died. A previous attempt in 2013 at getting C. neglecta larvae to accept C. florida similarly failed, with all

larvae preferring to starve rather than to eat C. florida. An earlier 2012 observation of ovipositional

behavior by captive females also found that C. neglecta females refused to lay eggs on C. florida. This

finding demonstrates that Cornus florida is not acceptable to C. neglecta , and the plant likely has toxic

properties to certain Celastrina species.

CONCLUSION

Based on observations made during this study and reported here, the following statements are

presented as clear succinct conclusions:

(1) The three taxonomic entities Celastrina ladon , C. lucia and C. neglecta behave as distinct full

species in sympatry in northern Virginia.

(2) The following species arrangement is hereby confirmed:

Celastrina ladon (Cramer, 1780)

Celastrina lucia (W. Kirby, 1837)

Celastrina neglecta (W. H. Edwards, 1862)

ACKNOWLEDGMENT

I wish to sincerely thank David M. Wright who critically reviewed multiple drafts of this manuscript and

offered several helpful suggestions.

LITERATURE CITED

Clark, A. H. and L. F. Clark. 1951. The Butterflies of Virginia. Smithsonian Miscellaneous Collections

116(7): vii +239 pp.

Iftner, D. C., J. A. Shuey and J. A. Calhoun. 1992. Butterflies and Skippers of Ohio. Ohio Biological

Survey Bulletin, New Series 9(1): xii + 212 pp.

The North American Butterfly Association (NAB A). 2001. Checklist & English Names of North

American Butterflies, Second Edition. B. Cassie, J. Glassberg, A. Swengel and G. Tudor, editors.

The North American Butterfly Association, Inc. 60 pp.

Pavulaan, H. & D. M. Wright, 2005. Celastrina serotina (Lycaenidae: Polyommatinae): A New Butterfly

Species from the northeastern United States and eastern Canada. The Taxonomic Report 6(6): 1-

19.

Pavulaan, H. & D. M. Wright, 2000. The biology, life history, and taxonomy of Celastrina neglectamajor

(Lycaenidae: Polyommatinae). The Taxonomic Report 2(5): 1-19..

Pelham, J. P. 2008. A Catalogue of the Butterflies of the United States and Canada. J. Res. Lepid. 40:1-

658.

Wright, D. M. and H. Pavulaan. 1999. Celastrina idella (Lycaenidae: Polyommatinae): A New Butterfly

Species from the Atlantic Coastal Plain. The Taxonomic Report 1(9): 1-11.

Fig. 28: Map showing known county distribution of Celastrina lucia in Virginia, as of June 16, 2014.

The Taxonomic Report

is a publication of

The International Lepidoptera Survey (TILS)

(a tax exempt non-profit scientific organization )

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10

Volume 7 Number 8

17 June 2014

The Taxonomic Report

OF THE INTERNATIONAL LEPIDOPTERA SURVEY

Cercyonis pegala agawamensis (Satyridae):

A new butterfly subspecies from the coastal salt marshes

of the northeastern United States of America

me Ringlet (Coenonympha nipisiquit McDunnough) whk

a tullia inomata W. H. Edwards), and the Dos Passos C

Copper (L. dorcas claytoni A. E. Brower) - known only from a few calcareous b

in adjacent New Brunswick.

OVERVIEW OF NAMED TAXA OF “WOOD NYMPHS”

Ill

in of the concept of in

n & Sternburg (2001) -'

le of Klots (1951), a.

ilications referred to;

:ist in BOTH cases.

sArey and Grkovich, news.

ETYMOLOGY.

Male: FW length 25-26 mm. Upperside: Ground color brown. FW apex typically somewhat more poir

other pegala subspecies. FW eye patch fairly broad (width of patch 10 mm along costal margin), yellow tc

in light blue. HWe;

k line prominent; occasional second smaller eyespot above the main HW eyespot somet

yroaLnd'briVyenow 8 eyespotlas^n th^above s^acJ^^pflS^blnc 6 Smal]

n C. p. agawamensis and C. p. n

idian dark line; the eyespots are smaller but at least 4 or 5 are present and fairly well-defii

s are less noticeably two-toned on the hindwing, the postmedian line is less distinct and

broad (12-15

soar- 1 '*

ative phenology for C. pegala dope or maritima are very late Ju

ig from 12 July to 30 July. Extreme dates for C. pegala dope a

larity to the recent taxas.

caria), Joe-Pye-Weed (.

(A. incamata ), Red Clover (Ti

As noted elsewhere, agawamensis also differs from all eastern Cercyonis - except for unusual populations of C.

pegala “nephele ” occurring in northern Coos Co., New Hampshire - in that it nectars freely in its salt marsh habitat.

These “ nephele ” which we suspect should be further studied, also differ from typical nephele in that they also occur

in rather boggy wet meadows and roadsides, nectar freely and frequently on Joe Pye Weed, Everlastings, etc. and

also have a fast and rather straight-ahead “unbouncy” flight. We have observed these very blackish populations

with very conspicuous ocelli in early to late August at such elevated locations as Pinkham Notch (Mount

Washington, 2100 ft.) and at the Scott Bog, East Inlet Road, etc. (above 1900 ft.) near the New Hampshire/Quebec

border.

GEOGRAPHIC RANGE AND DISTRIBUTION. Range confirmation for this subspecies is so far restricted to

coastal estuarine habitat from southern Essex County, Massachusetts (vicinity of Salem) extending northward to

southern York County, Maine (near Ogunquit). The greatest and most continuous concentration of this insect

occurs in the more extensive coastal salt marshes north of Cape Ann in Massachusetts (Ipswich River, Parker River

and Merrimack River drainages) to the New Hampshire seacoast approximately 12 to 15 Km. south of the

Piscataqua River. This encompasses the coastal towns/cities of Essex, Ipswich, Rowley, Newbury, Newburyport

and Salisbury in Massachusetts (Essex Co.), and the salt marshes in Seabrook and Hampton (Rockingham Co.) of

the southernmost coastline in New Hampshire. This subspecies has also been confirmed in less extensive habitat

south of Cape Ann in the Danvers River / North River drainage in the vicinity of Pope’s Landing (Danvers) and the

tidal flats within the Forest River drainage system in Salem. Another population occurs in a more extensive

estuarine meadow located in Ogunquit, Maine (Piscataqua and Kennebunk River coastal watersheds), which is

speculated to be the northernmost range of agawamensis. Habitats south of Essex County, MA, especially in

Plymouth, Barnstable and Bristol Counties, and even coastal Rhode Island have yet to be explored. However, there

is a strong likelihood that they do occur in the more extensive estuaries and tidal river basins in much of this region.

Agawamensis should also be compared to the geographically restricted island populations of subspecies maritima

that occur specifically on the islands of Nantucket, Martha’s Vineyard, Block Island (R.I.) and Naushon Island

(MA.) located in Buzzards Bay. Maritima is also very common in dry weedy open field habitats on mainland

Rhode Island, on Long Island, and also near Gettysburg, PA.

TYPE LOCALITY AND TYPE SERIES. TL: Estuarine salt marsh (open), Boston Road, Newbury (Parker

River watershed), Essex Co., Massachusetts. The holotype specimen will be deposited in the McGuire Centre for

Lepidoptera and Biodiversity, Gainesville, Florida. Holotype (female), allotype paratype (male) and all paratypes

are identified as follows:

MASSACHUSETTS:

ESSEX CO.. Newbury (Parker River watershed) :

NB001 (female): Boston Road, estuarine salt marsh (open), 7/21/2007 (HOLOTYPE).

NB002 (male): Boston Road, estuarine salt marsh (open), 7/21/2007 (ALLOTYPE).

NB003-NB004 (2 males): Boston Road, estuarine salt marsh (open), 7/21/2007.

NB005-NB006 (2 females): Boston Road, estuarine salt marsh (open), 7/21/2007.

NB010 (female): U.S. Route 1, wet meadow/salt marsh (open), 7/17/2007.

NB011-NB012 (2 males): Boston Road, estuarine salt marsh (open), 7/20/2008.

NB013-NB014 (2 males): Boston Road, estuarine salt marsh (open), 8/2/2008.

NB015 (female): Boston Road, estuarine salt marsh (open), 7/20/2008.

NB016-NB018 (3 females): Boston Road, estuarine salt marsh (open), 7/28/2013.

NB019-NB020 (2 females): Boston Road, estuarine salt marsh (open), 8/2/2008.

NB021 (1 male): Plum Island, salt marsh, 8/14/1999.

NB022-NB023 (2 females): Plum Island, salt marsh, 8/14/1999.

NB024-NB029 (6 males): Boston Road, estuarine salt marsh (open), 7/28/2010.

NB030-NB033 (4 females): Boston Road, estuarine salt marsh (open), 7/28/2010.

NB034-NB038 (5 males): Boston Road, estuarine salt marsh (open), 7/24/2011.

NB039-NB042 (4 females): Boston Road, estuarine salt marsh (open), 7/24/2011.

NB043-NB045 (3 females): U.S. Route 1, salt marsh, 8/12/2011.

NB046-NB050 (5 males): U.S. Route 1, salt marsh, 8/8/2013.

NB051-NB053 (3 females): U.S. Route 1, salt marsh, 8/8/2013.

NB054-NB055 (2 females): Plum Island, salt marsh, 7/24/2003.

ESSEX CO., Essex (Essex River watershed) :

EX001-EX004 (4 males): Route 133, salt marsh, 7/26/2008.

EX005-EX007 (3 females): Route 133, salt marsh, 7/26/2008.

ESSEX CO., Ipswich (Ipswich River watershed) :

IW003-IW004 (2 males): Argilla Road, estuarine salt marsh (open), 7/22/2007.

IW005 (female): Argilla Road, estuarine salt marsh (open), 7/22/2007.

IW006 (female): Argilla Road, old field (coastal), 7/22/2007.

IW007 (male): Argilla Road, estuarine salt marsh (open), 7/9/2013.

IW008 (female): Argilla Road, estuarine salt marsh (open), 7/9/2013.

IW009-IW011 (3 males): Argilla Road, estuarine salt marsh (open), 8/5/2013.

IW012 (female): Argilla Road, estuarine salt marsh (open), 8/5/2013.

ESSEX CO., Newburyport (Merrimack River watershed) :

NP001 (female): Plum Island Turnpike, coastal salt marsh (open), 7/31/1983.

ESSEX CO., Rowley (Rowley River watershed) :

RW001 (male): Route 1A, estuarine salt marsh (open), 7/27/2007.

RW002-RW005 (4 males): Route 1A, estuarine salt marsh (open), 7/19/2013.

ESSEX CO., Salem (Forest River watershed) :

SA001-SA002 (2 males): Forest River Park, estuarine salt marsh - tidal river basin, 7/22/2007.

ESSEX CO., Salisbury (Merrimack River watershed) :

SB001-SB002 (2 males): U.S. Route 1 A, estuarine salt marsh (open), 7/24/2007.

SB003 (female): U.S. Route 1A, estuarine salt marsh (open), 7/20/2008.

SB004 (female): U.S. Route 1A, estuarine salt marsh (open), 7/31/2008.

SB005-SB007 (3 males): U.S. Route 1 A, estuarine salt marsh (open), 7/30/2009.

SB008-SB011 (4 females): U.S. Route 1 A, estuarine salt marsh (open), 7/30/2009.

SB012-SB013 (2 females): U.S. Route 1A, estuarine salt marsh (open), 8/3/2009.

MAINE:

YORK CO.; Ogunquit (Stevens Brook watershed) :

OQ001 (1 male): Furbish Road, estuarine salt marsh (open), 8/1/2007.

OQ002 (1 female): Furbish Road, estuarine salt marsh (open), 8/1/2007.

OQ003 (1 male): Furbish Road, estuarine salt marsh (open), 8/1/2007.

NEW HAMPSHIRE:

ROCKINGHAM CO.; Seabrook (Blackwater River watershed) :

SK001-SK004 (4 males, dark type): Route 286, estuarine salt marsh (open), 7/24/2007.

SK005-SK007 (3 males): Route 286, estuarine salt marsh (open), 8/6/2007.

SK008-SK0010 (3 females): Route 286, estuarine salt marsh (open), 8/6/2007.

SK011 (male): Route 286, estuarine salt marsh (open), 8/6/2007.

SK012 (female): Route 286, estuarine salt marsh (open), 8/6/2007.

SK013-SK015 (3 males): Route 286, estuarine salt marsh (open), 8/6/2007.

SK016-SK017 (2 females): Route 286, estuarine salt marsh (open), 8/6/2007.

Additional locations where C. p. agawamensis have been vouchered:

MASSACHUSETTS : ESSEX CO.: Danvers. MAINE : WASHINGTON CO.: Addison (possible intermediate with C. p.

nephele). NEW HAMPSHIRE : ROCKINGHAM CO.: Hampton.

Locations where C. p. alope have been vouchered for comparison to C. p. agawamensis :

MASSACHUSETTS : ESSEX CO.: Boxford, Georgetown, Groveland, Ipswich, Newbury, Rowley, Topsfield. NORFOLK

CO.: Norfolk. BERKSHIRE CO.: Sheffield, Ashley Falls. MAINE : CUMBERLAND CO.: Gorham. YORK CO.: Buxton,

Hollis, Saco, York. VERMONT : WINDHAM CO.: Mount Snow, Green River, Guilford.

Locations where C. p. maritima have been vouchered for comparison to C. p. agawamensis:

MASSACHUSETTS : WORCESTER CO: Wachusett Mountain. ESSEX CO.: Danvers, Peabody, North Andover, Boston

Hill. MIDDLESEX CO.: Melrose (Middlesex Fells), Woburn (Horn Pond Mountain). DUKES CO. (MARTHA’S

VINEYARD): Oak Bluffs (TL of maritima), Edgartown. PLYMOUTH CO.: Plymouth, Carver, Middleboro. RHODE

ISLAND : PROVIDENCE CO.: Central Falls. WASHINGTON CO.: Westerly, West Kingston. MAINE :

ANDROSCOGGIN CO.: Lewiston, Auburn. OXFORD CO.: Oxford.

CO.: West Hawley. MAINE : AROOSTOOK CO.: Sherman Mills Twp. KENNEBEC

ID CO.: Streaked Mountain. PENOBSCOT CO.: Dixmont. WALDO CO.: Belfast,

WASHINGTON CO.: Addison, Cherryfield, Columbia, Columbia Falls, Harrington. NEW HAMPSHIRE :

Pinkham Notch, Pittsburg, Scott Brook Road, East Inlet Road, Clarkston. GRAFTON CO.: Franconia Notch.

CO.: Pittsford. GRANDE ISLE CO.: C

OF THE EVOLUTIONARY ORIGINS OF S'

WITHIN toe typical agawamensis salt marsh habitat. A lope k described as having a yellow FW eye patch and a

the tension zone" between the salt marshes and the adjacent wet meadow habitats appear to be chiefly if not

edby anotl

d in the h

s later in the early 17 th Century, came the ai

:s in Eastern North A

ACKNOWLEDGMENTS

Figure 1. Holotype of Cercyonis pegala agawamensis. Female, wingspan 54 mm.

HOLOTYH:;.

C ercyonis pegala Fabr.

j agawamensis Arey & Grkovich $

L .1 Salt Marsh-Boston Rd. j

Newbury, Essex Co. MA

» iul 21,2007 M. Arey leg. 1

Salt Marsh-Boston Rd.

Newbury, Essex Co. MA

Jul 21, 2007 M. Arey leg.

11

Figure 5. Male C. pegala nephele, wingspan 47 mm

Dry meadow - Rt. 220 E Troy, Waldo Co. Maine, August 11, 2007 M. Arey leg.

Ww

Figure 6. Male C. pegala alope, wingspan 49 mm

Witch Hollow Farm, Boxford, Essex Co. MA, July 4, 1998 M. Arey leg.

Figure 7. Female C. pegala alope, wingspan 53 mm

Powerline cut - Rt 97 S Georgetown, Essex Co. MA, July 23, 2002 M. Arey leg

Figure 8. A large female C. pegala agawamensis, wingspan 60 mm

Salt Marsh - Salisbury Beach S.P., Salisbury, Essex Co. MA July 30, 2008 M. Arey leg.

12

Figure 9. Possible female intergrade between nephele and agawamensis, wingspan 54 mm

Along tidal river - Columbia Rd. W Addison, Washington Co. ME, August 6, 2006 M. Arey leg.

13

Figure 12. Habitat at type locality - Boston Rd. Newbury, Essex Co. MA; June 6, 2014

Figure 13. Habitat at type locality - Boston Rd. Newbury, Essex Co. MA; June 6, 2014

14

sr

The Taxonomic Report

is a publication of

The International Lepidoptera Survey (TILS)

(a tax exempt non-profit scientific organization )

The Taxonomic Report is published for the purpose of providing a public and permanent scientific record.

It appears on printed paper in sequential issues, is regularly disseminated to institutional and individual

subscribers, and is also available as separate issues free of charge upon request at the discretion of authors

and/or the editor. Contents are peer-reviewed but not necessarily through the anonymous review and

comment process preferred by some publishers of serial literature.

TILS Purpose

TILS is devoted to the worldwide collection of Lepidoptera for the purpose of scientific discovery,

determination, and documentation, without which there can be no preservation.

TILS Motto

“As a world community, we cannot protect that which we do not know”

Articles for publication are sought

They may deal with any area of research on Lepidoptera, including faunal surveys, conservation topics,

methods, etc. Taxonomic papers are especially welcome. Before sending a manuscript, simply write to

TILS editor, Harry Pavulaan, P.O. Box 1124, Herndon, VA 20172 to set up discussion on how to best

handle your material for publication; or email harrypav@hotmail.com

Donations are needed

to support and further our efforts

to discover and protect butterflies worldwide.

All donations are US tax deductible. Please help generously.

Donations should be mailed to: TILS, c/o Harry Pavulaan, P.O. Box 1124, Herndon, VA 20172.

Checks should be made payable to: TILS. Please indicate if you need an individual receipt.

Visit The International Lepidoptera Survey on the World Wide Web at:

http://lepsurvev.carolinanature.com/

Join the discussion at our list serve on YahooIGroups at:

TILS-Leps-T alk

https://groups.vahoo.coni/neo/groups/TILS-leps-talk/info