J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018 1

Journal of the

Entomological Society of British Columbia

Volume 115 December 2018 ISSN#0071-0733

Directors of the Entomological Society of British Columbia 2017-2018....................0.. 2

First records of Baetis vernus Curtis (Ephemeroptera: Baetidae) in North America, with

morpholovical notes. >. 0c Mer Seamer ee oan pane erty Neer ee ee 3

Corrections for the Hemiptera: Heteroptera of Canada and Alaska ........................05. 25

The bees of British Columbia (Hymenoptera: Apoidea, Apiformes) ......................065 44

Efficacy of diamide, neonicotinoid, pyrethroid, and phenyl pyrazole insecticide seed

treatments for controlling the sugar beet wireworm, Limonius californicus (Coleoptera:

Elateridae), in spring wheat vabasit A. eck eortabie pemete A tal wd toch cea gk ye ogee 86

SCIENTIFIC NOTES

A pheromone-baited pitfall trap for monitoring Agriotes spp. click beetles (Coleoptera:

Elateridaé) arid: other soil-surtace ms eeta so a. ateuena sa icdmryas sb) 15h sen aairycis ham aitleeuae sienna 101

Identifying larval stages of Orgyia antiqua (Lepidoptera: Erebidae) from British Columbia,

eave) Bais a Skate CURES SAR dics EVE aR ies 6 AC eee eae 104

NATURAL HISTORY AND OBSERVATIONS

New records of Hymenoptera from British Columbia and Yukon .......................0065 110

First Record of Culex tarsalis (Diptera: Culicidae).in the Yukon: nips cick. iindstesasetverenans 123

An updated list of the mosquitoes of British Columbia with distribution notes ........... 126

NOTICE TOC NTR Th ead Wee hee ee re al a a ad Ne ia Inside Back Cover

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 Ds

DIRECTORS OF THE ENTOMOLOGICAL SOCIETY

OF BRITISH COLUMBIA FOR 2017-2018

President:

Lisa Poirier (president@entsocbc.ca)

University of Northern B.C., Prince George

Ist Vice President:

Tammy McMullan

Simon Fraser University, Burnaby

2nd Vice-President:

Wim van Herk

AAFC, Agassiz

Past-President:

Jenny Cory

Simon Fraser University, Burnaby

Treasurer:

Ward Strong (membership@entsocbc.ca)

BC Ministry Forests, Lands and Natural Resource Operations and Rural Development, Vernon

Secretary:

Tracy Hueppelsheuser (secretary@entsocbc.ca)

B.C. Ministry of Agriculture, Abbotsford

Directors:

Tamara Richardson, Grant McMillan

Graduate Student Representative:

Dan Peach

Regional Director of National Society: Editor, Boreus:

Brian Van Hezewijk Gabriella Zilahi-Balogh(boreus@entsocbc.ca)

Canadian Forest Service, Victoria Canadian Food Inspection Agency

; Web Editor: Editor, Emeritus:

Brian Muselle (webmaster@entsocbc.ca) Peter & Elspeth Belton

University of British Columbia, Okanagan Simon Fraser University

Campus

Society homepage: http://entsocbe.ca Journal homepage: http://journal.entsocbe.ca

Editorial Committee of the Journal of the Entomological Society of British Columbia:

Editorial Board: Marla Schwarzfeld, Bo Staffan

Lindgren, Katherine Bleiker, Lisa Poirier, Lee

Humble, Bob Lalonde, Lorraine Maclauchlan,

Robert McGregor, Steve Perlman, Joel

Gibson, Dezene Huber

Editor-in-Chief:

Katherine Bleiker (journal@entsocbc.ca)

Canadian Forest Service, Victoria

Copy Editor: Monique Keiran Technical Editor: Jesse Rogerson

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 3

First records of Baetis vernus Curtis (Ephemeroptera:

Baetidae) in North America, with morphological notes

STEVEN K. BURIAN!, DANIEL J. ERASMUS’, CLAIRE M.

SHRIMPTON?, DOUGLAS C. CURRIE?, DONNA J. GIBERSON%,

DEZENE P.W. HUBER?

ABSTRACT

The Baetis vernus group (Ephemeroptera: Baetidae) — which includes B. brunneicolor

McDunnough, B. bundyae Lehmkuhl, B. hudsonicus Ide, B. jaervii Savolainen, B.

liebenauae Keffermiiller, B. macani Kimmins, B. subalpinus Bengtsson, B. tracheatus

Keffermiiller & Machel, and B. vernus Curtis — is both diverse and taxonomically

tangled. Some members of the group — B. brunneicolor, B. bundyae, and

B. hudsonicus — have been previously found in North America. The remainder of the

group is known to be only of Palearctic distribution, including B. vernus, which has a

wide trans-Palearctic distribution. We report the collection of specimens from the

Northwest Territories and British Columbia that we have identified as B. vernus using

DNA barcoding and morphological examination and provide characters to assist

separation of the North American members of the group from B. vernus. A genetically

cohesive Holarctic clade for B. vernus likely relates to a Beringian dispersal event.

This substantial expansion of the known range of B. vernus adds new phylogeographic

and ecological complexity, but it may also help to provide further clues to the

evolutionary history of this group.

INTRODUCTION

Mayflies of the Baetis vernus group (Savolainen et al. 2007; Stahls and Savolainen

2008; Drotz et al. 2012) are widespread across the Holarctic, but distributions of its

members are challenging to determine because many are difficult to separate in the most

commonly collected larval stage using morphological characters (Stahls and Savolainen

2008; Drotz et al. 2012). This is due both to similarity of characters among group

members and to high levels of variation relating to environmental conditions

(Bauernfeind and Humpesch 2001; Stahls and Savolainen 2008). Stahls and Savolainen

(2008) stressed the importance of combining molecular and morphological data to sort

out species distributions in this group. |

Until recently, only three species in this group were known in North America

(McCafferty and Jacobus 2017). Baetis brunneicolor McDunnough is widespread in the

Nearctic: it is reported from across Canada, including Arctic and Sub-Arctic zones

(Harper and Harper 1981; Cordero et al. 2017; Giberson and Burian 2017) and is found

in the northeastern, northwestern, and southeastern United States (USA; McCafferty and

Jacobus 2017). Baetis bundyae Lehmkuhl has a generally northern distribution in

Nearctic and Palearctic: in North America, it is widespread across the north but also

extends into the northern USA (Giberson ef a/. 2007; Giberson and Burian 2017). Baetis

hudsonicus Ide has so far been reported only in northern and far northern Canada

(Cordero et al. 2017; Giberson and Burian 2017; McCafferty and Jacobus 2017).

Recent collecting efforts in northern British Columbia (Huber ef a/. 2019) and in the

Northwest Territories (Cordero et al. 2017) revealed four specimens whose cytochrome

! Southern Connecticut State University, New Haven, CT 06515

* University of Northern British Columbia, Prince George, BC, V2N 4Z9

3 Royal Ontario Museum, Toronto, ON, M5S 2C6

4 Corresponding author: University of Prince Edward Island, Charlottetown, PE, CLA 4P3;

giberson@upel.ca

4 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

oxidase I (COI) barcode matched Palearctic specimens of Baetis vernus Curtis, a species

not previously reported in North America. Baetis vernus specimens showed many

morphological similarities to B. brunneicolor, potentially causing confusion when

determining the distribution of the two species in North America. Recently, Webb et al.

(2018) recommended that B. brunneicolor and B. vernus be treated as a species complex

(the B. vernus complex) and not identified further if identifying larvae using current

morphologically-based keys. Here, we describe DNA barcode data of the Canadian B.

vernus specimens, demonstrating their genetic similarity to Palearctic B. vernus and

distance from other B. vernus group members, as well as the relevant morphology of

those same specimens compared to North American B. brunneicolor characteristics.

METHODS AND MATERIALS

Mayfly larvae examined in this study resulted from recent aquatic insect sampling in

river and lake habitats in northern British Columbia (BC), Yukon (YT) and Northwest

Territories (NT), plus examination of archived specimens in the Canadian National

Collection (CNCI) in Ottawa (Giberson and Burian 2017; Huber et al. 2019). Collection

locality, voucher, and DNA sequence data for specimens that were collected and/or

analyzed in this study are described in Table 1. The cytochrome oxidase I (COI) barcode

region (Hebert et al. 2003; Ball et al. 2005) of the Crooked River, BC, specimen was

sequenced at the Biodiversity Institute of Ontario, and other barcode sequence data were

extracted from public databases. North American B. vernus group specimens were

compared to other described members of the B. vernus for which sequence data were

available (exceptions: B. jaervii Savolainen and B. tracheatus Keffermiiller & Machel). A

FASTA file of all sequences used in this study, including sequence ID and accession

information, is available as supplemental data. All sequence data are publicly available as

listed in Table 1 and as BOLD IDs (most sequences) or an NCBI accession number

(Yellowknife specimen sequence) in Figure 1. Barcode sequences were aligned with

ClustalW and visualized with FigTree v.1.4.3.

Specimens were observed for morphological characters, colouration, and colour

patterns under Wild MSA stereoscopic and Bausch & Lomb phase contrast compound

light microscopes (up to 1000x magnification). Mouth and body parts of the larvae were

dissected in 80% alcohol and slide mounted in Hoyer's Mounting Media. Specimens

were photographed using a Nikon D300s DSLR and the Nikon Camera Control Pro2®

software. All measurements were made using a calibrated ocular micrometer (nearest

0.10 mm). Measurements were made from entire specimens and/or parts (not mounted on

slides) that were held as flat as possible (without inducing distortion) using sections of

broken glass microscope slides and coverslips.

Specimens were determined to species by comparing morphological characters to all

pertinent descriptions and morphological studies of members of the Baetis vernus species

group on a global basis, as well as Nearctic keys to species of Baetis (Ide 1937; Leonard

1950; Macan 1957; Keffermiiller and Machel 1967; Miiller-Liebenau 1969; Lehmkuhl

1973; Morihara and McCafferty 1979a, b; Jacob 2003; Wiersema et al. 2004; Savolainen

2009; Jacobus et al. 2014). In addition, reared specimens of B. brunneicolor from the

USA and voucher specimens of larvae of B. macani and B. jaeverii (provided by E.

Savolainen) from Finland were used to evaluate characters observed on B. vernus

specimens from northern Canada.

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J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

6 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

RESULTS AND DISCUSSION

DNA barcodes for B. vernus specimens collected by us (Cordero et al. 2017; Huber et

al. 2019) and others from British Columbia and the Northwest Territories were virtually

identical to each other and to sequences of B. vernus collected in Finland. The sequences

were substantially different [much greater than 2% (Zhou et al. 2009; Webb et al. 2012;

Cordero et al. 2017)] from other B. vernus group members, including group members

found in North America (Fig. 1). Morphological examination of the Northwest Territories

and Crooked River, BC, specimens revealed traits similar to B. brunneicolor, such that

the specimens keyed to B. brunneicolor in the most recent key to Baetis spp. in North

America (Wiersema et al. 2004, updated with recent couplet patches found in Jacobus et

al. 2014).

B.subaipinus|EPHF1044|BOLD:AAC4082|Norway

B.liebenauae|FBAQU770-10|/BOLD:AAB9706|Germany

B.bundyae|CUMAY028-09|BOLD:AAB6788|Churchill, MB

B. hudsonicus|CUMAY027-09|/BOLD:AAD2388|Churchill, MB

B.macani|GBA14627-14|BOLD:AAB1423|Finland

B. vernus|GBMIN37275-13|BOLD:AAB 1424 /Finland

B.vernus}GBMIN37274-13/BOLD:AAB 1424 Finland

B.vernus|BBGCO1179-15|BOLD:AAB1424/Canyon Ck, BC

B.vernus|KJ675352|No BIN|Yellowknife, NT

B.vernusi}CREPH018-16/BOLD:AAB1424|Crooked River, BC

B.vernusiBBGCO1178-15|BOLD:AAB1424|Canyon Ck, BC

B.brunneicolorK CUMAY062-09|BOLD:AAA5478|Churchill. MB

0.02

Figure 1: A DNA barcode comparison of Palearctic and Nearctic specimens of B. vernus and

other closely related baetid species derived from our own collections (Yellowknife and

Crooked River B. vernus specimens) and from the BOLD database (Ratnasingham and Hebert

2017). The sequences were aligned with Clustal W and visualized with FigTree 1.4.3.

Approximate collection locations in Canada and Europe are listed next to each specimen

along with that specimen’s BIN (Ratnasingham and Hebert 2013). Sequence data are publicly

available using the BOLD IDs (most specimens) or NCBI accession number (Yellowknife

specimen) associated with each specimen.

These combined results prompted us to look at other B. vernus group specimens

collected in northwest Canada. These consisted of specimens labeled "B. brunneicolor",

"B. bundyae", and "Baetis n. sp. (vernus group)" collected from northern Yukon

(Porcupine River drainage), southern Yukon (streams along the Alaska Highway), and

streams in the Mackenzie Mountains west of the Mackenzie River Valley in the

Northwest Territories (Table 1). Baetis bundyae and B. hudsonicus specimens were

usually easy to distinguish from B. brunneicolor and B. vernus due to the presence of

narrow gills on the abdomen. A third group of specimens collected from northern Yukon

and from the Mackenzie Mountains west of the Mackenzie River (Table 1) appeared to

show characteristics of both groups, with narrow gills like B. bundyae but other

characters more consistent with B. vernus. Due to the age and/or storage conditions of

these latter specimens, DNA barcoding was not possible, so the primary concern was

distinguishing the larvae of the widespread Nearctic B. brunneicolor and the newly

discovered B. vernus. Morphological features are described for each species below,

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 7

including a summary of characters that can be used to distinguish the two species in

northwestern Canada. .

Baetis brunneicolor — General Morphological Description of Larva (Figs. 2—13):

Head: Frons — General shape subtriangular with blunt apex, lateral edges straight or

slightly concave (Fig. 2). Antennae — Scape and pedicel with many small hair-like setae,

no robust setae present. Small hair-like setae seem to be restricted to distal half of scape

(Figs. 3a, 3b). No apparent pattern of small setae on pedicel.

Figure 2. Baetis brunneicolor: dorsal

view of frons.

Figure 3. Baetis brunneicolor: antennal scape

and pedicel; (b) is enlargement of lower

section of (a).

Mouthparts: Labrum — Lateral edges tend to be rounded, never appearing straight

(Fig. 4a). Dorsal setal pattern 1 long median pair, a gap then row of 4-5 smaller setae

(Fig. 4b) extending to edge of anterior margin (i.e., 1 + 4—5). Dorsal surface with many

setae, most concentrated near posterior corners. Middle of dorsal surface with somewhat

rounded raised area surrounded with small surface setae, no obvious dark marking

associated with raised medial area. Right Mandible — First tooth of outer incisor larger

than second tooth and with squared-off outer edge; second tooth larger than third tooth,

and with blunt outer edge; and third tooth smallest of three with rounded outer edge (Fig.

5a, left). This is the “new” condition after moulting, worn teeth are much more similar in

size and shape (Fig. 5a, right). Prostheca with pectinate tip, most apical setae about same

size and equally spaced (Fig. 5b). Left Mandible — One or two small auxiliary teeth

present between molar teeth and large apical projection on anterior margin (Fig. 6a).

Outer incisor with first tooth slightly larger than second tooth and with squared-off edge;

second tooth distinctly larger than third tooth and both teeth with irregularly pointed

apices (Fig. 6b, left). This is the “new” condition after moulting, worn teeth are much

more similar in size and shape (Fig. 6b, right). Maxillae — Four maxillary canines present

that lack serrations (Fig. 7a, 7b). Dense brush of long setae along anterior margin of

galea-lacinia below canines; margin below setae slightly concave (Fig. 7b). Maxillary

palpi two segmented and both segments with many small hair-like setae (Fig. 7a, 7c).

Segment 1 of maxillary palpi as long as segment 2. Tips of maxillary palpi extend about

one-third of their total length above the tips of canines. Labium — Paraglossae broad

with mostly straight margins approaching the apices (Fig. 8a, 8b). Apices of paraglossae

with 12—15 long setae in two rows. Glossae with broadly pointed apices, ventral surface

with single row of about nine long setae located along medial edge (Fig. 8b). Segment 2

of labial palpi with either well developed inner apical lobe and distinctly concaved

8 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

margin below lobe (Fig. 8a) or moderately developed inner apical lobe and only slightly

concave margin below lobe (Fig. 8c).

Zi LLL Zi

Figure 4. Baetis brunneicolor. two views of the labrum — (a) entire labrum cleared and slide-

mounted; (b) anterior area enlarged to show setal pattern.

Figure 5: Baetis brunneicolor: right mandible — (a) the difference in wear on new and old

incisors (new incisors of next instar visible in cleared mandible); (b) entire mandible with

prostheca enlarged in inset.

Figure 6. Baetis brunneicolor: left mandible — (a) entire mandible with insets showing the

auxiliary teeth; (b) the difference in wear on new and old incisors (new incisors of next instar

visible in cleared mandible).

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 9

Figure 8. Baetis brunneicolor: \abium — (a) entire labium; (b) glossa and paraglossae of

labium; (c) labial palp.

Forelegs: Femora broadest near midpoint of segment (Fig. 9a). Outer edge with two

staggered rows of long, blunt setae that have uniform width from base to tip, setae

become more widely spaced and fewer in number near apex of segment (Fig. 9a). Tibia

and tarsus generally seem to be much stouter (i.e., wider and shorter) compared to those

of B. vernus (compare Fig. 9b to Fig. 21b). Foreleg claw with about nine denticles that

progressively become larger from base toward apex, apex of claw appears slightly

attenuated (i.e., narrowed) (Fig. 9b).

Abdomen: Abdominal Tergite V — Shape typical for abdomen with outer edge of

tergite slightly tapering posteriorly (Fig. 10, top). Posterior lateral corners with minimal

dark brown colour in fresh specimens at gill insertion. Numerous scale setae present and

few scattered hair-like setae between scale setae. Surface with faint cuticular ridges (i.e.,

weakly grainy) (Fig. 10, bottom). Posterior margins with spinules, but not darkly

pigmented compared to rest of tergite (Fig. 10, top). Abdominal Gills 4 and 5 — Gill 4

larger than gill 5, but both have same basic oval shape with smoothly curved dorsal edge

and outer margin (Fig. 11). Both gills have marginal teeth; at 20X magnification gill 4

has about 8 teeth/0.05 mm of edge and gill 5 has 8—9 teeth/0.05 mm of edge. Both gills

have distinct central trachea with one or two smaller side branches visible (Figs. 11). Gill

4 length slightly less than twice the width (i.e., width x 1.8=length). Gill 5 has same

length/width relationship as gill 4. Paraprocts — Inner apical edges with regular, large

spines (Fig. 12). Surface with scattered long, hair-like setae that are more or less

uniformly distributed over surface (Fig. 12). No other distinctive surface features or

textures.

10 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Colour Pattern of Body: Overall body colour uniform brown with less distinct

contrasting lighter areas (Fig. 13a). Pronotum lacking bi-lobed brown spots, but large

somewhat “c-shaped” diffuse blotches are sometimes present (Fig. 13b). Meso- and

metanotum of thorax mostly brown with some lighter streaks and spots, especially on the

mesonotum. Abdominal terga with submedian paired brown spots; these are faint on

some specimens (Fig. 13a). A medial pale spot with paired lateral pale spots separated by

brown background colour seems a common pattern on terga. Posterolateral edges of terga

pale compared to brown medial part of terga.

Figure 9. Baetis brunneicolor: foreleg — (a) entire foreleg, with numbers denoting areas of the

femur with setal patterns shown at right; (b) tibia and tarsus, with inset showing denticles on

claw.

General Shape of Abdomen: The overall shape of the abdomen, viewed dorsally, 1s

one of a gradually tapering cylinder that is widest at segment I and narrowest at segment

X (Fig. 13a). The shape results from a change in the width/length ration from anterior

segments to posterior segments. Segment I is about three times as wide as long and

segment X is almost as wide as long.

Baetis vernus — General Morphological Description of Larva (Figs. 14—26):

Head: Frons — General shape subtriangular with blunt apex, lateral edges either

straight or slightly concave (without slide mounting intact specimens can even appear

slightly convex) (Fig. 14). Antennae — Scape and pedicel with many small hair-like

setae, no robust setae present. Small hair-like setae seem to be restricted to distal part of

scape (Fig. 15). No apparent pattern of small setae on pedicel.

(b) enlargement

ite:

terg

ire

— (a) ent

1 tergite V

ina

icolor: Abdomi

J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

Figure 10. Baetis brunne

showing cuticular patterns

Figure 11. Baetis brunneicolor

‘gills 4 and 5.

1 of the lower

detai

ing

SS :

ht show

inset at rig

inse

ith

WwW

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

paraprocts,

Baetis brunneicolor

paraproct at left.

igure 12

showing

lour

ing co

iews of several larvae

ions of larvae showi

(a) dorsal v

rior sec

il of ante

body features —

(b) deta

SS)

Ss 0

Oo OU

ioe

bose 7

on —

om =O

fled CD

patterns on thorax

dorsal view of frons.

Figure 14. Baetis vernus

J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018 13

Figure 15. pe and el, wi

highlighting different features.

Mouthparts: Labrum — Lateral edges tend to be straight, or at most slightly rounded

(Fig. 16a), making the labrum appear somewhat rectangular. Dorsal setal pattern 1 long

medial pair, a gap then row of 3-4 smaller setae (Fig. 16b) extending to edge of anterior

margin (i.e., 1 + 3-4). Dorsal surface with relatively few scattered setae, most tend to be

concentrated near edges of somewhat triangular raised area that is flanked by dark bands

(bands faint on some specimens) (Fig. 16a). Right Mandible — First tooth of outer

incisor larger than second tooth and with squared-off outer edge; second tooth only

slightly larger than third tooth and both with irregularly pointed tips (Fig.17a, left). This

is the “new” condition after moulting, worn teeth are much more similar in size and

shape (Fig. 17a, right). Prostheca pectinate with a single row of setae along inner edge

near apex (Fig. 17b). Setae of variable lengths and some form a cluster near apex of

prostheca. Left Mandible — One or two small auxiliary teeth present between molar teeth

and large apical projection on anterior margin (Fig. 18a). Outer incisor with first tooth

slightly larger than second tooth and with squared-off edge; second tooth only slightly

larger than third tooth and both teeth with irregularly pointed apices (Fig. 18b, left). This

is the “new” condition after moulting, worn teeth are much more similar in size and

shape (Fig. 18b, right). Maxillae — Four maxillary canines present that lack serrations

(Figs. 19a, 19b). A dense brush of long setae along anterior margin of galea-lacinia below

canines; margin below setae straight or only slightly concave (Figs. 19a, 19b). Maxillary

palpi two segmented and both segments with many small hair-like setae (Fig. 19a).

Segment | of maxillary palpi as long as segment 2. Tips of maxillary palpi extend about

one-third of their total length above the tips of canines (Fig. 19a). Labium — Paraglossae

with broad curved apices (Figs. 20a, 20b). Apices of paraglossae with 11—12 long setae in

two rows (Fig. 20b). Glossae with narrowly pointed apices, ventral surface with single

row of about six long setae located along medial edge (Fig. 20b). Segment 2 of labial

palpi with moderately developed inner apical lobe and slightly concaved margin below

lobe (Fig. 20a).

2 ¢3 az

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with inset showing prostheca.

J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018 15

Figure 18. Baetis vernus: left mandible — (a) entire mandible with inset showing the auxiliary

teeth; (b) the difference in wear on new and old incisors (new incisors of next instar visible in

cleared mandible).

ntire maxilla; (b) canines.

Figure 19. Baetis vernus: maxilla — (a) e

16 J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

Figure 20. Baetis vernus: labrum — (a) entire labium; (b) glossa and paraglossa.

Forelegs: Femora about same width from base to apex (Fig. 21a). Outer edge with

two staggered rows of long, blunt setae, many of which have narrow bases and broad

ends, but some setae have uniform width from base to tip (Fig. 21a). Setae on outer edge

of femora are numerous near the base of the segment and gradually become fewer in

number approaching joint with tibia, stopping entirely just before the joint with the tibia

(Fig. 21a, right hand panels). Tibia and tarsus are thinner and more delicate compared to

those of B. brunneicolor (compare Fig. 21b to Fig. 9b). Foreclaw with 8-10 denticles,

small near base of claw and only gradually become larger toward apex making the row

appear more uniform over its length (Fig. 21b). Apex of claw thicker and not noticeably

attenuated as in B. brunneicolor (compare Figs. 9b and 21b)

Abdomen: Abdominal Tergite V — Shape typical for abdomen with outer edges of

tergite nearly parallel (Fig. 22). Posterior lateral corners with distinct dark brown colour

at gill insertions (Fig. 22). Numerous scale setae present, but widely spaced over surface

of cuticle and with few scattered hair-like setae between scale setae (Fig. 22). Surface

with distinct cuticular ridges (i.e., moderately grainy). Posterior margins with spinules,

pigmented darker brown compared to lighter brown colour of rest of tergite (Fig. 22).

Abdominal Gills 4 and 5 — Gill 4 is larger than gill 5 and shaped distinctly different

shape compared to gill 5 (Fig. 23). Gill 4 more subtriangular with a bulged dorsal

margin. Gill 5 closer to sub-oval shape typical of B. brunneicolor gills (Fig. 23). Both

gills have marginal teeth; at 20X magnification, gill 4 has about 8 teeth/0.05 mm of edge

and gill 5 has 8—9 teeth/0.05 mm of edge. Gill 4 has only faint traces of the central

trachea and gill 5 has no visible trachea (Fig. 23). Gill 4 almost exactly twice as long as

wide. Gill 5 length is slightly less than twice the width. Paraprocts — Inner apical edges

with irregular row of large spines, which become smaller around the apical corner

(Fig. 24). Surface with few scattered hair-like setae and dense cluster of small cuticular

scales near outer apical edge (Fig. 24).

Colour Pattern of Body: Overall body colour of Northwest Territories (NT)

specimen is much more contrasting compared to the BC specimen (Figs. 25, 26).

Generally, body somewhat brown with large pale areas. Pronotum with distinct paired

medial brown spots or blotches, lateral edges dark brown, but rest of surface pale (Figs.

25, 26). Thorax with several large pale areas and smaller distinct brown spots or blotches

(BC specimen seems closer in thoracic colour patterning to B. brunneicolor than NT

specimen) (compare Figs. 25, 26 to Fig. 13). Abdominal terga I-IV of NT specimen

mostly brown with large paired pale spots and a smaller medial spot (Fig. 25). Tergite V

mostly white with limited brown marks at anterior margin and laterally. Terga VI-IX

similar in colour to preceding terga. Tergite X white. The BC specimen had a much less-

contrasting overall colour pattern but almost the same pattern of marks and spots (Fig.

J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018 17

26). However, tergite VI on the BC specimen was not pale, but patterned similar to other

terga. Also, tergite X was uniformly light brown, not pale as in the NT specimen.

General Shape of Abdomen: The overall shape of the abdomen, viewed dorsally,

seemed to change more gradually over its length, not appearing distinctly tapered as in B.

brunneicolor (compare Figs. 25, 26 to Fig. 13). The change in width/length relationship

was less per segment, which resulted in the appearance of a more uniformly shaped

abdomen. Edges of individual tergites seemed less tapered compared to B. brunneicolor.

On the BC specimen (Fig. 26), where gills were lost, it was clear that the abdomen did

taper from anterior to posterior, segment I was approximately 2.3 times as wide as long.

Segment X was slightly wider than long.

Figure 21. Baetis vernus: foreleg — (a) left: entire foreleg, with numbers denoting areas for

femur setal patterns; panels at right show the same view at different focus settings to show

setal patterns; (b) tibia and tarsus, with insets showing denticles on claw.

Figure 22. Baetis vernus: Abdominal tergite V ~ (a) entire tergite; (b) enlargement showing

cuticular patterns.

t different focus settings to

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

iP)

oped

cD)

area

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se} 5

= 5

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i S.

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: =f

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= =|

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a fy sa

show setal patterns.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

fe in

iews of two larvae collected near Yellowkn

lour patterns and body shape

dorsal and lateral v

is vernus:

Figure 25. Baet

Northwest Terr

ing co

showi

tories,

ia,

h Columbi

Hiern Br

in no

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showing colour patterns and body shape.

: dorsa

vernus

Figure 26. Baet

20 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

Diagnosis of larvae of Baetis vernus group species in North America:

In North America, the Baetis vernus group includes: B. brunneicolor, B. bundyae, B.

hudsonicus, and B. vernus. Larvae of B. bundyae and B. hudsonicus can be separated

from those of B. brunneicolor and B. vernus by the elongate abdominal gills, which are

distinctly longer than twice their width. Baetis bundyae can be separated from B.

hudsonicus by the presence of a short terminal filament, usually much shorter than

lengths of adjacent cerci, whereas B. hudsonicus has a long terminal filament that is

about equal to the length of adjacent cerci (secondarily, populations of B. hudsonicus

seem to be completely parthenogenetic, no males have been detected).

Larvae of Baetis vernus can be separated from those of B. brunneicolor by the

presence of the following combination of characters:

(1) Dorsal surface of body with distinctive contrasting colour pattern of brown with large

pale spots and marks (Fig. 27), especially on abdominal terga, terga V and X mostly pale

but other terga brown with large paired pale submedian spots,

(2) General shape of abdomen from dorsal perspective somewhat cylindrical eyeing to

very gradually taper from segments I to X,

(3) Dorsal surface of labrum with subtriangular raised area flanked by two brown bands

that converge medially near base of notch in anterior margin, dorsal setal formula 1+3—4,

(4) Prostheca of right mandible with cluster of setae along inner edge near apex,

(5) Paraglossae of labium with apices distinctly curved inward,

(6) Femora with large blunt setae along outer edge with narrow bases and broad ends,

(7) Foretibia and -tarsus slender,

(8) Foreclaw with 8-10 denticles that gradually enlarge from base of claw toward tip,

(9) Tip of claw not attenuated (i.e., narrowed) beyond denticles,

(10) Abdominal terga with distinctive dark brown shading around gill insertions and

spinules along posterior margins dark brown,

(11) Cuticle of abdominal terga moderately grainy with many distinct cuticular ridges

among bases of scale setae, and

(12) Abdominal gills 2-4 with only faint traces of the medial trachea and trachea not

visible on other gills.

Mature larvae of B. brunneicolor can usually be separated from those of B. vernus by

the presence of the following combination of characters:

(1) Dorsal surface of body relatively uniformly brown, lacking large distinct contrasting

pale spots or marks, especially on abdominal terga where some small paired dark marks

are present (Fig. 27), Tergite X is usually a uniform light brown on mid-instar larvae but

can be pale on black wing pad larvae,

(2) General shape of abdomen from dorsal perspective conical appearing to taper more

distinctly from segments I to X,

(3) Dorsal surface of labrum with rounded raised area with no associated dark bands,

dorsal setal formula 1+4—5,

(4) Prostheca of right mandible with single row of anaiee spaced setae along inner

edge near apex,

(5) Paraglossae of labium with apices straight or only slightly curving inward,

(6) Femora with large blunt setae that usually have uniform width from base to tip,

(7) Foretibia and -tarsus stout,

(8) Foreleg claw with only about nine denticles that appear to change more abruptly in

size from base of claw toward tip,

(9) Tip of claw attenuated (i.e., narrowed) beyond denticles,

(10) Abdominal terga with only small areas of brown shading around gill insertions and

spinules along posterior margins not darker that rest of surface,

(11) Cuticle of abdominal terga weakly grainy with few widely spaced cuticular ridges

among bases of scale setae, and

(12) Abdominal gills 2—6 with distinct medial trachea, lateral trachea also visible on

larger gills.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 21

Figure 27. Comparison of B. vernus and B. brunneicolor larvae at approximately the same

stage of development. The B. vernus specimens were collected from near Yellowknife in

Northwest Territories and from the Crooked River in northern British Columbia, and the B.

brunneicolor \arva is from Wisconsin.

Distribution of B. vernus in Canada

The distribution of B. vernus in Canada is still unclear, as there are currently only

four verified specimens of B. vernus from North America. Their distribution ranges from

central British Columbia to the south-central Northwest Territories (Fig. 28, Table 1).

However, B. vernus overlaps in distribution with B. brunneicolor (Fig. 28), so some

specimens previously identified as B. brunneicolor from this region may be B. vernus,

which could extend the distribution considerably. The specimens mapped in Fig. 4 (and

listed in Table 1) were confirmed through comparison with confirmed specimens of B.

brunneicolor and descriptions of B. vernus from the Palearctic. Another potential source

of confusion stems from a group of specimens from northern Yukon and the Mackenzie

Mountains that show characters of both species and may represent hybrid forms or a new

species in the B. vernus group (Table 1). We show that a combination of DNA barcoding

and morphological examination can resolve the two species, but targeted collecting

should occur through northern Canada to obtain specimens for molecular examination to

assess distribution patterns for species within the entire Baetis vernus group.

Phylogeography

Morphological analyses and DNA barcoding now confirm the Nearctic presence of B.

vernus, and the locations of the currently known specimens of B. vernus indicate a

widespread distribution in North America. Such’a Holarctic distribution is not surprising

as it is analogous to the distribution of B. bundyae (Giberson et al. 2007; Savolainen ef

al. 2014) and other Ephemeroptera (Kjerstad et al. 2012) and because Baetis spp. may

be particularly pre-adapted for rapid dispersal due to their wing-loading characteristics

(Corkum 1987). These results may also imply a widespread northern Asian distribution

for B. vernus — or a common ancestor of it and other members of the B. vernus group —

leading to a Beringian dispersal event. Members of the B. vernus group seem to be

variably tolerant of a range of lentic (standing water) and lotic (running water) habitats

pp J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

(Bauernfeind and Humpesch 2001; Giberson et al. 2007; Savolainen et al. 2007; Drotz et

al, 2012), and differential use of such habitats may be a driver of structured populations

or speciation (Drotz et al. 2012; Stahls and Savolainen 2008). From our direct knowledge

of collections (Cordero et al. 2017; Huber eft al. 2019) or extrapolation from GPS

coordinates (BOLD specimens in this study), the North American B. vernus specimens

were collected in a range of situations, including a marshy area (Cordero ef al. 2017), a

slow-moving outflow of a lake (Huber ef al. 2019), and seemingly typical lotic

environments (BOLD specimens). Baetis vernus in Europe is mostly — but not

exclusively — known from lotic systems (Savolainen et a/. 2007). This seeming ability to

reproduce and survive in a variety of habitats may have also aided B. vernus’ dispersal

ability.

DNA barcoding was vital for the initial detection of this species in North America and

remains a valuable tool for distinguishing between B. brunneicolor and B. vernus (as

well as potential new species in the group) in northern Canada. Morphological work on

these specimens has revealed new questions regarding B. vernus group taxonomy and

phylogeograhy, and these results highlight the need for substantial further collection of

the B. vernus group in northern Canada. The growing use of eDNA surveys of likely

habitat will be important for extending our knowledge of this and other mayfly species.

0 250 500

kilometres

: ook r + ;

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cei ea a SF 3 P ae y

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Figure 28: Confirmed record locations for B. vernus and B. brunneicolor across northern

Canada. Red symbols = B. vernus, with different symbol shapes denoting different collections

[red inverted triangle: Cordero et al. (2017); red circle: Huber et al. 2019; red square: BOLD-

mined B. vernus data (two specimens)]. Blue symbols = B. brunneicolor, with different

symbol shapes denoting different collections [blue circles: Giberson and Burian (2017),

confirmed through comparison with eastern B. brunneicolor specimens; blue inverted

triangles: Cordero et al. (2017); blue triangles: adult specimens reported in Harper and Harper

1981].

J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018 a3

ACKNOWLEDGEMENTS

Aspects of this research were funded by the University of Northern British Columbia,

the Canada Research Chairs Program, the Canada Foundation for Innovation, the British

Columbia Knowledge Development Fund, and the Natural Sciences and Engineering

Research Council of Canada.

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J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 23

Corrections for the Hemiptera: Heteroptera of Canada

and Alaska

G.G.E. SCUDDER!

ABSTRACT

A total of 175 changes to the current checklist of Hemiptera: Heteroptera of Canada

and Alaska are reported. Eighty deletions, eighty-eight nomenclature changes, and

seven spelling corrections are detailed. In addition, comments are given on Anthocoris

tomentosus Péricart, Orius diespeter Herring, O. tristicolor (White), and Tupiocoris

agilis (Uhler).

Key words: Changes, checklist, Heteroptera, Canada, Alaska

INTRODUCTION

Maw et al. (2000) published a checklist of the Hemiptera of Canada and Alaska,

giving details of the occurrence of the species of Heteroptera. Since then, there have been

a large number of taxonomic changes that have resulted in deletions and nomenclature

modifications for many of the taxa. In addition, a few spelling errors have been noted.

Details of the 175 changes are outlined here, and comments on four taxa are given.

The order of taxa follows Maw et al. (2000), but species are listed in alphabetical

order in each family.

Museum abbreviations are as follows:

CNC Canadian National Collection of Insects, Agriculture and Agri-Food Canada,

Ottawa, Ontario

RBCM _ Royal British Columbia Museum, Victoria, B.C.

UAM University of Alaska Museum, Fairbanks, Alaska.

UBCZ — Spencer Entomological Collection, Beaty Biodiversity Museum (formerly

Spencer Entomological Museum, Department of Zoology) University of

British Columbia, Vancouver, B.C.

USNM National Museum of Natural History (formerly United States National

Museum), Washington, D.C.

SYSTEMATIC TREATMENT

I. Deletions

Family CORIXIDAE

Glaenocorisa quadrata Walley

This corixid was originally described by Walley (1930) from Quebec. Jaczewski and

Lansbury (1961) followed Ossianilsson (1960) and considered G. guadrata a synonym of

G. cavifrons (Thomson), and stated that G. cavifrons was at most a subspecies of G.

propinqua (Fieber). Although doubted by Brown (1946), this was accepted by Jansson

(1986), who concluded that there were two subspecies of G. propinqua, with G.

propinqua cavifrons occurring in North America. However, as noted by Jansson (2002),

G. cavifrons was raised to specific status by Jansson (2000), because the two subspecies

are sympatric in Scotland and northern Finland. Hence, G. quadrata Walley should be

1 Corresponding author: Spencer Entomologist Collection, Beaty Biodiversity Museum, University of

British Columbia, 2212 Main Mall, Vancouver, B.C. V6T 1Z4; scudder@zoology.ubc.ca

26 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

deleted, and all occurrence records in North America placed under G. cavifrons

(Thomson).

Sigara modesta (Abbott)

This species was recorded from British Columbia by Downes (1934) as Arctocorisa

modesta, with a listing of material from Vernon, 26.1x.1919 (W. Downes), determined by

G.S. Walley. This record was accepted by Polhemus ef al. (1988) and repeated in Maw et

al. (2000).

Sigara modesta was not listed from British Columbia by Hungerford (1948),

Lansbury (1960), or Scudder (1977). Scudder (1977) excluded S. modesta (Abbott) from

the British Columbia list of Corixidae, noting that there were no other records of this

species in the province. He also noted that other specimens in the Downes collection that

were labelled “modesta”’ were in fact S. grossolineata Hungerford and that the Vernon

determination was probably incorrect. Unfortunately, he overlooked the Vernon record of

S. washingtonensis listed in Hungerford (1948), even though there were specimens with

the appropriate date in the Downes collection that had been donated to the UBCZ

collection in 1958. However, Scudder (1977) did quote a Vernon record in Lansbury

(1955), although the date in the latter was printed as 26.1x.1929 (W. Downes).

Hungerford (1948) lists a male specimen with data Vernon, 26.1x.1919 (W. Downes),

under his new species S. washingtonensis. In the UBCZ collection, with collection

numbers COR3139-COR3141, I have located one male and three females with data

“Vernon, 26.1x.1919 (W. Downes)’. These are from the Downes collection donated to

UBCZ in 1958; it is assumed that these are the specimens mentioned by Downes (1934).

Hence, the record of S. modesta (Abbott) from British Columbia should be deleted.

Trichocorixa verticalis fenestrata (Walley)

This is treated the same as T. verticalis verticalis (Fieber) by Jansson (2002). Hence,

T: verticalis fenestrata should be deleted and records should be placed under 7° verticalis

verticalis.

Family SALDIDAE

Salda anthracina Uhler

This saldid was recorded from Alaska and British Columbia by Polhemus (1988) and

from Alaska, British Columbia, Northwest Territories, and the Yukon Territory in Maw et

al. (2000). Specimens in the UBCZ collection from Alaska, Northwest Territories, and

the Yukon were initially determined by me as S. anthracina using the key in Schuh

(1967), with particular attention paid to the fact that this key noted that the second

antennal segment in S. anthracina was pale. At that time, I had not seen specimens of S.

anthracina from elsewhere. This led me to record S. anthracina in Maw et al. (2000).

These specimens were as follows:

AK: Donnelly Cr., Richardson Hwy., 15.vii.1985 (S.G. Cannings) [UBCZ].

NT: 15 km WN of BC border, Liard Hwy., 26.vi.1985 (E. Bijdemast) [UBCZ].

YK: Dawson, 31 km E, 26.vi.1980 (Bruce Gill) [UBCZ].

Kluane N.P., Slims River flats, 21.vi1.1979 (G.G.E. Scudder) [UBCZ].

Kluane, Slims R. delta, 7.viii.1986, (S.G. Cannings) [UBCZ].

Mi 1059 Alaska Hwy., Kluane L., 5.vii.168 (Campbell-Smetana) [CNC].

Moose Cr., 68°31'N 137°O1'W, 26.vi1.1982 (G.G.E. Scudder) [UBCZ].

Von Wilczek L., 2.v1i.1980 (Bruce Gill) [UBCZ].

After receiving one male and one female determined by the late J.T. Polhemus as S.

provancheri Kelton & Lattin, and noting that these specimens from Colorado, Weld

County, Pawne National Grasslands, July 1970 (R.T. Bell), had the second antennal

segment mostly pale and the first segment dorsally flavescent and ventrally fuscous, I

redetermined my western specimens as S. provancheri and not S. anthracina. As a result,

in Scudder (1997), I recorded S. provancheri from Dawson (31 km E), Moose Cr., Slims

R. delta and von Wilczek Lks. The specimens from Alaska and the Northwest Territories

were also determined as S. provancheri, and not S. anthracina.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 of

I note that Schuh (1967) stated that S. anthracina is quite variable morphologically

and lives in situations similar to those preferred by S. provancheri, which was recorded

by Schuh (1967) as S. bouchervillei. | have been unable to trace the original record of S.

anthracina from Alaska, although this record is reported by Drake and Hoberlandt

(1950), Drake and Hottes (1950), and Drake (1952). D. Sikes (in litt., 15 March 2018)

informs me that there are no specimens under S. anthracina in the University of Alaska

Museum.

All the specimens I have seen from Alaska that might be S. anthracina are, in fact, S.

provancheri.

Drake and Hottes (1950) cite a record of S. provancheri as S. bouchervillei, from

Alaska (Rampart), noting that his species is quite variable in size and degree of wing

development. Salda provancheri was also recorded from Cook Inlet, Valdez Bay, in

Alaska by Bahr and Schulte (1976). Polhemus (1988) recorded S. provancheri from

Alaska, British Columbia, and the Northwest Territories.

Salda anthracina was recorded from British Columbia by Downes (1927) as

Lampracanthia anthracina, with the observation that the British Columbia material was

in the CNC. I have been unable to locate specimens of S. anthracina from British

Columbia in the CNC, and this absence has been confirmed by H.E.L. Maw (in litt., 22

Feb. 2018). However, there are specimens of S. provancheri from British Columbia in

the CNC, RBCM, and UBCZ collections, with some of the latter being recorded by

Downes (1927) as S. coriacea, a synonym of S. provancheri.

Hence, it is evident that the records of S. anthracina from British Columbia, the

Northwest Territories, and the Yukon in Maw et al. (2000) should be deleted. The

occurrence of 8. anthracina in Alaska needs to be confirmed.

Family ANTHOCORIDAE

Tetraphleps uniformis Parshley

Lattin (2006) has shown that 7: uniformis Parshley is a synonym of 7: canadensis

Provancher, and restored 7: furvus Van Duzee as a valid species in its place. Hence, 7.

uniformis should be deleted and replaced by 7. furvus Van Duzee.

Xylocoris umbrinus Van Duzee

Lattin (2005) has shown that _X. umbrinus Van Duzee is a synonym of X. californicus

(Reuter). Thus, X. umbrinus Van Duzee should be deleted and replaced by X. californicus

(Reuter).

Family NABIDAE

Pagasa fusca (Stein)

After Kerzhner (1993a) raised P. nigripes Harris to specific status and recorded this

species from Alberta, Saskatchewan, Quebec and Alaska, Scudder (2008) showed that all

the specimens of P. fusca (Stein) reported from the Yukon and the Northwest Territories,

and some specimens from British Columbia, were P. nigripes. Thus, P. fusca should be

deleted from the Yukon and Northwest Territories.

Family MIRIDAE

Adelphocoris superbus (Uhler)

Schwartz and Scudder (2003) concluded that A. superbus (Uhler) is a synonym of A.

rapidus (Say). Hence, A. superbus and all three provincial records should be deleted.

Agnocoris pulverulentus (Uhler)

This species was first reported from Alaska (Fort Yukon) by Moore (1955). Moore

(1956) did not list the Alaska (Fort Yukon) material when he recorded A. rubicundus

(Fallén) in the New World, but considered this latter species as Holarctic. Wheeler and

Henry (1992) also did not record A. rubicundus from Alaska, although they stated that

this species may have survived in an Alaska refugium. Maw ef al. (2000), while

recording A. pulverulentus in Alaska following Moore (1955), also noted A. rubicundus

28 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

from Alaska. This was based on specimens from Alaska in the CNC determined by M.D.

Schwartz as A. rubicundus. Included was material labelled ‘Alaska, Fort Yukon, 900',

4.vili.1951 (H.C. Severin)’. T.J. Henry informs me (in litt., 15 February 2018), that he

could not locate Alaska specimens of Agnocoris in the USNM, although Moore (1955)

recorded one male and three females as A. pulverulentus from Alaska, Fort Yukon, July

18, 1951 (R.I. Sailer) [USNM].

As a result, I hereby delete the record of A. pulverulentus from Alaska, assuming it is

in fact A. rubicundus.

Aoplonema uhleri (Van Duzee)

Forero (2008) has shown that Hadronema uhleri Van Duzee is a synonym of A.

princeps (Uhler). Aoplonema uhleri should be deleted and replaced by A. rubrum Forero.

Capsus ater (Linnaeus)

This species was reported from Alberta (Edmonton) by Blatchley (1926), quoted from

Alberta by Henry and Wheeler (1988), and reported by Maw et al. (2000). The record

was doubted by Wheeler and Henry (1992), and it was not included from Alberta in

Kelton (1980). All the specimens of Capsus that I have examined from Alberta are C.

cinctus (Kolenati). This latter species, recorded as C. simulans (Stal), was first reported

from Banff and Lethbridge in Alberta by Knight (1926) and subsequently were recorded

under this name from Alberta by Strickland (1953), MacNay (1953), and Kelton (1980).

Hence, it is assumed that the record of C. ater from Alberta should be deleted.

Vinokurov (1977) synonymized C. simulans (Stal) with C. cinctus (Kolenati) and noted

that this species occurred in North America from Alaska to Iowa in the United States.

Chlamydatus pullus (Reuter)

Many of the records formerly placed under C. pullus (Reuter) by Kelton (1965),

Scudder (1997), and Maw ef al. (2000) are now placed under the species C. keltoni

Schuh & Schwartz (Schuh and Schwartz 2005). Chlamydatus pullus (Reuter) as noted by

Schuh and Schwartz (2005) is found only in Quebec, Saskatchewan, and the Yukon. The

latter were recorded as “Chlamydatus sp. near auratus Kelton” by Scudder (1997).

The result is that all records of C. pullus in Canada, except those from Quebec,

Saskatchewan, and the Yukon, should be deleted.

Coquillettia insignis (Uhler)

Wyniger (2011) revised the genus Coquillettia Uhler and found that C. insignis Uhler

is confined to California. The species in Alberta and Saskatchewan was described as a

new species C. schwartzi Wyniger and the specimens from British Columbia as being

either of two new species, described as C. pergrandis Wyniger or C. thomasi Wyniger.

Hence, C. insignis Uhler, in Maw et al. (2000), should be deleted and replaced by the

species listed above.

Dacota hesperia Uhler

This species was recorded from British Columbia in Maw et al. (2000), based on a

single female specimen from B.C., Fraser, 29.vii.1982 (G.G.E. Scudder). This specimen

was subsequently determined in 2010 by M.D. Schwartz as Pinophylus rolfsi (Knight)

and recorded as AMNH_PBI00394201. Pinophylus rolfsi is now P. alpinus (Van Duzee),

according to Schwartz (2013).

Thus, the record of D. hesperia Uhler from British Columbia should be deleted.

Dicyphus vestitus Uhler

Dicyphus vestitus was recorded from British Columbia by Parshley (1919) and

Downes (1927). Parshley (1919) cited specimens from B.C., Saanich Dist., V.I., Apr. 30,

Sept. 14, 1918 (W. Downes), and Downes (1927) cited specimens from Goldstream,

Sept. 9, 1923 (K.F. Auden), Vernon, May 6", 1920 (H.R. Ruhmann), and Victoria, Sept.

7, 1920 (W. Downes).

Based on these records, D. vestitus was recorded from British Columbia by Henry

and Wheeler (1988), and this record was repeated by Maw et al. (2000).

In the UBCZ collection, which now contains the late W. Downes collection, there are

specimens of D. discrepans Knight that are labelled B.C., Saanich Dist., 14.1x.1918 (W.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 29

Downes) and B.C., Goldstream, 9.1x.1923 (K.F. Auden): these are evidently specimens

listed by Parshley (1919) and Downes (1927), respectively. Although Knight (1923)

described D. discrepans and distinguished it from D. vestitus, D. discrepans was not

listed by Downes (1927). It is evident that the early records of D. vestitus in Parshley

(1919) and Downes (1927) should be assigned to D. discrepans.

Hence, the D. vestitus Uhler record from British Columbia in Maw et al. (2000)

should be deleted. Henry (1999a) gives a recent key to D. discrepans and D. vestitus.

Lopidea confluenta (Say)

Maw et al. (2000) recorded L. confluenta (Say) from Alberta, Manitoba, Ontario, and

Quebec.

Lopidea confluenta is not recorded from Alberta by Strickland (1953) nor from the

prairie provinces by Kelton (1980). However, it is listed from Ontario and Quebec by

Asquith (1991) and Wheeler and Henry (1988). It was recorded from Manitoba (Aweme)

by Criddle (1921), and this was the basis for its inclusion in Scudder (2014).

It is evident that the Alberta record is an error and should be deleted. The Manitoba

record needs to be confirmed.

Lopidea nigridea serica Knight

This was reported from Alaska in Maw et al. (2000), based on one female specimen

from Tok, 22.vii.1982 (L.A. Kelton) [CNC]. However, M.D. Schwartz has since

determined that this specimen is L. dakota (Knight).

Hence, L. nigridea serica Knight should be deleted for Alaska, as noted by Scudder

and Sikes (2014).

Megalopsallus lycii (Knight)

Europiella lycii Knight 1968 was transferred to the genus Megalopsallus Knight by

Schuh et al. (1995) and synonymized with M. humeralis (Van Duzee) by Schuh (2000).

The latter species does not occur in Canada and should therefore be deleted. The Alberta

and Saskatchewan records under M. lycii in Maw et al. (2000) should be assigned to M.

sparsus (Van Duzee) (Schuh 2000).

Megalopsallus montanae (Knight)

Europiella montanae Knight 1968 was transferred to the genus Megalopsallus Knight

by Schuh ef al. (1995) and synonymized with M. nigrofemeratus (Knight) by Schuh

(2000). Hence, M. montanae (Knight) can be deleted.

Melanotrichus concolor (Kirschbaum)

This European species was reported from Quebec as Orthotylus concolor

(Kirschbaum) by Moore (1980), Larochelle (1984), and Roch (2008). This record as M.

concolor (Kirschbaum) was reported from Quebec by Henry and Wheeler (1988) and

repeated by Maw ef al. (2000).

However, Henry (1991) could not confirm the identity of Quebec specimens and

believed that they actually are M. virescens (Douglas & Scott). As a result, the record of

M. concolor from Quebec should be deleted and replaced by M. virescens.

Microphylellus elongatus Knight

Microphylellus elongatus Knight was cited as a synonym of Plagiognathus flavipes

(Provancher) by Schuh (2001) (see below). Hence, M. elongatus Knight should be

deleted.

Orectoderus salicis Knight

This species has been synonymized with O. montanus Knight by Wyniger (2010).

Hence, it can be deleted.

Orthotylus candidatus Van Duzee

Scudder (2008) reported that the earlier records of O. candidatus Van Duzee from

Ontario and Saskatchewan were referable to O. nyctalis Knight. Hence, the records of O.

candidatus Van Duzee from Ontario and Saskatchewan should be deleted.

Paradacerla downesi (Knight)

This species was recorded by Downes (1934) from B.C., Jordan Meadows on

Vancouver Island, at 1700 feet (W. Downes) det Downes. However, specimens from

30 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

Jordan Meadows are not in the late W. Downes collection donated to UBCZ in 1958 and

are not in the RBCM. Specimens from British Columbia were not listed in Kelton and

Knight (1959), and currently P downesi is unknown in British Columbia. Hence, P.

downesi should be deleted from the Canadian list.

Pilophorus clavatus (Linnaeus)

This European species was listed from Alberta, British Columbia, Manitoba, Nova

Scotia, Ontario, Quebec, and Saskatchewan by Henry and Wheeler (1988), and these

records were repeated in Maw eft al. (2000). Pilophorus clavatus was first reported from

Newfoundland in 2005 by Wheeler et al. (2006).

Downes (1927) recorded P. clavatus determined by H.H. Knight, from British

Columbia, Victoria, 17.1x.1924 (W. Downes) and Mission, 22.1x.1925 (W. Downes),

while Kelton (1980) noted this species from British Columbia, Alberta, Saskatchewan,

Manitoba, Ontario, and Nova Scotia. Pilophorus clavatus was recorded from Quebec by

Moore (1950) and Larochelle (1984), but not by Roch (2008), who queried the

occurrence in Ontario and New Brunswick.

Schuh and Schwartz (1988) noted that they were unable to confirm all the earlier

records of P. clavatus in Canada, except for the records from Manitoba and Nova Scotia.

Schuh and Schwartz (1988) considered that other records of P. clavatus in Canada could

either be P. neoclavatus Schuh & Schwartz or misidentified other species. These

comments were repeated by Wheeler and Henry (1992).

In the late W. Downes collection in the UBCZ, I found one female with the data,

B.C., Victoria, 17.1x.1924 (W. Downes), which is evidently the specimen recorded by

Downes (1927) from British Columbia. This specimen was determined by M.D.

Schwartz in 1998 as P. vicarius Poppius, so the British Columbia record of P. clavatus

should be deleted.

It would thus appear that all records of P. clavatus from Canada, except those for

Manitoba, Nova Scotia, and Newfoundland, should be deleted.

Pilophorus uhleri Knight

This species was recorded from Alberta, British Columbia, Manitoba, Ontario, and

Saskatchewan by Henry and Wheeler (1988), while Schuh and Schwartz (1988)

considered P. uhleri an eastern North American species, occurring west to Alberta. Schuh

and Schwartz (1988) gave records for Alberta, Manitoba, New Brunswick, Nova Scotia,

Prince Edward Island, and Saskatchewan, but not British Columbia.

Downes (1927) reported P. uhleri Knight, determined by H.H. Knight, from British

Columbia, Victoria, 15 Sept. 1924 (W. Downes), on Pinus contorta. This record was

accepted by Henry and Wheeler (1988) and Maw ef al. (2000).

However, a female specimen in the late W. Downes collection at UBCZ, with the

data, B.C., Victoria, Pinus contorta, 15.1x.1924 (W. Downes), is evidently the specimen

listed by Downes (1927). It was determined by M.D. Schwartz in 1998 as P. americanus

Poppius.

Hence, P. uhleri from British Columbia, should be deleted. It may be noted that

Schuh and Schwartz (1988) reported that P uhleri most closely resembles P. americanus.

Plagiognathus albatus vittiscutis Knight

Treated as P. albatus (Van Duzee) by Schuh (2001). Hence, P albatus vittiscutis

Knight can be deleted.

Plagiognathus albonotatus Knight

Synonymized with P. fuscosus (Provancher) by Schuh (2001). Hence, P. albonotatus

Knight should be deleted.

Plagiognathus caryae Knight

Synonymized with P. albatus (Van Duzee) by Schuh (2001). Hence, P. caryae Knight

should be deleted.

Plagiognathus cuneatus Knight

This variety of P. annulatus Uhler, established by Knight (1923), was synonymized

with P. obscurus Uhler by Schuh (2001). Hence, PR. cuneatus Knight should be deleted.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 3]

Plagiognathus fumidus Uhler

Considered a synonym of Europiella decolor (Uhler) by Schuh (2001). Hence, P

fumidus Uhler should be deleted. ?

Plagiognathus fusciflavus Knight

Synonymized with P. verticalis (Uhler) by Schuh (2001). Hence, P. fusciflavus Knight

should be deleted and Plagiognathus verticalis (Uhler) added to the B.C. listing.

Plagiognathus moerens (Reuter)

According to Schuh (2001), this species is not known to occur in Alberta and British

Columbia. Records for these two provinces should be transferred to P. shoshonea Knight.

Thus, the records of P. moerens (Reuter) for Alberta and British Columbia should be

deleted.

Plagiognathus nigritus Knight

Synonymized with P. brevirostris Knight by Schuh (2001). Hence, P. nigritus Knight

should be deleted and the Alberta record transferred to P. brevirostris Knight.

Plagiognathus obscurus albocuneatus Knight

Treated as P. obscurus Uhler by Schuh (2001). Thus, P obscurus albocuneatus

Knight should be deleted.

Plagiognathus politus flaveolus Knight

Treated as P. politus Uhler by Schuh (2001). Thus, P. politus flaveolus Knight should

be deleted.

Plagiognathus repletus Knight

Synonymized with P albatus (Van Duzee) by Schuh (2001). Hence, P. repletus

Knight should be deleted.

Plagiognathus similis Knight

Synonymized with P. albatus (Van Duzee) by Schuh (2001). Hence, P. similis Knight

should be deleted.

Psallus variabilis (Fallén)

Psallus variabilis (Fallén) was reported from Ontario by Blatchley (1926) and Henry

and Wheeler (1988), and this record was repeated in Maw et al. (2000). However,

Wheeler and Henry (1992) reported that this Ontario record was incorrect, Knight (1927)

having noted that early records of P. variabilis in North America were incorrect and that

specimens were misidentified. Knight (1927) said that these early records refer to

Lepidopsallus rubidus var atricolor Knight, which Wheeler and Henry (1992) call

Atractotomus atricolor (Knight). However, Stonedahl (1990) does not record A. atricolor

(Knight) from Ontario, although Stonedahl (1990) reported A. rubidus (Uhler) from

Ontario. Nevertheless, valid records for P. variabilis (Fallén) in North America were

given by Wheeler and Hoebeke (1982) and Wheeler and Henry (1992): these did not

include Ontario. Larochelle (1984) synonymized L. rubidus var atricolor Knight with L.

rubidus (Uhler).

It is evident that the record of P. variabilis (Fallén) from Ontario and Canada in Maw

et al. (2000) should be deleted.

Sixeonotus insignis Reuter

This species was recorded from Quebec by Larochelle (1984). However, Quebec

specimens from Knowlton, 4.v11.1929 (G.S. Walley), Knowlton, 8.vii.1929 (L.J. Milne),

and Otter Lake, 24.vii.1958 (L.A. Kelton) in the CNC have been determined by M.D.

Schwartz in 2000 as S. deflatus Knight. Hence, the Larochelle (1984) record from

Quebec probably refers to S. deflatus. Thus, the S. insignis Reuter record from Quebec

should be deleted.

Slaterocoris robustus (Uhler)

This species was recorded from Alberta in Maw et al. (2000), but it was not cited by

Strickland (1953), Kelton (1968, 1980), Henry and Wheeler (1988), or Schwartz (2011).

Evidently, this record for Alberta was a mistake and should be deleted. The record of S.

robustus (Uhler) from British Columbia was confirmed by Schwartz (2011).

Trigonotylus americanus Carvalho

32 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

In the original description of 7. americanus, in Carvalho and Wagner (1957),

paratypes were listed from British Columbia, Vernon, vi-i-47 (H.B. Leech). Based on

determinations by the late L.A. Kelton, 7’ americanus was recorded from Alaska (Hope)

and the Yukon by Scudder (1997), and so recorded in Maw et al. (2000). As noted by

Scudder and Sikes (2014), a male specimen with the data ‘Alaska, Hope, Kenai Pen.,

15.11.1951 (W.J. Brown)’ in the CNC has been determined by M.D. Schwartz as T. viridis

(Provancher). Hence, Scudder and Sikes (2014) stated that ZT) americanus Carvalho

should be removed from the list of Heteroptera from Alaska, because no other specimens

of the species are known from the state. :

Similarly, M.D. Schwartz has dissected males of the Yukon specimens listed as 7.

americanus by Scudder (1997) and found all of these to be 7: viridis (Provancher). Golub

(1989) resurrected 7: viridis (Provancher), which Kelton (1971) considered a synonym of

T: ruficornis (Geoffrey). All other specimens of TJrigonotylus Fieber from the Yukon

appear to be 7! viridis. Hence, the record of TZ. americanus Carvalho from the Yukon

should be deleted.

Trigonotylus tenuis Reuter

Henry and Wheeler (1988) reported T. doddi (Distant) from Alberta, Manitoba, and

Saskatchewan. Since Golub (1989) showed that ZT. doddi was a junior synonym of 7.

tenuis Reuter, Maw et al. (2000) reported the Henry and Wheeler (1988) records as T.

tenuis Reuter. However, Wheeler and Henry (1992) have noted that the original Henry

and Wheeler (1988) records undoubtedly refer to other species of 7rigonotylus Fieber.

Perhaps they refer to 7’ canadensis Kelton, described from Alberta, Manitoba, and

Saskatchewan by Kelton (1970).

Hence, it is reasonable to delete the record of 7. tenuis Reuter from the prairie

provinces and Canada. It is not included in Kelton (1980).

Family ARADIDAE

Aradus lugubris nigricornis Reuter

This taxon, treated as a subspecies by Froeschner (1988), was said by Parshley (1921)

not to be of geographic significance, because it occurs throughout the range of the

species A. /ugubris Fallén in North America. Hence, it should be deleted.

Family ORSILLIDAE

Nysius groenlandicus (Zetterstedt)

This species in North America was recorded from Alaska, Manitoba, Newfoundland,

Ontario, Prince Edward Island, and Quebec by Ashlock and Slater (1988). These records

were repeated in Maw et al. (2000), with the addition of the Yukon and Labrador. All of

these occurrence records were based on the published literature, although not all authors

cited the diagnostic characters used in their identification.

The published records were: Alaska (many cited by Slater 1964); Yukon (Scudder

1997); Manitoba (Churchill) (Barber 1947a, 1947b); Newfoundland (presumably Brown

1934); Ontario (Muskoka Lake District) (Van Duzee 1889); Prince Edward Island.

(Barber 1947a, 1947b); Quebec (Bradore Bay) (Brown 1934; Barber 1947a; Moore 1950;

Béique and Robert 1964; Larochelle 1984); Labrador (Nain) (Brown 1934).

Ashlock (1967) questioned whether N. groenlandicus (Zett.) occurred in North

America, and this was noted by Bocher (1976). Bocher (1978) observed that N.

groenlandicus seems to be absent in North America, and this was repeated by Danks

(1981).

At present, the record of N. groenlandicus from Prince Edward Island should be

deleted, although the identity of this material still must be determined.

Family RHYPAROCHROMIDAE

Perigenes constrictus (Say)

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 35

This species was reported from Alaska by Van Duzee (1919), and this record was

repeated by Slater (1964), Ashlock and Slater (1988), Maw et al. (2000), and Lattin

(2008). However, Scudder and Sikes (2014) noted that the female specimen in the CNC

from Ketchikan, on which the Alaska record is based, is actually a specimen of

Ligyrocoris sylvestris (Linnaeus). Hence, the record of P. constrictus (Say) from Alaska

should be deleted, as noted by Scudder and Sikes (2014).

Scolopostethus atlanticus Horvath

This species was reported from British Columbia, Manitoba, Ontario, Quebec, and

Newfoundland by Ashlock and Slater (1988), and these records were repeated in Maw et

al. (2000). These provincial records were evidently based on earlier reports, namely those

for Manitoba (Winnipeg) by Gibson (1912), for Ontario (Ottawa) by Gibson (1915), and

for Quebec by Béique and Robert (1964); Roch (2008) also reported S. atlanticus from

Ontario and Quebec. The records for Newfoundland were from Torre-Bueno (1917) and

Slater (1964), and Torre-Bueno (1946). However, it may be noted that neither Parshley

(1919) nor Downes (1927) gave records for S. diffidens Horvath.

Sweet (1964) gave a detailed description of the distinguishing characters of S.

atlanticus and considered this species an eastern Nearctic taxon. He thought that most of

the distribution records for S. atlanticus from the northern part of North America were

incorrect, and he specifically noted that the records for British Columbia in Parshley

(1919) referred to S. thomsoni Reuter. He also noted that the late H.G. Barber had

frequently mistakenly named specimens of S. thomsoni and S. diffidens in the USNM as

S. atlanticus.

I examined and photographed the male lectotype of S. atlanticus Horvath in Budapest

in February 1965 and have not seen similar material in all the numerous specimens of

Scolopostethus Fieber from Canada that I have examined over the past 60 years. In fact,

in the late W. Downes collection donated to UBCZ in 1958, there is one short-winged

female from B.C., Agassiz, 25.vi1.1921 (W. Downes). This is obviously the specimen

listed by Downes (1927), but it is S. diffidens. The same collection contains a

macropterous female from B.C., Enderby, 14.x.1920 (W. Downes). This was listed by

Downes (1927) as S. atlanticus, but is actually S. thomsoni. Furthermore, a short-winged

female from B.C., Colquitz, 4.1v.1919 (W. Downes), and a macropterous female from

B.C., Cowichan, 24.viii.1918 (W. Downes), both listed by Brown (1934) as S. atlanticus

and now in the late W. Downes collection at UBCZ, are in fact S. thomsoni.

Hence, I conclude that S. atlanticus should be deleted from the list of species in

Canada.

II. Nomenclature Changes

Family ANTHOCORIDAE

Orius minutus (Linnaeus)

Lewis and Lattin (2010) have noted that this introduced species in British Columbia

is actually O. vicinus Ribaut. Hence, this name should be replaced with O. vicinus.

Family NABIDAE

Kerzhner and Henry (2008) have rearranged the checklist of the Nabidae in North

America. This has resulted in a large number of nomenclatural changes. Nabicula Kirby

and Omanonabis Asquith & Lattin are treated as subgenera of Nabis Latreille, and

Anaptus Kerzhner is considered a subgenus of Himacerus Wolff. These changes result in

nine nomenclatural changes in the Nabidae as follows:

* —Anaptus major (Costa): change to Himacerus (Anaptus) major (Costa).

* Nabicula (Dolichonabis) americolimbata (Carayon): change to Nabis

(Dolichonabis) americolimbatus (Carayon).

° Nabicula (Dolichonabis) limbata (Dahlbom): change to WNabis

(Dolichonabis) limbatus Dahlbom.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Nabicula (Dolichonabis) nigrovittata nearctica Kerzhner: change to Nabis

(Dolichonabis) nigrovittatus nearctica (Kerzhner).

Nabicula (Limnonabis) propinqua (Reuter): change to Nabis (Limnonabis)

propinquus Reuter.

Nabicula (Nabicula) flavomarginata (Scholtz): change to Nabis (Nabicula)

flavomarginatus Scholtz.

Nabicula (Nabicula) subcoleoptrata Kirby: change Nabis (Nabicula)

subcoleoptratus (Kirby).

Nabicula (Nabicula) vanduzeei (Kirkaldy): change to Nabis (Nabicula)

vanduzeei (Kirkaldy).

Omanonabis lovetti (Harris): change to Nabis (Omanonabis) lovetti Harris.

Family MIRIDAE

Coniferocoris pinicolus (Coniferocoris Schwartz & Schuh)

This genus Coniferocoris Schwartz & Schuh was synonymized with Plesiodema

Reuter by Schwartz (2006). Thus, this species, listed in Maw et al. (2000) as C.

pinicolus, should be changed to Plesiodema pinicolus (Schwartz & Schuh).

Icodema nigrolineatum (Knight)

Henry (1999b) has shown that Plagiognathus nigrolineatum Knight should be placed

as the type species of a new genus that he named Americodema. Hence, the name J.

nigrolineatum (Knight) should be changed to Americodema nigrolineatum (Knight).

Genus Lygocoris, subgenus Neolygus Knight

Neolygus Knight was raised to generic status by Yasunaga ef al. (2002). This results

in 29 name changes as listed below:

Lygocoris alni (Knight): change to Neolygus alni (Knight).

Lygocoris atricallus Kelton: change to Neolygus atricallus (Kelton).

Lygocoris atritylus (Knight): change to Neolygus atritylus (Knight).

Lygocoris belfragii (Reuter): change to Neolygus belfragii (Reuter).

Lygocoris caryae (Knight): change to Neolygus caryae (Knight).

Lygocoris clavigenitalis (Knight): change to Neolygus clavigenitalis

(Knight).

Lygocoris communis (Knight): change to Neolygus communis (Knight).

Lygocoris contaminatus (Fallén): change to Neolygus contaminatus (Fallén).

Lygocoris fagi (Knight): change to Neolygus fagi (Knight).

Lygocoris geneseensis (Knight): change to Neolygus geneseensis (Knight).

Lygocoris hirticulus (Van Duzee): change to Neolygus hirticulus (Van

Duzee).

Lygocoris inconspicuus (Knight): change to Neolygus inconspicuus

(Knight).

Lygocoris invitus (Say): change to Neolygus invitus (Say).

Lygocoris johnsoni (Knight): change to Neolygus johnsoni (Knight).

Lygocoris knighti Kelton: change to Neolygus knighti (Kelton).

Lygocoris laureae (Knight): change to Neolygus laureae (Knight).

Lygocoris omnivagus (Knight): change to Neolygus omnivagus (Knight).

Lygocoris ostryae (Knight): change to Neolygus ostryae (Knight).

Lygocoris parrotti (Knight): change to Neolygus parrotti (Knight). _

Lygocoris parshleyi (Knight): change to Neolygus parshleyi (Knight).

Lygocoris piceicola (Kelton): change to Neolygus piceicola (Kelton).

Lygocoris quercalbae (Knight): change to Neolygus quercalbae (Knight).

Lygocoris semivittatus (Knight): change to Neolygus semivittatus (Knight).

Lygocoris univittatus (Knight): change to Neolygus univittatus (Knight).

Lygocoris viburni (Knight): change to Neolygus viburni (Knight).

Lygocoris vitticollis (Reuter): change to Neolygus vitticollis (Reuter).

Lygocoris walleyi (Kelton): change to Neolygus walleyi (Kelton).

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 36

Melanotrichus elongatus Kelton

Orthotylus leonardi was proposed by Kerzhner and Schuh (1995) for O. elongatus

(Kelton 1980), M. elongatus Kelton, a junior secondary homonym of O. elongatus

Wagner 1965.

It would seem that the name M. leonardi (Kerzhner & Schuh) should replace M.

elongatus Kelton.

Microphylellus adustus binotatus Knight

This species was synonymized with Reuteroscopus falcatus Van Duzee by Schuh

(2001). However, R. falcatus Van Duzee was made the type species of the new genus

Vanduzeephylus by Schuh and Schwartz (2004). Hence, the name M. adustus binotatus

Knight should be replaced by Vanduzeephylus falcatus (Van Duzee).

Microphylellus flavipes (Provancher)

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it should now be called Plagiognathus flavipes (Provancher).

Microphylellus longirostris (Knight)

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it should now be called Plagiognathus longirostris (Knight).

Microphylellus maculipennis Knight

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it is now called Plagiognathus maculipennis (Knight).

Microphylellus modestus Reuter

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it is now called Plagiognathus modestus (Reuter).

Microphylellus tsugae Knight

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, this is now Plagiognathus tsugae (Knight).

Microphylellus tumidifrons Knight

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it is now Plagiognathus tumidifrons (Knight).

Parapsallus vitellinus (Scholtz)

This introduced species was transferred to the genus Plagiognathus Fieber by Schuh

(2001). Hence, it should be called Plagiognathus vitellinus (Scholtz).

Pinophylus rolfsi (Knight)

This is now P. alpinus (Van Duzee) according to Schwartz (2013), as noted under

Dacota hesperia Uhler above. Hence, a nomenclature change is necessary.

Platylygus Van Duzee

Pappus Distant has been shown to be the senior synonym of Platylygus Van Duzee by

Henry Renne). Thus, all five species of Platylygus should be transferred to Pappus:

Platylygus luridus (Reuter): change to Pappus luridus (Reuter).

Platylygus piceicola Kelton: change to Pappus piceicola (Kelton).

Platylygus pseudotsugae Kelton: change to Pappus pseudotsugae (Kelton).

Platylygus rolfsi Knight: change to Pappus rolfsi (Knight).

Platylygus rubripes Knight: change to Pappus rubripes (Knight).

Plesiodema sericeum (Heidemann)

Plesiodema sericeum Heidemann has been placed as the type species of the new

genus /zyaius by Schwartz (2006). Hence, the name P. sericeum should be changed to

Izyaius sericeum (Heidemann).

Psallus alnicenatus Knight

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it should now be called Plagiognathus alnicenatus (Knight).

Psallus morrisoni Knight

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it should now be called Plagiognathus morrisoni (Knight).

Psallus parshleyi Knight

36 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it should now be called Plagiognathus parshleyi (Knight).

Psallus physocarpi Henry

This species has been transferred to the genus Plagiognathus Fieber by Schuh (2001).

Hence, it should now be called Plagiognathus physocarpi (Henry).

Sthenarus cuneotinctus Van Duzee

Schuh and Schwartz (2004) have made this species the type of the new genus

Aurantiocoris. Hence, the species should now be cited as Aurantiocoris cuneotinctus

(Van Duzee).

Teleorhinus brindleyi Knight

This species was synonymized with 7: cyaneus Uhler by Wyniger (2010). Hence, the

name should be changed to T. cyaneus Uhler.

Family TINGIDAE

Dictyonota tricornis (Schrank)

The genus Kalama Puton was recognized by Péricart (1982), with Kalama tricornis

(Schrank) being one of the included species (Froeschner 2001). This latter species was

recorded as introduced into Canada and the United States by Drake and Ruhoff (1965)

under the name D. (Alcletha) tricornis (Schrank), a fact reiterated by Froeschner (2001).

Hence, this tingid should now be recorded as an introduction under the name K. tricornis

(Schrank).

Family OXYCARENIDAE

Crophius ramosus Barber

Henry et al. (2015) resurrected the genus Mayana Distant and cited Mayana ramosa

(Barber) as one of the included species. Hence, C. ramosus Barber should be changed to

M. ramosa (Barber).

Family PIESMATIDAE

Piesma cinereum (Say)

Péricart (1974) made Tingis cinerea Say the type species of a new subgenus that he

named Parapiesma, and Parapiesma Péricart was raised to generic status by Heiss and

Péricart (1997). Hence, Parapiesma cinereum (Say) is the current name for what was

previously called Piesma cinereum (Say).

Piesma explanatum McAtee

This piesmatid was included in the subgenus Parapiesma by Péricart (1974). Since

Parapiesma Péricart was raised to generic status by Heiss and Péricart (1997), the current

name for this taxon, previously called Piesma explanatum McAtee, is Parapiesma

explanatum (McAtee).

Family PENTATOMIDAE

Genus Acrosternum, subgenus Chinavia Orian

Chinavia Orian was treated as a distinct genus by Ahmad ef al. (1996). This results in

the following changes:

e A. hilare (Say): change to C. hilaris (Say).

e A. pensylvanicum (Gmelin): change to C. pensylvanica (Gmelin).

Apateticus bracteatus (Fitch)

Thomas (1992) recognized the genus Apoecilus Stal separate from Apateticus Dallas

and keyed Apoecilus bracteatus Fitch. This is the name that should be recognized for this

species.

Apateticus cynicus (Say)

This species should now be called Apoecilus cynicus (Say), as noted above.

Codophila remota (Horvath)

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 37

Kerzhner (1993b) and Rider (1998) have noted that the correct name for this taxon is

Anthemia eurynota remota (Horvath).

Cosmopepla bimaculata (Thomas)

Rider and Rolston (1995) have noted that the correct name for the species is C.

lintneriana Kirkaldy.

Holcostethus piceus (Dallas)

Rider and Rolston (1995) proposed the new name H. macdonaldi for this species.

III. Spelling Errors

Family CORIXIDAE

Hesperocorixa harrisi (Uhler)

Should be Hesperocorixa harrisii (Uhler), according to Jansson (2002).

Hesperocorixa kennicotti (Uhler)

Should be Hesperocorixa kennicottii (Uhler), according to Jansson (2002).

Family MIRIDAE

Actinocoris Reuter

This should be spelt Actitocoris Reuter.

Atractotomus cerocarpi Knight

This should be spelt Atractotomus cercocarpi Knight.

Closterotomus norvegicus (Gmelin)

This should be spelt Closterotomus norwegicus (Gmelin), according to Kerzhner and

Josifor (1999).

Family TINGIDAE

Alveotingis grossocerata Osborne & Drake

Should be A/veotingis grossocerata Osborn & Drake.

Family LYGAEIDAE

Melanopleurus pyrropterus (Stal)

This should be spelt Melanopleurus pyrrhopterus (Stal).

IV. Other Comments

Family ANTHOCORIDAE

Anthocoris tomentosus Péricart

Lewis and Horton (2012) have shown that many of the occurrence records listed from

A. tomentosus from the Yukon by Scudder (1997) are a new species that was described as

A. aquilivenis Lewis. Lewis and Horton (2012) also gave records of A. aquilivenis for

Alaska and British Columbia that had previously been determined as A. tomentosus.

However, A. tomentosus still has valid occurrence records from Alaska, Yukon and

British Columbia.

Orius diespeter Herring

_The Yukon records for O. diespeter Herring given in Scudder (1997) are in fact the

species O. sibericus Wagner (Lewis et al. 2015). Hence, the Yukon record for O.

diespeter in Scudder (1997) should be deleted and replaced by O. sibericus. However, O.

diespeter Herring does occur in the Yukon (Lewis and Horton 2010), although it is

recorded as O. tristicolor (White) by Scudder (1997) (see below).

Orius tristicolor (White)

Lewis and Horton (2010) have shown that all records from the Yukon listed by

Scudder (1997) as O. tristicolor are in fact a colour variation of O. diespeter Herring.

Lewis and Horton (2010) also suggest that all records of O. tristicolor in eastern Canada

38 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

actually refer to O. diespeter. Hence, O. tristicolor is deleted for Saskatchewan to

Newfoundland, and replaced by O. diespeter.

Lewis and Horton (2010) updated the known distribution of O. diespeter to include

Alberta, British Columbia, Nova Scotia, Ontario, Quebec, the Yukon, and Alaska: O.

tristicolor was recorded from Alberta and British Columbia but not Alaska.

Lewis informed me on January 30, 2018 (in litt.) that a male specimen in the UBCZ

collection, number ANTHO0784, with data “Firth R., 69°08'N 140°14'W, 23.v1.1984 (S.G.

Cannings)” that she determined in 2010 is in fact O. tristicolor. This was originally

reported in Lewis and Horton (2010) as O. diespeter. However, this has been clarified

since by Lewis (in litt., 23 February 2018). Hence, O. tristicolor is still recorded from the

Yukon but not Alaska.

Family MIRIDAE

Tupiocoris agilis (Uhler)

Tupiocoris agilis was first reported from British Columbia by Parshley (1919) as

Dicyphus agilis Uhler with records for Saanich Dist., V.I., Apr. 30, Sept. 14, 1918 (W.

Downes). It was also reported from British Columbia by Downes (1927) under the same

name, with specimens recorded from Agassiz, Sept. 1921 (R. Glendenning), Duncan,

Aug. 4, 1921 (W. Downes) and Saanich, June 18, 1918 (W. Downes). Kelton (1980)

writing under D. confusus Kelton, concluded that the early records of what is now 7:

agilis (Uhler) probably refer to what is now 7: confusus (Kelton), 7: similis (Kelton), or

some other species.

However, new records of 7: agilis (Uhler) for British Columbia were published by

Schwartz and Scudder (2001). Although I have been unable to trace the earlier specimen

listed by Parshley (1919) and Downes (1927), these can be ignored, as the recent records

by Schwartz and Scudder (2001) validate the species in British Columbia.

ACKNOWLEDGEMENTS

I thank Dr. M.D. Schwartz for accurately naming some of the specimens listed, and

H.E.L. Maw for checking data on specimens in the CNC. Claudia Copley (RBCM),

Derek Sikes (UAM) and T.J. Henry (USNM) kindly checked collections for me. Dr. D.

Rider (North Dakota State University) provided information on the pentatomid genus

Chinavia Orian.

R.G. Foottit (CNC) and H.E.L. Maw (CNC) made useful suggestions on the

manuscript, which was prepared by Launi Lucas (UBC).

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44 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

The bees of British Columbia (Hymenoptera: Apoidea,

Apiformes)

C. S. SHEFFIELD! AND J. M. HERON?

ABSTRACT

British Columbia is the most biologically diverse province in Canada, and its wide

range of landscapes — particularly the dry valley bottoms and basins of the Columbia,

Kootenay, Okanagan, Kettle, and Similkameen River systems — make it ideal for many

groups of Hymenoptera, including bees. With the exceptions of some generic- or

family-level treatments, no comprehensive account of the bees of British Columbia

has been published, although recent studies have indicated that more than half of

Canada’s bee species may be found in the province, with many of these found

nowhere else in the country.

Here, we summarize the province’s bee fauna by providing a comprehensive annotated

checklist of species. For each species, we indicate the ecozone(s) in which they are

presentently known to occur, and we provide summary statistics and analyses to

compare ecozones. We also summarize the growth in knowledge of the province’s bee

species over time, and all species accounts for the province are accompanied by a list

of supporting literature or data. Although we feel this list is comprehensive, it is likely

that we have overlooked some published accounts, and additional undocumented

species will show up.

In total, we record 483 bee species from British Columbia, 37 of which are considered

new to the province. Among these, 20 species (or subspecies) are recorded as new to

Canada, including: Andrena (Euandrena) misella Timberlake, Panurginus

cressoniellus Cockerell [Andrenidae], Lasioglossum (Dialictus) obnubilum

(Sandhouse), L. (Evylaeus) argemonis (Cockerell), L. (Hemihalictus) glabriventre

(Crawford), L. (Hemihalictus) kincaidii (Cockerell) [Halictidae], Osmia (Melanosmia)

laeta Sandhouse, O. (Melanosmia) malina Cockerell, O. (Melanosmia) pulsatillae

Cockerell, O. (Melanosmia) raritatis Michener, Anthidium (Anthidium) formosum

Cresson, Dianthidium (Dianthidium) plenum plenum Timberlake, D. (Dianthidium)

singulare (Cresson), Stelis (Stelis) ashmeadiellae Timberlake, S. (Stelis) calliphorina

(Cockerell), Dioxys pomonae pomonae Cockerell, Megachile pugnata pomonae

Cockerell [Megachilidae], Nomada crotchii Cresson, Melissodes (Eumelissodes)

saponellus Cockerell, and Habropoda miserabilis (Cresson) [Apidae].

Key words: Andrenidae, Apidae, Colletidae, Halictidae, Megachilidae, Melittidae,

diversity

INTRODUCTION

British Columbia is a vast landscape with variable topography, geology, and climate

that enable the largest total biodiversity of any province or territory in the country

(Cannings and Cannings 2015; Canadian Endangered Species Conservation Council

2016). Approximately 80,000 species are estimated to live in Canada (Canadian

Endangered Species Conservation Council 2016), with more than 50,000 species

occurring in British Columbia alone (Cannings and Cannings 2015). However, precise

knowledge comes only from fully documenting the species that have been recorded via

faunal checklists. In addition to providing data for increasing faunistic knowledge,

species checklists provide important baselines for assigning a species’ conservation status

and enabling the prioritization of habitat protection, management, conservation and land

| Corresponding author: Royal Saskatchewan Museum, 2340 Albert Street, Regina, Saskatchewan S4P

2V7; cory.sheffield@gov.sk.ca

2 British Columbia Ministry of Environment and Climate Change Strategy, Vancouver, British Columbia

V6T 1Z]1

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 45

use decisions. For many invertebrate groups, species checklists do not exist or are

incomplete, although the completion of Wild Species 2015 (Canadian Endangered

Species Conservation Council 2016) has enabled a better understanding of the provincial

and territorial diversity across the country for many taxa, including in British Columbia.

In the last decade pollinators, particularly bees, have come to the forefront of

conservation importance due to their integral link to pollination, food supply and overall

ecosystem health. A key component to assessing the conservation status of bee

communities begins with understanding the species present, their respective range

extents, and potential habitat associations according to the ecosystem mapping

throughout the species’ range. The range extents for many bee species recorded from

British Columbia are unclear, and more inventories are needed to better define their

limits (Heron and Sheffield 2015). The inventory for bee species in British Columbia is

incomplete, and most past efforts to compile species lists have focused on documenting a

narrow range of taxa (e.g., Buckell 1949, 1950, 1951; Cannings 2011 - Bombus), or have

not been comprehensive (e.g., Viereck et al. 1904a—d, 1905a, b, 1906). More recently,

studies providing species information have been ecological in nature and have focussed

within a limited geography (e.g., Elwell et al. 2016). However, bee diversity estimates

for British Columbia have been treated in a more general sense: the province is known to

have the highest bee diversity in Canada, with estimates ranging from 369 (Sheffield ef

al. 2014) to possibly more than 600 species (Sheffield et a/. 2017), the latter estimate

being based on DNA barcoding results.

Numerous factors likely contribute to this high biodiversity. For example, bees are

closely associated with plant diversity and habitat type. Approximately 2,500 native

vascular plants have been recorded from British Columbia (Douglas et a/. 2002; British

Columbia Conservation Data Centre 2018), some of which are part of rare ecosystems

and plant communities unique to Canada (Straley et a/. 1985). The southern part of the

province is also the northernmost extension of numerous unique southern ecosystems,

allowing numerous bee species to range into these same areas. Many of these bee species

are solitary and depend on specific soil and climate variables that define or seemingly

restrict their range (Sheffield et a/. 2014); the Western Interior Basin for example, though

by far the smallest ecozone in Canada, contains a significant number of the country’s bee

species, some of which occur nowhere else in Canada (Sheffield et a/. 2014). Though no

comprehensive checklist of British Columbia bee species has been previously completed

(although see Sheffield and Heron 2017), some components of the province’s bee fauna

were covered, as indicated above. In addition, Tepedino and Griswold (1995) provided a

list of species for the Columbia Basin, which included some specimens from British

Columbia.

Our objective here is to provide the first published, comprehensive list of the bees of

British Columbia, correcting, updating, and validating occurrence data in lists previously

provided to the Canadian Endangered Species Conservation Council (2016) and E-Fauna

(Sheffield and Heron 2017). Species occurences in the province are fully documented

with references to literature, and links to datasets are provided. This project also

contributes to the overall knowledge of apoid wasps in the province; the Spheciformes

treated recently by Ratzlaff (2015) and Ratzlaff et al. (2016) and all studies building on

the provincial summary of Apoidea provided by Cannings and Scudder (2001).

MATERIALS AND METHODS

Most of the data presented here were compiled from published literature, ranging

from published taxonomic treatments, species lists, ecological studies, and unpublished

graduate theses. In addition, data were also mined from websites and non-peer-reviewed

or unpublished studies (i.e., grey literature) and verified with specimen or photographic

evidence. Our list builds on previous faunistic work that has focused on northwestern

North American, including British Columbia (Viereck et al. 1904a—d, 1905a, b; 1906),

46 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

and later works specific to the province (Buckell 1949, 1950, 1951), much of which was

compiled for the Wild Species 2015 national assessment (Canadian Endangered Species

Conservation Council 2016). In cases where records for “BC” were recorded in the

literature (e.g., Hurd 1979) without accompanying data, the “species x British Columbia”

were entered as search terms in Biodiversity Heritage Library (https://

www.biodiversitylibrary.org/); however, in a few instances, no supporting literature/data

could be found. References and notes supporting the presence of each species in the

province are given next to each taxon in Supplemental Material.

Data were also compiled from many past and more recent collection efforts in the

province, including studies being conducted by the British Columbia Ministry of

Environment and Climate Change Strategy (JMH), Royal Saskatchewan Museum (CSS),

and past studies conducted out of York University (Toronto, ON). Much of this recent

material was used in the Barcodes of Life campaign for the bees of Canada (Sheffield et

al. 2017). In addition, many specimens were examined from the Royal British Columbia

Museum (Victoria, BC), the Spencer Entomology Museum, University of British

Columbia (Vancouver, BC), the Royal Saskatchewan Museum (Regina, SK), York

University (Toronto, ON), and the Canadian National Collection of Insects, Arachnids,

and Nematodes (Ottawa, ON). The complete species list has been added to Canadensys

(http://www.canadensys.net/) at https://doi.org/10.5886/NKZFXC, and has been

registered with GBIF [assigned the following GBIF UUID: 7b944cc6-1ffa-49de-

aab8-2a5ab543422b]. Occurrence data from species recorded as new to the province and/

or country have also been added to Canadensys [https://doi.org/10.5886/INGA8Z] and is

also registered with GBIF [GBIF UUID: f9c49aed-ba4b-454e-b88a-cbe1 fff5b2b6]. An

updated version of the list will also be maintained at the Bees of Canada website: http://

www.beesofcanada.com/home.

Although some of the literature sources examined (e.g., Mitchell 1960, 1962; Hurd

1979; Cannings 2011) list a species as only occurring in the province, we specifically

tried to mine data that would provide geographic information to allow us to assign each

species to the Canadian ecozones represented in the province (see Ecological

Stratification Working Group 1995; Environment and Climate Change Canada 2016).

Canada’s terrestrial land base is classified into 15 ecozones that are part of a broad

ecological framework for North America (Ecological Stratification Working Group 1995;

Wilken et al. 1996; Commission for Environmental Cooperation 1997) that classify a

geographic area of the country with similar physiography, hydrology, climate, wildlife

potential and vegetation. The attributes of each ecozone promote classification based on

unique assemblages of plant and animal communites based on climate zones and soils.

The ecozones in which each bee species occurs provides additional ecological

information that may provide conservation value. The six Canadian ecozones represented

in British Columbia are the Pacific Maritime [PacM], Western Interior Basin [WIB],

Montane Cordillera [MonC], Boreal Plains [BorPl], Boreal Cordillera [BorC], and Taiga

Plains [TaiP1]. Information on each ecozone in British Columbia is summarized from the

references above.

The Pacific Maritime [PacM] ecozone has an area of 195,000 km’, and occurs along

the west coast (including coastal islands) from the United States (Washington) border in

the south, northwards to the Alaska Panhandle. This ecozone is the wettest in the Canada,

with extensive areas of temperate old-growth coniferous forests (1.e., western redcedar

(Thuja plicata Donn ex D. Don), yellow-cedar (Cupressus nootkatensis D. Don), western

hemlock (7suga heterophyla (Raf.) Sarg.), mountain hemlock (7suga mertensiana

(Bong.) Carr.), Douglas-fir (Pseudotsuga menziesii (Mirb.) Franco), Pacific silver fir

(Abies amabilis Douglas ex J. Forbes), and Sitka spruce (Picea sitchensis (Bong.) Carr.),

with high mountains with alpine tundra and glacial, and lowland estuary and valley-

bottom floodplain habitats (Fig. 1). It contains numerous rare and endangered

ecosystems, including Garry Oak (Quercus garryana Douglas ex Hook.) and associated

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 47

ecosystems, sparsely vegetated coastal sand ecosystems, bog and wetland habitats, and

the lowland riparian forests of the Fraser Valley.

Figure 1. Pacific Maritime [PacM] ecozone. A) subalpine coastal coniferous forests, Greig

Ridge, Strathcona Provincial Park. Photo J. Heron; B) coastal sand ecosystem on south side of

Savary Island. Photo J. Heron.

The Western Interior Basin [WIB, also called the Semi-Arid Plateau] is the smallest

ecozone in Canada (previously classification considered this ecozone part of the Montane

Cordillera), and all 56,466 km/? are restricted to the south—central part of the province.

The boundary of this ecozone is comparable to the Southern Interior Ecoprovince of the

province’s Ecoregion Classification System. The ecozone (Fig. 2) represents the

northernmost extension of the Great Basin Sagebrush Desert Biome that stretches from

British Columbia through the Midwestern United States to Mexico. Approximately 2% of

the land area of this ecozone is classified as native grasslands and 73% as forests. There

are a number of species at risk that are confined to the WIB and, more specifically, to the

low-elevation plant communities of this ecozone. The cumulative effects from multiple

threats, such as natural habitat conversion, fragmentation, recreational use and invasive

species, have led to these species being at risk. In particular, the antelope-brush

48 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

(Purschia tridentata (Pursh) DC.) plant communities in the south Okanagan Valley have

significantly declined in quality and spatial area since the 1800s (Schluter et al. 1995;

Lea 2001, 2008; Iverson and Haney 2012; Iverson 2012). More specifically, the antelope-

brush/needle-and-thread Grass plant community has declined from 9,863 ha in 1800 to

3,217 ha in 2009, a loss of 67.4% of the original extent of this ecosystem (Iverson 2012).

More broadly across the WIB, approximately 16% of grasslands (1188km/7) have been

converted to urban and agricultural development since 1850 (Wikeem and Wikeem 2004;

Grasslands Conservation Council of British Columbia 2004; B.C. Ministry of

Environment 2007). Habitat loss continues with high development pressure on

undesignated provincial Crown land and natural areas into housing, commercial and

agricultural use. Livestock overgrazing is also a threat within provincial Crown lands —

both grassland and forested areas.

Figure 2. Western Interior Basin [WIB] ecozone. A) lower Okanagan Valley. Photo C.S.

Sheffield; B) antelope-brush plant community at Osoyoos Desert Centre, west of Osoyoos.

Photo J. Heron.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 49

The British Columbia portion of the Montane Cordillera [MonC] ecozone — about

90% of the total for Canada (Scudder and Smith 2011) — comprises 389,000 km/?. It

covers the eastern portion of the province and spans the Rocky Mountains into western

Alberta. The ecozone ranges from the United States border to the Skeena Mountains in

north-central British Columbia, and includes a broad range of ecosystems, from dense

conifer forests to alpine tundra, grasslands, and rugged mountains (Fig. 3): it is likely the

most complex ecozone in the province (Scudder and Smith 2011). Approximately 70% of

the area is forested, 27% is non-forested, and 3% is water (Scudder and Smith 2011). The

climate is characterized by wet winters and dry summers, with mild climate overall

throughout the year. The Kootenay region of the province includes the western slopes of

the Rocky Mountains, small portions of arid sagebrush and grasslands, fir and cedar

forests, large rivers, and numerous valleys that extend southwards into the United States

and bring a number of species to their northernmost limits.

ow Ze S 3

Figure 3. Montane Cordillera [MonC] ecozone. A) view south from Cristina Creek. Photo J.

Heron; B) Flathead Valley, east of Fernie. Photo J. Heron.

50 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Approximately 5% of the Boreal Plains [BorPl] ecozone occurs in the province

(37,940 km) and exists across a small portion of north-eastern British Columbia (Fig. 4).

More than half of this ecozone (60%) comprises forests and, in British Columbia, the

ecozone’s other habitats are shrublands and wetlands, as well as native grasslands that

have been converted to agricultural areas. Forests grow slowly in the Boreal Plains due to

low-nutrient and poorly drained soils and discontinuous permafrost (ESTR Secretariat

2014).

Figure 4. The Boreal Plains [BorP1] ecozone. A) along the Peace River west of Fort St. John

enar Hudson’s Hope. Photo J. Heron; B) at Pink Mountain, looking northwest at the Rocky

Mountains. Photo S. Cannings.

The portion of the Boreal Cordillera [BorC] in British Columbia spans a large portion

of the northern half of the province, and stretches into the Yukon. The BorC ecozone

(Fig. 5) covers 189,000 km? and is dominated by forests of black spruce [Picea mariana

(Mill.) Britton, Sterns & Poggenburg] and white spruce [P glauca (Moench) Voss],

lodgepole pine (Pinus contorta Douglas), trembling aspen (Populus tremuloides Michx.),

balsam poplar (P. balsamifera L.) and white birch (Betula papyrifera Marshall), with

higher-elevation areas of subalpine-fir [Abies lasiocarpa (Hooker) Nuttall].

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 on

Less than 10% of the Taiga Plains [TaiPl] ecozone occurs in British Columbia —

approximately 70,000 km*. Much of this ecozone (Fig. 6) is boreal spruce forest (68%),

wetland, and peatland habitats, with extensive shrub cover (20%). The ecozone also

contains some elements of subarctic habitats (ESTR Secretariat 2013).

Figure 5. Boreal Cordillera [BorC] ecozone. A) alpine country east of Atlin. Photo S.

Cannings; B) North Tetsa River, Stone Mountain Provincial Park. Photo S. Cannings.

The bee fauna of the ecozones in British Columbia were compared both by tallying

the species known to occur in each and based on the number of species per 1000 km/;

this latter calculation was done to highlight the diversity of bee species based on the size

of each ecozone specifically to draw attention to bee biodiversity hot spots and areas of

high conservation value. In addition, a presence/absence matrix of bee species by

ecozone was created, and a single link cluster analysis of incidence-based similarity (i.e.,

Jaccard’s index) was performed using Biodiversity Pro (McAleece et al. 1997) to explore

faunistic similarity of the ecozones occurring in the province.

32 J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

Figure 6. Taiga Plains [TaiPl] -ecozone. AD at Fort Nelson, looking west to the Rocky

Mountains. Photo S. Cannings; B) Grayling River Hot Spring. Photo C.S. Sheffield.

RESULTS AND DISCUSSION

The first published record of a bee in British Columbia was that of Smith (1861), who

described the cuckoo bumble bee, Apathus (=Bombus) insularis (Smith), from the

province. Knowledge of the bee fauna of British Columbia has increased dramatically

over the last ca. 160 years, with several major published contributions adding greatly to

the list of species recorded for the province throughout this period (Fig. 7). The most

significant years of contributions (1.e., additions of 20 or more species per year resulting

from a single published study or series of related published studies) occurred in the early

1900s (Viereck et al. 1904a-d, 1905a, b, 1906; see “A” on Fig. 7), 1924 (Criddle et al.

1924; “B” on Figure 7), and 1925 (Sandhouse 1925a, b; “C” on Fig. 7). Yearly increases

did not exceed 20 species again until 2010 (Gibbs 2010; “D” on Fig. 7). More recently,

Elwell et al. (2016) added another 20 species to the provincial list (“E” on Fig. 7). In the

present study, we add an additional 37 species (“F” on Fig. 7), 20 of which are new for

Canada, for a cumulative provincial count of 483 bee species (Fig. 7). The Megachilidae

is the family most well represented, with more than 150 species found in the province,

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 aa

followed by Andrenidae (largely the genus Andrena Fabricius) and Apidae, both with

more than 100 species, and Halictidae (Fig. 8).

ome buns

& Mia SOLE

Number of Species

Cumulative Number of Species

& Be

‘ees

Figure 7. The year by year addition and cumulative total of bee species in British Columbia

based on published literature records and other data from 1861 to present (see links to data

sets above). Black bars show the number of new species records for each year (1.e., based on

the earliest recorded occurrence in the province, see Supplemental Material) (left axis); red

line shows the cumulative number of species (right axis) based on these additions. Letters

adjacent to bars represent published studies where 20 or more species were added as the result

of one publication or group of related publications. A=Viereck et al. 1904a-d (32 species) +

Vachal 1904 (6 species); B=Criddle et al. 1924 (56 species) + Sandhouse 1924 (4 species) +

Viereck 1924 (11 species); C=Sandhouse 1925a-b (20 species); D=Gibbs 2010 (27 species) +

Rightmyer 2010 (1 species); E=Elwell et al. 2016 (20 species); F=present study (37 species).

The rapid growth in species numbers observed in the past 10 years has largely been

facilitated through surveys (Heron and Sheffield 2015; Elwell et a/. 2016), taxonomic

revisions (Gibbs 2010; Sheffield et a/. 2011), and DNA barcoding (Sheffield et a/. 2017),

with a large proportion of species occurring in British Columbia having sequences in the

Barcodes of Life Data system (BOLD) with specimens contributed by the Royal

Saskatchewan Museum, York University, Simon Fraser University, and the Royal British

Columbia Museum, and collecting efforts of the authors and associated researchers at

these institutions. These efforts have verified many previous records of others (see

Supplemental Material) and have added new records to the province (Gibbs 2010; Heron

and Sheffield 2015; Elwell et al. 2016). The DNA barcoding efforts have also highlighted

the fact that there is still much taxonomic work to do with the British Columbia bee

fauna, especially with the cleptoparasitic genera Sphecodes (Halictidae) and Nomada

(Apidae) (Sheffield et al. 2017). A recent estimate (Sheffield et a/. 2017) suggests there

could be upwards of 600 species in the province — almost three-quarters of the total for

54 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

Canada — with the vast majority of these found in the WIB ecozone (Fig. 9). This is

supported by previous estimates of bee diversity in the Columbia Basin in the adjacent

USA, which suggests almost 650 species (Mayer eft al. 2000; Niwa et al. 2001), with

estimates as high as 1,000 species (Tepedino and Griswold 1995).

103

Number of Species

Melittidae Andrenidae Halictidae Colletidae | Megachilidae Apidae

Bee Family

Figure 8. The number of species currently recorded for each bee family in British Columbia.

Although not as diverse as some other North American bee hot spots (Carril ef al.

2018), the WIB ecozone is the most diverse for bees, with 411 confirmed species (Fig. 9)

— almost half of those known from Canada — with 176 of these not occurring in the

province’s other ecozones (and most of these species are not found anywhere else in

Canada). The PacM and MonC ecozones are also diverse with respect to bees, with 207

and 204 confirmed species in each, respectively (Fig. 9). The PacM ecozone has 26 bee

species not yet reported elsewhere in the province, with an additional 18 seemingly

restricted to the MonC ecozone within the province. These southern ecozones (..e.,

PacM, WIB, MonC) have higher levels of similarity to each other than to more northerly

ecozones (i.e., TaiPl, BorC, BorPl; Fig. 10), although the bee fauna of the WIB shared

less than 37% of its species with the MonC and PacM. This low level of similarity 1s due

to the large number of species endemic to the WIB within Canada, supporting the

suggestion that this small area has very high conservation value (South Okanagan

Similkameen Conservation Program 2012), especially for bees in Canada (Fig. 9). The

British Columbian segments of the three other ecozones are much less speciose, with no

bee species seemingly restricted to any one specific ecozone; all three ecozones share

more than 50% of their species (Fig. 10). The BorPl ecozone has 88 recorded bee

species, the BorC has 73 bee species, and the TaiP1 contains 70 bee species. The northern

Bombus occidentalis mckayi Ashmead, with a national conservation status by the

Committee on the Status of Endangered Wildlife in Canada (COSEWIC) of Special

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 55

Concern in Canada (COSEWIC 2014; Sheffield et al. 2016), is seemingly restricted to

the BorC ecozone within the province. |

For the overall checklist structure below, we follow Sann ef a/. (2018) for family

placement and, for convenience, we follow Michener (2007) for within-family

classification, except for the genus Lasioglossum Curtis, which follows Gibbs ef al.

(2013). New records for the province are indicated with an “*”, new records for Canada

are indicated with an “t+”. These specimens are usually supported by material in the

Barcodes of Life Data (BOLD) System (and see Sheffield et a/. 2017), although notes are

also provided in the supplementary links provided above. Species notes and other

annotations are provided for some species to clarify their status in the province.

500 8

450 ®

400 PAA tain! rt

” 6 =

® L 5 oO

~*~ Soo

ihepesza Hews:

O 250 4 YM

be 0

~”

_ 200 7 o

=z 150 & ©

z= 9 2.

100 ~”

0 0

PacM WIB MonC BorP| BorC TaiPl

Ecozone

Figure 9. The number of species (bars, left Y-axis) and species/1000km/ (dots, right Y-axis)

for each ecozone in British Columbia. PacM=Pacific Maritime; WIB=Western Interior Basin;

MonC=Montane Cordillera; BorPl=Boreal Plains; BorC=Boreal Cordillera; TaiPl=Taiga

Plains.

56 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

TaiPl

56.04

: BorC

55.44

BorPl

36.91

WIB

Ecozone

37.67

MonC

39.12

PacM

0 50 100

Percent Similarity

Figure 10. Incidence-based similarity (i.e., Jaccard’s) of the bee fauna of each ecozone in

British Columbia. The X-axis and numbers on the graph indicate percent similarity of each

ecozone or group of ecozones, the right Y-axis indicates the ecozone: PacM=Pacific

Maritime; WIB=Western Interior Basin; MonC=Montane Cordillera; BorPl=Boreal Plains;

BorC=Boreal Cordillera; TaiPl=Taiga Plains.

ANNOTATED CHECKLIST OF THE BEES OF BRITISH

COLUMBIA

PacM WIB MonC BorPl BorC TaiPl

FAMILY MELITTIDAE

Subfamily Melittinae

Genus Macropis Panzer, 1809

Subgenus Macropis Panzer, 1809

Macropis nuda (Provancher, 1882) PacM - MonC - — —

Species notes: Although Kline (2017) reported the family Melittidae (the genus Macropis) from

British Columbia, presumably for the first time, specimens of M. nuda from Agassiz in the

Canadian National Collection (Ottawa) were collected in 1914 (see Sheffield and Heron 2018).

Macropis nuda, the likely species photographed by L.R. Best (see Kline 2017) based on the shiny

terga (see Michez and Patiny 2005), is considered transcontinental (Snelling and Stage 1995) and is

known to occur across most of southern Canada (Michez and Patiny 2005) and into Montana

(Michener 1938a) in the United States. Michener (1938a) was the first to record the genus in

western North America (presumably he did not examine material in the Canadian National

Collection) — M. nuda (as M. morsei Robertson) from Colorado, and M. steironematis Robertson

from Washington (Morgan’s Ferry), Yakima River is the type locality for Macropis steironematis

J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018 i

opaca Michener, although Michener’s subspecies is considered rare (Snelling and Stage 1995). It is

possible that M. steironematis 1s also in the province.

FAMILY ANDRENIDAE

Subfamily Andreninae

Andrena Fabricius, 1775

Subgenus Andrena Fabricius, 1775

Andrena aculeata LaBerge, 1980 - WIB MonC - ot aa

Andrena buckelli Viereck, 1924 PacM WIB MonC — me a

Andrena ceanothifloris cretata _ a) MonC. = vat es

LaBerge, 1980

Andrena clarkella (Kirby, 1802) ~ -

Andrena edwardsi Viereck, 1916 - WIB

Andrena frigida Smith, 1853 PacM WIB MonC -BorPl. .Bor€,. , TaiPl

Andrena hemileuca Viereck, 1904 PacM = WIB - ~ — —

Andrena laminibucca - WIB MonC -— — ~

Viereck & Cockerell, 1914

Andrena macoupinensis a WIB = _ = or

Robertson, 1900

Andrena milwaukeensis PacM WIB ~ ~ BorC ~

Graenicher, 1903

Andrena perarmata Cockerell, 1898 PacM WIB ~~ -- --

Andrena rufosignata Cockerell, 1902 PacM WIB MonC BorPl BorC _ TaiPl

Andrena saccata Viereck, 1904 PacM - _

Andrena schuhi LaBerge, 1980 PacM WIB MonC

Andrena thaspii Graenicher, 1903 PacM WIB MonC . BorP! Bor® TaiPl

Andrena topazana Cockerell, 1906 — WIB MonC — BorC —

Andrena vicinoides Viereck, 1904 PacM WIB MonC - BorC —

Andrena washingtoni Cockerell, 1901 PacM WIB MonC — BorC -

Subgenus Cnemidandrena Hedicke, 1933

Andrena colletina Cockerell, 1906 _ WIB — — _ _

Andrena columbiana Viereck, 1917 PacM WIB MonC BorPl ~~ BorC TaiPl

*Andrena costillensis - WIB - — _ ~

Viereck & Cockerell, 1914

Andrena nubecula Smith, 1853 oa WIB MonC -— - _

Andrena runcinatae Cockerell, 1906 — - MonC — _ ~

Andrena scutellinitens Viereck, 1917 — WIB MonC -— — —

Andrena surda Cockerell, 1910 — WIB MonC —- _ ~

Species notes: Although Buckell (1949) reported A. colletina Cockerell from Chilcotin, Donovan

(1977) indicated that the collection date (16 April 1921) was too early for this species; members of

the subgenus Cnemidandrena are typically summer-flying species. However, Criddle (1924)

examined specimens collected in September from Penticton and Cranbrook, so we include it in the

list only from the WIB.

Subgenus Dactylandrena Viereck, 1924

Andrena berberidis Cockerell, 1905 — WIB — a ie me

Andrena porterae Cockerell, 1900 - WIB - 2 itd wn

Subgenus Dasyandrena LaBerge, 1977

Andrena cristata Viereck, 1917 -- WIB — Bs es

Subgenus Diandrena Cockerell, 1903

Andrena cuneilabris Viereck, 1926 — WIB ~ a he uf

Andrena evoluta a WIB ie mh int va

Linsley & MacSwain, 1961 |

Andrena nothocalaidis “ WIB we mn KY ye

Cockerell, 1905

58 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Subgenus Euandrena Hedicke, 1933

Andrena astragali — WIB - ~ _ ~

Viereck & Cockerell, 1914

Andrena auricoma Smith, 1879 PacM — - —

Andrena caerulea Smith, 1879 PaeM -— — — = “=

Andrena chlorura Cockerell, 1916 PaM - — — 2 =

*4Andrena geranii Robertson, 1891 — WIB ~ - - -

Andrena lawrencei ~ WIB - ~ - —

Viereck & Cockerell, 1914

+Andrena misella Timberlake, 1951 — WIB - - - ~

Andrena nigrihirta (Ashmead, 1890) PacM WIB MonC BorPl ~ BorC TaiPl

Andrena nigrocaerulea PacM WIB - - - ~

Cockerell, 1897

Andrena segregans Cockerell, 1900 — — MonC — - ~

Species notes: Although Linsley (1951b) reported A. chlorura Cockerell from the province, no

specific details were provided. Ecozone information is provided from confirmed material at the

Spencer Entomology Museum, University of British Columbia

Subgenus Geissandrena LaBerge & Ribble, 1972

Andrena trevoris Cockerell, 1897 PacM WIB MonC

Subgenus Holandrena Pérez, 1890

Andrena cressonii infasciata PacM WIB ~ — = a

Lanham, 1949

Subgenus Larandrena LaBerge, 1964

Andrena miserabilis Cresson, 1872 PacM WIB _ a ms &

Subgenus Leucandrena Hedicke, 1933

Andrena barbilabris (Kirby, 1802) PacM WIB MonC BorPl BorC TaiPl

Subgenus Melandrena Pérez, 1890

Andrena carlini Cockerell, 1901 - WIB — _ ~ =

Andrena cerasifolii Cockerell, 1896 — — MonC -— = _

Andrena commoda Smith, 1879 - WIB — — ~ a

Andrena lupinorum Cockerell, 1906 — WIB — - —

Andrena nivalis Smith, 1853 PacM WIB MonC BorPl — BorC TaiPl

Andrena pertristis Cockerell, 1905 — WIB MonC "= — _

Andrena regularis Malloch, 1917 - WIB MonC BorPl —- --

Andrena sola Viereck, 1917 —- WIB ~ ~ — —

Andrena transnigra Viereck, 1904 PacM WIB MonC = BorC —

Andrena vicina Smith, 1853 PacM WIB MonC - — —~

Subgenus Micrandrena Ashmead, 1899

Andrena candidiformis — WIB ~ ~ ~ _

Viereck & Cockerell, 1914

Andrena chlorogaster Viereck, 1904. — WIB - ~ ~ ~

Andrena illinoiensis Robertson, 1891 — WIB _ - ~ ~

Andrena melanochroa PacM WIB MonC —- — ~

Cockerell, 1898

Andrena microchlora Cockerell, 1922 — WIB — — ~ —

Andrena piperi Viereck, 1904 ~ WIB - — _ ~

Andrena salictaria Robertson, 1905 — WIB MonC BorPl —- _

Subgenus Parandrena Robertson, 1897

Andrena andrenoides (Cresson, 1878) — WIB ~ - be si

Andrena concinnula Cockerell, 1898 — WIB - _ - a

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Andrena gibberis Viereck, 1924 -

Andrena nevadensis Cresson, 1879 ~~

Subgenus Plastandrena Hedicke, 1933

Andrena crataegi Robertson, 1893 PacM

Andrena prunorum prunorum PacM

Cockerell, 1896

Subgenus Scaphandrena Lanham, 1949

Andrena chapmanae Viereck, 1904. —

Andrena merriami Cockerell, 1901 —

Andrena montrosensis -

Viereck & Cockerell, 1914

Andrena scurra Viereck, 1904 PacM

Andrena sladeni Viereck, 1924 ~

Andrena walleyi Cockerell, 1932 ~

WIB

MonC

Species notes: Ribble (1974) considered A. montrosensis (recorded in the province by Buckell

1949) synonymous with a hybrid of A. scurra x arabis x capricornis, although Lanham (1984,

1987, 1993) later considered it a valid species, which is followed here.

Subgenus Simandrena Pérez, 1890

Andrena angustitarsata Viereck, 1904 PacM

Andrena pallidifovea Viereck, 1904 —

Andrena subtrita Cockerell, 1910 -

Andrena wheeleri Graenicher, 1904. —

Subgenus Thysandrena Lanham, 1949

Andrena candida Smith, 1879 PacM

Andrena knuthiana Cockerell, 1901 —

Andrena medionitens Cockerell, 1902 PacM

Andrena trizonata (Ashmead, 1890) —

Andrena vierecki Cockerell, 1904 PacM

Andrena w-scripta Viereck, 1904 PacM

Subgenus Trachandrena Robertson, 1902

Andrena amphibola (Viereck, 1904) | PacM

Andrena cleodora (Viereck, 1904) PacM

Andrena cupreotincta Cockerell, 1901 PacM

Andrena cyanophila Cockerell, 1906 PacM

Andrena forbesii Robertson, 1891 PacM

Andrena fuscicauda (Viereck, 1904) —

Andrena hippotes Robertson, 1895 PacM

Andrena mariae Robertson, 1891 PacM

Andrena miranda Smith, 1879 PacM

Andrena quintiliformis Viereck, 1917 —

Andrena salicifloris Cockerell, 1897 PacM

Andrena sigmundi Cockerell, 1902 §PacM

Andrena striatifrons Cockerell, 1897 PacM

Subgenus 7ylandrena LaBerge, 1964

Andrena erythrogaster PacM

(Ashmead, 1890)

Andrena perplexa Smith, 1853 PacM

Andrena subaustralis Cockerell, 1898 PacM

Andrena subtilis Smith, 1879 PacM

Unplaced Species

Andrena angustifovea Viereck, 1904 —

WIB

MonC

TaiPl

60 J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

Andrena excellens Viereck, 1924 PaM - _ — _

Andrena fulvicrista Viereck, 1924 PacM WIB _ — ~ ~

Andrena lillooetensis Viereck, 1924 PacM = — —

Andrena revelstokensis Viereck, 1924 — — MonC - —

Andrena singularis Viereck, 1924 PaeM — MonC — - a

Species notes: Though Viereck et al. (1904c) included A. augustifovea in their key to male

Andrena in a treatment of bees of the Pacific North West, no specific information was provided in

that work on the type material(s), including the number of specimens in the type series or the type

locality. Cresson (1928) reviewed non-Cresson type material at the ANSP, including a specimen of

A. augustifovea [ANSP no. 10,286] from Moscow, Idaho. Linsley (1951) and subsequent

catalogues (i.e., Hurd 1979) have subsequently included British Columbia in the range of this

species, suggesting other type material exists, even though we can find no further mention of this

species in the literature. Although Linsley (1951) and Hurd (1979) did not assign this species to a

subgenus, Ascher and Pickering (2018) place it within subgenus Simandrena Pérez and indicate

three specimens (from Oregon, Idaho, and British Columbia); however, LaBerge (1989) did not

include A. angustifovea as a valid species or as a synonymy in his revision of the subgenus. As

such, we place it here until further work is done.

Subgenera not confirmed in British Columbia: Two male specimens identified by W. LaBerge

as Andrena (Anchandrena) angustella Cockerell are in the Spencer Entomology Collection at the

University of British Columbia — one from Vaseux Lake; the other from the north end of Galiano

Island. Although LaBerge (1986) proposed and revised this subgenus, with this as the type species,

both of the British Columbia specimens have an entirely black clypeus. A yellow clypeus (or

yellow in part) is diagnostic for the subgenus (LaBerge 1986). As such, we consider these

specimens misidentified. No specimens of Anchandrena have yet been reported from Canada

(LaBerge 1986).

Criddle et al. (1924) reported Andrena (Taeniandrena) wilkella (Kirby) from British Columbia

(Saanich), but this is well outside the known range of this introduced species establishment in

North America. However, this could also represent another introduction event for this species in

another major port area.

Subfamily Panurginae

Tribe Protandrini

Genus Pseudopanurgus Cockerell, 1897

Pseudopanurgus didirupa — WIB ~ - -- _

(Cockerell, 1908)

Tribe Panurgini

Genus Panurginus Nylander, 1848

*Panurginus atriceps (Cresson, 1878) — WIB MonC -—- BorC -

+Panurginus cressoniellus — WIB — ~~ — —

Cockerell, 1898

*Panurginus ineptus Cockerell, 1922 PacM WIB — ~ BorC _—

Tribe Perditini

Genus Perdita Smith, 1853

Subgenus Perdita Smith, 1853

Perdita fallax Cockerell, 1896 — WIB ~ ~ — —

Subgenus Pygoperdita Timberlake, 1956

Perdita nevadensis Cockerell, 1896 PacM WIB — _ _ St

Tribe Calliopsini

Genus Calliopsis Smith, 1853

Subgenus Nomadopsis Ashmead, 1898

Calliopsis scitula Cresson, 1878 ~ WIB ~ -- — —

FAMILY HALICTIDAE

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 61

Subfamily Rophitinae

Genus Dufourea Lepeletier, 1841

*Dufourea dilatipes Bohart, 1948 - WIB MonC — —- —

Dufourea holocyanea — WIB MonC — - ~

(Cockerell, 1925)

Dufourea marginata (Cresson, 1878) — WIB -- - - ~

Dufourea maura (Cresson, 1878) — WIB MonC -—- — ~

Dufourea trochantera Bohart, 1948 — WIB - - - -

Other records: Dufourea oryx (Viereck) was recorded in British Columbia (Salmon Arm,

Naramata) by Criddle et al. (1924), but it is likely that these are misidentified specimens of D.

holocyana.

Subfamily Nomiinae

Genus Nomia Latreille, 1804

Subgenus Acunomia Cockerell, 1930

Nomia melanderi Cockerell, 1906 ~ WIB — _ a —

Species notes: This species was introduced to British Columbia (Ashcroft, Kamloops) from the

western United States for alfalfa pollination (Bohart 1970; Hurd 1979), but there is no evidence

that it became established in these areas. However, Stephen (1959) suggests that this species likely

occurs naturally in southern parts of the interior valleys of the province.

Subfamily Halictinae

Tribe Halictini

Genus Agapostemon Guérin-Méneville, 1844

Subgenus Agapostemon Guérin-Meéneville, 1844

Agapostemon femoratus — WIB — — —

Crawford, 1901

Agapostemon obliquus PacM — - — - -

(Provancher, 1888)

Agapostemon texanus Cresson, 1872 PacM WIB MonC - _ =

Agapostemon virescens — WIB — — - -

(Fabricius, 1775)

Genus Halictus Latreille, 1804

Subgenus Nealictus Pesenko, 1984

Halictus farinosus Smith, 1853 — WIB - - ~

Subgenus Odontalictus Robertson, 1918

Halictus ligatus Say, 1837 — WIB — ~ ~ -

Subgenus Protohalictus Pesenko, 1984

Halictus rubicundus (Christ, 1791) PacM WIB ~ — — TaiP1

Subgenus Seladonia Robertson, 1918

Halictus confusus arapahonum ~ WIB ~ - - -

Cockerell, 1906

Halictus confusus confusus PacM WIB MonC BorPl — --

Smith, 1853

Halictus tripartitus Cockerell, 1895 — WIB ~ - = -

Halictus virgatellus Cockerell, 1901 — WIB - ~ — TaiPl

Genus Lasioglossum Curtis, 1833

Subgenus Dialictus Robertson, 1902

Lasioglossum abundipunctum _ WIB - — m= <

Gibbs, 2010

Lasioglossum albipenne PacM WIB MonC — S “

(Robertson, 1890)

62 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Lasioglossum albohirtum PacM WIB MonC - _ ~

(Crawford, 1907)

Lasioglossum brunneiventre PacM WIB - - — ~

(Crawford, 1907)

Lasioglossum cressonii PacM WIB MonC BorPl — ~

(Robertson, 1890)

Lasioglossum dashwoodi — WIB ~ ~ ~ =

Gibbs, 2010

Lasioglossum hyalinum PacM WIB - ~~ - -

(Crawford, 1907)

Lasioglossum imbrex Gibbs, 2010 ~ WIB - - — a

Lasioglossum incompletum PacM WIB MonC - ~ --

(Crawford, 1907)

Lasioglossum knereri Gibbs, 2010 PacM WIB MonC - ~

Lasioglossum laevissimum PacM WIB MonC BorPl -—- TaiPl

(Smith, 1853)

Lasioglossum lilliputense Gibbs, 2010 — — MonC -—- ~ _

Lasioglossum macroprosopum = WIB CS — _ =

Gibbs, 2010

Lasioglossum marinense PacM WIB — — — _

(Michener, 1936)

Lasioglossum nevadense PacM WIB - - ~ -

(Crawford, 1907)

Lasioglossum nigroviride = WIB MonC BorPl -— —-

(Graenicher, 1910)

Lasioglossum novascotiae PacM WIB MonC BorPl — BorC TaiPl

(Mitchell, 1960)

+Lasioglossum obnubilum _ — MonC -— - —

(Sandhouse, 1924)

Lasioglossum pacatum PacM WIB ~ - - ~

(Sandhouse, 1924)

Lasioglossum planatum PacM WIB MonC. BorPl — TaiPl

(Lovell, 1905)

Lasioglossum prasinogaster — WIB MonC -— — —

Gibbs, 2010

Lasioglossum pruinosum PacM WIB MonC — — —

(Robertson, 1892)

Lasioglossum punctatoventre — WIB — — — _

(Crawford, 1907)

Lasioglossum reasbeckae Gibbs, 2010 PacM = WIB ~ - ~ —

Lasioglossum ruidosense PacM WIB MonC BorPl — BorC TaiPl

(Cockerell, 1897)

Lasioglossum sagax _ WIB MonC BorPl — -

(Sandhouse, 1924)

Lasioglossum sandhousiellum PacM WIB — — ~ ~

Gibbs, 2010

Lasioglossum sedi (Sandhouse, 1924) — WIB -- a - —

Lasioglossum subversans PacM WIB MonC BorPl - -

(Mitchell, 1960)

Lasioglossum tenax — WIB MonC BorP! — BorC TaiPl

(Sandhouse, 1924)

Lasioglossum testaceum — WIB — — — —

(Robertson, 1897)

Lasioglossum tuolumnense — ~~ WIB _ — _ ~

Gibbs, 2009

Lasioglossum yukonae Gibbs, 2010 PacM — — - BorC -

Other records: Lasioglossum atriventre (Crawford) was declared nomen dubium by Gibbs (2010);

the type locality is within British Columbia (Goldstream).

J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

Subgenus Evylaeus Robertson, 1902

+Lasioglossum argemonis PacM

(Cockerell, 1897)

Subgenus Hemihalictus Cockerell, 1897

Lasioglossum diatretum PacM

(Vachal, 1904)

+Lasioglossum glabriventre -

(Crawford, 1907)

Lasioglossum inconditum PacM

(Cockerell, 1916)

+Lasioglossum kincaidii ~

(Cockerell, 1898)

Lasioglossum macoupinense PacM

(Robertson, 1895)

Lasioglossum ovaliceps PacM

(Cockerell, 1898)

Lasioglossum pectoraloides —

(Cockerell, 1895)

Subgenus Lasioglossum Curtis, 1833

Lasioglossum anhypops PacM

McGinley, 1986

Lasioglossum athabascense _

(Sandhouse, 1933)

Lasioglossum colatum (Vachal, 1904) —

Lasioglossum egregium PacM

(Vachal, 1904)

Lasioglossum mellipes PacM

(Crawford, 1907)

Lasioglossum olympiae PacM

(Cockerell, 1898)

Lasioglossum pacificum PacM

(Cockerell, 1898)

Lasioglossum sisymbrii PacM

(Cockerell, 1895)

Lasioglossum titusi (Crawford, 1902) —

Lasioglossum trizonatum =

(Cresson, 1874)

Subgenus Leuchalictus Warncke, 1975

Lasioglossum zonulum (Smith, 1848) PacM

Subgenus Sphecodogastra Ashmead, 1899

Lasioglossum arctoum (Vachal, 1904) —

Lasioglossum boreale PacM

Svensson, Ebmer & Sakagami, 1977

Lasioglossum comagenense —

(Knerer & Atwood, 1964)

Lasioglossum cooleyi _

(Crawford, 1906)

Lasioglossum cordleyi PacM

(Crawford, 1906)

Lasioglossum nigrum (Viereck, 1903) PacM

Lasioglossum quebecense PacM

(Crawford, 1907)

Genus Sphecodes Latreille, 1804

WIB

BorPl

TaiPl

64 _ J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

+Sphecodes arvensiformis PacM WIB _ - - -

Cockerell, 1904

* Sphecodes clematidis _ PacM WIB — — - -

Robertson, 1897

*Sphecodes pecosensis pecosensis PacM WIB ~ - _ -

Cockerell, 1904

*Sphecodes prosphorus — WIB — - - ~

Lovell & Cockerell, 1907

*Sphecodes solonis Graenicher, 1911 — WIB = — _ TaiPl

FAMILY COLLETIDAE

Subfamily Colletinae

Tribe Colletini

Colletes Latreille, 1802

Colletes compactus hesperius — WIB — — ~ ~

Swenk, 1906

Colletes consors pascoensis — WIB — — - ~

Cockerell, 1898

Colletes fulgidus fulgidus PacM WIB MonC -— ~ ~

Swenk, 1904

Colletes gypsicolens Cockerell, 1897 — WIB — — -

Colletes hyalinus oregonensis - WIB ~ - _ -

Timberlake, 1951

Colletes impunctatus lacustris — WIB — BorPl = BorC TaiPl

Swenk, 1906

Colletes kincaidii Cockerell, 1898 PacM WIB MonC -—- ~ -

Colletes mandibularis Smith, 1853 — WIB — _ = zs

Colletes phaceliae Cockerell, 1906 — WIB ~ ~ BorC -

Colletes simulans nevadensis ~ WIB — — ~ -

Swenk, 1908

Colletes slevini Cockerell, 1925 — WIB — _ =

Other records: As indicated by Stephen (1954), the record of Colletes angelicus Cockerell from

British Columbia (Pentiction, Walhachin) by Criddle et al. (1924) is likely based on a

misidentification, so is not included here.

The same is likely also true for C. gilensis Cockerell, recorded by Gibson (1914) (Similkameen,

Okanagan), because the species distribution also seems restricted to the southern United States.

Subfamily Hylaeinae

Aylaeus Fabricius, 1793

Subgenus Cephalylaeus Michener, 1942

Hylaeus basalis (Smith, 1853) PacM WIB MonC BorPl -—- TaiPl

Subgenus Hylaeus Fabricius, 1793

Hylaeus annulatus (Linnaeus, 1758) PacM WIB MonC BorPl ~~ BorC TaiPl

Hylaeus leptocephalus - WIB - _ - ~-

(Morawitz, 1871)

Hylaeus mesillae (Cockerell, 1896) = — WIB - - - ~

Hylaeus rudbeckiae - WIB ~ a - -

(Cockerell & Casad, 1895)

Hylaeus verticalis (Cresson, 1869) _ WIB a BorPl — ~

Species notes: Elwell (2012) recorded H. rudbeckia from the Western Interior Basin, but this was

not indicated in the follow-up publication (Elwell et al. 2016). This species was also recorded on

Discover Life (Ascher and Pickering 2018) from material in the AMNH [Cache Creek].

Subgenus Paraprosopis Popov, 1939

Hylaeus coloradensis WIB ~ on a4. 5.

(Cockerell, 1896)

J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018 __. 65

Hylaeus nevadensis (Cockerell, 1896) — WIB _ — = ws

Hylaeus wootoni (Cockerell, 1896) = — WIB ~ “= he ee

Other records: Criddle et al. (1924) reported H. cookii (Metz) from British Columbia (Kaslo), but.

this was likely a misidentification; Snelling (1970) indicates that, until 1970, the species was

known only from the type specimen (from New Mexico), and suggests that Metz's original

description was not helpful for recognizing this species. Therefore, we do not include this species

here.

Subgenus Prosopis Fabricius, 1804

Hylaeus affinis (Smith, 1853) _ WIB MonC -—- — =

Hylaeus episcopalis (Cockerell, 1896) — WIB — a = ss

Hylaeus modestus citrinifrons PacM WIB MonC —- ss om

(Cockerell, 1896)

Species notes: Gibson and Criddle (1920) recorded H. modestus Say from British Columbia

(Kaslo), but here we assume it was the subspecies H. modestus citrinifrons.

FAMILY MEGACHILIDAE

_ Subfamily Megachilinae

Tribe Osmiini

Genus Ashmeadiella Cockerell, 1897

Subgenus Ashmeadiella Cockerell, 1897

Ashmeadiella bucconis denticulata = — WIB ~ ~ = ~

(Cresson, 1878)

Ashmeadiella cactorum cactorum — WIB ~ ~- - ~

(Cockerell, 1897)

Ashmeadiella californica californica — WIB ~ -- -

(Ashmead, 1897)

Ashmeadiella cubiceps ~ WIB ~ - - ~

(Cresson, 1879)

Other records: Hurd and Michener (1955) showed a range map indicating that Ashmeadiella

(Argochila) foxiella Michener was likely in British Columbia (Western Interior Basin), but no

locality data were provided. Therefore, it is not included in the list above.

Genus Atoposmia Cockerell, 1935

Subgenus Afoposmia Cockerell, 1935

Atoposmia abjecta (Cresson, 1878) — — MonC —- — =

Species notes: Hurd and Michener (1955) showed a range map indicating that Atoposmia oregona

(Michener) was likely in southern British Columbia, but no locality data were provided. Therefore,

it is not included in the list above.

Subgenus Hexosmia Michener, 1943

Atoposmia copelandica copelandica — WIB -- _ in =

(Cockerell, 1908)

Genus Chelostoma Latreille, 1809

Subgenus Foveosmia Warncke, 1991

Chelostoma minutum Crawford, 1916 — WIB — a is s

Chelostoma phaceliae Michener, 1938 — WIB - - ~ --

Genus Heriades Spinola, 1808

Subgenus Neotrypetes Robertson, 1918

Heriades carinata Cresson, 1864 - WIB — 4 “ =

Heriades cressoni Michener, 1938 -- WIB - - i -

Heriades variolosa variolosa - WIB — a sis ed

(Cresson, 1872)

66 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Genus Hoplitis Klug, 1807

Subgenus Alcidamea Cresson, 1864

Hoplitis albifrons albifrons ~ ~ MonC BorPl ~~ BorC TaiPl

(Kirby, 1837)

Hoplitis albifrons argentifrons PacM WIB MonC - — --

(Cresson, 1864) |

Hoplitis fulgida fulgida PacM WIB MonC BorPl - -

(Cresson, 1864)

Hoplitis grinnelli septentrionalis WIB MonC — a -

Michener, 1947

Hoplitis hypocrita (Cockerell, 1906) PacM WIB MonC — £ a

Hoplitis louisae (Cockerell, 1934) PacM WIB MonC —- = me

Hoplitis producta subgracilis PacM WIB MonC —- — —

Michener, 1947

Hoplitis sambuci Titus, 1904 PacM WIB MonC — ~ -

Hoplitis spoliata (Provancher, 1888) — WIB MonC BorPl -—- TaiPl

Species notes: Michener (1947b) and Hurd and Michener (1955) indicate that H. albifrons

albifrons occurrs across Canada, including in northern British Columbia, being replaced by H.

albifrons argentifrons in the southern part of the province. Michener (1947a) indicates that

separation of the subspecies (based on hair colour) in some areas would likely be difficult, but

DNA barcoding suggests there is much variation within this species in the province (i.e., three

clusters with no apparent geographic pattern) all sharing a single Barcode Index Number.

Incidentally, there are three subspecies in North America (Michener 1947a, b; Hurd and Michener

1955; Rowe 2017).

Subgenus Formicapis Sladen, 1916

Hoplitis robusta robusta PacM WIB MonC BorPl BorC _ TaiPl

(Nylander, 1848)

Species notes: Michener (1938c) recorded this species from MonC (Field); Hurd (1979) recorded

this Holarctic species from British Columbia, but no specific localities were provided. However, it

is likely found in all ecozones in the province.

Genus Osmia Panzer, 1806

Subgenus Cephalosmia Sladen, 1916

Osmia californica Cresson, 1864 - WIB ~ a - -

Osmia marginipennis Cresson, 1878 — WIB ~ - — —

Osmia montana montana -- WIB MonC — - —

Cresson, 1864

Osmia subaustralis Cockerell, 1900 — - MonC BorPl — -

Subgenus Helicosmia Thomson, 1872

Osmia caerulescens caerulescens PacM WIB MonC -—- - —

(Linnaeus, 1758)

Osmia coloradensis Cresson, 1878 - WIB MonC -—- ~ -

Osmia texana Cresson, 1872 PacM WIB — ~ - —

Subgenus Melanosmia Schmiedeknecht, 1885

Osmia albolateralis Cockerell, 1906 PacM WIB MonC —- _- _

*Osmia aquilonaria — — MonC. = — ~

Rightmyer, Griswold & Arduser, 2010

*Osmia atriventris Cresson, 1864 ~ — MonC — 2 =

Osmia atrocyanea Cockerell, 1897 — WIB - = ta “a

Osmia austromaritima # WIB me if = ma

Michener, 1936

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 67

Osmia bella Cresson, 1878 PacM WIB MonC — — —

Osmia brevis brevis Cresson, 1864 PacM WIB — — ~ _

Osmia bruneri Cockerell, 1897 - WIB MonC -— _ ———

Osmia bucephala Cresson, 1864 PacM WIB MonC BorPl BorC TaiPl

Osmia calla Cockerell, 1897 _ WIB ~ — _ =

Osmia cobaltina Cresson, 1878 — WIB _ ~ — _

Osmia cyanella Cockerell, 1897 WIB - ~ a

Osmia cyaneonitens Cockerell, 1906 — WIB — — ~

Osmia densa densa Cresson, 1864 PacM WIB MonC — fa ag

Osmia dolerosa Sandhouse, 1939 PacM WIB — = a ma

Osmia ednae Cockerell, 1907 = WIB 7 = = ~

Osmia enixa Sandhouse, 1924 ~ WIB — — ~ ~

Osmia exigua Cresson, 1878 — WIB — ~ ~ -

Osmia giliarum Cockerell, 1906 — WIB MonC -— - -

Osmia inermis (Zetterstedt, 1838) — — MonC BorPl — BorC TailPl

Osmia integra Cresson, 1878 — WIB MonC - > TaiP]

Osmia inurbana Cresson, 1878 PacM WIB ~ - — —

Osmia juxta juxta Cresson, 1864 — WIB MonC -—- ~ —

Osmia juxta subpurpurea — WIB — _ — —

Cockerell, 1897

Osmia kincaidii Cockerell, 1897 PacM WIB MonC -— - —

+Osmia laeta Sandhouse, 1924 - WIB MonC — - -

Osmia lignaria propinqua PacM WIB MonC — - — ~

Cresson, 1864

Osmia longula Cresson, 1864 ~ WIB MonC -— BorC -

+Osmia malina Cockerell, 1909 — WIB ~ ~ ~ ~

Osmia melanopleura Cockerell, 1916 — WIB ~ = -- ~

Osmia mertensiae Cockerell, 1907 PacM WIB _ — ~ Ee

Osmia nanula Cockerell, 1897 PacM WIB ~ ~ —~ _

Osmia nemoris Sandhouse, 1924 - WIB — - — ~

Osmia nifoata Cockerell, 1909 ~ WIB ~ _ ~ --

Osmia nigrifrons Cresson, 1878 _ WIB MonC - — ~

Osmia nigriventris (Zetterstedt, 1838) PacM WIB MonC BorPl — BorC ~-

Osmia obliqua White, 1952 _ WIB _ ~ -

Osmia odontogaster Cockerell, 1897 — WIB MonC — _ _

*Osmia paradisica Sandhouse, 1924 — WIB ~ ~ - —

Osmia pentstemonis Cockerell, 1906 — WIB ~ a - ~

Osmia pikei Cockerell, 1907 — WIB ~ - _ -

Osmia proxima Cresson, 1864 PacM WIB MonC BorPl — BorC TaiP|

+ Osmia pulsatillae Cockerell, 1907 — WIB MonC — ~ -

Osmia pusilla Cresson, 1864 PacM ss — WIB — - ~ ~

+Osmia raritatis Michener, 1957 “ WIB - ~ _ ~

Osmia regulina Cockerell, 1911 — WIB — _ - -

Osmia sedula Sandhouse, 1924 — WIB — ~ oe ~

Osmia simillima Smith, 1853 — WIB MonC BorPl -— —

Osmia tersula Cockerell, 1912 ~ — MonC BorPl -— -

Osmia trevoris Cockerell, 1897 — WIB — — — —

Osmia tristella tristella PacM WIB — BorPl -— ~

Cockerell, 1897

Osmia unca Michener, 1937 - WIB _ a a —

Species notes: Osmia mertensiae Cockerell and Osmia inurbana Cresson (as O. eutrichosa

Cockerell) were recorded from British Columbia by Sandhouse (1925b) so are listed here, but Hurd

(1979) considers the species questionable from British Columbia.

Tribe Anthidiini

Genus Anthidiellum Cockerell, 1904

Subgenus Loyolanthidium Urban, 2001

Anthidiellum robertsoni WIB - - — —

68 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

(Cockerell, 1904)

Species notes: Based on distinct differences in the cytochrome c oxidase I (COI) gene that

resulted in two distinct Barcode Index Numbers (BINs) (see Sheffield et a/. 2017), we agree with

Urban (2001) and consider this a separate species from the eastern 4. notatum (Latreille).

Genus Anthidium Fabricius, 1804

Subgenus Anthidium Fabricius, 1804

Anthidium atrifrons Cresson, 1868 — WIB — a — oe

Anthidium clypeodentatum — WIB MonC - - TaiPl

Swenk, 1914

Anthidium emarginatum (Say, 1824) — WIB MonC — — —

+ Anthidium formosum Cresson, 1878 — WIB — - - -

Anthidium manicatum PaeM = -— MonC -— - ~

(Linnaeus, 1758)

Anthidium mormonum Cresson, 1878 — WIB MonC -— ~ —

Anthidium palliventre Cresson, 1878 — — MonC — - ~

Anthidium psoraleae Robertson, 1902 — WIB ~ — ~ ~

Anthidium tenuiflorae Cockerell, 1907 — WIB — -- _ TaiPl

Anthidium utahense Swenk, 1914 - WIB MonC -— - _

Species notes: Although Michener (1951) and Hurd (1979) recorded A. porterae Cockerell from

“BC” (no specific locality), we have found reference to this species in Canada only from Alberta

(Calgary) by Cockerell (1912). Gonzalez and Griswold (2013) and Griswold et a/. (2014) did not

record this species from Canada in their revision and compilation of occurrence records for the

genus in the Western Hemisphere, respectively.

Genus Dianthidium Cockerell, 1900

Subgenus Dianthidium Cockerell, 1900

Dianthidium curvatum sayi = WIB - - — ~

Cockerell, 1907

tDianthidium plenum plenum - WIB ~ - — ~

Timberlake, 1943

Dianthidium pudicum pudicum He WIB - ~ — —

(Cresson, 1879)

{Dianthidium singulare ~ WIB ~ — - --

(Cresson, 1879) |

Dianthidium subparvum PacM WIB ~- _ — —

Swenk, 1914

Dianthidium ulkei ulkei — WIB a ~ — -

(Cresson, 1878)

Genus Stelis Panzer, 1806

Subgenus Stelis Panzer, 1806 |

tStelis ashmeadiellae PacM — ~ ~ -- -

Timberlake, 1941 .

+Stelis calliphorina (Cockerell, 1911) — WIB a ~ ~ “

Stelis callura Cockerell, 1925 — WIB — me a ee

Stelis carnifex Cockerell, 1911 — WIB ~ ~ — —

*Stelis coarctatus Crawford, 1916 - WIB ~ ~ _ ~

Stelis elegans Cresson, 1864 - WIB — — - ~

Stelis lateralis Cresson, 1864PacM — = ve us =a

Stelis maculata (Provancher, 1888) PacM —

Stelis montana Cresson, 1864 ub ~ WIB 24 2 ie ye)

Stelis monticola Cresson, 1878 bes WIB = ms oy fas

Stelis occidentalis _ WIB uy as a i

Parker & Griswold, 2013

Stelis ricardonis (Cockerell, 1912) PacM WIB —

Stelis rubi Cockerell, 1898 - WIB MonC -—- - —

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Tribe Dioxyini

Genus Dioxys Lepeletier & Serville, 1825

+Dioxys pomonae pomonae —

Cockerell, 1910

WIB

Species notes: This is the first record of this species from Canada; however, Sheffield et al. (2017)

recorded this genus (this species, based on this single barcoded specimen) from British Columbia,

Canada.

Tribe Megachilini

Genus Coelioxys Latreille, 1809

Subgenus Boreocoelioxys Mitchell, 1973

Coelioxys banksi Crawford, 1914 PacM

Coelioxys funeraria Smith, 1854 PacM

Coelioxys moesta Cresson, 1864 PacM

Coelioxys novomexicana —

Cockerell, 1909

Coelioxys octodentata Say, 1824 --

Coelioxys porterae Cockerell, 1900 PacM

Coelioxys rufitarsis Smith, 1854 PacM

Coelioxys sayi Robertson, 1897 —

Subgenus Coelioxys Latreille, 1809

Coelioxys hirsutissima =

Cockerell, 1912

Coelioxys sodalis Cresson, 1878 PacM

Subgenus Cyrtocoelioxys Mitchell, 1973

Coelioxys deani Cockerell, 1909 -

Subgenus Synocoelioxys Mitchell, 1973

Coelioxys alternata Say, 1837 PacM

Coelioxys apacheorum PacM

Cockerell, 1900

Coelioxys erysimi Cockerell, 1912 -

Subgenus Xerocoelioxys Mitchell, 1973

Coelioxys edita Cresson, 1872

Coelioxys grindeliae Cockerell, 1900 PacM

Genus Megachile Latreille, 1802

Subgenus Argyropile Mitchell, 1934

Megachile parallela Smith, 1853 PacM

Subgenus Chelostomoides Robertson, 1901

Megachile angelarum Cockerell, 1902 PacM

Species notes: Although Megachile (Chelostomoides) subexilis

British Columbia (Kaslo, Penticton) by Gibson (1917), it is

misidentified specimens. Gibson (1917) reported this species

WIB

MonC

BorPl BorC TailP1

BorPl BorC TaiPl

— _ TaiPl

_ BorC TaiPl

Cockerell was recorded from

likely that this is based on

in both Ontario and British

Columbia, but he likely confused it with M. campanulae (Robertson) and M. angelicus found in

each of those provinces, respectively (see Sheffield et a/. 2011). Interestingly, Criddle et al. (1924)

also record it from Alberta, Saskatchewan, Manitoba, and Fort Norman (Northwest Territories),

supporting that these records were misidentified.

Subgenus Eutricharaea Thomson, 1872

Megachile apicalis Spinola, 1808 PacM

WIB

70 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Megachile rotundata PacM WIB — 4g ss pa

(Fabricius, 1793)

Subgenus Litomegachile Mitchell, 1934

Megachile brevis Say, 1837 — WIB - — ~- -

Megachile cleomis Cockerell, 1900 = - WIB a ~ - ~

Megachile coquilletti Cockerell, 1915 — WIB - — - -

Megachle gentilis Cresson, 1872 PacM WIB -- — - -

Megacile lippiae Cockerell, 1900 - WIB “- a ~ —

Megachle mendica Cresson, 1878 - WIB ~ a ~ -

Megachile onobrychidis — WIB a — ~ -

Cockerell, 1908

Megachile snowi Mitchell, 1927 - WIB - - -

Megachile texana Cresson, 1878 PacM WIB ~ — a ~

Subgenus Megachile Latreille, 1802

Megachile centuncularis PacM WIB MonC — oa -

(Linnaeus, 1758)

Megachile inermis Provancher, 1888 — WIB MonC BorPl —

Megachile lapponica Thomson, 1872 PacM — Mont *BorPl-. ‘BorC:.TaiPl

Megachile montivaga Cresson, 1878 PacM WIB MonC — -

Megachile relativa Cresson, 1878 PacM WIB MonG>“BorPi << BorC TaiPl

Subgenus Megachiloides Mitchell, 1924

Megachile subnigra Cresson, 1879 — WIB - -- - -

Megachile umatillensis — WIB ~ ~ = _

(Mitchell, 1927)

Megachine wheeleri Mitchell, 1927 — WIB - ~ - -

Subgenus Sayapis Titus, 1906

Megachile fidelis Cresson, 1878 PacM WIB ~ — — -

Megachile mellitarsis Cresson, 1878 — WIB o - -~ -

+ Megachile pugnata pomonae — WIB - — _ ~

Cockerell, 1916

Megachile pugnata pugnata Say, 1837 PacM ~=>WIB MonC BorPl — TaiPl

Subgenus Xanthosarus Robertson, 1903

Megachile circumcincta (Kirby, 1802) — WIB MonC BorPl BorC TaiPl

Megachile frigida Smith, 1853 PacM WIB MonC BorPl — BorC TaiP]

Megachile gemula Cresson, 1878 PacM WIB MonC BorPl -—- TaiPl

Megachile melanophaea Smith, 1853 PacM WIB MonC BorPl BorC TaiPl

Megachile perihirta Cockerell, 1898 PacM WIB MonC BorPl — BorC TaiPl

Subgenera not confirmed in British Columbia: Criddle et al. (1924) recorded Megachile

(Pseudocentron) pruina Smith from western Canada (including Summerland, British Columbia),

but this subgenus has not been recorded in Canada (Sheffield et a/. 2011). It is suspected these

records are misidentified specimens of M. parallela Smith.

FAMILY APIDAE

Subfamily Xylocopinae

Tribe Ceratinini

Genus Ceratina Latreille, 1802

Subgenus Zadontomerus Ashmead, 1899

Ceratina acantha Provancher, 1895 PacM WIB MonC -—- — “

Ceratina nanula Cockerell, 1897 PacM WIB MonC -—- - —

Ceratina pacifica Smith, 1907 PacM WIB - ~ — -

Subfamily Nomadinae

Tribe Nomadini

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 71

Genus Nomada Scopoli, 1770

Nomada aldrichi Cockerell, 1910 _ WIB — — — ™

Nomada articulata Smith, 1854 — _ MonC -— o a

Nomada bella Cresson, 1863 PacM WIB MonC — ~ ~

Nomada citrina Cresson, 1878 PaeM -—- _— - - _

Nomada civilis Cresson, 1878 _ WIB - ~- — ~

Nomada corvallisensis - WIB - - - ~

Cockerell, 1903

+Nomada crotchii Cresson, 1878 PacM WIB - ~ — ~

Nomada edwardsii Cresson, 1878 PacM WIB ~ _ -- -

Nomada grayi Cockerell, 1903 PaeM - — — — —

Nomada pascoensis Cockerell, 1903 — WIB - ~ ~ -

Nomada perbella (Viereck, 1905) PacM — ~ - - -

Nomada rhodomelas Cockerell, 1903 PacM —

Nomada sayi Robertson, 1893 — WIB — — — —

Nomada scita Cresson, 1878 — WIB — — _ —

Nomada superba Cresson, 1863 PacM — MonC — - -

*Nomada texana Cresson, 1872 - WIB — — ~

Nomada ultima Cockerell, 1903 PaM -— _ _ — _

Nomada valida Smith, 1854 — - WIB ~ —_ —_ _

Nomada vernonensis Cockerell, 1916 — WIB — — — —

Species notes: Nomada proxima Cresson was recorded from British Columbia (Vernon) by

Viereck (1926), but that species is known only from type material from Maine. We presume

Viereck’s record to be misidentified.

Mitchell (1962) and Hurd (1979) recorded Nomada valida Smith from British Columbia, but

provided no specific localities. However, N. nigrocincta Smith, recorded from Penticton by Criddle

et al. (1924), is considered an unpublished synonymy of N. valida (opinion of Snelling, as cited by

Ascher and Pickering 2018).

Tribe Epeolini

Genus Epeolus Latreille, 1802

Epeolus americanus (Cresson, 1878) — WIB MonC — ~ -

Epeolus compactus Cresson, 1878 - WIB MonC — _ -

Epeolus minimus (Robertson, 1902) PacM WIB MonC - BorC -

Epeolus olympiellus Cockerell, 1904. PacM WIB ~ ~ oe bi

Genus Triepeolus Robertson, 1901

Triepeolus occidentalis — WIB _ ~ ~ -

(Cresson, 1878)

Triepeolus paenepectoralis PacM - - ~ — ~

Viereck, 1905

Triepeolus subalpinus Cockerell, 1910 — WIB - ~ - -

Triepeolus texanus (Cresson, 1878) = — WIB — — — —

Tribe Biastini

Genus Neopasites Ashmead, 1898

Neopasites aff. fulviventris ~ WIB — — - —

(Cresson, 1878)

Tribe Emphorini

Genus Diadasia Patton, 1879

*Diadasia australis (Cresson, 1878) — WIB MonC - _ ~-

Diadasia diminuta (Cresson, 1878) WIB MonC — — -

Tribe Eucerini

Genus Eucera Scopoli, 1770

Subgenus Synhalonia Patton, 1879

Eucera acerba (Cresson, 1879) — WIB — -- —

72 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Eucera actuosa (Cresson, 1878) PacM WIB ~ — — —

Eucera cordleyi (Viereck, 1905) - WIB - - - ~

Eucera douglasiana (Cockerell, 1906) — WIB - _ ~ -

Eucera edwardsii (Cresson, 1878) — WIB — ~ - a

Eucera frater lata (Provancher, 1888) PacM — — — — —

Eucera fulvitarsis fulvitarsis ~ WIB — ~ - -

(Cresson, 1878)

Eucera hirsutissima (Cockerell, 1916) PacM ss — ~ — - -

Eucera hurdi (Timberlake, 1969) — ~ — — _ os

Eucera virgata (Cockerell, 1905) - WIB - - ~ -

Species notes: Eucera hirsutissima (Cockerell) was recorded from “British Columbia” by

Cockerell (1916b), Gibson (1918), and Hurd (1979) though no specific localities were provided.

He (Cockerell 1916b) indicated that a second label, “Toba” was on the type specimen at the British

Museum, suggesting Toba Inlet on Powell River. Thus, we include the PacM in the list above.

Similarly, E. hurdi was recorded from the province by Hurd (1979), but no other literature records

are known. Thus, we do not specify ecozone information for this species.

Genus Melissodes Latreille, 1825

Subgenus Eumelissodes LaBerge, 1956

Melissodes agilis Cresson, 1878 _ - MonC — _ _

Melissodes bimatris LaBerge, 1961 — WIB - — — —

Melissodes lutulentus LaBerge, 1961 — WIB — - ~ ~

Melissodes menuachus Cresson, 1868 — WIB a = _ ot

Melissodes microstictus PacM WIB — — - ~

Cockerell, 1905

Melissodes pallidisignatus - WIB ~ - ~

Cockerell, 1905

1 Melissodes saponellus — WIB —~ — — —

Cockerell, 1908

Melissodes semilupinus = WIB ~ ~ - _

Cockerell, 1905

Species notes: Although Michener (195le) recorded Melissodes illatus Lovell and Cockerell from

the province, no additional information was provded. However, LaBerge (1961) did not record it

from British Columbia in his revision, so it is not included here.

Subgenus Heliomelissodes LaBerge, 1956

Melissodes rivalis Cresson, 1872 ~ WIB ~ = Et i

Subgenus Melissodes Latreille, 1825

Melissodes communis alopex 7 WIB - 2 Jes aif

Cockerell, 1928

Tribe Anthophorini

Genus Anthophora Latreille, 1803

Subgenus Clisodon Patton, 1879

Anthophora terminalis Cresson, 1869 PacM WIB MonC BorPl — BorC TaiPl

Subgenus Lophanthophora Brooks, 1988

Anthophora pacifica Cresson, 1878 — WIB - 2 ie ee

Anthophora porterae Cockerell, 1900 PacM WIB — ut 2 es

Anthophora ursina Cresson, 1869 ~ WIB MonC — ss =

Subgenus Melea Sandhouse, 1943

Anthophora bomboides Kirby, 1838 PacM WIB MonC BorPl — BorC TaiPl

Anthophora occidentalis PacM WIB ~ - -

Cresson, 1869

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Subgenus Micranthophora Cockerell, 1906

Anthophora peritomae — WIB

Cockerell, 1905

Subgenus Mystacanthophora Brooks, 1988

*Anthophora urbana Cresson, 1878 PacM WIB

Subgenus Pyganthophora Brooks, 1988

Anthophora crotchii Cresson, 1878 — WIB

Anthophora edwardsii Cresson, 1878 — WIB

Genus Habropoda Smith, 1854 |

Habropoda cineraria (Smith, 1879) PacM WIB

| Habropoda miserabilis PaceM —

(Cresson, 1878)

Habropoda murihirta ~ WIB

(Cockerell, 1905)

Species notes: Stainer (1959) and Hurd (1979, likely based on Stainer’s publication) recorded

Habropoda murihirta (Cockerell) from British Columbia, likely Okanagan Mission. We have not

been able to locate this material (17 specimens) in the CNC and, although we assume that these

were likely specimens of H. cineraria, we leave it in the list.

Tribe Melectini

Genus Melecta Latreille, 1802

Subgenus Melecta Latreille, 1802

Melecta pacifica fulvida PacM WIB

Cresson, 1878

Melecta pacifica pacifica PacM WIB

Cresson, 1878

Melecta separata callura — WIB

(Cockerell, 1926)

Melecta thoracica Cresson, 1875 — WIB

Genus Xeromelecta Linsley, 1939

Subgenus Melectomorpha Linsley, 1939

Xeromelecta californica — WIB

(Cresson, 1878)

Tribe Bombini

Genus Bombus Latreille, 1802

Subgenus Alpinobombus Skorikov, 1914

Bombus kirbiellus Curtis, 1835 - -

Bombus neoboreus Sladen, 1919 PaM -

Bombus polaris Curtis, 1835 — —

MonC

BorPl BorC

— BorC

rs BorC

TaiPl

TalP1

Species notes: Although B. natvigi Richards (= North American B. hyperboreus Schénherr) was

listed from “British Columbia” by Cannings (2011), no records were recorded by Williams ef al.

(2014; as B. hyperboreus). Although it is likely that this species does occur at high elevations and/

or latitudes in the province, we have not yet found any records supporting this, so we do not

include it here.

Subgenus Bombias Robertson, 1903

Bombus nevadensis Cresson, 1874 PacM WIB

Subgenus Bombus Latreille, 1802

Bombus cryptarum (Fabricius, 1775) — WIB

Bombus occidentalis mckayi — —

Ashmead, 1902

MonC

BorPl BorC

ag BorC

TaiPl

74 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Bombus occidentalis occidentalis PacM WIB MonC __ BorPl BorC TaiPl

Greene, 1858

Bombus terricola Kirby, 1837 ~ WIB MonC BorPl — BorC TaiPl

Subgenus Cullumanobombus Vogt, 1911

Bombus griseocollis (DeGeer, 1773) — WIB ~ - - -

Bombus morrisoni Cresson, 1878 PacM WIB — a

Bombus rufocinctus Cresson, 1863 PacM WIB MonC BorPl = - ~

Subgenus Psithyrus Lepeletier, 1833

Bombus bohemicus (Seidl, 1837) WIB MonC — BorC TaiPl

Bombus flavidus Eversmann, 1852 PacM WIB MonC BorPl — BorC TaiPl

Bombus insularis (Smith, 1861) PacM WIB MonC BorPl BorC TaiPl

Bombus suckleyi Greene, 1860 PacM WIB MonC BorPl BorC TaiPl

Subgenus Pyrobombus Dalla Torre, 1880

Bombus bifarius Cresson, 1878 PacM WIB MonC BorPl — BorC TaiPl

Bombus caliginosus (Frison, 1927) PacM — aa — — —

Bombus centralis Cresson, 1864 PacM WIB MonC BorPl — BorC -

Bombus flavifrons Cresson, 1863 PacM WIB MonC BorPl BorC TaiPl

Bombus frigidus Smith, 1854 “ - MonC BorPl BorC TaiPl

Bombus huntii Greene, 1860 = WIB MonG..<iBorPl = =

Bombus impatiens Cresson, 1863 PacM = _ oa . =

Bombus jonellus (Kirby, 1802) MonC BorC TaiPl

Bombus melanopygus Nylander, 1848 PacM WIB MonC BorPl BorC _ TaiPl

Bombus mixtus Cresson, 1878 PacM WIB MonC BorPl — BorC TaiPl

Bombus perplexus Cresson, 1863 PacM WIB — BorPl ..BorC.. -—

Bombus sitkensis Nylander, 1848 PacM WIB MonC BorPl BorC TaiPl

Bombus sylvicola Kirby, 1837 PacM WIB MonC BorPl BorC TaiPl

Bombus ternarius Say, 1837 — — MonC BorPl — -

Bombus vagans vagans Smith, 1854 — WIB -- -- a ~

Bombus vandykei (Frison, 1927) _ WIB MonC —- - —

Bombus vosnesenskii PacM WIB MonC = ~ -

Radoszkowski, 1862

Species notes: Bombus impatiens was first recorded as an established species in British Columbia

by Ratti and Colla (2010; but see Ratti 2006), but it has been used as a commercial pollinator in the

province for much longer (see Van Westendorp and McCutcheon 2001).

Although B. sandersoni Franklin was recorded from British Columbia by Williams et a/. (2014), it

is likely that this specimen is misidentified, and is thus removed from the provincial list until it can

be confirmed.

Subgenus Subterraneobombus Vogt, 1911

Bombus appositus Cresson, 1878 PacM WIB MonC - ~ ~

Bombus borealis Kirby, 1837 oe — - BorPl — TaiPl

Subgenus Thoracobombus Dalla Torre, 1880

Bombus fervidus (Fabricius, 1798) PacM WIB MonC BorPl BorC -

Other records: Venables (1914) recorded Bombus pensylvanicus (De Geer) from British

Columbia, but it is likely that these specimen(s) were of the dark form of B. nevadensis or possibly

B. terricola (see Williams et al. 2014). Earlier authors (e.g., Viereck et al. 1904a) considered this

name synonymous with B. fervidus. During research for a recent Committee on the Status of

Endangered Wildlife in Canada (COSEWIC) assessment of B. pensylvanicus in Canada, all records

from west of southern Ontario in Canada were found to be misidentified (C.S.S., unpublished).

Stephen (1957) did not record B. pensylvanicus (as B. sonorus Say) from British Columbia.

Similarly, Buckell [1951; and later Hurd (1979) and Cannings (2011)] recorded Bombus auricomus

Robertson from British Columbia (Centurian, and Departure Bay [Vancouver Island]), but it is

J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018 75

likely that these specimens and possibly other specimens of this species recorded from western

Canada are the dark form of B. nevadensis.

Tribe Apini

Genus Apis Linnaeus, 1758

Subgenus Apis Linnaeus, 1758

Apis mellifera Linnaeus, 1758 PacM WIB MonC BorPl BorC —

ACKNOWLEDGEMENTS

Much time and effort went into compiling this species list, and many of the references

were searchable and accessable through the Biodiversity Heritage Library (https://

www.biodiversitylibrary.org/), and other sources. The list has also grown, thanks to

assistance with field collection and habitat information provided by Orville Dyer, Dawn

Marks, Cara Dawson, Mark Weston, David Fraser, Grant Furness, Lea Gelling, Leah

Ramsay, Syd Cannings, Rob Cannings, Geoff Scudder, David Holden, Pascale Archibald,

Babita Bains, Claudia Copley, Darren Copley, Bonnie Zand, Erica McClaren, Kristen

Peck, Sara Bunge, Rob Stewart, Lora Neild, Jayme Brooks, Brenda Costanzo, Karen

Needham, the Elizabeth Elle Lab (Simon Fraser University), Lindsay Anderson, Hannah

Flagg, Kyle Grant, Kella Sadler, Andrea Tanaka, Megan Harrison, Nick Page, Denis

Knopp, Dennis St. John, Jamie Leatham, Ted Leischner, and Jayme Brooks. Some

species records reported here came from DNA barcoded specimens collected by Lincoln

Best during graduate fieldwork at York University under the direction of Laurence

Packer; many of these were identified by CSS, Jason Gibbs, Lincoln Best, and others,

and were accessed at the Packer Collection and/or via BOLD. Some of the ecozone

photos were provided by Syd Cannings. Funding for this project is from BC Ministry of

Environment and Climate Change Strategy, Federal Habitat Stewardship Program

Prevention Stream, Royal Saskatchewan Museum, Young Canada Works, Environment

and Climate Change Canada, and BC Ministry of Forests, Lands, Natural Resource

Operations and Rural Development. Lastly, thanks to Carole Sinou (Research Officer in

Biodiversity with Canadensys) for assistance with making the checklist and occurrence

datasets publically available.

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86 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Efficacy of diamide, neonicotinoid, pyrethroid, and phenyl

pyrazole insecticide seed treatments for controlling the

sugar beet wireworm, Limonius californicus (Coleoptera:

Elateridae), in spring wheat

W.G. VAN HERK|, T.J. LABUN?, AND R.S. VERNON?

ABSTRACT

Four classes of insecticide applied on seed were evaluated for managing high

populations of the sugar beet wireworm, Limonius californicus (Coleoptera:

Elateridae), in spring wheat in southern Alberta, Canada. Three separate field trials

were conducted, and assessments made for stand protection, yield, and wireworm

survival. Imidacloprid and thiamethoxam applied at 10-30 g AI and cyantraniliprole

applied at 10-40 g AI provided initial stand protection, but did not protect seedlings

until harvest and did not decrease wireworm populations. A-cyhalothrin applied at 30 g

AI provided stand protection that persisted until harvest, but yields were considerably

lower than observed in fipronil treatments and there was little (23%) decrease in

populations relative to controls. Fipronil applied at 0.6, 1.0, and 5.0 g AI, either singly

or in blend with thiamethoxam at 10 g AI, provided stand protection until harvest and

significantly reduced numbers of wireworms larger than 10 mm (range: 74-96%).

Very low numbers of small (<11 mm) wireworms were observed in all trials. These

results are compared to data from laboratory and field studies for this and other

wireworm species. The relation between crop stand protection and wireworm

mortality, the potential of insecticide blends, and the importance of seed type,

wireworm species, and activity periods for managing wireworms with seed treatments

are discussed.

Keywords: Limonius californicus, wireworm, pest management, thiamethoxam,

fipronil, insecticide blend

INTRODUCTION

Wireworms have long been important insect pests in cereal, sugar beet, and potato

production in southern Alberta (AB) (Strickland 1927). Historically, the main pest

species were the prairie grain wireworm, Selatosomus destructor (Brown) and Hypnoidus

bicolor (Esch.) (Strickland 1927; Arnason 1931). Recent surveys indicate S. destructor

and H. bicolor remain the most commonly occurring elaterid pests in AB and

Saskatchewan (SK), while the sugar beet wireworm, Limonius californicus (Mann.), is of

more regional importance (van Herk and Vernon 2014). Described as an occasional pest

new to AB in the 1950s (MacNay 1954), and historically found only in low numbers

alongside S. destructor and H. bicolor (Doane 1977), L. californicus is currently the third

most prevalent wireworm species in arable land in the Prairie Provinces (van Herk and

Vernon 2014). In southern AB, where it is often the predominant species in continuously

cropped cereals, high L. californicus populations can cause complete stand destruction of

spring wheat, even if treated with insecticides (T.J. Labun, personal observation). The

‘relatively recent emergence of this species as a pest in this region may stem from changes

in cultivation practices, including the implementation of minimal tillage practices in

recent decades which have increased soil moisture retention. Limonius californicus 1s

' Corresponding author: Agassiz Research and Development Centre, Agriculture and Agri-Food Canada,

P.O. Box 1000, Agassiz, British Columbia, Canada, VOM 1A0; wim.vanherk@canada.ca

2 Syngenta Crop Protection (Canada) Inc., 6700 Macleod Trail Southeast, Calgary, Alberta, T2H 0OM4

3 Sentinel IPM Consulting, 4430 Estate Drive, Chilliwack, B.C., Canada V2R 3B5

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 87

known to prefer moist soils (e.g., irrigated land) and is typically not found on dry land

(van Herk and Vernon 2014). Little else is known about the ecology and life history of

this species, other than what was described by Stone (1941) for California, which

suggests it is similar to the dusky wireworm, Agriotes obscurus L., and has a three- to

five-year larval stage in the field.

In cereal production, wireworms have historically been managed by seed treatments,

particularly chlorinated hydrocarbons (Toba et al. 1988; Grove et a/. 2000). Treating seed

with lindane decreased wireworm damage in cereal crops in the Canadian prairies by

90% and pest populations by 70% in the 1940s (Arnason and Fox 1948), and led to

further decreases in damage between 1954 and 1961 (Burrage 1964). Similar results were

obtained with other species, including L. californicus, in spring wheat in the Pacific

Northwest (Toba et al. 1985, 1988). The effectiveness of lindane as a seed treatment

stemmed from its ability to kill multiple pest species and all wireworm instars of these

species, including neonates emerging from eggs laid after the seed is planted (Vernon ef

al. 2009). As a result of the latter property, wireworms would not repopulate fields to

economic levels for several years after treatment, and farmers typically treated their

cereal crops every 3-4 years (Arnason and Fox 1948). The reduction of wireworm

populations achieved by planting lindane-treated seeds also protected high-value

rotational crops such as sugarbeet, potato, and canola planted in subsequent years.

Following lindane’s de-registration (Canada in 2004; USA in 2006), there has been a

gradual but continual increase in the incidence of wireworm damage in Canadian

agricultural land. As a result, there is now a pressing need to identify and register new

wireworm control measures for cereal crops. Such measures should be cost effective,

pose negligible risk to humans and the environment, and offer similar efficacy to lindane

by providing both stand and yield protection and reduction of wireworm populations

(including controlling neonates) (Vernon ef al. 2013a).

Initial results from laboratory and field evaluations of candidate insecticides to

replace lindane as cereal seed treatments indicated neonicotinoid insecticides applied to

wheat seed at 10-30 g AI/100 kg seed provide excellent stand and yield protection of

spring wheat in the field in the presence of moderate to high populations of A. obscurus,

but they did not decrease populations relative to control treatments (Vernon ef a/. 2009,

2013a). This disconnect between crop protection and lack of wireworm mortality was

due to these insecticides inducing rapid and prolonged periods of morbidity, during

which wireworms are unable to feed and after which they generally recover (Vernon ef

al. 2008, 2009). In contrast, the phenyl-pyrazole fipronil applied at 60 g AI/100 kg seed

(a rate similar to that formerly registered for lindane) provided excellent stand and yield

protection and eliminated A. obscurus populations (including neonate larvae) in the field

(Vernon et al. 2009, 2013a). Laboratory studies indicated that dermal exposure of A.

obscurus to fipronil causes rapid and irreversible morbidity, leading to complete

mortality at higher rates. At low rates of fipronil exposure, wireworms showed no

morbidity symptoms for several months, after which latent morbidity symptoms became

apparent and mortality followed (Vernon ef a/. 2008). Exposing wireworms to wheat seed

treated with low rates of fipronil permits them to feed normally until they succumb to

latent mortality (Vernon ef al. 2013a).

Based on these observations, we hypothesized that applying low rates of both

thiamethoxam and fipronil to wheat seed would both provide stand and yield protection

equivalent to lindane, and significantly reduce wireworm (including neonate) populations

in the field (Vernon ef al. 2013a). Specifically, thiamethoxam would provide early-season

crop protection, while fipronil, even at very low doses, would cause late-season

wireworm mortality. This approach would require low amounts of chemical, thereby

reducing the environmental and human risk posed by these insecticides. Subsequent

studies with A. obscurus demonstrated that blends of thiamethoxam at 10 g AI/100 kg

seed and fipronil at | g Al/100 kg seed provided plant protection and wireworm control

equivalent to lindane (Vernon ef al. 2013a). Similarly, Morales-Rodriguez and Wanner

88 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

(2015) found that blends of these insecticides provide plant protection and reduce

numbers of L. californicus and H. bicolor, but their field studies evaluated a single rate of

these chemicals and under low pest pressure.

Here, we present results from three trials conducted in southern AB in fields with

very high populations of L. californicus to determine the efficacy of these blends and

other candidate insecticides. As wireworm species differ in their susceptibility to

insecticides, the results presented here constitute an important extension to the efficacy

data previously reported for other species.

MATERIALS AND METHODS

Plot layout and preparation. All three trials were conducted in 2012 near Granum,

AB, on a commercial field (approx. 240 ha) that had been planted to barley, peas, and

wheat in 2009, 2010, and 2011, respectively, and that had a recent history of wireworm

damage. No insecticides had been applied to crops planted in this field since ca. 2000.

Experimental design. All trials were randomized complete block designs with four

replicates. Each trial contained seed not treated with insecticide as a control treatment,

and included a combined thiamethoxam (Cruiser 5FS at 10 g AI/100 kg seed) and

fipronil (Regent 5OOFS at 1 g AI/100 kg seed) as a common insecticide seed treatment

(hereafter referred to as ‘Standard T+F Blend’) to permit between-trial comparisons.

Individual treatment plots in all trials consisted of seven 6.0-metre-long rows of wheat

oriented due West to East, with 0.20 m spacing between treatment rows, 1.6 m between

adjacent treatment plots, and 2.0 m between replicates.

Seed treatments. Seeds (hard red spring wheat: Syngenta, WR859CL) were treated

with a Hege 11 liquid seed treatment applicator (Wintersteiger Inc., Salt Lake City, UT)

by technicians at a Syngenta Crop Protection (Canada) seed treatment facility in Portage

la Prairie, Manitoba, with the following insecticides:

Trial 1: Cyantraniliprole and i-cyhalothrin: Cyantraniliprole (Fortenza 600FS) at 10,

20, 30, and 40 g AI/100 kg seed, A-cyhalothrin (Demand 100CS) at 30 g AI,

thiamethoxam (Cruiser 5FS) at 30 g AI, fipronil (Regent SOOFS) at 5 g AI, and the

Standard T+F Blend. All treatments also contained the fungicide Dividend XLRTA at 13

g Al (containing 3.21% difenoconazole and 0.27% mefenoxam).

Trial 2: Fipronil, alone and blended with thiamethoxam: Thiamethoxam (Cruiser

5FS) at 10 g AI/100 kg seed, fipronil (Regent SOOFS) at 0.6, 1, and 5 g AI, and blends of

thiamethoxam at 10 g AI + fipronil at 0.6, 1, and 5 g AI. All treatments also contained the

fungicides Proseed at 2.5 g AI (containing 40.3% fludioxonil) and Vibrance XL at 17.5 g

AI (containing 1.2% sedaxane, 5.9% difenoconazofe, and 1.5% metalaxyl-M).

Trial 3: Imidacloprid and thiamethoxam: \midacloprid (Stress Shield 480SC) at 10,

20, and 30 g AI, thiamethoxam (Cruiser SFS) at 10, 20, and 30 g AI, and Standard T+F

Blend. The imidacloprid treatments also contained the fungicide Raxil MD at 3.5 g Al;

all other treatments also contained the fungicides Proseed 480FS at 2.5 g and Vibrance

XL at 17.5 gAl.

Planting: All plots were planted on 8 May 2012 with a seven-row double disc drill,

no till planter (Fabro Enterprises Ltd., Swift Current, SK) directly into the wheat stubble

from the previous year’s crop. No tillage was done in the previous fall nor immediately

prior to planting. Seeds were planted approx. 2.5 cm deep, at 285 seeds/m?. As rows were

spaced 20 cm apart, this seeding rate was equivalent to approx. 57 seeds per 1 m of row,

or 100 kg seed/ha.

Stand assessment. Plant survival (hereafter “stand”) was determined by counting the

number of wheat seedlings alive in the central two-metre sections of the middle three

rows of each plot at 14 and 29 days after planting (DAP) (22 May and 6 June,

respectively) in all three trials, and measuring the plant reflective index (NDVI; Crop

Circle ACS-430, Holland Scientific, Lincoln NE) at 21, 29, and 37 DAP (29 May, 6 and

14 June, respectively).

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 89

Plot maintenance: Plots were kept weed free by treating with glyphosate on 4 May

prior to seeding, and no further weed control was deemed necessary. After harvest, the

remaining wheat stubble was left intact over winter to prevent disturbance of surviving

wireworm populations, which were assessed by trapping the following spring.

Harvest. All trials were harvested on 28 August 2012 (112 DAP) using a small plot

combine (Wintersteiger Inc., Salt Lake City, UT) that calculated both the moisture

percentage in the seed and per hectare yield. Some plots (e.g., neonicotinoid treatments

in Trial 3) were not harvested due to the lack of surviving plants.

Wireworm trapping. To determine the longer-term effects of the various treatments

on wireworm mortality, wireworms were sampled in the spring of the following year

using a bait-trapping procedure similar to that described in Vernon ef al. (2009). Bait

traps were installed in the plots (three per plot) on 1-2 May, 2013, and removed on 13

May. Bait trap locations were spaced | m apart along the middle of each plot, so that the

traps were 2, 3, and 4 m from the front and 75 cm from the outer rows of each plot. Each

bait trap consisted of a 450-ml plastic flower pot filled with coarse-grade vermiculite and

100 ml untreated hard red spring wheat placed in a layer in the middle of the pot. Traps

were soaked to run-off with lukewarm water twice several hours before placement in

circular holes (10 cm diameter, 15 cm deep) cored into the ground. Soil was carefully and

consistently packed around and on top of the bait traps, and a 20-centimetre-diameter

inverted tray placed 5 cm above the trap and level with the ground. To reduce variability

in data, considerable effort was taken to ensure each trap was prepared and installed

identically. After removal, bait traps were immediately transported to the Agassiz

Research and Development Centre (AAFC, Agassiz, BC), where they were placed in

Tullgren funnels on 15 May for 2 weeks to extract wireworms. Extracted wireworms

were counted, measured to the nearest millimetre, and identified to species. As

wireworms shrink when they desiccate after extraction, 200 living L. californicus larvae

were individually placed directly under the funnel heat source (25W incandescent light

bulbs) for 48 h, and measured and weighed to 0.1 mg (Sartorius CP64 analytic balance;

Sartorius AG, Goettingen, Germany) both before and after desiccation. Simple linear

regression of desiccated to living wireworm length yielded the relation, living length =

(desiccated length + 0.5391) / 0.6655; R* = 0.81, which was subsequently used to convert

the lengths of desiccated wireworms to the corresponding size of living ones. For

analyses, larval lengths were combined in three millimetre categories, since binning into

two millimetre categories or showing each size separately would produce artifacts due to

sizes calculated from desiccated lengths being rounded to the nearest | mm, which

causes underestimations of the number that were 6, 9, 12 mm, etc. long. Wireworms were

considered small, or neonate, if equal or less than 10 mm long, and large (or resident) if

greater than 10 mm.

Statistical Analysis. All data analyses were conducted using SAS (SAS 9.2, SAS

Institute, Cary, NC). Treatment means were compared by ANOVA (Proc GLM), followed

by mean separation with Tukey’s standardized range honestly significant difference

(HSD) test at a = 0.05. Where data could not be easily normalized using a power

transformation (Trial 3, reflective index and yield only), the Kruskal—Wallis test (Proc

NPAR1 WAY) was used, after which normalized rank values were assigned to treatments

(Proc RANK) and the standard ANOVA and the Tukey procedures performed on the rank

values. The relationship between the amount of stand reflectivity and plant stand counts

recorded on the same day was analyzed with linear regression (Proc GLM). Count data

were analyzed with chi-square tests (Proc FREQ).

RESULTS

Wireworm sampling

Post-treatment bait-trapping results confirmed the trial areas had very high wireworm

populations, with 403 larvae collected from the combined control treatments in the three

90 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

trials (12 plots, 36 traps), and 2,234 from the combined treated plots (88 plots). Of the

latter, only 190 wireworms were in plots with seeds treated with fipronil alone or in

blend with another insecticide (36 plots). Similar numbers were found in the control plots

of all three trials (range of means: 8.8—13.4/trap, Tables 1-3), suggesting a fairly

homogeneous population in the study area. Wireworm populations were predominantly

L. californicus (97.3%), with very low numbers of H. bicolor (2.0%), S. destructor

(0.7%), and Aeolus mellillus (Say) (<0.1%) — the latter species are included in the totals

presented in Tables 1-3. To compare the age structures of wireworm populations

retrieved from the various treatments, the distribution of larval lengths (range: 3—28 mm)

were compared for wireworms retrieved from control plots, plots seeded to treatments

containing fipronil, and plots seeded to treatments containing other insecticides (Fig. 1).

Chi-square analyses indicated significant differences in population structures (i.e., in the

relative number in each of the size classes), both between control and fipronil treatments

(Chi=1089.3, df=7, P<0.0001) and between control and other treatments (Chi=144.9,

df=7, P<0.0001). Comparison of the age structures (Fig. 1A—C) indicates control

treatments had significantly lower numbers of small (3-10 mm) wireworms per plot

(1.08) than fipronil (1.94) and non-fipronil (2.92) insecticide treatments (Chi=104.3,

df=1, P<0.0001; Chi=8.85, df=1, P=0.0016; respectively). In contrast, the control and

non-fipronil treatments had a similar number of large (~10 mm) wireworms per trap

(32.5 and 36.4, respectively per plot), while very low numbers (3.3 per plot) were

retrieved from treatments containing fipronil (Fig. 1A—C).

Relation between plant reflectivity and plant stand

A direct, highly significant relationship was observed between plant reflectivity index

(RI) and plant stand when the two were measured on the same day (29 DAP). This was

true for trials with cyantraniliprole (t=7.95, df=1,34, P<0.0001, R?=0.64), fipronil

(t=11.17, df=1,30, P<0.0001, R? = 0.80), and imidacloprid and thiamethoxam (t =7.06,

df=1,30, P<0.0001, R*=0.61), and indicates that plant RI is an acceptable metric for

assessing plant stand (e.g., at 37 DAP, when individual plant counts were not conducted).

Trial 1: Cyantraniliprole and 4-cyhalothrin

Stand protection and yield

Greatest stand protection was provided by fipronil at 5 g AI and Standard T+F Blend

treatments. These treatments had higher stand counts than the control at 14 DAP (1.55x)

and 29 DAP (6.03x, 5.61x, respectively). However, RI readings at 37 DAP indicate better

stand protection in fipronil (5 g AJ) than the Standard T+F Blend (2.08x vs 1.77x control,

respectively), which resulted in higher yields at harvest (respectively, 18.3 vs 11.5x the

control; Table 1). Thiamethoxam applied at 30 g AI provided good initial plant protection

(respectively, 1.54x and 3.14x control at 14 and 29 DAP), but the RI at 37 DAP and yield

at harvest were similar to control and significantly less than fipronil (5 g AI) and

Standard T+F Blend treatments. Similarly, A-cyhalothrin at 30 g AI provided stand

protection (respectively, 1.80x and 4.60x control at 14 and 29 DAP) that resulted in

similar yield to the Standard T+F Blend, but yield was significantly lower than observed

for fipronil at 5 g AI (Table 1).

Cyantraniliprole applied at 10-40 g AI provided stand protection equivalent to or

greater than thiamethoxam at 30 g AI (i.e., 1.60-1.80x control at 14 DAP, 3.02-3.85x

control at 29 DAP), which resulted in numerically higher yields (1.73-2.56x

_thiamethoxam). Stand protection with cyantraniliprole was equivalent to that provided by

A-cyhalothrin and fipronil (5 g AI) at 14 DAP, but this had diminished by 29 DAP

(0.50-0.64x fipronil), and the RI at 37 DAP and yields at harvest were significantly lower

than fipronil (5 g AI) (Table 1). There were no significant differences in stand protection

or yield between rates of cyantraniliprole (Table 1).

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

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94 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Wireworm survivorship

Significantly fewer large (>10 mm) wireworms were collected in bait traps in both

the fipronil (0.06x control) and Standard T+F Blend (0.21x control) treatments (Table 1),

indicating high mortality in these treatments. In contrast, there were no significant

reductions in large wireworms caught in the thiamethoxam (30 g AJ), A-cyhalothrin (30 g

AJ), and cyantraniliprole treatments relative to the control treatment (Table 1). Relatively

few small (neonate) wireworms were collected in all insecticide treatments, and this was

similar to numbers taken in the control treatment (Table 1).

Trial 2: Fipronil, alone and blended with thiamethoxam

Stand protection and yield

Higher stand protection was observed in fipronil (0.6, 1.0, and 5.0 g AI) treatments

relative to the untreated control (range: 1.25—1.62x stand at 14 DAP, 2.67—3.69x stand at

29 DAP) and the thiamethoxam (10 g AJ) treatments (1.08—1.39x stand at 14 DAP, 1.62—

2.24x stand at 29 DAP). Stand protection increased with the rate of fipronil applied. As in

the other trials, thiamethoxam failed to provide lasting plant protection, leading to very

low yields at harvest (Table 2). In contrast, all rates of fipronil provided significantly

higher yields than either the thiamethoxam or untreated control treatments (13.76—17.92x

control; Table 2). No significant differences in yield were observed between the fipronil

rates. Combining thiamethoxam at 10 g AI with fipronil at 0.6, 1.0, or 5 g AI provided

similar stand protection than the fipronil treatments alone at the same rates, and did not

significantly increase yields (13.31—19.11x control; Table 2).

Wireworm survivorship

Populations of large wireworms were significantly reduced in the fipronil (0.6, 1.0,

and 5.0 g Al) (range: 0.03—0.23x control) and combined thiamethoxam (10 g AJ) and

fipronil (0.6, 1.0, and 5.0 g AI) treatments (0.03—0.26x control) (Table 2). Mortality was

highest in treatments with the 5 g AI rate of fipronil. Although not statistically

significant, there was notably higher mortality in the Standard T+F Blend than in the

treatment with fipronil at 1 g AI alone (Table 2). In contrast, more (1.19x control) large

wireworms were collected from the thiamethoxam than the control treatment (Table 2).

Low and similar numbers of neonate wireworms were collected from all treatments.

Trial 3: Imidacloprid and thiamethoxam

Stand protection and yield

Both imidacloprid (10, 20, and 30 g AI) and thiamethoxam (10, 20, and 30 g Al)

provided initial stand protection (1.33—1.71x, 1.23-1.55x control at 14 DAP,

respectively; 2.44-3.45x, 1.31-2.58x control at 29 DAP). For each rate tested,

imidacloprid provided numerically greater protection than thiamethoxam, with protection

increasing with rate for both chemicals (Table 3). Stand protection disappeared after 37

DAP, leading to complete destruction of the plots and no harvestable plants.

Good initial plant protection was observed in the Standard T+F Blend (1.58x and

4.88x control at 14 and 29 DAP, respectively). The effect of fipronil in the Standard T+F

Blend was evident when compared to thiamethoxam applied alone at 10 g AI (1.28x and

3.74x thiamethoxam at 14 and 29 DAP, respectively). Plant stand protection in the

Standard T+F Blend persisted throughout the season, and this was the only treatment

with harvestable plants. Yields at harvest were similar to that observed for the same

treatment evaluated in the other two trials (respectively, 2305, 3542, and 2824 kg/ha,

Tables 1-3).

Wireworm survivorship

Populations of large wireworms were not reduced in any of the imidacloprid or

thiamethoxam treatments (range: 0.75—1.15x, 0.79-1.31x control, respectively), and

highest numbers were collected from plots seeded to the highest rates of these chemicals

(Table 3). In contrast, very low numbers of large wireworms (0.04x control) were

collected from the Standard T+F Blend treatment, indicating high mortality. Low and

similar numbers of neonate larvae were collected from all treatments (Table 3).

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 95

A. Control plots

Wireworms / plot

iW)

<5 5-7 . 8-10 11-13 14-16 17-19 20-22 23-25 es

Wireworms / plot

o |

<5 5-7 8-10 11-13 14-16 17-19 20-22 23-25 >25

C. Plots treated with other insecticides

Wo pan nn ene nn nn nn nn nn nn nn rn ne nnn nnn een neers

Wireworms / plot

14-16 17-19 20-22 23-25 >25

Wireworm length (mm)

Figure 1. Size distribution of wireworms (predominantly Limonius californicus) collected

from three insecticide efficacy trials conducted in Claresholm, Alberta. Mean (SD) number of

wireworms retrieved from bait traps placed in control plots (A., N = 12 plots), in plots treated

with fipronil alone or in blend with another insecticide (B., N = 36), and in plots treated with

an insecticide other than fipronil (C., N = 52). Note the differences in vertical axes between B

and A, C. )

96 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

DISCUSSION

Neonate versus resident wireworm mortality

The number of small (neonate) wireworms that would have been produced during this

study was low (approx. 10%) in all treatments relative to the number of large (resident)

wireworms that would have been present at the time of planting. This is in contrast to

field studies with A. obscurus in which higher numbers of neonates were trapped in

control plots and plots treated with neonicotinoids relative to fipronil-containing plots

(cf. Vernon et al. 2009). There are a number of possible reasons for the differences in

neonate catches between the previous and current studies. In the current study, plant

stand in some (e.g., control, neonicotinoids; Tables 1—3) treatments was poor to non-

existent, which would have reduced oviposition and food availability relative to

treatments with higher stands (e.g., fipronil-containing treatments). This is partially

substantiated by the cyantraniliprole treatments in Trial 1, where stand and yield were

higher than in the control treatment, and neonate numbers were numerically higher (4.0—

4.8 per plot) than in the other treatments (1.5—2.7 per plot) (Table 1). This also suggests

cyantraniliprole may not be lethal to neonate wireworms.

In plots containing fipronil, which had excellent stand protection, low neonate

numbers were likely due to the residual and toxic effect of this chemical. Numbers of

resident wireworms were also very low in these treatments, and fipronil has previously

been shown to be highly toxic to both resident and neonate A. obscurus (Vernon et al.

2009, 2013a, 2016). The effect of the pyrethroid, A-cyhalothrin, in reducing neonate

populations in the current study is more difficult to ascertain. Because stand protection

and yield were similar to the Standard T+F Blend, the reduction in resident populations

was not significantly different from thiamethoxam or the control (Table 1), and the

neonate numbers were low, it appears that A-cyhalothrin is persistent and toxic to this

stage and/or that the presence of this insecticide in plots reduced egg laying due to

repulsion of female beetles. We have previously shown that residues of another

pyrethroid, bifenthrin, are repulsive to A. obscurus larvae >200 d after an in-furrow

application to soil in potatoes (van Herk et a/. 2013). While the overall low number of

neonates in this study might be attributed to low click beetle emergence and egg-laying,

this typically occurs in fields treated with an insecticide (e.g., that induces prolonged

morbidity and prevents late-instar larvae from feeding sufficiently to pupate in the fall),

whereas no insecticides had been applied to the study field since approx. 2000 (T.J.

Labun, unpublished data).

It is interesting that the lack of food in certain plots did not appear to affect the

survival and retention of resident wireworms, with high numbers of larvae trapped from

plots with little or no plant survival (e.g., neonicotinoid treatments, Table 3). This

supports the concern that later instars of some pest species can survive with minimal food

for prolonged periods of time (Vernon and van Herk 2013). Also worth noting is that

none of the fungicide treatments used in these trials appeared to negatively affect

wireworm populations. This is consistent with results from lab and field studies with both

A. obscurus and L. canus LeC. (Vernon et al. 2009, 2013a; van Herk et al. 2008, 2015).

Crop protection vs. wireworm mortality, and benefits of blended treatments

The above results underscore the importance of evaluating wireworm mortality

(inferred here from the difference in wireworm numbers collected from treatment vs

control plots) in field efficacy studies. While wireworm mortality could be deduced from

crop protection in earlier insecticide efficacy studies with OP and OC insecticides, this is

usually not possible with newer chemistries (Vernon et al. 2009), as exposure to

neonicotinoid insecticides generally induces prolonged, reversible morbidity during

which time wireworms are unable to feed (Vernon et a/. 2008). Hence, these insecticides

may protect plants from feeding damage without decreasing wireworm populations

(Vernon et al. 2009, 2013a). A similar result was seen in efficacy studies with potatoes,

where neonicotinoid treatments applied at planting reduced feeding damage to daughter

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 97

tubers without decreasing wireworm numbers (Vernon ef al. 2013b). Pyrethroid

insecticides also protect wheat and potatoes from wireworm feeding damage without

reducing populations, but here the mechanism is mainly repellency (van Herk ef al. 2008,

2015). Conversely, exposure to an insecticide that induces morbidity and mortality

latently can result in wireworm population reductions without providing adequate stand

protection (Vernon ef al. 2013a).

Contrary to results with A. obscurus in BC, high rates of imidacloprid and

thiamethoxam failed to protect wheat seedlings from L. californicus past 29 DAP in these

trials. This could result from differences in insecticide susceptibility between species or

from the very high wireworm populations in the field. In southern Alberta, high

populations of L. californicus can cause complete crop destruction in fields of spring

wheat treated with a high (39 g AI) rate of thiamethoxam (T.J. Labun, personal

observation). The observed failure of high rates of these commonly used insecticides to

reduce populations of L. californicus is similar to findings by Esser et al. (2015) with L.

californicus and L. infuscatus Mots., and likely explains why damage in wheat from

these species is increasing in severity and frequency across the region.

Both cyantraniliprole and A-cyhalothrin provided greater protection at the rates tested

than either imidacloprid or thiamethoxam, although this was likely through different

mechanisms. While A-cyhalothrin and other pyrethroids (e.g., tefluthrin, bifenthrin)

induce repellency and thereby reduce feeding (van Herk ef al. 2008, 2015),

cyantraniliprole is not repulsive and likely induces morbidity after feeding (van Herk et

al. 2015). Considering the high wireworm populations in these trials, the partial plant

protection observed is encouraging, and cyantraniliprole may be a potential candidate for

blending with low rates of a lethal insecticide. It should be noted that at the rates tested,

cyantraniliprole and A-cyhalothrin by themselves did not cause significant wireworm

mortality in either this study or in previous work with A. obscurus (Vernon et al. 2013b;

van Herk et al. 2015).

Combining a non-lethal insecticide that rapidly induces morbidity with a low rate of a

chemical that causes mortality latently can provide both stand protection and long-term

population reductions in the field (Vernon ef a/. 2013a). Since wireworms live for up to

4—5 years in the soil, one application with an insecticide lethal to all wireworm stages can

remove the economic threat of wireworms for three or more years. This blended

treatment concept was evaluated in numerous lab and field studies with A. obscurus, A.

sputator, and L. canus, which demonstrated that combinations of thiamethoxam at 5 or 10

g Al with fipronil at rates as low as | g AI will provide both acceptable crop protection

and high neonate and resident wireworm mortality for these species (Vernon ef al. 2009,

2013a). These results provided the basis for the current study with L. californicus and

allowed the concept to be extended to using insecticide-blended wheat seed as an in-

furrow treatment that both protects potato tubers from damage and reduces wireworm

populations (Vernon ef al. 2016).

In the work reported here, both the fipronil and various thiamethoxam + fipronil

blend treatments provided significant stand protection and reduction in populations of

resident wireworms, relative to the untreated control and all other treatments tested. Of

note is that, in Trial 2, combining thiamethoxam at 10 g AI with fipronil at 0.6, 1.0, and

5.0 g AI did not improve stand protection and yield, nor increase resident wireworm

mortality relative to the corresponding fipronil treatments. This suggests that L.

californicus may respond differently to neonicotinoid and fipronil insecticide blends than

A. obscurus, where the presence of thiamethoxam considerably improved stand and yield

(Vernon et al. 2013a). Also of note is that stand, yield, and mortality were notably higher

at the 5.0 g than 1.0 g and 0.6 g AI rates of fipronil. Similarly, in Trial 1, fipronil at 5 g Al

provided 1.6x greater yield and 3.6x higher mortality than the Standard T+F Blend. This

suggests that where fipronil is used alone as a seed treatment to control high populations

of L. californicus, it should be applied at a rate higher than | g AI, and that (unlike for A.

98 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

obscurus) there is no additional benefit from combining fipronil with a neonicotinoid

such as thiamethoxam.

Neonicotinoid and fipronil insecticide blends on wheat seed have been evaluated for

wireworm management elsewhere. Morales-Rodriguez and Wanner (2015) observed high

(>70%) mortality in L. californicus and H. bicolor exposed in laboratory assays to wheat

seed treated with fipronil at 1 and 5 g AI/100 kg seed but low mortality (<30%) if

exposed to thiamethoxam at 39 g AI. In field trials, seed treated with thiamethoxam at 39

g AI provided plant protection but resulted in higher wireworm populations than control

plots, while seed treated with both thiamethoxam at 39 g AI and fipronil at 5 g Al

significantly reduced populations. Combining thiamethoxam at 39 g AI with fipronil at 1

g AI/100 kg seed caused less mortality in lab studies than either insecticide alone, and we

suggest that the high rate of thiamethoxam in this blend may have induced morbidity

before sufficient fipronil was ingested. Higher rates of thiamethoxam decrease the

duration of feeding in L. canus (van Herk et al. 2008), and in lab studies mortality is

greater when wireworms are exposed to fipronil at 1 g AI alone than in combination with

thiamethoxam at 10 g AI (van Herk et al. 2015). However, when larvae were exposed to

a blend of thiamethoxam at 10 g AI and a higher rate of fipronil (e.g., 5 g AI), enough of

the latter chemical was ingested to cause high mortality (van Herk ef al. 2015). Under

field conditions, high mortality of A. obscurus was observed with blends of

thiamethoxam at 5 or 10 g AI and fipronil at both 1 and 5 g AI (Vernon ef al. 2013b),

likely because of longer exposure to the seeds than in laboratory studies and because

other factors (1.e., desiccation, predation on moribund wireworms) contribute to mortality

in the field (Vernon et al. 2009).

Potential of seed treatments for controlling wireworms in cereals

In a recent review of insecticides for controlling wireworms in cereals, it was

observed that, in general, the most effective chemistries appear to be those that target

GABA-gated chloride channels (e.g., fipronil, lindane) (van Herk ef al. 2015). As noted

by Lange et al. (1949), the efficacy of seed treatments also depends on “the species of

wireworms involved, wireworm activity at the time the seed is planted, the proportion of

the population attracted to the seed, the type of seed, and the time of planting.” Some of

these observations are briefly considered here.

Time of planting and wireworm activity

Seed treatments are most likely to be effective when seed is planted shortly before

larvae become active (Vernon and van Herk 2013). Many pest wireworm species have

two main periods of feeding activity (spring and fall), between which they burrow

downwards to avoid desiccation (Traugott et al. 2015). Planting seed treated with a non-

residual insecticide after wireworms have fed would therefore reduce exposure and

resultant mortality. This would be a concern where cropping practices (e.g., continuous

cropping, minimal tillage) provide alternative food sources before or after the seeds are

planted (e.g., roots and decaying plant matter from the previous year’s crop). Under these

conditions, wireworms would presumably feed less on the treated seeds, if at all, and

therefore ingest less insecticide (Vernon ef a/. 2013b). Early season planting, before

wireworms become active in the spring, may not be feasible, as wireworms can cause

considerable feeding damage even at low soil temperatures (van Herk and Vernon 2013).

Determining when wireworms become active in the spring has been the focus of

considerable research (reviewed in Traugott et al. 2015 and Vernon and van Herk 2013),

and the high mortality observed in the fipronil treatments reported here suggests the

spring activity period of L. californicus coincides with spring wheat planting in southern

Alberta.

Differences between species

Insecticide seed treatment efficacy may vary between wireworm species due to

differences in species phenology (e.g., when they begin to feed) and different

susceptibilities to insecticides (Vernon et al. 2008). Lange et al. (1949) noted that L.

canus is more susceptible to lindane than L. californicus, possibly because of differences

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 99

in the activity levels of these species. In eastern Washington State, repeated exposure to

thiamethoxam-treated spring wheat resulted in no observed changes in populations of L.

californicus, whereas at a nearby site it appeared to reduce L. infuscatus populations

(Esser et al. 2015; Milosavljevic et al. 2016). Hence, it is critically important to know

what species are present in the field before applying a management approach,

particularly as pest species frequently co-occur.

Differences between cereals

In laboratory studies, Edwards and Evans (1950) observed no difference in wheat and

oat (Avena sativa L.) seedling survival when exposed to Corymbites cupreus Fabr.,

Agriotes spp., or Athous (=Hemicrepidius) niger L. larvae, but slightly higher survival of

barley (Hordeum vulgare L.) than wheat and oat seedlings exposed to Agriotes spp. and

C. cupreus. In contrast, recent work suggests both oat and barley seedlings may be less

susceptible to L. infuscatus and L. californicus feeding (respectively) than wheat

(Higginbotham et al. 2014, Rashed et al. 2017). Recent field studies in Alberta suggest

insecticides (e.g., fipronil) applied on barley cause lower mortality in L. californicus than

when applied to spring wheat seed (van Herk ef a/., unpublished data). This may be due

to the barley seed hull absorbing some of the seed dressing, or to the susceptibility of the

seed itself to wireworm feeding (cf. Higginbotham ef a/. 2014). While more data is

required to determine if these results are real or result from the usual sources of

variability that plague wireworm field studies (e.g., patchy distributions in the field),

insecticides used as seed treatments may need to be applied at higher rates on barley than

wheat to achieve the same level of population reduction, but at lower rates to achieve the

same level of stand protection.

ACKNOWLEDGEMENTS

This work would not have been possible without the expert assistance of Joshua Spies

and crew from Syngenta who assisted with planting, plot maintenance, harvesting, and

bait trapping, and of the summer students of RSV and WVH who processed wireworms

extracted from bait traps.

REFERENCES

Arnason, A.P. 1931. A morphological study of the immature states of Cryptohypnus nocturnus

Eschscholtz and a study of some ecological factors concerning wireworms. M.S. thesis, University

of Saskatchewan, Saskatoon.

Arnason, A.P. and Fox, W.B. 1948. Wireworm control in the prairie provinces. Dominion of Canada,

Science Service, Division of Entomology, Publication No. 111, Ottawa.

Burrage, R.H. 1964. Trends in damage by wireworms (Coleoptera: Elateridae) in grain crops in

Saskatchewan, 1954-1961. Canadian Journal of Plant Science, 44: 515-519.

Doane, J.F. 1977. The flat wireworm, Aeolus mellillus: studies on seasonal occurrence of adults and

incidence of the larvae in the wireworm complex attacking wheat in Saskatchewan. Environmental

Entomology, 6: 818-822.

Edwards, E.E. and Evans, J.R. 1950. Observations on the biology of Corymbites cupreus F. (Coleoptera,

Elateridae). Annals of Applied Biology, 37: 249-259.

Esser, A.D., Milosavljevié, I., and Crowder, D.W. 2015. Effects of neonicotinoids and crop rotation for

managing wireworms in wheat crops. Journal of Economic Entomology, 108:1786—1794.

Grove, I.G., Woods, S.R., and Haydock, P.P.J. 2000. Toxicity of 1, 3—dichloropropene and fosthiazate to

wireworms (Agriotes spp.). Annals of Applied Biology, 137: 1-6.

Higginbotham, R.W., Froese, P.S., and Carter, A.H. 2014. Tolerance of wheat (Poales: Poaceae) seedlings

to wireworm (Coleoptera: Elateridae). Journal of Economic Entomology, 107: 833-837.

Lange Jr, W.H., Carlson, E.C., and Leach, L.D. 1949. Seed treatments for wireworm control with

particular reference to the use of lindane. Journal of Economic Entomology, 42: 942-955.

MacNay, C.G. 1954. New records of insects in Canada in 1952: a review. The Canadian Entomologist,

86: 55—60.

100 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

Milosavljevic, I., Esser, A.D., and Crowder, D.W. 2016. Effects of environmental and agronomic factors

on soil-dwelling pest communities in cereal crops. Agriculture Ecosystems and Environment, 225:

192-198.

Morales—Rodriguez, A. and Wanner, K.W. 2015. Efficacy of thiamethoxam and fipronil, applied alone

and in combination, to control Limonius californicus and Hypnoidus bicolor (Coleoptera:

Elateridae). Pest Management Science, 71: 584-591.

Rashed, A., Rogers, C.W., Rashidi, M., and Marshall, J.M. 2017. Sugar beet wireworm Limonius

californicus damage to wheat and barley: evaluations of plant damage with respect to soil media,

seeding depth, and diatomaceous earth application. Arthropod Plant Interactions, 11: 147-154.

Stone, M.W. 1941. Life history of the sugarbeet wireworm in southern California. USDA Technical

Bulletin No. 744.

Strickland, E.H. 1927. Wireworms of Alberta, a preliminary report. University of Alberta, Bulletin 2,

Edmonton, Alberta.

Toba, H.H., O'Keeffe, L.E., Pike, K.S., Perkins, E.A., and Miller, J.C. 1985. Lindane seed treatment for

control of wireworms (Coleoptera: Elateridae) on wheat in the Pacific Northwest. Crop Protection, 4:

372-380.

Toba, H.H., Pike, K.S., and O’Keeffe, L.E. 1988. Carbosulfan, fonofos, and lindane wheat seed

treatments for control of sugarbeet wireworm. Journal of Agricultural Entomology, 5: 35-43.

Traugott, M., Benefer, C.M., Blackshaw, R.P., van Herk, W.G., and Vernon, R.S. 2015. Biology, ecology,

and control of elaterid beetles in agricultural land. Annual Review of Entomology, 60: 313-334.

van Herk, W.G. and Vernon, R.S. 2013. Wireworm damage to wheat seedlings: effect of temperature and

wireworm state. Journal of Pest Science, 86: 63-75.

van Herk, W.G. and Vernon, R.S. 2014. Click beetles and wireworms (Coleoptera: Elateridae) of Alberta,

Saskatchewan, and Manitoba. Jn Arthropods of Canadian Grasslands, vol. 4. Edited by D.J. Giberson

D. J. and H.A. Carcamo. Biological Survey of Canada, Ottawa. Pp. 87-117.

van Herk, W.G., Vernon, R.S., Moffat, C., and Harding, C. 2008. Response of the Pacific Coast

wireworm, Limonius canus, and the dusky wireworm, Agriotes obscurus (Coleoptera: Elateridae), to

insecticide-treated wheat seeds in a soil bioassay. Phytoprotection, 89: 7-19.

van Herk, W.G., Vernon, R.S., and McGinnis, S. 2013. Response of the dusky wireworm, Agriotes

obscurus (Coleoptera: Elateridae), to residual levels of bifenthrin in field soil. Journal of Pest

Science, 86: 125-136.

van Herk, W.G., Vernon, R.S., Vojtko, B., Snow, S., Fortier, J., and Fortin, C. 2015. Contact behaviour

and mortality of wireworms exposed to six classes of insecticide applied to wheat seed. Journal of

Pest Science, 88: 717-739.

Vernon, R.S. and van Herk, W.G. 2013. Wireworms as pests of potato. Jn Insect pests of potato: global

perspectives on biology and management. Edited by A. Alyokhin C. Vincent, and P. Giordanengo.

Academic Press, Amsterdam, the Netherlands. Pp. 103-164.

Vernon, R.S., van Herk, W.G., Tolman, J., Ortiz Saavedra, H., Clodius, M., and Gage, B. 2008.

Transitional sublethal and lethal effects of insecticides following dermal exposures to five economic

species of wireworms (Coleoptera: Elateridae). Journal of Economic Entomology, 101: 367-374.

Vernon, R.S., van Herk, W.G., Clodius, M., and Harding, C. 2009. Wireworm management I: stand

protection versus wireworm mortality with wheat seed treatments. Journal of Economic Entomology,

102: 2126-2136.

Vernon R.S., van Herk, W.G., Clodius, M., and Harding, C. 2013a. Crop protection and mortality of

Agriotes obscurus wireworms with blended insecticidal wheat seed treatments. Journal of Pest

Science, 86: 137-150.

Vernon, R.S., van Herk, W.G., Clodius, M., and Harding, C. 2013b. Further studies on wireworm

management in Canada: damage protection versus wireworm mortality in potatoes. Journal of

Economic Entomology, 106: 786—799.

Vernon, R.S., van Herk, W.G., Clodius, M., and Tolman, J. 2016. Companion planting attract-and-kill

method for wireworm management in potatoes. Journal of Pest Science, 89: 375-389.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 101

SCIENTIFIC NOTE

A pheromone-baited pitfall trap for monitoring Agriotes

spp. click beetles (Coleoptera: Elateridae) and other soil-

surface insects

W.G. VAN HERK!, R.S. VERNON’, and J.H. BORDEN?

Pheromone traps have been developed specifically for the survey, research and

management of click beetles (Coleoptera: Elateridae) in temperate North America (NA),

Europe and Asia (Ritter and Richter 2013; Vernon and van Herk 2013; Traugott ef al.

2015). These include: ‘Estron Traps’ for survey of Agriotes species in the former USSR

(Oleshchenko et al. 1987); ‘Yatlor Traps’ for survey and scientific study of Agriotes in

Europe (Furlan et al. 2001), and; ‘Vernon Beetle Traps’ for survey and integrated pest

management (IPM) of invasive Agriotes in NA, 1.e., A. obscurus (AO), A. lineatus (AL)

and A. sputator (AS; Vernon 2004). Although effective, these traps are no longer

available commercially, although the Yatlor Trap has been re-designed as a funnel trap to

better intercept various flying Agriotes in Europe (Csalomon, Budapest, Hungary). The

loss of the Vernon Beetle Trap (VBT) and customized lures for AO, AL and AS [formerly

produced by Contech Enterprises Inc., Delta, British Columbia (BC), Canada]

necessitated the development of a new trap for use in Agriotes IPM program

development in Canada. Based on the authors’ experience with earlier Agriotes traps, the

new trap was designed to: provide trapping efficacy comparable to the VBT; reduce the

time required for assembly, installation and inspection; exclude insectivorous vertebrates

and water, and; be consistent, reliable, inexpensive, small, easy to transport, and durable.

The new trap, named the Vernon Pitfall Trap® (VPT) (Fig. 1), is constructed of

durable polypropylene, and is formed from three custom injection molds (Exact Molds

Ltd, Abbotsford, BC). Two essential components are an in-ground pitfall chamber for

specimen collection (Fig. 1A) and a protective cover containing a pheromone-bait holder

and vertebrate-exclusion cage (Fig. 1B). The pitfall chamber forms a tapered cup that is

10 cm high from base to apex of the trap, with a 5.8-cm-diameter base (inside diameter,

ID) and a 9-cm-diameter opening (ID) (Fig. 1A). The inside of the cup, three centimetres

from the apex, is molded to receive a commercially available specimen cup (specifically,

Fisherbrand™ 4.5-o0z. Polypropylene Graduated Specimen Container). These removable

containers, which can be filled with a preserving liquid such as propylene glycol or used

without, and accompanying lids are used for labelling and storing collected specimens.

Surrounding the apex of the chamber is a rounded collar that slopes gradually away from

the opening (3 cm outward and 1 cm downward), with a steeper decline 0.5 cm from the

outermost edge. The collar has raised ridges (0.1 mm high) spaced 1—2 mm apart to

enable climbing by walking insects (Fig. 1A and D). Beneath the collar are four evenly

spaced supports that link the collar to the chamber to provide rigid stability to the trap. At

the apex of the collar are two 1.2-cm-diameter (outside diameter, OD) x 2-cm-high

hollow wells, spaced 8.5 cm apart, which receive and secure the trap lid (Fig. 1A and D).

The shape of the pitfall chamber is similar to typical hand-held or upright bulb planters,

which can be used to quickly remove exact soil cores for tight trap insertion. Moreover,

overlapping traps can be conveniently stacked for transport. When the base is inserted

into the cored soil, foot pressure on the reinforced collar seals the base tightly to the

' Corresponding author: Agassiz Research and Development Centre, Agriculture and Agri-Food Canada,

P.O. Box 1000, Agassiz, B.C., Canada VOM 1A0; wim.vanherk@canada.ca

2 Sentinel IPM Consulting, 4430 Estate Drive, Chilliwack, B.C., Canada V2R 3B5

3 JHB Consulting, 6552 Carnegie St., Burnaby, B.C. Canada V5B 1Y3

102 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

ground (Fig. 1D). This process does not require the clearing of surface grass or

excavation, as is typically required for other pitfall traps, and the raised collar helps

reduce water entry into the base.

Figure 1. Views of Vernon Pitfall Trap®, showing the bottom, pitfall component (A), the

underside of the cover with inserted lure and vertebrate exclusion fence (B), the assembled

trap (C), the trap installed in soil with collected A. sputator (D), the optional cover for

winterizing (E), and the assembled trap with winterizing cover (F). Photo credits: Warren

Wong. Individual, high-resolution images are available as supplementary files on the journal

website.

The second component is an easily detached cover, 16.5 cm in diameter, that tapers

slightly downward as a shallow cone 0.5 cm from base to apex to shed rain (Fig. 1B).

The cover contains a circular (1.2 cm diameter by 3 cm high, OD) downward-projecting

well that is centred on the underside to hold 0.75-cm-diameter cylindrical Agriotes spp.

pheromone baits (AO, AL and AS lures available from Csalomon, Budapest, Hungary),

and two pegs (0.75-cm-diameter (OD) by 3 cm high) that are located 8.8 cm apart to fit

into the corresponding wells on the base (Fig. 1A, B, and C). On the underside of the lid

(Fig. 1B), six evenly spaced supports (0.5 cm high) that radiate from the projected

pheromone lure well to the outside of the lid provide stability and prevent warping of the

cover. To exclude insectivorous vertebrates (e.g., mice, voles, shrews, snakes) a circular

fence of downward-projecting pins (3 mm diameter) is present on the underside of the

lid. The pins are spaced 0.5 cm apart and range in length from 2.5 cm (30 pins) to 3 cm

(4 pins). The longest pins just touch the base’s collar section at four sites when base and

lid are joined, lending stability to the assembled trap (Fig. 1C). The shorter pins leave a

0.5-cm-high passage above the collar to permit entry by click beetles or other walking

insects.

The traps are manufactured in brown (used in Canada for AL), black (used for AO)

and green (used for AS) to help avoid pheromone cross-contamination between species.

An optional trap component is a 16.5-cm-diameter winter lid (Fig. 1E) that replaces

the main lid at the end of the trapping season. The winter lid is designed to snugly fit

flush with the trap base (Fig. | F), so that the trap can remain in situ overwinter,

protected from entry of debris, insects and water.

Should a need arise in the future, the cover’s downward-projecting well for holding

Agriotes pheromone lures could be replaced by a ring for hanging lures or a removable

lure-holding basket, as in the Unitrap, for deploying lures for other target soil-surface

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 103

species. This would require the mold to be restructured. Alternatively, such lure-holders

could be constructed as separate components that fit into the well.

The new trap offers a number of improvements to monitoring AO, AL and AS,

relative to the former Vernon Beetle Trap. The VPT requires less time to assemble, install

and inspect, and is more durable and transportable than the VBT. The VPT is similar to

the VBT in catch of AO, AL and AS (van Herk, unpublished data). It has proven highly

effective, with or without pheromone baits, in monitoring programs for AO and AL in BC

and for AS in Prince Edward Island (PEI) (Table 1). The highest catch recorded to date

for a single trap is 6,955 AS over a 5-day period in Orwell, PEI (27 May—1 June, 2015)

(Fig. 1D). The trap has also been used, without pheromone baits, to successfully trap

other elaterid species in other provinces of Canada, including A. mancus (Say), Aeolus

mellillus (Say), Hypnoidus abbreviatus (Say), H. bicolor (Eschscholtz), Limonius

californicus (Mannerheim), Melanotus communis (Gyllenhal), and Selatosomus

destructor (Brown) (van Herk, unpublished data). It has also been used successfully to

trap other walking insects, including carabids and weevils (e.g., Sitona lineatus L.; St.

Onge et al. 2018).

Table 1

Catch of three Agriotes species in baited versus unbaited Vernon Pitfall Traps® (VPT) in field

as die in BC (AO and AL) and PEI (AS). N = number of traps.

|Year | Agriotes ' Trapping period _| Baited VPT | Unbaited VPT

| | Species’ | nee |

| od i sao toca -|N |Mean(SD) N | Mean (SD)

2015 | AO | 26 Mar-16 July | 22 977.7 (451.5) | 33 | 10.2 (12.2)

2016 | AL | 21 Mar-11 July 22 | 171.0 (92.7) 33 0.8 (1.0) |

2015 [aS | 20May-13 Aug | 44 | 7,797 1 (2,783.9) 38 | 71.6 (53.0) |

TAO = “A. obscurus; AL= = A. lineatus; AS = =A. Kewanee

REFERENCES

Furlan, L., Toth, M., Parker, W.E., Ivezic, M., Panéi¢, S., Brmez, M., Dobrincic, R., Baréic, J.I.,

Muresan, F., Subchev, M., Toshova, T., Molnar, Z., Ditsch B., and Voigt, D. 2001. The efficacy of the

new Agriotes sex pheromone traps in detecting wireworm population levels in different European

countries. Jn Proceedings of XXI IWGO Conference (Venice, Italy). Edited by D. Carollo. Veneto

Agricollo, Legonaro, Italy. Pp.293-—303.

Oleshchenko, I. N., Ismailov, V.Y., Soone, J.H., Laats, K.V., and Kudryavtsev, I.B. 1987. A trap for pests

(in Russian). USSR Author’s Cer. No. 1233312. Byulleten' Izobretenii, 11: 299.

Ritter, C. and Richter, E. 2013. Control methods and monitoring of Agriotes wireworms (Coleoptera:

Elateridae). Journal of Plant Disease Protection, 120: 4-15.

St. Onge, A., Carcamo, H.A., and Evenden, M.L. 2018. Evaluation of semiochemical-baited traps for

monitoring the pea leaf weevil, Sitona lineatus (Coleoptera: Curculionidae) in field pea crops.

Environmental Entomology, 47: 93-106

Traugott, M., Benefer, C.M., Blackshaw, R.P., van Herk, W.G., and Vernon, R.S. 2015. Biology, ecology

and control of Elaterid beetles (in agricultural land). Annual Review of Entomology, 60: 313-34.

Vernon, R.S. 2004. A ground-based pheromone trap for monitoring Agriotes lineatus and A. obscurus

(Coleoptera: Elateridae). Journal of the Entomological Society of British Columbia, 101: 141-142.

Vernon, R.S. and van Herk, W.G. 2013. Wireworms as pests of potato. Jn Insect Pests of Potato: Global

Perspectives on Biology and Management. Edited by P. Giordanengo, C. Vincent, and A. Alyokin.

Academic Press, Amsterdam, The Netherlands. Pp. 103-164.

104 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

SCIENTIFIC NOTE

Identifying larval stages of Orgyia antiqua (Lepidoptera:

Erebidae) from British Columbia, Canada

BRIAN VAN HEZEWLJK!', JESSICA MACLEAN|!, and RACHEL MCMAHON!

The rusty tussock moth, Orgyia antiqua (Linnaeus, 1758) (Lepidoptera: Erebidae) 1s

being used as an ecological surrogate to measure the impact of native natural enemies on

the establishment of European gypsy moth, Lymantria dispar (Linnaeus, 1758), in British

Columbia, Canada. To measure stage-specific mortality rates, one must be able to

identify accurately different life stages of the species under study, ideally with

characteristics that can be used in the field. The existing literature describing the number,

size, and colouration of larval instars for O. antiqua is highly inconsistent (Table 1). The

number of reported larval instars varies from 5—6 in males and 5—7 in females. Only two

papers report the width of larval head capsules, with substantial disagreement between

them (Dyer 1893; Payne 1917). Later instars of O. antiqua are characterised by dense

tufts of setae on the dorsal surface of segments 4—7. These have been variously described

as white, yellow, rusty brown, dark grey or black, and have been proposed by some

authors to be aposematic warnings (Sandre et al. 2007a) that vary among instars.

Through careful rearing of individual larvae and consistent measurements of head

capsule width, we sought to clarify the number of larval instars and identify unique

morphological characters that would facilitate the determination of instar in the field.

Table 1

Published descriptions of larval O. antiqua with respect to the colour patterns of the four

dorsal tufts and the corresponding head capsule widths. Listed colours should be read as tuft

colour from anterior to posterior starting on the first abdominal segment. B=Black, Y=Yellow,

W=White, G=Grey, Br=Brown, ?=not reported.

Dyer 1893 ~Gentner Hardy 1945 Payne 1917 Sandre Sandre

eg ss 2007a 2007b

Instar Colour Pattern of Dorsal Tufts

3 B-B-Y-Y G-G-W-W _B-B-W-W _G-G-W-W —_?-?-?-? 2-2-2-?

4 B-B-Y-Y 9-2-Y-Y Y-Y-Y-Y G-G-Y-Y B-B-Y-Y B-B-Y-Y *

(Pied)

» B-B-Y-Y W-W-W-W — ?-?-2-2 W-W-W-W_ Y-Y-Y-Y Y-Y-Y-Y *

(Bright)

6 W-W-W-W_ - 7 W-W-W-W _ Br-Br-Br-Br_ Br-Br-Br-Br

(Dull) _

qs W-W-W-W> - - - ~

Head Capsule Width (mm)

1 0.55 0.518 -

0.3317

2 OFF5 0.812 -

| 0.875

3 ey 1161.35

4 |e 1.80 - 2.02

5 2.1 2.24 - 2.64

6 - 3.0 - 3.5

7 ~

*Sandre 2007b reports that this is the typical pattern but that “nearly all other combinations were also present.”

| Natural Resources Canada, Canadian Forest Service, Pacific Forestry Centre, 506 W Burnside Road,

Victoria, British Columbia V8Z 1M5, Canada; brian. vanhezewijk@canada.ca

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 105

In May 2017, 33 O. antigua egg masses were collected from a small population in

Burnaby, BC, Canada (49.258821 N, 123.009661 W), that was feeding on an isolated

Colorado spruce (Picea pungens Engelm) planted as a landscaping tree. Larvae were

reared for one generation on Alnus rubra. In May 2018, 40 newly eclosed larvae from the

second generation were reared individually in 50-mm plastic Petri dishes (20°C, 18L:6D)

and fed fresh foliage of locally collected Himalayan blackberry (Rubus armeniacus)

every 1—3 days. Head capsule widths for each larval instar were measured, using a Leica

MSS dissecting microscope with an ocular micrometer with a precision of 0.012 mm, on

live larvae that had been chilled for approximately 10 minutes at 5°C. Shed head

capsules were retained for each individual larva in order to confirm the number of

moults.

In a separate trial, a small sample of 10 newly eclosed O. antiqua larvae were reared

on coastal Douglas-fir (Pseudotsuga menziesii var. menziesii) foliage to determine if they

could complete development on this host. The head capsule widths for these larvae were

measured for 4" and successive instars only. Representative photographs were taken of

individual larvae from each instar using a Nikon D7000 digital camera equipped with a

Nikon Speedlight SB-700 flash unit. After pupation and subsequent emergence, the

gender of adults was recorded.

Of the 40 larvae reared on blackberry, eight died of unknown causes before the 3"4

instar and were excluded from the analysis. Males (n = 17) invariably had five instars,

whereas females (n = 15) typically had six instars, with the exception of one female that

pupated after the 5" instar. For the first four instars, the head capsule widths of the larvae

grew exponentially, closely following Dyar’s rule (Fig. 1). For male and female larvae,

5th instar head capsules were smaller than expected, based on the progression of the first

four instars. Similarly, head capsules of 6 instar females were also smaller than

expected. There was no overlap in the head capsule widths of successive instars for either

sex through 1‘ to 6" instar (Fig. 2), and our measurements closely matched those of

Payne (1917). Larvae reared on Douglas-fir foliage had very similar head capsule widths

to those reared on blackberry (Fig. 2). Consistent with the blackberry-reared larvae, the

male larvae reared on Douglas-fir (n=5) had five instars, and the female larvae (n=5)

predominantly had six instars with the exception of one female, which pupated following

the 5" instar.

The morphological appearance of the first three instars closely matched the

descriptions previously published in the literature (Table 1, Fig 3). First instar larvae are

characterised by the absence of orange tubercles on the 6" and 7" abdominal segments.

These tubercles are present in the 2"¢ instar larvae, but this stage lacks lateral pencils on

the 1’ thoracic segment. The 3" instar is characterised by distinct lateral pencils on the

1st thoracic segment, as well as by the appearance of dorsal tufts on the 1‘ to 4%

abdominal segments. We found that the dorsal tufts of 4 instar larvae always had a

“pied” (Table 1) or two-toned colouration that varied considerably between individuals

(e.g., larvae 6 and 7 in Fig. 3). Fifth instar males had four monochromatic tufts that

ranged in colour from white to yellow to a rusty brown. In females that had six instars,

the 5 instar was pied (e.g., Fig. 3, larvae 6 and 28). Sixth instar females looked the same

as 5‘ instar males, with four monochromatic tufts ranging from white to yellow and rusty

brown.

In conclusion, it is difficult to unambiguously discriminate between 4", 5", and 6%

instars in the field based solely on the colouration of the dorsal tufts. An individual with

tufts of different colours could be a 4" instar of either sex or a 5" instar female that will

eventually moult into a 6" instar. An individual with tufts of a single colour could be a 5"

instar of either sex or a 6"" instar female. Head capsule width, however, could be used to

discriminate unambiguously between each of the instars. In our sample, there was no

overlap in the size distributions for each instar, even when we reared the larvae on

Douglas-fir, which we considered a sub-optimal host based on previously observed

slower growth rates.

106 J. ENTOMOL. Soc. BRIT. COLUMBIA 115, DECEMBER 2018

It is interesting to note that when females had an ‘extra’ instar, it was not a typical

supernumerary instar as has been reported in other lepidopteran larvae (Leonard 1970;

Retnakaran, 1973; Hatakoshi et al. 1988) but rather a repetition of the 4" instar; the

additional instar was morphologically similar to the 4" instar, only larger. Only one of

the 15 females reared on blackberry did not have a 6" instar and that individual had the

largest head capsule of all the female larvae from the 3" to 5" instars. This suggests to us

that the physiological trigger for an extra instar is related to size and that this is triggered

at some point during or before the 4" instar.

4.0

3.5

3.0 : |

So. +F

25 ran

or,

E 2.0

= hs

eee?” So

= |

= 15

0.5 A

1 2 3 4 5 6

instar

Figure 1. Average head capsule widths for female (circles) and male (triangles) Orgyia

antiqua larvae according to instar number. The linear regression line was fitted to the first

four instars only as the head capsule widths for the final two instars deviated significantly

from a linear relationship logio9pHCW) = 0.185 x Instar - 0.475, (R* = 0.999, Fi,6=7620,

P<0.001). Vertical grey bars represent the range of measurements for each instar.

107

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

= Douglas-Fir

Ge OF Sc O0¢- StL OL § O 8 9 v G 0

Aouanbel4 Aduenbe

3.5

3.0

2.9

2.0

1.0

0.5

Head Capsule Width (mm)

Figure 2. Distribution of head capsule sizes according to instar and sex when reared on

fir (bottom panel). Instars were assigned

based on the number of observed moults. Vertical dashed lines indicate proposed cut-off

points to discriminate field collected larval instars.

Himalayan blackberry (top panel) and Douglas

108 J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

5th instar 4" instar 3" Instar 2"4 instar 1* instar

6' Instar

Figure 3. Representative images of Orgyia antiqua larvae reared on Himalayan blackberry

leaves. Larvae in instars 1-3 (first three rows) exhibited little variation in colouration. The

dorsal tufts were always uniformly coloured in the final instar, which was 5 in males and

either 5 or 6 in females. Arrows between images indicate successive images of the same

individual.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 109

ACKNOWLEDGEMENTS

Thanks to Dave Holden for locating the population of rusty tussock moth on which

this investigation is based. The work was supported by Natural Resources Canada A-base

funding to BVH.

REFERENCES

Dyar, H.G. 1893. Orgyia badia and other notes, with a table to separate the larvae of Orgyia. Psyche: A

Journal of Entomology, 6:419-421.

Gentner, L.G.O. 1915. Thesis on the antique or rusty tussock moth. BSc in Agriculture. Oregon

Agricultural College, Corvalis, Oregon. Available from https://ir.library.oregonstate.edu/downloads/

kh04dt341 [Accessed 15 August 2018]

Hardy, G.A. 1945. Notes on the life history of the vapourer moth (Notolophus antiqua badia) on

Vancouver Island (Lepidoptera, Liparidae). Journal of the Entomological Society of British

Columbia, 42:3—6. Available from https://journal.entsocbc.ca/index.php/journal/article/view/747/755

Hatakoshi, M., Nakayama, I. and Riddiford, L.M. 1988. The induction of an imperfect supernumerary

larval moult by juvenile hormone analogues in Manduca sexta. Journal of Insect Physiology,

34:373-378. doi:10.1016/0022-1910(88)90106-0

Leonard, D.E. 1970. Effects of starvation on behaviour, number of larval instars, and developmental rate

of Porthetria dispar. Journal of Insect Physiology, 16:25—31. doi:10.1016/0022-1910(70)90109-5

Payne, H. 1917. The rusty tussock moth (Notolophus antiqua) Linn. Proceedings of the Nova Scotia

Entomological Society, 3:54—-61.

Retnakaran, A. 1973. Hormonal induction of supernumerary instars in the spruce budworm,

Choristoneura fumiferana (Lepidoptera: Tortricidae). The Canadian Entomologist, 105:459-461.

doi:10.4039/Ent105459-3

Sandre, S.L., Tammaru, T., and Mand, T. 2007a. Size-dependent colouration in larvae of Orgyia antiqua

(Lepidoptera: Lymantriidae): A trade-off between warning effect and detectability? European Journal

of Entomology, 104:745—752. doi:10.14411/eje.2007.095

Sandre, S.L., Tammaru, T., Esperk, T., Julkunen-Tiitto, R., and Mappes, J. 2007b. Carotenoid-based

colour polyphenism in a moth species: search for fitness correlates. Entomologia Experimentalis et

Applicata. 124:269—277. doi:10.1111/j.1570-7458.2007.00579.x

110 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

NATURAL HISTORY AND OBSERVATIONS

New records of Hymenoptera from British Columbia

and Yukon

C.G. RATZLAFF'

ABSTRACT— Thirty species of Hymenoptera are recorded for the first time from

British Columbia and Yukon, including nine with records representing the first for

Canada, with specimens from the families Bethylidae, Braconidae, Chrysididae,

Crabronidae, Diapriidae, Figitidae, and Thynnidae. A description of the male of

Diodontus spiniferus (Mickel) [Crabronidae], a correction to the distribution of

Dryudella elegans (Cresson) [Crabronidae], and a correction to the locale for the

holotype of Aspicera mirieiae Ros-Farré & Pujade- Villar [Figitidae] are also provided.

Key words: Hymenoptera, wasps, new, Canada, British Columbia, Yukon

INTRODUCTION

The diverse habitats of British Columbia and Yukon provide homes for a large

number of insect species, including many that, in Canada, are found only in this area.

Among the Hymenoptera, this is especially true, and new species are being recorded

every year (Heron and Sheffield 2015; Ratzlaff 2015; Ratzlaff et al. 2016). Many groups

of bees and wasps have been fairly well studied in British Columbia, while the last

significant study of Yukon wasp fauna was Finnamore’s chapter on aculeate wasps in the

1997 publication, /nsects of the Yukon. Large swathes of remote wilderness cover much

of the province and territory and, undoubtedly, many more known and unknown species

have yet to be discovered. Even just recently, a new bumblebee species, Bombus

kluanensis Williams & Cannings, was discovered in Yukon (Williams et al. 2016).

Recent field collecting trips, along with study of existing museum specimens at the

Spencer Entomological Collection, have resulted in 30 species of wasps being newly

identified from British Columbia and Yukon. These records are presented here.

Collection abbreviations used are as follows: CGR — Author’s personal collection;

CNCI — Canadian National Collection of Insects, Arachnids, and Nematodes, Ottawa,

ON; RBCM — Royal British Columbia Museum, Victoria, BC; SEM — Spencer

Entomological Collection, Beaty Biodiversity Museum, University of British Columbia,

Vancouver, BC. All specimens examined are located at the SEM with exception of two in

the CGR and one in the RBCM. Unless otherwise indicated, all scale bars shown are

equivalent to 1 mm.

FAMILY BETHYLIDAE

Anisepyris occidentalis (Ashmead)

First species records for Canada. Previously recorded from the western USA and

Mexico (Gordh and Moczar 1990).

BRITISH COLUMBIA: 19, Galiano I., north end, 20.vii.1986 (G. G. E. Scudder)

[SEM]; 12, Osoyoos, Haynes Ecological Reserve, 14.vi.—3.vili.1987, pan trap, Purshia/

Aristida steppe (S. G. Cannings) [SEM]; 1¢, Penticton, West Bench, 11.viii.1988, rose

thicket/grassland boundary (S. G. Cannings) [SEM]; 24, Kalamalka Lake Prov. Pk.,

21.viii.1987 (S. G. Cannings) [SEM] (Fig. 1); 12, Osoyoos, East Bench, 28.v.2000,

biting person (J. Scudder) [SEM]; 14, Tsawwassen, Boundary Bay Regional Pk.,

49.0176°N, 123.0422°W, 10.viii.2015 (C. G. Ratzlaff) [SEM]

| Corresponding author: Spencer Entomological Collection, Beaty Biodiversity Museum, 2212 Main

Mall, Vancouver, BC V6T 1Z4; chris.ratzlaff@gmail.com

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 111

Figure 1. Male Anisepyris occidentalis, from Kalamalka Lake Provincial Park, “rycen _

Epyris clarimontis Kieffer

First species records for Canada. Recorded as being widespread in the USA and

Mexico (Gordh and Moczar 1990).

BRITISH COLUMBIA: 29, Osoyoos, Haynes Ecological Reserve, 20.v.—14.vi.

1987, pan trap, Purshia/Aristida steppe (S. G. Cannings) [SEM]; 3°, Osoyoos, Haynes

Ecological Reserve, 14.vi.—3.vili.1987, pan trap, Purshia/Aristida steppe (S. G.

Cannings) [SEM]; 12, Osoyoos, Haynes Ecological Reserve, 13.vii—17.viii.1988, pitfall

trap, Purshia/Aristida steppe (S. G. Cannings) [SEM]; 12, Osoyoos, Haynes Ecological

Reserve, 23.vii.—26.viii.1989, pitfall trap, rose thicket (S. G. Cannings) [SEM]; 16,

Osoyoos, Haynes Ecological Reserve, 26.viii.—23.1x.1989, pitfall trap, Rosa clump at

edge of wetland (S. G. Cannings) [SEM]

FAMILY BRACONIDAE

Ascogaster borealis Shaw

First species record for Yukon. Previously recorded from BC, SK, ON, QC, NS, WA,

ID, MT, WI, and ME (Shaw 1983).

YUKON: 1, Million Dollar Falls, 60.1082°N, 136.9466°W, 26.vi.2017 (SEM

Team) [SEM]

Meteorus vulgaris (Cresson)

First species record for Yukon. Previously recorded from all of southern Canada and

much of the USA (Muesebeck 1923).

YUKON: 19, Carcross, Montana Mt., 60.1341°N, 134.7195°W, 28.vi.2017, 1075 m

(SEM Team) [SEM]

FAMILY CHRYSIDIDAE

Chrysurissa densa (Cresson)

First species records for Canada. Previously recorded from the western half of the

USA (Kimsey 2005).

BRITISH COLUMBIA: 13, SOCAP Site #28, 15.v.1990 (H. Knight) [SEM]; 1<3,

Osoyoos, Haynes Ecological Reserve, 1.vi.2000 (G. G. E. Scudder) [SEM]

it? J. ENTOMOL. SOc. BRIT. COLUMBIA 115, DECEMBER 2018

Pseudospinolia neglecta Shuckard

First species record for British Columbia. Previously recorded from AB, WA, CO,

MT, MN, NE, and NY. It is also found in the Palearctic region (Bohart and Kimsey

1982).

BRITISH COLUMBIA: 1°, Attachie, Don Phillips Way (Hwy. 29), 10V 599090

6233848 (56.23917°N, 121.40123°W), 22.vi.2015, 631 m (C & D Copley, J. Heron, H.

Gartner & K. Ovaska) [RBCM] (Fig. 2)

Figure 2. Female Pseudospinolia neglecta, from Attachie, BC.

FAMILY CRABRONIDAE

Crabro nigrostriatus Bohart

First species record for Yukon. Previously recorded from BC, OR, NV, and CA

(Bohart 1976).

YUKON: 14, Kookatsoon L., 60.5587°N, 134.8758°W, 29.vi.2017 (SEM Team)

[SEM] (Fig. 3)

Diodontus argentinae Rohwer

First species records for Yukon. Previously recorded from BC, WA, OR, WY, CA,

CO, UT, DC, and Mexico (Eighme 1989).

YUKON: 1.3, Kluane Nat. Pk., Sheep Mt., 5.vii.1979 (S. G. Cannings) [SEM]; 1,

Pelly Crossing, 2.vii.1985 (E. Bijdemast) [SEM]; 14, Dawson, 13.vii.1985, steep

Artemesia slope (S. G. Cannings) [SEM]

Diodontus bidentatus Rohwer

First species records for Yukon. Previously recorded from BC, AB, QC, NB, AK, ID,

MT, CO, NE, NY, MI, ND, and PA (Krombein 1979; Eighme 1989; Finnamore 1994;

Buck 2004; Ratzlaff 2015).

YUKON: 1, Duke River, Burwash Landing, 9.vii.1979 (S. G. Cannings) [SEM]; 16,

Kluane L., Emerald I., 61.0209°N, 138.4893°W, 24.v1.2017 (SEM Team) [SEM]

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 113

Figure 3. Male Crabro nigrostriatus, from Kookatsoon Lake, YT.

Diodontus leguminiferus Cockerell

First species records for Yukon. Previously recorded from BC, AB, ID, CA, MT, CO,

UT, AZ, NM, MO, and IA (Eighme 1989; Ratzlaff 2015).

YUKON: 1, Carcross, sand dunes, 20.vii.1987 (S. G. Cannings) [SEM]; 1¢,

Carcross Desert, 60.1876°N, 134.6899°W, 30.vi.2016 (C. G. & N. A. Ratzlaff) [SEM];

24, Carcross Desert, 60.1876°N, 134.6899°W, 28.vi.2017 (SEM Team) [SEM]

Diodontus occidentalis Fox

First species records for Yukon. Previously recorded from BC, AB, AK, ID, CA, NV,

UT, WY, CO, AZ, MI, NY, and ND (Eighme 1989; Finnamore 1994; Ratzlaff 2015).

YUKON: 19, Silver City, 23.vii.1979 (G. G. E. Scudder) [SEM]; 19, Pelly

Crossing, 26.vii.1980 (R. J. Cannings) [SEM]; 1%, Tenas Creek, 5 km East on North

Canol Rd., 62°02’N 132°14’W, 11.vi.1981 (C. S. Guppy) [SEM]; 19, Haines Junction,

Pine Cr., 25.vi.1981 (C. S. Guppy) [SEM]; 12, Porcupine R., Blue Bluffs, 67°38’N

138°38’W, 11.vii.1981 (C. S. Guppy) [SEM]; 1419, Old Crow, 6 km E, 67°34’N

139°41’°W, 13.vii.1981 (C. S. Guppy) [SEM]; 12, Whitehorse, Wolf Cr., 17.vii.1981 (C.

S. Guppy) [SEM]; 192, Slims R. delta, 21.vi.1982 (R. D. Wilkie & S. G. Cannings)

[SEM]; 12, Kluane, Sheep Mt., 24.vi.1982 (S. G. Cannings, R. D. Wilkie, L. Vasington

& R.A. Moore) [SEM]; 19, Carmacks, 30 km E, 62°02’N 135°51’W, 10.vii.1982 (S. G.

Cannings, L. Vasington & R. A. Moore) [SEM]; 19, Old Crow, 30.vi.1983, top of open

S-facing bluff, malaise trap (R. A. Cannings) [SEM]; 19, Old Crow, 2.vii.1983, top of

open S-facing bluff, malaise trap (R. A. Cannings) [SEM]; 19, Old Crow, 4.vii.1983, top

of open S-facing bluff, malaise trap (S. G. Cannings) [SEM]; 19, Little Salmon L., 35

km E, 28.vi.1985 (E. Krebs & J. J. Robinson) [SEM]; 2%, Tatchun L., 29.vi.1985 (E.

Krebs & J. J. Robinson) [SEM]; 2, Pelly Crossing, 2.vii.1985 (S. G. Cannings) [SEM];

19, Dawson, Midnight Dome, 12.vii.1985 (E. Bijdemast) [SEM]; 1’, Carcross, Montana

Mt., 60.1341°N, 134.7195°W, 28.v1.2017, 1075m (SEM Team) [SEM]

114 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Diodontus spiniferus (Mickel)

First species records for British Columbia and Yukon. Previously recorded from AB,

ON, QC, CA, MT, CO, IA, NE, MD, MO, and MN (Eighme 1989; Buck 2004). The male

of the species has never been described, but a few key characters were provided by Buck

(2004) that are useful when comparing it to eastern specimens. Several other similar

species exist in western Canada, and the necessary characters for identification are

described here.

Male. (Fig. 4) Black. Mandible (except base and teeth), palps, apex of fore-femur,

fore- and mid-tibiae dorsally, and anterior half of tegula yellow. Hind-tibia brownish-

yellow dorsally, fading apically or nearly all brown in darker specimens. Fore and mid-

tarsi yellow, hind-tarsi brown, the last two segments of all tarsi darkened. Antenna with

small placoids on flagellomeres V—X, ranging from reddish-brown to brown. Frons with

numerous larger punctures, often with much reticulation, giving it a rough appearance.

Humeral angle prominent with close to a 90° angle. Propodeum reticulate. Wing veins

and stigma brown. Abdominal terga sparsely punctate, lightly reticulate.

Figure 4. Male Diodontus spiniferus, from Kluane National Park, YT.

Using Eighme’s (1989) key, males of D. spiniferus end up at couplet 16 with retiolus

and /eguminiferus but lack the strong reticulation on the abdomen found in retiolus. They

differ from /eguminiferus in having numerous large punctures and stronger sculpture on

the frons (Fig. 5a), a prominent, roughly 90° humeral angle (Fig. 5b), and reddish-brown

placoids on the antenna (Fig. 5c). Two other similar species are boharti and crassicornus,

but spiniferus differs from the former by having a dark pronotal lobe and from the latter

by having much less-inflated antenna and smaller placoids.

BRITISH COLUMBIA: 16, Pink Mt., 24 km S, 24.vi.1985 (S.G. Cannings) [SEM]

YUKON: 3429, Kluane Nat. Pk., Sheep Mt., 8.vi.1979 (S. G. Cannings) [SEM]

(Fig. 4; Fig. 5c); 1419, Carmacks, Mt. Nanson Rd., 62.0587°N, 136.3781°W, 26.vi.2016

(C. G. & N. A. Ratzlaff) [SEM] (Fig. 5a, b); 19, Ibex Valley Salt Flats, 60.8616°N,

135.7126°W, 23.vi.2017 (SEM Team) [SEM]

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 1S

Figure 5. The face (a) and posterolateral view of | the humeral angle on the pronotum (b) of a

male Diodontus spiniferus from Carmacks, YT. The antennal placoids (c) of a male D.

spiniferus from Kluane National Park, YT.

Diodontus vallicolae (Rohwer)

First species record for Yukon. Previously recorded from BC, AB, AK, ID, WY, CA,

CO, NV and UT (Eighme 1989).

YUKON: 16, Carcross, sand dunes, 20.vii.1987 (S. G. Cannings) [SEM]

Dryudella elegans (Cresson)

In Cresson’s (1881) original species description for D. elegans (as Astata elegans),

the holotype location is given as “Washington Territory” and the paratype locations as

‘“Vancouver’s Island”, Nevada, and Colorado. These locations appear again in Fox’s

(1893) synopsis of the North American Larridae and then disappear from all subsequent

publication on the species, with the exception of Nevada. Parker (1969) records D.

elegans from ID, WY, UT, NV, CA, and AZ, stating the holotype location only as “W. T.”

It appears that D. elegans should also be listed from BC (Vancouver Island), WA, and CO

even thouth it currently is not. Why these localities were not included in the subsequent

published species distributions is unknown, but additional British Columbian records are

presented here, confirming the original northern range.

116 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

BRITISH COLUMBIA: 1, Osoyoos, Haynes Ecological Reserve, 9.vii.—9.viii.

1996, BGxhl, pitfall trap (G. G. E. Scudder) [SEM]; 14, Oliver, McKinney Rd.,

49.19869°N, 119.49967°W, 26.vii.2017 (C. G. Ratzlaff) [CGR]

Philanthus pulcher Dalla Torre

First species record for Yukon. Finnamore (1997) expected this species to occur in the

territory, and it has been previously recorded from the western half of Canada and the

USA, including NT (Bohart and Grissell 1975).

YUKON: 103, Pelly Crossing, 2.vii.1985 (S. G. Cannings) [SEM]

Solierella albipes (Ashmead)

First species record for Canada. Previously recorded from ID, CO, and CA

(Krombein 1979).

BRITISH COLUMBIA: 19, Osoyoos, Strawberry Creek Rd., 49.0364°N,

119.5002°W, 9.vi1i.2016 (C. G. Ratzlaff) [SEM] (Fig. 6)

Solierella sayi (Rohwer)

First species records for Canada. Previously recorded from CO and CA (Krombein

1979).

BRITISH COLUMBIA: 2, Whipsaw Creek Forest Service Rd., 49.3536°N,

120.6097°W, 7—10.vii1.2016, 986m, blue pan (C. G. Ratzlaff) [CGR, SEM]

FAMILY DIAPRIIDAE

Ismarus halidayi Forster

First species record for British Columbia. Previously recorded in the Nearctic region

from AB, NB, NF, CA, and MO, and in the Palearctic region from England and Finland

(Masner 1976).

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 7

BRITISH COLUMBIA: 19, Sidney I., Dragonfly Pond, 49.6033°N, 123.3046°W,

14.vi1.2016 (SEM Team) [SEM]

FAMILY FIGITIDAE

Alloxysta halterata (Thomson)

First species records for Canada. Previously recorded in the Nearctic region from CO,

and in the Palearctic region from England, Finland, Germany, Scotland, and Sweden

(Ferrer-Suay et al. 2014; Ferrer-Suay 2017). |

YUKON: 1, White Mts., “Erebia Cr.”, 67°58’N 136°29’W, 2.vii. — 9.vii.1987,

2500’, sandstone slope, pan trap (S. G. Cannings) [SEM] (Fig. 7); 19, Emerald L.,

60.2639°N, 134.7520°W, 29.v1.2017 (SEM Team) [SEM].

Figure 7. Male Al/oxysta halterata, from the White Mountains, YT.

Alloxysta obscurata (Hartig)

First species record for Yukon. Previously recorded in the Nearctic region from BC

and AK, and in the Palearctic region from Andorra, France, Germany, Hungary, Iceland,

Poland, Romania, and Scotland (Ferrer-Suay 2017).

YUKON: 19, Cottonwood Cr., 60°55’N 132°58’W, 2.viil.1981 (C. S. Guppy) [SEM]

Alloxysta pallidicornis (Curtis)

First species record for British Columbia. Previously recorded in the Nearctic region

from AB, QC, AK, and CO, and in the Palearctic region from Austria, England, Finland,

France, Germany, Norway, Spain, and Sweden (Ferrer-Suay 2017).

BRITISH COLUMBIA: 19, Saturna I., Gulf Islands National Pk. & Reserve,

A48.8084°N, 123.1856°W, 17.vii.2015 (C. G. Ratzlaff) [SEM]

Alloxysta postica (Hartig)

First species records for Canada. Previously recorded in the Nearctic region from AZ

and in the Palearctic region from Bulgaria, Czech Republic, and Germany (Ferrer-Suay

et al. 2014; Ferrer-Suay 2017).

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

118

2017 (SEM Team) [SEM]

19, Kookatsoon L., 60.5587°N, 134.8758°W, 29.vi.2017 (SEM Team) [SEM]

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BRITISH COLUMBIA

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Figure 8. Male Aspicera santamar

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 119

Omalaspis cavroi (Hedicke)

First species record for Yukon. Previously recorded from BC, AB, ON, QC, NB, AK,

MT, CA, AR, and ME (Ros-Farré & Pujade-Villar 2011b).

YUKON: 16, Carcross, sand dunes, 20.vii.1987 (S. G. Cannings) [SEM]

Paraspicera brandaoi Ros-Farré & Pujade-Villar

First species records for Yukon. Previously recorded from BC, AB, and ID (Ros-Farré

and Pujade-Villar 201 1a).

YUKON: 12, Old Crow, 1 km E, 16.vii.1981 (C.S. Guppy) [SEM]; 1, Old Crow,

2.vii.1983, top of open S-facing bluff, malaise trap (S. G. Cannings) [SEM]

Phaenoglyphis gutierrezi Andrews

First species record for Yukon. Previously recorded from BC, SK, and MT (Andrews

1978).

YUKON: 19°, Cottonwood Cr., 60°55’N 132°58’W, 2.viii.1981 (C. S. Guppy) [SEM]

Phaenoglyphis pilosus Andrews

First species record for Yukon. Previously recorded from BC, AB, ID, CA and CO

(Andrews 1978).

YUKON: 19, Emerald L., 60.2639°N, 134.7520°W, 29.vi.2017 (SEM Team) [SEM]

Phaenoglyphis ruficornis (Forster)

First species record for Yukon. Previously recorded in the Nearctic region from BC,

SK, ON, QC, and CA, and in the Palearctic region from Germany and Israel (Ferrer-Suay

2017).

YUKON: 19, Tagish, 22.vii.1981 (S. G. Cannings) [SEM]

Phaenoglyphis villosa (Hartig)

First species record for Yukon. A very widespread species that has been recorded

from every continent except Antarctica (Ferrer-Suay 2017).

YUKON: 19, Kluane Nat. Pk., S end of Kluane L., 60.9930°N, 138.4674°W, 24.vi1.

2017 (SEM Team) [SEM]

Sarothrus nasoni Ashmead

First species record for Canada. Previously known only from IL (Burks 1979).

BRITISH COLUMBIA: 19, Pink Mt., 57.0487°N, 122.8687°W, 2.vii.2016, 1715m

(C. G. & N. A. Ratzlaff) [SEM] (Fig. 9)

FAMILY THYNNIDAE

Lalapa lusa Pate

First species records for Canada. Goulet and Huber (1993) suspected that this species

occurred in southern BC, and it has been previously recorded from WA, ID, OR, and CA

(Johnson et al. 1995).

BRITISH COLUMBIA: 1°, Osoyoos, Haynes Ecological Reserve, The Throne,

10.vii.—14.viii.1986, under sage brush, pitfall trap (S. G. Cannings) [SEM]; 19,

Penticton, West Bench, 3.viii.1987 (S. G. Cannings) [SEM]; 1°, Penticton, West Bench,

23.viii.1987 (S. G. Cannings) [SEM] (Fig. 10); 19, Osoyoos, Haynes Ecological

Reserve, 13.vii.—17.viii.1988, Purshia/Aristida steppe, pitfall trap (S. G. Cannings)

[SEM]; 12, Osoyoos, Haynes Ecological Reserve, 9.vili.1995, (G.G.E. Scudder) [SEM];

12, Osoyoos, Haynes Ecological Reserve, 15.viii—11.ix.2004, BGxhl, AN Recovery

after 1993 fire, Pitfall trap ER2-4 (G. G. E. Scudder) [SEM]

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

faa)

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J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 121

CONCLUSION

Bioblitzes have become an important part of the study of flora and fauna in British

Columbia, the Yukon Territories, and the rest Canada. These events facilitate a

concentrated effort to document the species present in areas where often not much has

previously been done. This is particularly true in places with regulated research access,

such as national parks and, as a result, the known range of many species has been

expanded. Much of this new material, however, unfortunately ends up unidentified in

different natural history collections, alongside many other unexamined specimens.

Undoubtedly, study of these specimens will yield new information about many species in

British Columbia and the Yukon.

ACKNOWLEDGEMENTS

Thank you to Syd Cannings, Parks Canada, and the other organizers of the 2016

Carmacks Bioblitz and the 2017 Kluane National Park Bioblitz for the invitation and the

opportunity to visit these unique Yukon habitats. Thank you to Athena George, Parks

Canada, and the other organizers of the 2015 Saturna Island Bioblitz and the 2016 Sidney

Island Bioblitz for the invitation and opportunity to visit the Gulf Islands National Park

and Reserve.

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species (Hymenoptera: Cynipidae). Occasional Papers in Entomology 25:1—128.

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Smith, and B. D. Burks, eds. Catalog of Hymenoptera in America north of Mexico: Volume 1,

Symphyta & Apocrita (Parasitica). Smithsonian Institution Press, Washington, DC. 1198 pp.

Bohart, R. M. 1976. A review of the Nearctic species of Crabro (Hymenoptera: Sphecidae). Transactions

of the American Entomological Society 102:229-287,

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Sphecidae). Bulletin of the California Insect Survey 19:1—92.

Bohart, R. M., and L. S. Kimsey. 1982. A synopsis of the Chrysididae in America north of Mexico.

Memoirs of the American Entomological Society 33:1—266.

Buck, M. 2004. An annotated checklist of the Spheciform wasps of Ontario (Hymenoptera: Ampulicidae,

Sphecidae and Crabronidae). Journal of the Entomological Society of Ontario 134:19—84.

Cresson, E. T. 1881. Descriptions of new Hymenoptera in the collection of the American Entomological

Society. Transactions of the American Entomological Society 9: Proceedings of the Monthly

Meetings of the Entomological Section of the Academy of Natural Sciences, Philadelphia tii—vi.

Eighme, L. E. 1989. Revision of Diodontus (Hymenoptera: Sphecidae) in America north of Mexico.

Annals of the Entomological Society of America 82:14—28.

Ferrer-Suay, M. 2017. Interactive Charipinae Worldwide Database. http://www.charipinaedatabase.com/

Ferrer-Suay, M., J. Selfa, and J. Pujade-Villar. 2014. First records, new species, and a key of the

Charipinae (Hymenoptera: Cynipoidea: Figitidae) from the Nearctic region. Annals of the

Entomological Society of America 107:50—73.

Finnamore, A. T. 1994. Hymenoptera of the Wagner Natural Area, a boreal spring fen in central Alberta.

Memoirs of the Entomological Society of Canada 169:181—220.

Finnamore, A. T. 1997. Aculeate wasps (Hymenoptera: Aculeata) of the Yukon, other than Formicidae.

pp. 868-900 in H.V. Danks & J.A. Downes (Eds.). Insects of the Yukon. Biological Survey of

Canada Monograph Series 2. 1034 pp.

Fox, W. J. 1893. The North American Larridae. Proceedings of the Academy of Natural Sciences of

Philadelphia 45:467—551.

Gordh, G., and L. Moczar. 1990. A catalog of the world Bethylidae (Hymenoptera: Aculeata). Memoirs

of the American Entomological Institute 46:1—364.

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Goulet, H., and J. T. Huber (Eds.) 1993. Hymenoptera of the world: An identification guide to families.

Agriculture Canada, Ottawa. 668 pp.

Heron, J., and C. S. Sheffield. 2015. First record of the Lasioglossum (Dialictus) petrellum species group

in Canada (Hymenoptera: Halictidae). Journal of the Entomological Society of British Columbia

112: 88-91.

Johnson, J. B., T. D. Miller, and W. J. Turner. 1995. Lalapa lusa Pate (Hymenoptera: Tiphiidae): new

localities and new floral associations in the Pacific Northwest. Pan-Pacific Entomologist 71:64—-65.

Kimsey, L. S. 2005. California cuckoo wasps in the family Chrysididae (Hymenoptera). bret noes of

California Publications in Entomology 125:1-311.

Krombein, K. V. 1979. Superfamily Sphecoidea, pp. 1573-1740 in K.V. Krombein, P.D. Hurd, Jr., D.R.

Smith, and B.D. Burks eds. Catalog of Hymenoptera in America north of Mexico: Volume 2,

Apocrita (Aculeata). Smithsonian Institution Press, Washington, D.C. 1101 pp.

Masner, L. 1976. A revision of the Ismarinae of the New World (Hymenoptera, Proctotrupoidea,

Diapriidae). The Canadian Entomologist 108:1243-1266.

Muesebeck, C. F. W. 1923. A revision of the North American species of ichneumon-flies belonging to the

genus Meteorus Haliday. Proceedings of the United States National Museum 63:1—44.

Parker, F. D. 1969. On the subfamily Astatinae. Part VI. The American species in the genus Dryudella

Spinola (Hymenoptera: Sphecidae). Annals of the Entomological Society of America 62:963—976.

Ratzlaff, C. G. 2015. Checklist of the Spheciform wasps (Hymenoptera: Crabronidae & Sphecidae) of

British Columbia. Journal of the Entomological Society of British Columbia 112:19-46.

Ratzlaff, C. G., K. M. Needham, and G. G. E. Scudder. 2016. Notes on insects recently introduced to

Metro Vancouver and other newly recorded species from British Columbia. Journal of the

Entomological Society of British Columbia 113:79-89.

Ros-Farré, P., and J. Pujade-Villar. 2011a. Revision of the genus Paraspicera Kieffer, 1907 (Hym.:

Figitidae: Aspicerinae). Zootaxa 2801:48—56.

Ros-Farré, P., and J. Pujade-Villar. 2011b. Revision of the genus Omalaspis Giraud, 1860 (Hym.:

Figitidae: Aspicerinae). Zootaxa 2917:1—28.

Ros-Farré, P., and J. Pujade-Villar. 2013. Revision of the genus Aspicera Dahlbom, 1842 (Hym.:

Figitidae: Aspicerinae). Zootaxa 3606: 1—110.

Shaw, S. R. 1983. A taxonomic study of Nearctic Ascogaster and a description of a new genus

Leptodrepana (Hymenoptera: Braconidae). Entomography 2:1—54.

Williams, P. H., S. G. Cannings, and C. S. Sheffield. 2016. Cryptic subarctic diversity: a new bumblebee

species from the Yukon and Alaska (Hymenoptera: Apidae). Journal of Natural History 50:2881—

2893.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 123

NATURAL HISTORY AND OBSERVATIONS

First Record of Culex tarsalis (Diptera: Culicidae) in the

Yukon

DANIEL A. H. PEACH!

ABSTRACT— The first record of Culex tarsalis in the Yukon is reported from a larva

collected in Kluane National Park in 2017. Details on the location and the specimen

are provided, and background information on the biology of Cx. tarsalis and its role in

arbovirus transmission are discussed.

Key words: Culex tarsalis, Western encephalitis mosquito, Culicidae, Yukon,

mosquito distribution

The western encephalitis mosquito, Culex tarsalis Coquillett, is a medium-sized

mosquito (wing length 4.0-4.4 mm) with bands of white scales on the tarsi and a broad

ring of white scales on the proboscis at mid-length (Belton 1983). Adult females

overwinter in sheltered areas such as caves, rodent burrows, and under rock piles (Wood

et al. 1979), and larvae are found in a wide variety of habitats including ponds, marshes,

ditches, and irrigation water (Belton 1983). Adults have been observed feeding from

flowers of goldenrod (Solidago spp.) (Sandholm and Price 1962) and common tansy

(Tanacetum vulgare), from which they carry pollen (Peach and Gries 2016). Females take

blood from birds and mammals (Wood et al. 1979).

Cx. tarsalis is an important vector of several viruses in southern Canada, including

West Nile virus (Roth et a/. 2010, Kulkarni et a/. 2015), western equine encephalitis

(McLintock et al. 1970), and St. Louis encephalitis (Hammon and Reeves 1943),

although these are not known from the Yukon (Artsob 1990). Snowshoe hare virus, in the

California Encephalitis (CE) group, is endemic in the Yukon (McLean ef a/l.1973) but is

not reported to have been isolated from Cx. tarsalis. However, CE itself has been isolated

from Cx. tarsalis in California (Hammon ef a/. 1952). Northway virus is also endemic to

the Yukon (McLean and Lester 1983), but little is known about this virus or its vectors.

The known range of Cx. tarsalis extends throughout much of central and western

North America (Darsie and Ward 2005), including southern British Columbia (Belton

1983) and southern Alberta (Wood ef al.1979). It has also been found in Norman Wells,

(65°N) in the Northwest Territories (Freeman 1952) and Belton and Belton (1990)

believed the species was likely to occur in the Yukon as well, based on its inclusion in a

list of Yukon mosquitoes by Nelson (1977). Nelson cites a personal communication from

D. M. Wood to support this, but Wood et al. (1979) show no records of Cx. tarsalis in the

Yukon.

A Culex sp. larva was collected in a shallow pond in Kluane National Park, Yukon,

Canada just outside the Slims River Flats (60°59'23.6"N, 138°29'31.9"W) on 24 June,

2017 as part of the Kluane Park bioblitz (research permit number KLU-2017-25041).

The larva was successfully reared to adulthood, and the female was identified as Cx.

tarsalis (Fig. 1) using the key of Wood et al. (1979). This specimen represents the first

confirmed record of this species in the Yukon. Of note is the incomplete white-scaled

ring at midpoint of the proboscis of this specimen as it possesses dark scales dorsally,

possibly due to phenotypic plasticity related to thermal melanism (Trullas et a/. 2007) or

poor larval habitat conditions (Talloen et al. 2004). The pond was adjacent to the Alaska

Highway, approximately 10 metres in diameter, shallow, and contained clear water.

! Corresponding author: Department of Biological Sciences, Simon Fraser University, 8888 University

Drive, Burnaby, British Columbia, VSA 1S6; (778) 782 5939, dap3@sfu.ca

124 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Nearby vegetation included willow (Salix spp.), spruce (Picaea sp.), fireweed

(Chamaenerion sp.), and patches of unidentified grass. Larvae of Anopheles earlei,

Aedes excrucians, and Culiseta alaskaensis were also collected from the pond, and adults

captured nearby included Ae. campestris, Ae. cataphylla, Ae. communis, Ae. excrucians,

Ae. fitchii, and Ae. implicatus. The Cx. tarsalis specimen has been deposited for

reference in the Beaty Biodiversity Museum at the University of British Columbia,

Vancouver, British Columbia. Due to the short summer season and the likeliness that

only small populations may be present it seems unlikely that Cx. tarsalis currently poses

a major human health risk in the Yukon. However, if temperatures rise these limiting

conditions may no longer apply (Chen ef al. 2013).

Figure 1. Antero-dorsal (A) and lateral (B) views of the Cx. tarsalis specimen collected in the

Yukon. Note incomplete band of white scales at mid-proboscis in (A), indicated by an arrow.

ACKNOWLEDGEMENTS

I thank Dr. Peter Belton of Simon Fraser University, and Karen Needham of the UBC

Spencer Entomology Collection, for their support and assistance, as well as Chris

Ratzlaff of the UBC Spencer Entomology Collection for the use of his photographs. I

would also like to thank three anonymous reviewers for their comments on this

manuscript. |

REFERENCES

Artsob, H. 1990. Arbovirus activity in Canada. Jn Hemorrhagic Fever with Renal Syndrome, Tick and

Mosquito-Borne Viruses. Archives of Virology Supplement, vol 1. Edited by C.H. Calisher. Springer,

Vienna, Austria. Pp. 249-258. doi: 10.1007/978-3-7091-9091-3 28.

Belton, E.M. and Belton, P. 1990. A review of mosquito collecting in the Yukon. Journal of the

Entomological Society of British Columbia, 87: 35-37.

Belton, P. 1983. The Mosquitoes of British Columbia. British Columbia Provincial Museum, Handbook

41. Victoria, British Columbia, Canada.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 125

Chen, C.C., Jenkins, E., Epp, T., Waldner, C., Curry, P.S., and Soos, C. 2013. Climate change and West

Nile virus in a highly endemic region of North America. International Journal of Environmental

Research: Public Health, 10: 3052-3071.

Freeman, T.N. 1952. Interim report on the distribution of the mosquitoes obtained in the Northern Insect

Survey. Defence Research Board of Ottawa. Technical Report 1.

Hammon, W.M. and Reeves, W.C. 1943. Laboratory transmission of St. Louis encephalitis by three

genera of mosquitoes. Journal of Experimental Medicine, 78: 241-253.

Hammon, W. M., Reeves, W.C., and Sather, G. 1952. California encephalitis virus, a newly described

agent: IT. Isolations and attempts to characterize the agent. Journal of Immunology, 69: 493-510.

Kulkarni, M.A., Berrang-Ford, L., Buck, P.A., Drebot, M.A., Lindsay, L.R., and Ogden, N.H. 2015.

Major emerging vector-borne zoonotic diseases of public health importance in Canada. Emerging

Microbes and Infections, 4: e33. doi: 10.1038/emi.2015.33.

McLean, D.M., Clarke, A.M., Goddard, E.J., Manes, E.S., Montalbetti, C.A., and Pearson, R.E. 1973.

California encephalitis virus endemicity in the Yukon Territory, 1972. Journal of Hygiene (London),

71: 391-402.

McLean, D.M. and Lester, S.A. 1983. Isolation of snowshoe hare virus from Yukon mosquitoes.

Mosquito News, 44: 200-203.

McLintock, J.A., Burton, N., McKiel, J.A., Hall, R.R., and Rempel, J.G. 1970. Known mosquito hosts of

Western encephalitis virus in Saskatchewan. Journal of Medical Entomology, 7: 446-454.

Nelson, J. 1977. Mosquito control in the Yukon Territory, Canada. MPM thesis, Simon Fraser University,

Canada.

Peach, D.A.H. and Gries, G. 2016. Nectar thieves or invited pollinators? A case study of tansy flowers

and common house mosquitoes. Arthropod Plant Interactions, 10: 497-506. doi: 10.1007/

s11829-016-9445-9.

Roth, D., Henry, B., Mak, S., Fraser, M., Taylor, M., Li, M., Cooper, K., Furnell, A., Wong, Q., Morshed,

M. et al. 2010. West Nile virus range expansion into British Columbia. Emerging Infectious

Diseases, 16: 1251-1258. doi: 10.3201/eid1608.100483

Sandholm, H.A. and Price, R.D. 1962. Field observations on the nectar feeding habits of some Minnesota

mosquitoes. Mosquito News, 22: 346-349.

Talloen, W., Van Dyck, H., and Lens, L. 2004. The cost of melanisation: butterfly wing coloration under

environmental stress. Evolution, 58: 360-366.

Trullas, S.C., van Wyk, J.H., and Spotila, J.R. 2007. Thermal melanism in ectotherms. Journal of

Thermal Biology, 32: 235-245.

Wood, D.M., Dang, P.T., and Ellis, R.A. 1979. The Insects and Arachnids of Canada 6. The Mosquitoes

of Canada (Diptera: Culicidae). Agriculture Canada, Ottawa, Ontario, Canada.

126 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

NATURAL HISTORY AND OBSERVATIONS

An updated list of the mosquitoes of British Columbia with

distribution notes

DANIEL A.H. PEACH!

Since “The Mosquitoes of British Columbia’, originally published by Dr. Peter

Belton 35 years ago, there have been only sporadic and incomplete updates on the

mosquito fauna of British Columbia (BC). Darsie and Ward’s (2005) “Identification and

Geographical Distribution of the Mosquitoes of North America, North of Mexico”

reported the presence and distribution of many species within BC but was continent-wide

in scope and did not provide BC-specific information. It also disregarded the presence or

distribution of several species.

Belton (1983) recognized 46 mosquito species as occurring within British Columbia,

discounting previous records of Culex restuans but including Aedes nevadensis due to

specimens he had collected from the BC Interior. Darsie and Ward (2005) recognized 45

species, discounting records of Ae. nevadensis from Belton (1983) as well as previous

records of Cx. restuans. Since 2005, several additional species records have been made

for BC, and a new record of Cx. restuans from southern Vancouver Island supports its

inclusion as part of BC’s mosquito fauna, bringing the total number of species known

from the province to 50. In several instances, the distribution of various species within

BC has also been extended, due to new collection records in previous unsurveyed or

undersurveyed areas.

List of the mosquito species known from British Columbia

Aedes (Ochlerotatus) aboriginis Dyar

Aedes (Ochlerotatus) aloponotum Dyar (Updated distribution)

Aedes (Ochlerotatus) campestris Dyar & Knab

Aedes (Ochlerotatus) canadensis (Theobald)

Aedes (Ochlerotatus) cataphylla Dyar

Aedes (Aedes) cinereus Meigen (Updated distribution)

Aedes (Ochlerotatus) communis (De Geer)

Aedes (Ochlerotatus) diantaeus Howard, Dyar & Knab

Aedes (Ochlerotatus) dorsalis (Meigen)

Aedes (Ochlerotatus) euedes Howard, Dyar & Knab

Aedes (Ochlerotatus) excrucians (Walker)

Aedes (Ochlerotatus) fitchii (Felt & Young)

Aedes (Ochlerotatus) flavescens (Miieller)

Aedes (Ochlerotatus) hendersoni Cockerell

Aedes (Ochlerotatus) hexodontus Dyar

Aedes (Ochlerotatus) impiger (Walker)

Aedes (Ochlerotatus) implicatus Vockeroth

Aedes (Ochlerotatus) increpitus Dyar

Aedes (Ochlerotatus) intrudens Dyar

Aedes (Finlaya) japonicus japonicus (Theobald) (Jackson et al. 2016) (Updated

distribution)

Aedes (Ochlerotatus) mercurator Dyar

! Corresponding author: Department of Biological Sciences, Simon Fraser University, 8888 University

Drive, Burnaby, B.C. V5A 186; dap3@sfu.ca

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 127

Aedes (Ochlerotatus) melanimon Dyar

Aedes (Ochlerotatus) nevadensis Chapman & Barr (Updated distribution, first formal

record)

Aedes (Ochlerotatus) nigripes (Zetterstedt)

Aedes (Ochlerotatus) pionips Dyar

Aedes (Ochlerotatus) provocans (Walker)

Aedes (Ochlerotatus) pullatus (Coquillett)

Aedes (Ochlerotatus) punctor (Kirby)

Aedes (Ochlerotatus) riparius Dyar & Knab

Aedes (Ochlerotatus) schizopinax Dyar (Jackson et al. 2013)

Aedes (Ochlerotatus) sierrensis (Ludlow)

Aedes (Ochlerotatus) spencerii spencerii (Theobald) (Updated distribution)

Aedes (Ochlerotatus) spencerii idahoensis (Theobald) (Updated distribution)

Aedes (Ochlerotatus) sticticus (Meigen)

Aedes (Ochlerotatus) togoi (Theobald)

Aedes (Aedes) vexans vexans (Meigen)

Aedes (Aedes) vexans nipponii (Theobald) (Belton 2015) (First formal record)

Anopheles earlei Vargas

Anopheles freeborni Aitken

Anopheles punctipennis (Say) (Updated distribution)

Culex pipiens L.

Culex restuans Theobald (McCann and Belton 2015)

Culex tarsalis Coquillett (Updated distribution)

Culex territans Walker (Updated distribution)

Culiseta alaskaensis (Ludlow)

Culiseta impatiens (Walker)

Culiseta incidens (Thomson)

Culiseta inornata (Williston)

Culiseta minnesotae Barr

Culiseta morsitans (Theobald)

Culiseta particeps (Adams) (Jackson et al. 2013) (Updated distribution)

Coquillettidia perturbans (Walker) (Updated distribution)

Notes on new species records and distribution updates

Anopheles punctipennis (Say) is previously known from both Vancouver Island and

the southern mainland of BC, but Darsie and Ward (2005) seem not to have recognized

records of this species from Vancouver Island. Surveys by Stephen ef al. (2006) found

this species to be widely distributed on Vancouver Island, re-confirming its presence

there.

Aedes aloponotum Dyar is known from the Fraser Valley and southern Vancouver

Island. The distribution shown in Darsie and Ward (2005) seems to erroneously display

the range of this species as extending up the Fraser Canyon and east to the interior of BC,

possibly due to a mis-citation of Gjullin and Eddy (1972), who reported this species as

occurring in the Fraser Valley. This may possibly be due to confusing the Fraser Valley

with the Fraser Canyon. Additionally, the author has found this species from the outskirts

of Whistler, extending the northern limits of its known range.

Aedes cinereus Meigen was reported from every part of BC by Belton (1983), but the

distribution displayed by Darsie and Ward (2005) does not include Vancouver Island.

Stephen et al. (2006) found Ae. cinereus in light traps on Vancouver Island,

demonstrating that this species does occur there.

128 J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018

Aedes japonicus japonicus (Theobald) was first reported in BC from samples

collected in Maple Ridge and Mission in 2014 (Jackson et al. 2016), and additional

specimens have been collected by Sean McCann in Langley and by the author in

Burnaby and Saanichton, with samples deposited in the Spencer Entomology Collection

at the UBC Beaty Biodiversity Museum. This species seems to have become established

in the Lower Mainland and southern Vancouver Island and may be spreading; if it is not

present throughout these regions yet, it may soon become so. Whether or not it can

become established in other parts of BC remains to be seen.

Aedes nevadensis Chapman and Barr was reported by Belton (1983) from larval

collections made in Castlegar. However, this record does not seem to have been

recognized by Darsie and Ward (2005), and its presence is recorded here to remove

ambiguity. Belton made further collections of this species just outside Manning Park, and

the author has collected them from Pemberton and from north of Princeton. The author’s

specimens have been deposited in the Spencer Entomology Collection at the UBC Beaty

Biodiversity Museum. This species is likely found in dry areas of much of the Southern

Interior of BC, although how far north its range extends is currently unknown. |

Aedes schizopinax Dyar was first reported in BC from a collection made in the

municipality of Sparwood, near the Alberta border (Jackson et al. 2013). An additional

specimen, collected in Williams Lake by C. Phippen, along with records from

Washington, suggest this species may exist in low numbers throughout the Interior of

BC.

Aedes spencerii spencerii (Theobald) was previously believed to be present in BC

only in the Peace River region (Belton 1983), with records from Kaslo of two specimens

— one collected by HG Dyar and one by RP Currie (Dyar 1904) — considered dubious

(Belton 1983). Examination of specimens in the Spencer Entomology Collection at the

UBC Beaty Biodiversity Museum have revealed additional specimens of Ae. spencerii

spencerii collected in the Southern Interior of BC, where it was previously believed only

the idahoensis subspecies was found (Belton 1983). These two subspecies probably

overlap in distribution throughout much of this region. I have also seen specimens in the

Royal BC Museum collected from the Chilcotin.

Aedes togoi (Theobald) is thought to be an invasive species from Asia; however, there

is evidence that this mosquito might be indigenous to rock pools along the-coast of BC

and adjacent Washington State (Sota eft al. 2015).

Aedes vexans nipponii (Theobald) is a subspecies of Ae. vexans from east Asia that

has recently been found in Ontario (Thielman and Hunter 2007). It is characterized by

the presence of a median longitudinal stripe of pale scales on the abdominal tergites,

which Ae. vexans vexans (Meighen) lacks (Tanaka et al. 1979). A specimen collected in

Cawston by P. Belton distinctly possesses this attribute and has been deposited in the

Spencer Entomology Collection at the UBC Beaty Biodiversity Museum.

Culex tarsalis Coquillett was previously thought to be found only in the southern half

of mainland BC (Belton 1983; Wood et al. 1979). However, this vector of West Nile

virus, Western equine encephalitis virus, and other viruses, has also recently been found

in man-made sites throughout Vancouver Island (Stephen ef a/. 2006) and as far north as

the Yukon (Peach 2018). It is likely to exist in suitable habitats throughout BC.

Culex territans Walker was reported as occurring across the south of BC by Belton

(1983) but has also been found as far north as the Yukon. (Belton and Belton 1990; Wood

et al. 1979). Recent records extend its range to Vancouver Island (Stephen et al. 2006).

These records imply that Cx. territans may be present throughout BC where suitable

habitat exists.

Culiseta particeps (Adams) was first reported by (Jackson et al. 2013) from locations

in Pitt Meadows and the Township of Langley. Additional specimens have also been

found in Vancouver, including an adult female collected in 1918 that was found in a

museum collection, and larvae that were found by the author in Prince Rupert. This

species is likely to be found all along the coast of BC.

J. ENTOMOL. SOC. BRIT. COLUMBIA 115, DECEMBER 2018 129

Coquillettidia perturbans (Walker) was previously known from suitable habitat

throughout mainland southern BC (Belton 1983). Recent work by Poirier and Berry

(2011) has revealed that this species is present as far north as Fort Nelson, and Stephen et

al. (2006) found it throughout much of Vancouver Island, as well. These new records

suggest it may be present in suitable habitat throughout most of BC, probably mirroring

the distribution of host plants such as cattails (Typha latifolia) (Poirier and Berry 2011).

ACKNOWLEDGEMENTS

I thank Karen Needham at the UBC Beaty Biodiversity Museum and Claudia Copley

at the Royal BC Museum for access to mosquito specimens, and the organizers of the

Whistler Bioblitz for mosquito-collecting opportunities. I also thank two anonymous

reviews for their constructive comments.

REFERENCES

Belton, E.M., and Belton, P. 1990. A review of mosquito collecting in the Yukon. Journal of the

Entomological Society of British Columbia, 87: 35-37.

Belton, P. 1983. The Mosquitoes of British Columbia. British Columbia Provincial Museum, Victoria,

British Columbia, Canada.

Belton, P. 2015. Mosquito species in BC and adjacent jurisdictions. Available from http://www.sfu.ca/

~belton/BCnames.pdf [accessed December 1, 2018]

Darsie, R.F.J. and Ward, R.A. 2005. Identification and Geographical Distribution of the Mosquitoes of

North America, North of Mexico. University Press of Florida, Gainesville, Florida, U.S.

Dyar, H. 1904. Notes on the mosquitoes of British Columbia. Proceedings of the Entomological Society

of Washington, 6: 7—14.

Gjullin, C. and Eddy, G. 1972. The Mosquitoes of the Northwestern United States. US Department of

Agriculture Technical Bulletin No. 1447 111.

Jackson, M., Belton, P., McMahon, S., McMahon, Hart, M, McCann, S., Azevedo, D., and Hurteau, L.

2016. The first record of Aedes (Hulecoeteomyia) japonicus (Diptera: Culicidae) and _ its

establishment in Western Canada. Journal of Medical Entomology, 53: 241-244.

Jackson, M., Howay, T., and Belton, P. 2013a. The first record of Culiseta particeps (Diptera: Culicidae)

in Canada. The Canadian Entomologist, 145: 115-116.

Jackson, M., Pyles, C., Breton, S., McMahon, T., and Belton, P. 2013b. British Columbia’s 50th

mosquito species, Aedes schizopinaz. Journal of the Entomological Society of British Columbia, 110:

38-39.

McCann, S. and Belton, P. 2015. A new record of Culex restuans Theobald (Diptera: Culicidae) in British

Columbia. Journal of the Entomological Society of British Columbia, 111: 13—14.

Peach, D.A.H. 2018. First record of Culex tarsalis Coquillett (Diptera: Culicidae) in the Yukon. Journal

of the Entomological Society of British Columbia In Press: 5 pgs.

Poirier, L.M. and Berry, K.E. 2011. New distribution information for Coquillettidia perturbans (Walker)

(Diptera, Culicidae) in northern British Columbia, Canada. Journal of Vector Ecology, 36: 461—463.

Sota, T., Belton, P., Tseng, M., Sen Yong, H., and Mogi, M. 2015. Phylogeography of the coastal

mosquito Aedes togoi across climatic zones: Testing an anthropogenic dispersal hypothesis. PLoS

ONE, 10: 1-13.

Stephen, C., Plamondon, N., and Belton, P. 2006. Notes on the distribution of mosquito species that

could potentially transmit West Nile virus on Vancouver Island, British Columbia. Journal of the

American Mosquito Control Association, 22: 553-556.

Tanaka, K., Mizusawa, K., and Saugstad, E.S. 1979. A revision of the adult and larval mosquitoes of

Japan (Including the Ryukyu Archipelago and the Ogasawara Islands) and Korea (Diptera:

Culicidae). Contributions of the American Entomological Institute, 16: 989.

Thielman, A. and Hunter, F. 2007. Photographic key to the adult female mosquitoes (Diptera: Culicidae)

of Canada. Canadian Journal of Arthropod Identification, 4: 1-117.

Wood, D.M., Dang, P.T., and Ellis, R.A. 1979. The Insects and Arachnids of Canada Part 6: The

Mosquitoes of Canada - Diptera: Culicidae. The Insects and Arachnids of Canada. Research Branch,

Agriculture Canada, Ottawa, Canada.

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NOTICE TO CONTRIBUTORS

The Journal of the Entomological Society of British Columbia is published online as submissions

are accepted. The JESBC provides immediate open access to its content on the principle that

making research freely available to the public supports a greater global exchange of

knowledge. Manuscripts dealing with all facets of the study of arthropods will be considered for

publication. Submissions may be from regions beyond British Columbia and the surrounding

jurisdictions provided that content is applicable or of interest to a regional audience. Review and

forum articles are encouraged. Authors need not be members of the Society. Manuscripts are peer-

reviewed, a process that takes about six weeks.

Submissions. The JESBC accepts only electronic submissions via the journal homepage: http://

journal.entsocbc.ca/. Style and format guidelines, open access fees, and other information for

authors can be found at the journal homepage.

Scientific notes. Scientific notes are an acceptable format for short reports. They must be two

journal pages maximum, about four manuscript pages. Scientific Notes do not use traditional

section headings, and the term "Scientific Note" precedes the title. A short abstract may be included

if desired. Notes are peer- reviewed in the same manner as regular submissions.

Natural History & Observations. Natural history observations are short — typically about two

pages maximum — pieces outlining entomological observations such as (but not exclusively) insect

outbreaks, population collapses, observations in unexpected locations, novel behaviors, etc. The

purpose of these papers will be to record potentially important phenomena — that may otherwise be

overlooked — for the use of future researchers. Papers of this sort were common in many of the

earlier years of the journal, and we feel that they still have a place, particularly as we move further

into the Anthropocene. Natural History and Observations pieces will be subject to peer review.

While not necessarily required, pieces that supply photographic, video, audio, two-witness,

voucher specimens, and/or other rigorous evidence of the observation will be more likely to be

accepted. Photographs will be included in the journal. Video, audio, or other such files should be

deposited in a DOI-based repository (such as figshare, Dryad, or an institutional repository) with

the DOI noted in the written piece. Voucher specimens should be deposited in recognized musuems

with a notation to that effect in the text. The JESBC encourages entomologists and other naturalists

working in British Columbia and the surrounding jurisdictions to consider submitting their

observations as part of the long term record of our regional entomological natural history.

Review and forum articles — Please submit ideas for review or forum articles for consideration to

the editor at journal@entsocbc.ca. Reviews should provide comprehensive, referenced coverage of

current and emerging scientific thought on entomological subjects. Forum articles of about 1000

words in length should provide opinions, backed by fact, on topics of interest to entomologists and

to the general public. Both review and forum submissions are only published in the journal

following full and rigorous peer review.

Electronic reprints. As an open access journal, article reprints from all issues of the journal can be

downloaded as PDFs from the journal homepage.

Back issues. Electronic back issues are available online free-of-charge.

Membership in the Society is open to anyone with an interest in entomology. Dues are $20 per

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receive electronic copies of Boreus (the newsletter of the Society), and when published, occasional

papers. Membership and membership renewals can be purchased at: http://entsocbc.ca/

membership/.

IAN LIB

WONT

020

Journal of the

Entomological Society of British Columbia

Volume 115 December 2018 ISSN#007 1-0733

Directors of the Entomological Society of British Columbia 2017-2018

First records of Baetis vernus Curtis (Ephemeroptera: Baetidae) in North America, with

morphological notes

Corrections for the Hemiptera: Heteroptera of Canada and Alaska

The bees of British Columbia (Hymenoptera: Apoidea, Apiformes)

Efficacy of diamide, neonicotinoid, pyrethroid, and phenyl pyrazole insecticide seed

treatments for controlling the sugar beet wireworm, Limonius californicus (Coleoptera:

Elateridae), in spring wheat

| SCIENTIFIC NOTES

A pheromone-baited pitfall trap for monitoring Agriotes spp. click beetles (Coleoptera:

Elateridae) and other soil-surface insects

Identifying larval stages of Orgyia antiqua (Lepidoptera: Erebidae) from British Columbia,

NATURAL HISTORY AND OBSERVATIONS

New records of Hymenoptera from British Columbia and Yukon

First Record of Culex tarsalis (Diptera: Culicidae) in the Yukon

An updated list of the mosquitoes of British Columbia with distribution notes

NOTICE TO CONTRIBUTORS Inside Back Cover