CONTENTS

I. FROM THE EDITOR 1-2

II. ARTICLES

J. L. RINGROSE, K. F. ABRAHAM and D. V. BERESFORD — New range records, and a

comparison of sweep netting and Malaise trap catches of horse flies and deer flies (Diptera:

Tabanidae) in northern Ontario 3-14

M. J. SKVARLA, J. L. LARSON and A. P. G. DOWLING — Pitfalls and preservatives: A

review. 15-43

D. B. LYONS, K. L. RYALL, S. M. PAIERO, G. C. JONES and L. VAN SEGGELEN — First

records of Ptosima walshii (Coleoptera: Buprestidae) in Canada 45-49

J. F. GIBSON, A. D. SLATOSKY, R. L. MALFI, T. ROULSTON and S. E. DAVIS — Eclosion

of Physocephala tibialis (Say) (Diptera: Conopidae) from a Bombus (Apidae: Hymenoptera)

host: A video record 51-60

S. M. PAIERO and S. A. MARSHALL — New Canadian and Ontario orthopteroid records,

and an updated checklist of the Orthoptera of Ontario 61-76

III. ESO OFFICERS AND GOVERNORS 2014-2015 77

IV. ESO OFFICERS AND GOVERNORS 2013-2014 inside front cover

V. FELLOWS OF THE ESO inside back cover

VI. APPLICATION FOR MEMBERSHIP inside back cover

VII. NOTICE TO CONTRIBUTORS

inside back cover

JOURNAL OF THE ENTOMOLOGICAL SOCIETY OF ONTARIO - VOLUME 145, 2014

JESO Volume 145, 2014

JOURNAL

of the

ENTOMOLOGICAL SOCIETY OF ONTARIO

VOLUME 145 2014

This is my last volume as editor and the first volume that is published in electronic

format only. I would have liked to see JESO continuing to publish hard copy up to and

including volume 150, in 2019, but that is not to be. Now, as soon as submitted manuscripts

are accepted for publication and edited technically they will be put up on the ESO website

for anyone to access freely. No need to wait until the end of the year to see your paper

published. Back issues of JESO are also available online, either through the website or the

Biodiversity Heritage Library. In time, JESO will have a greater and more useful online

presence because we expect to add all past journal articles to the ESO website as individual

searchable entities.

Whereas most, if not all, electronic journals have page charges for open access

publication JESO does not, so being able to publish without page charges and have anyone

access your paper immediately upon posting it on the ESO website should be a pretty good

incentive to publish there. Admittedly, people want their paper to appear in “high impact”

journals. But not every article is so brilliant or is of such general interest that it will be

accepted by the best journals. Rather, most articles are well-substantiated (hopefully) pieces

of research that add useful information, whether of interest to few or many, to the overall

store of sound scientific knowledge and are therefore worth publishing somewhere. JESO

is a good journal to consider. Papers on any aspect of entomology are welcome and the

scope need not be restricted to research done in Ontario or on Ontario insects. Taxonomists

wishing to publish in JESO should be aware of recent amendments to the International Code

of Zoological Nomenclature that detail the mandatory requirements needed to validate

new taxa published in e-journals. The essential point is that the work must be registered

in ZooBank before it is published. I will work with the technical editor and new editor to

determine how this is to be done.

This year’s volume is about half as long (75 pp.) as I had hoped but it is better than

the low of 55 pages in volume 140. It contains one scientific note, three scientific papers and

a review. Please check the ESO website and read/download the articles that interest you. It

would be very nice to see the number of published pages increase to a couple of hundred

pages each year.

It has been a pleasure to serve as JESO Editor for the past four years. I would

like to thank Tom Onuferko at York University, our new technical editor, for his help in

preparing the papers for this year’s volume, and the associate editors for looking after the

review process for manuscripts I passed on to them. Finally, I am pleased to introduce you

to Dr. Chris MacQuarrie in Sault Ste. Marie, who has graciously accepted the privilege

JESO Volume 145, 2014

and challenge of being the next JESO editor. Please send any manuscripts to him at Chris.

MacQuarrie@nrcan.rncan.gc.ca

I wish all of you the best for the coming year.

John T. Huber

Editor

New range records of tabanid flies in Northern Ontario

JESO Volume 145, 2014

NEW RANGE RECORDS, AND A COMPARISON OF SWEEP

NETTING AND MALAISE TRAP CATCHES OF HORSE FLIES

AND DEER FLIES (DIPTERA: TABANIDAE) IN NORTHERN

ONTARIO

J. L. RINGROSE 1 , K. F. ABRAHAM 2 , D. V. BERESFORE) 1 *

department of Biology, Trent University,

2140 East Bank Drive Peterborough, ON K9J 7B8

email, davidberesford@trentu.ca

Abstract J. ent. Soc. Ont. 145: 3-14

Horse flies and deer flies (Diptera: Tabanidae) were surveyed in northern

Ontario, Canada in 2011, at 11 sites, and 2012, at 12 sites using Malaise traps

and daily sweep netting. A total of 2168 tabanids representing 30 species:

10 Chrysops , 18 Hybomitra , and two Tabanus were collected. Malaise traps

caught fewer individuals than sweep netting but more species: 850 tabanids

of 28 species, eight of which were not caught by sweep netting. Sweep

netting caught 1318 tabanids of 22 species, with two not found in Malaise

trap samples. The first record of Hybomitra osburni (Hine) in Ontario, and

range extensions for several other species are given.

Published October 2014

Introduction

When habitats change, insect populations respond rapidly, up or down, depending

on the species characteristics (Niemela et al. 1993). These changes occur across a range

of temporal and spatial scales, and are unique for each species. This quality makes insect

diversity an efficient indicator for monitoring both short and long term environmental

changes (Danks 1992). The great variety of habitats occupied by insects in Ontario means

that studies that require tracking environmental change can benefit from using some insect

group for monitoring that change. For such work, up-to-date distributional data are needed

for the insect species of interest.

The eastern “Ring of Fire” region in the eastern area of Northern Ontario contains

large deposits of chromium and other minerals (Far North Science Advisory Panel 2010),

and anticipated large-scale extraction processes will alter insect diversity and distribution.

* Author to whom all correspondence should be addressed.

2 Wildlife Research and Development Section, Ontario Ministry of Natural Resources, 2140

East Bank Drive Peterborough, ON K9J 7B8

Ringrose et al.

JESO Volume 145,2014

To see the effect of such development, as well as possible effects of changing climates,

baseline distributional data for these areas are needed. With this purpose, the Ontario

Ministry of Natural Resources (OMNR) started a project in 2009 to survey insect diversity

and establish species distributions for Northern Ontario.

A widespread and easy to find and collect group are horse flies and deer flies

(Tabanidae). Pechuman et al. (1961) compiled the first comprehensive report on the

Tabanidae of Ontario. Teskey (1990) provided a more complete treatment of Tabanidae

in Canada. Since then, two pictorial keys, one on deer flies (Thomas and Marshall 2009)

and one on horse flies (Thomas 2011), added new range information to this group. Further

reports of sampling, especially from northern Ontario, continue to add distributional data to

our knowledge of Tabanidae (e.g., Beresford 2011).

Here we list the different species of Tabanidae caught in northern Ontario using

two different collecting methods and report on range extensions of several of them.

Materials and Methods

We sampled horse flies and deer flies in north-west Ontario in 2011 and north-east

Ontario in 2012 (Fig. 1, inset map) at 12 locations each year using two sampling methods,

sweep netting and Malaise trapping. Ringrose etal. (2013) provided detailed site descriptions

and locations. Generally, sampling took place within 1 km of remote field camps that were

accessed by helicopter. The 2011 sampling was completed in the western half of Ontario

boreal forest within a 150 km radius of the First Nations communities of Big Trout Fake and

Sandy Fake. The 2012 sampling occurred in the northeastern part of the province within a

150 km radius of the First Nations community of Fort Albany, Ontario. Sampling dates were

from 5 June to 17 July in 2011, and from 5 June to 15 July in 2012.

Tabanids were sampled each day by two methods, Malaise traps (6m trap model

no. 2877, BioQuip Products 2321 Gladwick Street, Rancho Dominguez, CA 90220, USA),

and sweep netting. The collecting heads of the Malaise traps were filled with 80% denatured

ethanol to kill and preserve captured tabanids. These were emptied and replaced each day

at 9 pm.

Sweep netting was completed at midday as the surveyor (JFR) walked slowly,

sweeping for 5 minutes any Tabanidae that assembled around the researcher. The netted

samples were killed by placing the end of the bag in large killing bottles charged with

acetone. Specimens were then removed from the net and stored in bottles filled with 80%

denatured ethanol. The ethanol in each storage bottle was replaced after 24 hours.

All tabanids were pinned and identified by JFR and DVB using the keys found in

Teskey (1990), Thomas and Marshall (2009) and Thomas (2011). The main pinned collection

is stored in insect cabinets at Trent University, Biology Department, Peterborough, Ontario.

A reference collection of voucher specimens is housed at the Canadian National Collection

of Insects, Ottawa.

Analysis

A list of the expected species was produced from the distribution records reported

in the keys listed above. For those species that did not have records in northern Ontario,

New range records of tabanid flies in Northern Ontario

JESO Volume 145, 2014

we reasoned that any species with records that straddled northern Ontario (either east and

west, or north and south of the sampling regions) was likely present in northern Ontario.

We compared this expected number of species to the predicted number which we calculated

using the lognormal distribution method (Preston’s method) as described in Ludwig and

Reynolds (1988). This approach allows one to predict the number of species present in an

area from sampling data. It is based on a general observation that most species are more or

less moderately abundant (the middle region of the lognormal distribution), a few species

are very abundant (forming the right tail of the lognormal distribution) and a few are very

rare (the left tail of the lognormal distribution). In practice, it enables one to predict the

number of rare species that are expected but which might have been missed. Parameters

for the lognormal model were fitted using the SOLVER function in MICROSOLT EXCEL

2007.

Catch data were analyzed using the online rarefaction calculator from the

University of Alberta (http://www.biology.ualberta.ca/jbrzusto/rarefact.php), to determine

the effects of collection size on the number of species collected, as well as to compare

trapping methods.

Results

Range records and extensions

From our assessment of published range maps, we expected to find 3 1 species of

Tabanidae: 23 with records from across northern Ontario in the regions where we conducted

our study (11 Hybomitra , 8 Chrysops , 2 Aty lotus, 2 Tabanus ), and 8 with ranges that straddle

our study regions (5 Hybomitra , 2 Aty lotus, and 1 Haematopota).

We collected 2168 tabanids from 30 species over the two years: 839 from 24

species in northwest Ontario (2011 sampling), and 1329 from 25 species in northeast Ontario

(2012) (Tables I and II, Fig. 1). We found 18 Hybomitra, 10 Chrysops, and 2 Tabanus, but

no Atylotus, or Haematopota.

The expected number of species was calculated to be 26 (lognormal fitted

parameters, a = 0.28, So = 4.09, yl = 7.93, p = 0.34, d.f.=7) in the western collections

(2011) and 28 (fitted parameters, a = 0.24, So = 3.73, yl = 9.89, p = 0.27, d.f. = 8) in the

eastern collections (2012), and 33 species for the combined data set (fitted parameters, a =

0.23, So = 4.27, x2 = 7.72, p = 0.56, d.f. = 9).

The three most abundant species caught in the northwest (2011) were Chrysops

excitans Walker (35%), Hybomitra epistates Osten Sacken (21%) and H lurida (Fallen)

(19%). In the northeast (2012) the most abundant were Hybomitra affinis (Kirby) (33%), C.

excitans (22%) and H lurida (19%).

New Ontario record

Our collection of three individuals of Hybomitra osburni (Hine) (two in 2011 and

one in 2012) are the first records of this species in Ontario. This species has been collected

in all western provinces and the Yukon Territory (Teskey 1990) but was previously not

known to occur east of Manitoba.

Ringrose et al. JESO Volume 145, 2014

TABLE 1. Tabanidae species and number of specimens collected in 2011 and 2012 using

Malaise traps and sweep netting, with abundance records.

Species

2011

2012

Total

Malaise netted

Malaise

netted

Chrysops ater Macquart

Chrysops cuclux Whitney

Chrysops dawsoni Philip

Chrysops excitans Walker

151

245

225

658

Chrysops frigidus Osten Sacken

Chrysops mitis Osten Sacken

Chrysops niger Macquart

Chrysops nigripes Zetterstedt

Chrysops venus Philip

Chrysops zinzalus Philip

Hybomitra affinis (Kirby)

268

354

Hybomitra arpadi (Szilady)

Hybomitra criddlei (Brooks)

Hybomitra epistates Osten Sacken

124

153

291

Hybomitra frontalis (Walker)

Hybomitra frosti Pechuman

Hybomitra hearlei (Philip)

Hybomitra illota (Osten Sacken)

Hybomitra lasiophthalma (Macquart)

Hybomitra lurida (Fallen)

104

131

121

415

Hybomitra minuscula (Hine)

Hybomitra nuda (McDunnough)

Hybomitra osburni (Hine)

Hybomitra pechumani Teskey & Thomas

Hybomitra tetrica (Marten)

Hybomitra trepida (McDunnough)

Hybomitra typhus (Whitney)

Hybomitra zonalis (Kirby)

Tabanus marginalis Fabricius

Tab anus vivax Osten Sacken

Total specimens

151

688

699

630

2168

Total species

TABLE 2. Tabanidae species listed for each sampling location and date. Only 11 sampling sites are included for 2011 as planned samples

were damaged by black bears.

New range records of tabanid flies in Northern Ontario

JESO Volume 145, 2014

Year 2011 2012

July 10 - July 16

92° E 43"

54° 9' 29"

July 10 - June 16

88° 54' 51"

53° 45' 34"

July 3 - July 9

89° 40' 42"

54° 25' 49"

July 3 - July 9

89° 6' 27"

53° 12' 8"

June 26 - July 2

82° 49' 1"

52° 28' 26"

June 26 - July 2

83° 17' 23"

51° 29' 53"

June 19 - June 25

82° 41' 2"

52° 53' 25"

June 19 - June 25

81° 39' 22"

51° 58' 8"

June 12 - June 18

81° 50' 56"

51° 39' 7"

June 12 - June 18

80° 23' 10"

51° 26' 40"

June 5 - June 1 1

82° 39' 13"

51° 55' 53"

June 5 - June 1 1

81° 57' 47"

52° 46' 34"

July 14 - July 21

92° 46' 3"

53° 44' 12"

July 6 - July 13

93° 32' 9"

53° 36' 8"

July 6 - July 13

91° 49' 8"

52° 27' 37"

June 28 - July 5

94° 13' 38"

52° 49' 27"

June 28 - July 5

93° 2' 32"

53° 27' 39"

June 16 - June 23

88° 33' 33"

54° 28' 18"

June 16 - June 23

90° 21' 37"

54° 27' 1"

June 8-15

92° E 43"

54° 9' 29"

June 8-15

88° 54' 51"

53° 45' 34"

May 3 1 - June 7

89° 40' 42"

54° 25' 49"

May 3 1 - June 7

89° 6' 27"

53° 12' 8"

Sampling dates

Longitude (West)

Latitude (North)

Species

C. ater

C. cuclux

C. dawsoni

C. excitans

TABLE 2 continued...

Ringrose et al.

JESO Volume 145,2014

Year 2011 2012

July 10 - July 16

July 10 - June 16

July 3 - July 9

June 26 - July 2

June 19 - June 25

June 12 - June 18

X X

June 5 - June 1 1

July 14 - July 21

July 6 - July 13

June 28 - July 5

June 16 - June 23

June 8-15

X X

May 3 1 - June 7

Sampling dates

Species

C. frigidus

C. mitis

C. niger

C. nigripes

C. venus

C. zinzalus

H. affinis

H. arpadi

H. criddlei

H. epistates

H. frontalis

H. frosti

H. hearlei

H. illota

TABLE 2 continued...

New range records of tabanid flies in Northern Ontario

JESO Volume 145, 2014

Year 2011 2012

July 10 - July 16

July 10 - June 16

July 3 - July 9

X X

June 26 - July 2

June 19 - June 25

June 12 - June 18

X X

June 12 - June 18

June 5 - June 1 1

July 14 - July 21

X X

July 6 - July 13

June 28 - July 5

June 16 - June 23

X X

June 16 - June 23

June 8-15

May 3 1 - June 7

Sampling dates

Species

H. lasiophthalma

H. lurida

H. minuscula

H. nuda

H. osburni

H. pechumani

H. tetrica

H. trepida

H. typhus

H. zonalis

T. marginalis

Ringrose et al.

JESO Volume 145,2014

Range extensions

We report nine new northern range records in Ontario. They are: Chrysops cuclux

Whitney, C. niger Macquart, C. venus Philip, Hybomitra criddeli (Brooks), H. epistates ,

H lasiophthalma (Macquart), H. pechumani Teskey & Thomas, H. tetrica (Marten), H.

trepida (McDunnough), and Tabanus vivax Osten Sacken. In addition, we provide three

new western records of Hybomitra for Ontario: H. minus cula (Hine), H. typhus (Whitney),

and H. frosti Pechuman.

Gap infill

Chrysops ater Macquart is described as an abundant species having a general

northern distribution in Canada south of the tree line (Teskey 1990). Our collection of a

single specimen in 2012 is therefore not a surprise; however, there has been little collection

in northern Ontario so our collection has filled a gap between previous collecting locations.

It is perhaps surprising that it was so rare in our collections. Our records of Chrysops

dawsoni Philip and C. frigidus Osten Sacken are consistent with known ranges.

Chrysops excitans , and C. mitis Osten Sacken, and to a lesser extent C. nigripes

Zetterstedt and C. zinzalus Philip, are found in Canada south of the tree line (Teskey 1990),

and have been reported from Polar Bear Provincial Park (Beresford 2011). Our records are

consistent with these reports.

We caught eight species of Hybomitra , consistent with known ranges: H. affinis ,

the most abundant and widely distributed Canadian species of Tabanidae (Teskey 1990;

Thomas 2011), H. arpadi (Szilady), H. frontalis (Walker), H. hearlei (Philip), H. illota

FIGURE 1 . Rarefaction analysis showing the expected number of species (y axis) for smaller

total catch sizes (x axis), for 2011 (closed circles) and 2012 (open circles). The inset map

shows sample locations in both years. Error bars represent standard deviations.

New range records of tabanid flies in Northern Ontario

JESO Volume 145, 2014

(Osten Sacken), H. lurida , H. nuda (McDunnough), and H. zonalis (Kirby).

Tabanus marginalis Fabricius has been collected from across Canada except on

the east coast (Teskey 1990). While the known range encompasses our sampling locations

(northern Manitoba and northern Quebec) our records are the northernmost from Ontario.

Trap comparison

The Malaise sampling caught fewer individuals than sweeping yet produced more

species. Malaise traps collected 850 specimens of 28 species (151 in 2011 and 699 in 2012);

sweep netting collected specimens 1318 of 22 species (688 in 2011 and 630 in 2012) (Fig.

2 ).

30 i

0 n 1 1 1 1

0 200 400 600 800

total number of specimens

FIGURE 2. Rarefaction analysis of 2011 and 2012 data, separated by trapping method.

Malaise traps (circles) and sweep netting (diamonds) in 2011 (closed) and 2012 (open).

Error bars represent standard deviations.

Ringrose et al.

JESO Volume 145,2014

Discussion

Comments on range extensions and new distributional locations are based on range

maps from Teskey (1990), Thomas and Marshall (2009) and Thomas (2011). Because of the

few intensive studies from northern Ontario we expected to add range records for many of

the species we collected.

Our range map assessment underestimated by four the number of species we

expected to catch, namely, 8 Chrysops , 16 Hybomitra , and two Tabanus; we caught 10

Chrysops , 18 Hybomitra , and two Tabanus. The lognormal prediction of 33 species was

three more than what we found. Our survey did not include catches from August, and we

expect there are more species in our study region that we did not manage to collect.

The two trapping methods collected different species (Table 1). Some species were

abundant in both trapping methods, e.g. C. excitans , H. epistates , and H. lurida. Hybomitra

zonalis was abundant in the Malaise collections, whereas H. affinis and C. mitis were

abundant in the netted samples. Two species, C. cuclux and H. nuda , were absent from

Malaise traps, and eight species, C. ater , C. frigidus , C. niger, C. venus , C. zinzalus , H

frosti , H. hearlei , and T. vivax were absent from the sweeps. When differences are examined

within each year, the effect of trapping method becomes even more pronounced: nine

species caught only by sweep netting and five only in Malaise traps in 2011; one species

caught only by sweep netting and 10 only in Malaise traps in 2012 (Table 1, Fig 2). These

results highlight the importance of collecting using a variety of methods in insect surveys to

overcome catch biases. Other methods used to sample Tabanidae include larval collection

(Philip 1928), chemical attractants (i.e., C0 0 or Octenol), baited traps such as the Nzi trap

(Mihok et al. 2007), traps designed to act as visual cues for host seeking Tabanidae such

as the unbaited Nzi traps (Mihok 2002), and Manitoba traps (Thorsteinson et al. 1964).

Any trap designed to work using visual or olfactory cues for host seeking tabanids would

produce high catches, most likely of host seeking females, but it is not known if these higher

catches would result in proportionately more or different species. Tarval collections do not

rely on adult flight or host seeking, but are limited by the habitat that is searched (Philip

1928).

Generally, the most abundant species were caught over the longest period, with

some exceptions. In 2011, H. affinis was the most abundant species (275 specimens), but

was only caught during five sampling sessions (Table 2) whereas H. arpadi (33 specimens)

and H. lasiophthalma (21 specimens), both relatively uncommon, were also captured during

five of the sampling sessions. In 2012, H affinis (79 specimens) was caught over 9 sessions,

but was less abundant than H lurida (252 specimens), which was caught over 7 sessions

(Tables I and II).

Acknowledgements

The authors would like to thank the Ontario Ministry of Natural Resources

Northeast Science and Information Section and Wildlife Research and Development Section

for project coordination and logistics, and Far North Branch for funding. Additional travel

support for JFR was provided by a Northern Scientific Training Program (NSTP) grant

New range records of tabanid flies in Northern Ontario

JESO Volume 145, 2014

through Trent University. We give special thanks to Dean Phoenix and the field crews of

the Far North Biodiversity Project in 2011 and 2012. We would also like to thank the First

Nations communities of Kitchenuhmaykoosib Inninuwug, Keewaywin and Fort Albany for

their hospitality and generosity.

References

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records. Journal of the Entomological Society of Ontario 142 : 19-27.

Danks, H. V. 1992. Arctic insects as indicators of environmental change. Arctic 45 : 159—

166.

FNSAP. 2010. Science for a Changing Far North. The Report of the Far North Science

Advisory Panel. A report submitted to the Ontario Ministry of Natural Resources.

Queen’s Printer for Ontario, Toronto, ON. 141 pp.

Gillis, M. D., Omule, A. Y. and Brierley, T. 2005. Monitoring Canada’s forests: the national

forest inventory. Forestry Chronicle 81 : 214-221.

Fudwig J. A. and Reynolds, J. F. 1988. Statistical Ecology: a primer on methods and

computing. John Wiley & Sons Inc., New York, NY. 337 pp.

Mihok, S. 2002. The development of a multipurpose trap (the Nzi) for tsetse and other

biting flies. Bulletin of Entomological Research 92 : 385-403

Mihok, S., Carlson, D. A. and Ndegwa, P. N. 2007. Tsetse and other biting fly response

to Nzi traps baited with octenol, phenols and acetone. Medical and veterinary

entomology 1 : 70-84.

Niemela, J., Fangor D. and Spence, J. R. 1993. Effects of clear-cut harvesting on boreal

ground-beetle assemblages (Coleoptera: Carabidae) in western Canada.

Conservation Biology 7: 551-561.

OMNR. 2012. Wildlife research: far north biodiversity. Ontario Ministry of Natural

Resources. Available online at: http://www.mnr.gov.on.ca/en/Business/Wildlife/

2ColumnSubPage/STDPROD_09995 5 .html.

Pechuman, F. F., Teskey, H. J. and Davies, D. M. 1961. The Tabanidae (Diptera) of Ontario.

Proceedings of the Entomological Society of Ontario 91 : 77-121.

Philip, C. B. 1928. Methods of collecting and rearing the immature stages of Tabanidae

(Diptera). The Journal of Parasitology 14 : 243-253.

Ringrose, J. F., Abraham, K. F. and Beresford, D. V. 2013. New range records of mosquito

species from northern Ontario. Journal of the Entomological Society of Ontario

144 : 3-14.

Teskey, H. J. 1990. The horse flies and deer flies of Canada and Alaska (Diptera: Tabanidae).

The Insects and Arachnids of Canada, Part 16: Agriculture Canada. Publication

1838. 381 pp.

Thomas, A. W. and Marshall S. A. 2009. Tabanidae of Canada, east of the Rocky Mountains

1 : a photographic key to the species of Chrysopsinae and Pangoniinae (Diptera:

Tabanidae). Canadian Journal of Arthropod Identification 8. Available online

at: http://www.biology.ualberta.ca/bsc/ejournal/tm_08/tm_08.html, doi: 10.3752/

cjai.2009.08.

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JESO Volume 145,2014

Thomas, A. W. 2011. Tabanidae of Canada, east of the Rocky Mountains 2: a photographic

key to the genera and species of Tabanidae (Diptera: Tabanidae). Canadian Journal

of Arthropod Identification 13 . Available online at: http://www.biology.ualberta.

ca/bsc/ejournal/t_13/t_l 3.html, doi:10.3752/cjai.2011.13.

Thorsteinson, A. J., Bracken, G. K. and Hanec, W. 1964. The Manitoba horse fly trap. The

Canadian Entomologist 96 : 166.

Pitfalls and preservatives: A review

JESO Volume 145, 2014

PITFALLS AND PRESERVATIVES: A REVIEW

M. J. SKVARLA 1 *, J. L. LARSON 2 , A. P. G. DOWLING 3

'^Department of Entomology, University of Arkansas,

Layetteville, Arkansas, 72701

email, mskvarla36@gmail.com

Abstract J. ent. Soc. Ont. 145: 15-43

An extensive review of the factors that affect the performance of arthropod

pitfall traps is given. Liquid preservatives are discussed in a separate section

because the choice affects the quality and composition of taxa collected in

pitfalls.

Published November 2014

Introduction

Pitfall traps are a popular method for collecting ground beetles, spiders, ants and

other epigeal arthropods (Westberg 1977; Niemela et al. 1992; Bestelmeyer et al. 2000;

Southwood & Henderson 2000; Phillips & Cobb 2005). While many shorter, general

overviews exist (e.g., general techniques: Balogh 1958; Duffey 1972; Bestelmeyer et al.

2000; Southwood and Henderson 2000; Woodcock 2005; issues with pitfalls: Adis 1979),

none have exhaustively examined the published literature recently. Herein we present such

a review with the hope it will provide a sound base for those incorporating pitfall traps into

research.

While the choice of preservative will affect the quality of specimens in any type of

trap, it is a critical decision in pitfalls for several reasons. Chiefly, preservatives differentially

attract and repel select arthropod taxa, which will affect the composition of taxa collected

(Weeks & McIntyre 1997). Additionally, pitfalls are often set without covers in open fields,

so lose more preservative through evaporation than other traps and are affected to a greater

degree by rain and dilution by rainwater (Porter 2005). Therefore, we include a section

detailing possible positives and negatives of preservatives used in pitfall traps.

* Author to whom all correspondence should be addressed.

2 Douglas-Sarpy County Extension, University of Nebraska, 8015 W. Center Rd, Omaha,

Nebraska, 68135

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Pitfall Traps

Pitfall traps were first described by Hertz (1927) and shortly thereafter by Barber

(1931) (Fig. 1) for collecting cave-inhabiting insects. A pitfall trap is simple in design,

consisting of a collecting container buried flush with the ground that passively collects

epigeal organisms that accidentally fall into the trap. It may be constructed from any

container large enough to hold the target organism, including a large bucket for reptiles

or small mammals (Ellis 2013), small plastic cup for larger insects such as Carabidae and

large Formicidae (Luff 1975; Abensperg-Traun & Steven 1995), or a glass test tube for

small insects such as most Formicidae and small Carabidae (Luff 1968; Abensperg-Traun

& Steven 1995). Pitfall traps are widely used in biodiversity surveys as they are cost-

effective, ecologically sensitive, collect large numbers of arthropods (Gist & Crossley

1973; Ekschmitt et al. 1997; Southwood & Henderson 2000; Work et al. 2002), and collect

nocturnal species missed by other methods (Tormala 1982; Samways 1983; Donnelly &

Gilmee 1985; Huusela-Veistola 1996).

Pitfall traps have been used to sample many arthropod groups, including

Scorpionida (Tourtlotte 1974; Margules et al. 1994); Isopoda (Hamner et al. 1969; Hayes

1970; Paoletti & Hassall 1999; Hornung et al. 2007); Diplopoda (Van der Drift 1963;

Kurnik 1988; Mesibov et al. 1995; Kime 1997; Snyder et al. 2006), Chilopoda (Kurnik

1988; Friind 1990; Adis 1992; Shear & Peck 1992; Voigtlander 2003), and Symphyla (Adis

1992; Shear & Peck 1992; Clark & Greenslade 1996); Araneae (Duffey & Millidge 1954;

Muma 1973; Uetz 1977; Corey & Taylor 1988; Bultman 1992; Koponen 1992; Bauchhenss

1995; Buddie et al. 2000); Acari (Zacharda 1993; Wickings 2007; Klosin'ska et al. 2009;

Mayoral & Barranco 2009; Wohltmann & Mqkol 2009; Lopez-Campos & Vazquez-Rojas

2010; Clark 2013); Collembola (Joosse-van Damme 1965; Pedigo 1966; Budaeva 1993;

Cole et al. 2001; Frampton et al. 2001); Coleoptera (Backhand & Marrone 1997; Simmons

et al. 1998; Arbogast et al. 2000) including Carabidae (Anderson 1985; Kalas 1985;

Cameron & Reeves 1990; Epstein & Kulman 1990; Togashi et al. 1990), Tenebrionidae

(Ahearn 1971), Staphylinidae (Anderson 1985; Braman & Pendley 1993; Ekschmitt et al.

1997), Scarabaeoidea (Young 1981; Peck & Howden 1985; Martinez et al. 2009; Anla§ et

al. 2011; Thakare et al. 2011), and certain Latridiidae (Hartley et al. 2007); Formicidae

(Van der Drift 1963; Greenslade 1973; Anderson 1991; Abensperg-Traun & Steven 1995;

Bestelmeyer et al. 2000); and even terrestrial Amphipoda (Craig 1973; Margules etal. 1994)

and Decapoda(Williams et al 1985; Smith et al. 1991; Hamr & Richardson 1994; McGrath

1994; Mclvor & Smith 1995). Of these taxonomic groups, ground-dwelling Araneae and

Coleoptera have been the most studied (Westberg 1977).

Variations on the basic trap have been developed, including more elaborate traps

for use under snow (Kronestedt 1968; Steigen 1973); live traps with a layer of gauze that

keeps trapped organisms from drowning in rainwater (Duffey 1972); modifications that

allow excess rainwater to drain before overflowing the trap (Duffey 1972; Porter 2005);

integrated internal funnel and rain cap (Fichter 1941); collecting cup integrated into a

larger structure with a base or ramp (Muma 1970); use of holes or slits in the side of a

container so an integrated cap can be used (Fig. 2) (Nordlander 1987; Lemieux & Lindgren

1999); modifications to facilitate emptying (Rivard 1962), including automated devices for

segregating trap catch over time (Williams 1958; Blumberg & Crossley, 1988; Buchholz

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2009); designs to reduce mortality of vertebrate bycatch including floating shelters and wire

mesh (Kogut & Padley 1997; Pearce etal. 2005); and inexpensive designs using commonly

discarded household materials (Morril 1975; Clark & Bloom 1992). Other techniques,

such as using an auger bit to drill placement holes for small diameter traps, and equipment,

such as a device that can pull traps out of placement holes without kneeling or disturbing

the surrounding soil, have been developed to make pitfall trapping easier (Vogt & Harsh

FIGURES 1-2. 1, Pitfall trap described by Barber for collecting cave- inhabiting insects.

After Barber (1931). 2, Pitfall trap modified with entrances in the side of the collection cup,

which discourages vertebrates from entering the trap and allows the use of an integrated rain

cap. Modified from Nordlander (1987) with permission.

Skvarla et al.

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2003).

Barrier fences have been employed, either with a single pitfall situated in the

middle of the fence or with pitfalls at the end of the fence (Fig. 3) (Haeck 1971; Meijer

1971; Reeves 1980; Durkis & Reeves 1982). Linear pitfall traps constructed from house

gutters have been employed with success in certain situations, such as investigating the

speed and timing of insect populations moving between habitats (Pamanes & Pienkowski

1965; Goulet 1974; Pausch et al. 1979).

Ramp traps collect arthropods similarly to pitfall traps, but rather than being

sunk into the ground target taxa are directs upwards into the trap via ramps; this allows

FIGURES 3-4. 3, Pitfall traps (modified from Nordlander 1987) on either side of a barrier

fence. 4, Ramp trap.

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them to be employed where conventional pitfalls cannot, such as where digging is difficult

(e.g., on rocks or in caves) or prohibited by law (Bouchard et al. 2000; Campbell et al.

2011). Bostanian et al. (1983) proposed the first ramp trap design, which is constructed

from metal, making it rather bulky and expensive and biased towards large ground beetles.

Bouchard et al. (2000) proposed a revised design that utilizes plastic sandwich containers

and plastic ramps, rendering it light-weight and inexpensive (Fig. 4). Ramp traps have

been successfully employed in caves (Campbell et al. 2011), areas polluted due to industrial

mining (Babin-Fenske & Anand 2010), orchards (Smith et al. 2004), and vineyards (Goulet

et al. 2004). Ramp traps capture a higher abundance and diversity of epigeal spiders than

conventional pitfall traps, though when comparing other taxa (e.g., beetles) they collect

a different species composition, thus making direct comparison between the trap types

difficult or impossible (Pearce et al. 2005; Patrick & Hansen 2013). Additionally, ramp

traps capture fewer vertebrates than conventional pitfall traps (Pearce et al. 2005).

Colored pan traps, sometimes referred to as water traps, are generally used

to collect flying insects via visual response to color cues (e.g. yellow, blue, purple or

red) (Kirk 1984; Aguiar & Sharkov 1997; Feong & Thorp 1999; Pucci 2008; Gollan et

al. 2011). While pan traps are generally set on or above the ground, they may be sunk

into it, effectively becoming pitfall traps that also attract and capture flying insects.

Issues with pitfall traps

Objections have been raised to the use of pitfall traps in ecological studies (Adis

1979; Majer 1997; South wood & Henderson 2000) because they do not evenly catch

different taxa for several reasons:

1 . Different taxa react differently at the lip of the trap. Gerlach et al. (2009) found

that millipedes show the most trap-avoidant behavior (20-60%) and carabids show the

least (10-25%); overall they found an average of 28% of taxa that encountered a trap were

caught, with a range of less than 5% ( Enantiulus nanus (Fatzel, 1884) (Julidae)) to 70%

(Pterostichus burmeisteri Herr, 1838 (Carabidae)). Fuff (1975) found approximately 75%

of Carabidae that encounter the edge of a pitfall are collected. In mark-recapture studies,

some species become trap-shy if they have been caught previously while other species do

not (Benest 1989).

2. Activity level (Ekschmitt et al. 1997), which is affected by variables such as

species-specific behavior (Greenslade 1964; Curtis 1980; Anderson 1991; Topping 1993;

Spence & Niemela 1994; Obrist & Duelli 1996); differences between gender and age (Hayes

1970; Benest 1989; Topping & Sunderland 1992; Thomas et al. 1998) including mate-

searching (Tretzel 1954), post-copulatory dispersal of females (Merrett 1967) and searching

for oviposition sites (Duffey 1956); weather (Williams 1940; Briggs 1961; Greenslade 1961;

Juillet 1964; Ericson 1979; Drake 1994); vegetation (Deseo 1959; Greenslade 1964; Novak

1969; Baars 1979), habitat structure (Melbourne 1999; Melbourne et al. 1997; Thomas et

al. 1998), and habitat type (Melbourne et al. 1997); size (Fuff 1975; Thiele 1977; den Boer

1981; Franke et al. 1 988) and speed (Braune 1 974; Adis 1 976); and hunger and prey density

(Gram 1971; Muller 1984; Henrik & Elcbom 1994), also affect the number of organisms

trapped, both within and between taxa (Southwood, & Henderson 2000) and are more

influential factors than population size (Briggs 1961) in determining trap catch.

3. Farger species are caught in significantly higher numbers than smaller species

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(Carabidae: Franke et al 1988; Spence & Niemela 1994). Several reasons have been

suggested for this. Larger, faster beetles are successfully caught a higher percentage of the

time than smaller, slower beetles (Braune 1974; Adis 1976) - though some authors have

found size and speed do not affect the ability to be caught (Luff 1975; Halsall & Wratten

1988). Smaller beetles may escape more readily from traps because scratches and soil

on trap walls may be enough to support their mass as they try to climb out whereas larger

beetles fall (Spence & Niemela 1994).

4. Species-specific morphology can affect escape ability; e.g., Demetrias

atricapillus (L.) has adhesive setae on the underside of the tarsi that allow it to climb out of

pitfalls more easily than other similarly sized carabids (Halsall & Wratten 1988).

5. Pitfall traps do not accurately reflect absolute density of the organisms sampled.

This has been demonstrated in the field (Gram 1959; Briggs 1961; Mitchell 1963; Marsh

1984; Topping & Sunderland 1992) and experimentally in a caged system (Lang 2000)

- though caution should be exercised interpreting caged results as they may be skewed by

“trap-happy” beetles that prefer dry pitfalls as refugia (Adis 1979, citing Thomas & Sleeper

1977) and may suffer from “Kreb’s effect” (Mac Arthur 1984). However, it should also be

noted that some studies have recorded 73-96% capture rates of marked beetles in caged

systems (Bonkowska & Ryszkowski 1975; Dennison & Hodkinson 1984; Desender et al.

1985; Desender & Maelfair 1986; Clark et al. 1995; Holland & Smith 1999) and one study

found no difference between population estimates of millipedes, spiders, and beetles based

on hand collecting or pitfalls in a caged system (Gist & Crossley 1973), suggesting such

systems may accurately reflect absolute density in certain situations with specific taxa.

In response to these criticisms, various calculations have been proposed to correct

for the differences between taxa collected and true population density based on locomotory

activity and motility range (Heydemann 1953; Tretzel 1955; Braune 1974; Thomas &

Sleeper 1977; Kuschka et al. 1987; Stoyan & Kuschka 2001; see also Seifert 1990), though

these have been rejected by others (Adis 1979; Muller 1984; Franke et al. 1988; Gerlach et

al. 2009).

Additionally, it has been argued that samples pooled over an entire season correctly

represent local species abundance as variations due to weather and other factors that affect

activity level are averaged out (Baars 1979; den Boer 1986; Luff 1982). Results of other

studies are conflicting, with some showing a large amount of variation between sampling

periods in similar habitat when the sampling periods are short (Niemela et al. 1986), and

others showing that traps set for short periods caught all species accumulated by longer

trapping periods (Niemela et al. 1990; Borgelt & New 2006). In addition, much of the cited

research has only examined carabids caught by pitfalls. When collecting other taxa, pitfalls

may estimate absolute population density relatively well (ants: Andersen 1991; Vorster et

al. 1992; Lindsey & Skinner 2001; cursorial spiders: Muma & Muma 1949; Duffey 1962;

Huhta 1971; Uetz & Unzicker 1976; tenebrionids: Thomas & Sleeper 1977).

Certain ecological questions, such as comparing taxa along a successional gradient

(Bultman & Uetz 1982) or between similar plots (Koivula et al. 1999), may be answered as

taxa will be equally biased to pitfall traps along the gradient or between plots.

Pitfalls can be used to answer non-ecological questions, such as investigating

the phenology (Maelfait & Baert 1975), seasonal and circadian activity (Williams

1959a, b; Williams 1962; Breymeyer 1966a, b; Doane & Dondale 1979), and lifespan

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(Goulet 1974) of commonly collected taxa, estimating the timing of movement of

epigeal species between habitats (Pamanes & Pienkowski 1965; Pausch et al. 1979),

and estimating dispersal using mark-release-recapture methods (Ericson 1977; Best et

al. 1981). They also can be employed in taxonomic surveys, though should be paired

with other sampling techniques that complement the deficiencies of pitfalls (Majer 1997)

Pitfall trap design

If pitfall traps are to be employed, several considerations must be made as there are

many factors that can affect the taxa collected.

Effects of shape, size, and material of receptacle . The shape of the trap affects the

composition and number of taxa collected (Cheli & Corley 2010). Pitfalls may be straight-

sided or round (Southwood & Henderson 2000), depending on the container used; however,

round and straight-edged traps with the same perimeter length catch different numbers of

specimens (Braune 1974; Luff 1975; Adis 1976; Spence & Niemela 1994).

Different diameters of pitfall trap collect different taxa at different rates. When

examining ants, larger diameter pitfalls catch more species, though differences are primarily

due to differential capture rates of rare species (Abensperg-Traun & Steven 1995). Work

et al. (2002) compared catch rates and species richness of Carabidae, Staphylinidae, and

Araneae across five diameters (4.5, 6.5, 11, 15, and 20 cm) of pitfall traps; they found that,

after standardizing circumference, small traps caught more small carabids and staphylinids

and large traps caught more wolf spiders. Luff (1975) found that small traps (2.5 cm dia.)

were the most efficient at catching small species of carabids, while large traps (10 cm dia.)

caught relatively more large beetles; however, their small traps were made of glass and large

traps made of metal, which probably had a confounding effect on the results. Brennan et al.

(1999) found the largest and second largest traps (17.4 and 11.1 cm dia.) they tested caught

the most diverse assemblage of species, though considered the smaller of the two traps more

appropriate for sampling spiders as it may decrease the potential of capturing non-target

species. One option when using larger traps is to add a funnel to the trap in order to increase

trap retention (Vlijm et al. 1961).

Another aspect of size is the depth of the trap. Shallow (8 cm) and deeper (15

cm) pitfalls do not effect ant diversity capture (Pendola & New 2007), therefore, when

targeting ants, shallow pitfalls are preferred as small vertebrates, such as skinks, may escape

more easily from them, thus reducing vertebrate bycatch. However, this has only been

demonstrated in ants and may not hold true for large insects, such as some carabids, which

are bigger than some small vertebrates.

Pitfall traps used to collect insects have been constructed out of glass (Briggs

1961; Greenslade 1964; Borgelt & New 2006; Pendola & New 2007), plastic (Luff 1973;

Morrill 1975; Clark & Blom 1992; Spence & Niemela 1994), or metal (Ahearn 1971; Hinds

& Rickard 1973; Clark & Blom 1992). Choice of material can affect the taxa sampled in

live traps as escape rates differ. One study on carabids found 0% escape from glass traps,

4% escape per day from plastic traps, and 10% escape per day from metal traps (Luff 1975).

Other studies have also found glass pitfalls retain more arthropods than plastic or metal

(Vennila & Rajagopal 2000), though one found no difference between glass and plastic

traps (Waage 1985). Similarly, Topping and Luff (1995) found plastic traps with rough

surfaces caught fewer linyphiid spiders than similar traps with smooth surfaces.

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Finally, color of the pitfall trap affects the taxa collected: white and yellow traps

catch higher numbers of Apidae, Araneae, Carabidae, Diptera, and Formicidae, while brown

and green traps catch higher numbers of Isopoda (Buchholz et al. 2010).

Effects of trap design, layout, and site selection . Some studies have found that

covers do not affect the composition of arthropods trapped by pitfall traps (Work et al.

2002; Buchholz & Hanning 2009; Cheli & Corley 2010) while others have found they

do (Briggs 1961; Baars 1979; Spence & Niemela 1994). Some of this may be due to the

material used as a cover. Man-made covers, such as metal or ceramic tile, are generally

used. Suggestions have been made to use natural material such as bark or rock for covers

(van der Berghe 1992), though this has not been systematically investigated.

Pitfall traps that have an integrated cap and circular entrances in the sidewall of the

trap (first proposed by Nordlander 1987) caught 80% of the same common carabid species

as conventional pitfalls in one study (Femieux & Findgren 1999), but otherwise have not

been thoroughly investigated and compared to conventional traps.

Pitfall traps must be level with the soil surface as excessive inclination of the

soil ringing the traps may direct some arthropods away from the trap (Heydemann 1953).

Similarly, a plastic disc surrounding the trap will influence sample size (Adis 1976).

Subterranean pitfall traps have been employed to trap hypogaeic ants (Yamaguchi

& Hasegawa 1996; Anderson & Brault 2010; Berghoff et al. 2003; Schmidt & Solar 2010),

though these preform no better than conventional pitfalls (Pacheco & Vasconcelos 2012).

Use of a barrier fence consistently increases the number of ground beetles collected

(Winder et al. 2001; Hansen & New 2005). However, the length of the fence influences

trap catch (Durkis & Reeves 1982; Morrill et al. 1990), with longer fences catching higher

diversity of families and species (Brennan et al. 2005), making it difficult to compare trap

catch between studies. Focation and number of the traps along the fence and fence material

may also affect trap catch, though these variables have not been specifically investigated.

Spacing between traps is an important consideration as populations, especially

of larger taxa such as carabids, can become locally depleted if traps are placed closely

together; this can affect trap catch and skew results. Snider and Snider (1986) found no

difference in trap catch between pitfalls spaced 0.5, 1, 2, and 4 meters apart. Similarly, Ward

et al. (2001) found no difference in trap catch between pitfalls spaced 1, 5, and 10 meters

apart. However, Digweed et al. (1995) found that carabid populations were depleted when

pitfalls were placed 10 meters apart but not 25 meters; in addition, traps spaced at 10 meters

had the most similar species assemblages and fewest rare species.

The optimum number of pitfall traps depends on the environment of the trapping

site. As few as five traps aresufficient in an arid steppe environment (Cheli & Corley

2010), whereas ten to twenty pitfall traps effectively collected the majority of species in

temperate areas (Formicidae: Santos et al. 2003; Coleoptera: Obrtel 1971; Isopoda Paoletti

and Hassall 1999; Araneae: Niemela et al. 1986), and at least twenty five are needed in

tropical areas (Vennila & Rajagopal 1999). Various non-parametric estimators have been

tested to estimate species richness based on as few as five traps per site (Brose 2002).

Finally, pitfall traps may not be the most efficient method for sampling epigeal

arthropods in environments with rugged, steep slopes and a high density of rocks or roots

in the soil where the traps are difficult to set or at high elevation where the mean body

size of taxa is generally smaller, and thus more difficult to trap (Nyundo & Yarro 2007).

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Additionally, some studies have found pitfalls trap more ants in drier areas and seasons

(Delsinne et al. 2008; Nunes et al. 2011), though others have found annual rainfall has no

effect (Delsinne et al. 2010).

Use of attractants in pitfall traps . The choice of preservative can affect the taxa

collected in pitfall traps (Weeks & McIntyre 1 997). For instance, bark beetles (Curculionidae:

Scolytinae), certain Staphylinidae, and Nitidulidae are caught in higher numbers in pitfalls

that use ethanol as the preservative (Drift 1963; Greenslade & Greenslade 1971). In one

study, some Carabidae, especially Bembidion , were caught in higher numbers in ethylene

glycol than water, though the effect varied by sex and time of year (Holopainen 1 990, 1 992);

another study, however, found no difference between ethylene glycol and water when trapping

four species of Diplopoda, one species of Chilopoda, and two species of Carabidae (Gerlach

et al. 2009), suggesting that any effect is species dependent. Formaldehyde has been found

to be repellant to Opiliones and Diplopodaand attractive to Carabidae and Staphylinidae

(Fuff 1 968; Pekar 2002; Gerlach et al. 2009), though one study found no difference between

water and formaldehyde when collecting Carabidae (Waage 1985). Differences have been

found between commercially available antifreeze and diluted ethylene glycol (Koivula et

al. 2003). Efficacy of preservatives can vary with trap size - one study found vinegar to

be more effective in large traps but propylene glycol more effective in small traps (Koivula

et al. 2003). Brine and an ethanol-glycerin mix have lower capture efficiency than other

fluids such as pure water, ethanol-water, and ethylene glycol-water, possibly due to the

high specific gravities of these fluids, which may allow captured arthropods to float and

escape (Schmidt et al. 2006). Brine is also attractive to Fepidoptera (Cheli & Corley 2010).

Additionally, attraction and repulsion to preservatives can vary due to sex (Adis 1976),

season (Dethier 1947; Adis & Kramer 1975; Adis 1976), and environment (Koivula et al.

2003). Thus, careful consideration should thus be used in order to avoid or account for the

influence of preservative on the taxa collected.

A drop of detergent is often used to break the surface tension of the preservative

in wet pitfalls. This does not seem to affect the rate of capture of most arthropods, though

Finyphiidae are caught in higher numbers (up to 1000%) in traps with detergent (Topping

& Fuff 1995; Pekar 2002), whereas Staphylinidae are caught in higher numbers in traps

without detergent (Pekar 2002).

Some Coleoptera naturally aggregate using pheromones to locate conspecifics

(Greenslade 1963; Wautier 1970, 1971; Ahearn 1971), which can affect trap catch

distribution as the first specimen captured may artificially attract others to the same trap

(Fuff 1968; Thomas & Sleeper 1977; Fuff 1986).

Digging- in effects have been recorded among Formicidae (Greenslade 1973),

Carabidae (Digweed et al. 1995; Schirmel et al. 2010) and other Coleoptera (Schirmel et

al. 2010), Collembola (Joosse-van Damme 1965; Joosse & Kapteijn 1968), Finyphiidae

and other Aranaea (Topping & Fuff 1995; Schirmel et al. 2010), and Isopoda (Schirmel

et al. 2010). These effects consist of high capture of certain taxa immediately after pitfall

traps are established followed by a subsequent decline. A variety of explanations - such

as an increased level of C02 (Collembola: Joosse & Kapteijn 1968), decreased barriers to

movement (Carabidae: Greenslade 1964), increased number of prey that attract predators

(Adis 1979), and decreasing number of foraging Formicidae workers (Romero & Jaffee

1989) - have been suggested, though no consensus has been reached. If digging- in effects

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are to be avoided, it has been suggested either to place pitfalls inverted for one week before

operating them as traps (Greenslade 1973; Schirmel et al. 2010) or to install a tube or

second container in which the pitfall can be placed in order to avoid disturbing the soil when

it is serviced (Schirmel et al. 2010). Alternatively, if the goal is to catch large numbers of

arthropods without regard to comparing between-trap catch, traps may be serviced more

frequently in order to take advantage of digging- in effects (Schirmel et al. 2010).

Disturbance of leaf litter and vegetation around the traps can cause increased

catch of highly mobile taxa, such as Gryllidae (Sperber et al. 2007). Areas around active

pitfalls should therefore not be visited unless the traps are being serviced. Alternatively,

regularly scheduled visits to the trap area will increase the catch of certain mobile taxa,

though care should be taken in designing and executing such visits in order to provoke the

same disturbance between traps (Sperber et al. 2007).

If attraction is desired, baits can be used to purposely affect the taxa collected

(Greenslade & Greenslade 1971). Dung and carrion can used to collect Scarabaeidae,

Staphylinidae, Silphidae, Ptiliidae, Histeridae, Hydrophilidae, and Leiodidae. Carnivore

and omnivore dung provide good results - with human dung being among the most effective

and readily available - while herbivore dung is generally poor (Newton & Peck 1975).

Meat, tuna, and honey can be used as baits for ants (Romero & Jaffee 1989). Though not

intentional, previously trapped insects may begin to rot in traps in which the preservative

is ineffective due to dilution from rain or large numbers of trapped insects, thus attracting

carrion feeding taxa (Holland & Reynolds 2005). Vegetable oils have been shown to

increase the catch of ants in the tropics (Pacheco & Vasconcelos 2012), especially army

ants (Weissflog et al. 2000; Berghoff et al. 2002; Berghoff et al. 2003), although this has

not been studied in temperate regions.

Pests of pitfall traps . Occasionally, traps will be regularly disturbed by mammals

between collections. Van der Berge (1992) presented three situations with the possible

culprits and associated solutions. For traps where the cup is still in the hole but pushed

up “just enough so that the rim is no longer flush with the soil” he suggests moles or voles

whose passage has been obstructed are to blame and moving the cup a short distance usually

resolves the problem. When one or a few cups, but not the entire trap line, are completely

out of the hole, spilled clean, but not chewed on he suggests squirrels are attempting to

burry or dig up nuts. Unfortunately, “one is helpless against squirrel disturbance”. The

third case is when many, and often the whole line, of cups are out of the hole and chewed or

mangled. This, he suggests, is the work of raccoons, opossums or deer that are interested in

consuming the preservative. Raccoons are intelligent and will continue to harass a line of

pitfall traps if they are reset, so it is best to abandon the line or add a distasteful substance

to the preservative. If deer are molesting the traps, it is best to switch from a salt-based

preservative which is probably drawing their attention.

Preservatives

Pitfall traps can be used to collect insects to be kept alive or killed in preservative. If

live specimens are required, such as for rearing experiments (as is common in parasitengone

mites to correlate life stages) or in cases where the taxon of interest is endangered, e.g.

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the American burying beetle (Nicrophorus americanus (Olivier, 1790)), traps are run dry

without preservative. In such cases, traps must be checked at least daily, and often more

frequently, so captured individuals do not succumb to heat, desiccate, drown in accumulated

rain water, or become predated on by other captured organisms (Mitchell 1963; Luff 1968;

Weeks & McIntyre 1997; Bestelmeyer et al. 2000; Moreau et al. 2013).

When collecting specimens to be killed, the choice of trap preservative is an

important consideration as it will affect the quality of specimens, cost of trap maintenance,

and how frequently traps must be serviced. Many authors have investigated the preservation

properties of different chemicals and solutions, which are summarized herein.

Ethylene glycol was once used as a preservative, especially in pitfall and pan traps,

as it has low volatility compared to ethanol and other alcohols (Martin 1977), is relatively

inexpensive, and is readily available as antifreeze. When used in the field it has been

reported to not preserve internal organs well and causes specimens to deteriorate to the

point of breaking when pinned (Aristophanous 2010), though other studies report sufficient

preservation (Sasakawa 2007; Cheli & Corley 2010). Because ethylene glycol is toxic to

vertebrates (Thrall e al. 1984) and is readily ingested due to its sweet taste (Grauer & Thrall

1982), its use has been discouraged (Hall 1991).

The addition of bitter agents, such as quinine, to ethylene glycol has been suggested

as a way to deter vertebrates from drinking the fluid (Hall 1991 ). Quinine added to ethylene

glycol, propylene glycol, and formalin has been shown to have no effect on the number of

spiders caught in pitfall traps; in addition, it improves the preservation quality of specimens

collected in ethylene glycol (Jud & Schmidt-Entling 2008). Alternatively, a red marking

flag placed next to the trap may deter large vertebrates from investigating the trap and

drinking the ethylene glycol (Cheli & Corley 2010).

An alternative to ethylene glycol but with similar characteristics is propylene glycol,

which is sold as recreational vehicle and boat antifreeze. It also has low volatility and is

inexpensive. Propylene glycol is nearly non-toxic as it is metabolized into constituents of

the Krebb’s cycle and extremely large quantities must be ingested over a short period of time

before acute toxicity is reached (Yu 2007). In the field, propylene glycol preserves insects

similarly to ethylene glycol (Jud & Schmidt-Engling 2008; Aristophanous 2010). However,

Moreau et al. (2013) found no detectable difference in the quality of DNA preservation

between propylene glycol and ethanol when undiluted chemicals were used in a lab setting.

One reason for the difference between field and lab studies may be due to the fact that

ethylene glycol and propylene glycol are hygroscopic; when humidity is moderate to high,

both substances will absorb water from the air and dilute naturally (Aristophanous 2010).

Salt brine and saturated borax solution are inexpensive and easy to make as the

constituent materials are readily available in grocery stores. The ability of these solutions to

preserve insects is extremely poor, however, and not outweighed by cost-savings (Lemieux

& Lindgren 1999; Sasakawa 2007; Aristophanous 2010) (though see Schmidt et al. 2006

for a counter opinion).

Carnoy’s fixative (60% ethanol, 30% chlorofonn, 10% acetic acid) and white

vinegar (10% acetic acid) do not preserve DNA and cause specimens to become brittle,

though they generally keep the specimens from rotting (Sasakawa 2007; Aristophanous

2010; Moreau et al. 2013). If DNA extraction is not intended, these may be acceptable

preservatives.

Skvarla et al.

JESO Volume 145, 2014

Methanol and chloroform do not preserve specimens in a way that allows DNA

extraction and amplification (Post et al. 1993; Fukatsu 1999). In addition, chloroform is

difficult to acquire, especially in the large quantities required for use as a trap preservative.

FAACC solution (fonnaldehyde 4%, acetic acid 5%, calcium chloride 1.3%)

and 4% phosphate buffered formaldehyde (4%PBF) both preserve internal organs well,

with 4%PBF being the superior of the two (Aristophanous 2010). However, specimens

become excessively stiff and although DNA can be extracted from specimens preserved

with formaldehyde solutions, DNA amplification is impossible with standard kits (such a

Qiagen DNEasy) because formaldehyde causes DNA to cross-link with proteins (Schander

& Halanych 2003). Protocols using prolonged extraction times (up to 7 days) (France &

Kocher 1996; Chatigny 2000; Schander & Halanych 2003) and chemical agents (Johnson

et al. 1995; Chatigny 2000) can be successful.

Amyl acetate is sometimes used in insect jars as the killing agent. This banana-

smelling liquid keeps specimens relaxed, unlike other killing agents such as chloroform

(Woodward 1951). It is commonly used as a water-removing solvent in industry and can

be purchased through specialized suppliers. Amyl acetate has been used for preservation

of anatomical dissections (Saunders & Rice 1944) and insects “may be kept stored almost

indefinitely between cotton- wool impregnated with this agent” (Woodward 1951), though it

has not been tested for DNA preservation (Nagy 2010). Additionally, it has not been tested

as a preservative in pitfall traps, can be a skin irritant, and is probably attractive to some

insect groups so other, more proven preservatives may be a better choice.

Ethanol is probably the most widely used preservative. It maintains the integrity

of internal organs and allows DNA to be easily extracted and amplified (Gurdebeke &

Maelfait 2002; Aristophanous 2010; Moreau et al. 2013). In the United States, price may

be prohibitive for individuals who do not qualify for ethanol tax exemption; however, fuel

ethanol has been shown to preserve specimens as well as pure ethanol, so this will provide

an alternative source as fuel ethanol becomes more widespread (Szinwelski et al. 2012). In

addition, ethanol is the most volatile commonly used preservative. In open containers such

as pitfall traps ethanol can lose % of its volume in fewer than 5 days (Aristophanous 2010).

Depending on the trap location this may have implications on how often the traps must be

serviced.

Isopropanol, commonly known as rubbing alcohol, is a cheap alternative to

ethanol. Similar to ethanol, it preserves DNA well (Rake 1972), so it can be extracted with

little difficulty. One drawback is that isopropanol often discolors specimens, which is a

hindrance to identification and morphological studies involving color.

Acetone has shown promise as a preservative. It is relatively inexpensive and

readily available as a paint solvent. DNA has been extracted and successfully amplified

from acetone-preserved Copepods (Goetze & Jungbluth 2013), pea aphid {Acyrthosiphon

pisum (Harris, 1776)) (Fukatsu 1999), and Zygoptera (Fogan 1999). Additionally, acetone

is used to preserve adult Odonata as it dissolves fat, dehydrates the specimen, and reduces

decomposition of enzymatic color pigments (Abbott 2008).

Other preservatives require more testing as contradictory results have been

reported. Fukatsu (1999) reported DNA amplification after specimens were stored in 2-

propanol, ethyl acetate, and diethyl ether, though Post et al. (1993) and Reiss et al. (1995)

reported poor results with 2-propanol and ethyl acetate, respectively.

Pitfalls and preservatives: A review

JESO Volume 145, 2014

Summary

Pitfall traps are often used to sample epigeal arthropods as they are inexpensive and

easy to use. However, many factors influence the taxa so collected. Abiotic factors, such

as weather, season, slope and aspect, degree of rockiness, and trap characteristics (color and

material of the trap, diameter of the opening, spacing between traps, and number of traps

at a site) affect the composition of collected taxa, often by affecting behavior of the target

arthropods. Biotic factors affecting trap catch include species-specific factors (activity

level, size, aggregation to conspecifics, and behavior at the edge of the trap), response to

digging-in effects, and habitat structure, including the density of low-growing vegetation.

The choice of preservative affects not only the level of preservation of specimens, but also

the composition of specimens collected because various compounds differentially repel

and attract different taxa. Taken together, these factors make comparisons between studies

difficult.

While there have been calls to standardize pitfall trapping, the design employed

in individual studies will continue to be based on the research question and materials

available. An effort, however, should be made to report all of the factors that might

influence the composition of specimens collected. While this may not be immediately

useful, comparisons may be made in the future after further studies elucidate the effects

various factors have upon trap catch.

Acknowledgements

We thank the reviewers for their helpful suggestions; this manuscript is a better

product because of them.

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First records of Ptosima walshii in Canada

JESO Volume 145, 2014

FIRST RECORDS OF PTOSIMA WALSHII (COLEOPTERA:

BUPRESTIDAE) IN CANADA

D.B. LYONS 1 *, K. L. RYALL 1 , S.M. PAIERO 2 , G.C. JONES 1 AND L. VAN

SEGGELEN 3 *

‘Canadian Forest Service, Natural Resources Canada,

1219 Queen St. East, Sault Ste. Marie, Ontario, Canada P6A2E5

email, Barry.Lyons@NRCan-RNCan.gc.ca

Scientific Note J. ent. Soc. Ont. 145: 45-49

Ptosima Dejean, 1833 (Coleoptera: Buprestidae) (Fig. 1) contains 10 extant

species worldwide (Bellamy 2008) with four species occurring in North American (Nelson

1978, Nelson et al. 2008). Nelson (1978) keyed and redescribed the North American

species. Plant genera records for Ptosima spp. include Crataegeus (Rosaceae), Cercis

(Fabaceae) and Quercus (Fagaceae) (Paiero et al. 2012). Ptosima idolynae Frost, 1923

and P. laeta Waterhouse, ! 882 are confined to south central North America, whereas P.

gibbicollis (Say, 1823) and P. walshii LeConte, 1863 are widely distributed in eastern

North America. Bright (1987) stated that P. gibbicollis “probably occurs in southern

Ontario”, but makes no mention of P. walshii. Paiero et al. (2012) mapped the distribution

of P. gibbicollis as occurring in Ontario. They also mapped the host range of P. walshii ,

as a potential distributional range for the beetle, encompassing Manitoba, Ontario and

Quebec, but no records were known for these Provinces. Ptosima walshii is considered

“rarely collected” (MacRae 2006) and “infrequently encountered” (Paiero et al. 2012).

Nelson (1978) stated that P. walshii had been reported from Illinois, Texas, Kansas

and California, suggested that the California record was doubtful, and added new State

distribution records for Iowa, Michigan, Minnesota, Mississippi, Missouri, Ohio and

Wisconsin. Westcott (1991) provided compelling evidence that the California record was

erroneous. Records for four states that border Canada — Michigan, Ohio, Wisconsin and

Minnesota — are represented by a total of eight collections (Nelson 1978). The Michigan

record was a single collection labeled “Ag. Coll. 20.V.1889” (Nelson 1978). Westcott

(1991) reported a specimen from Oklahoma. Thus, the literature records confirmed by

specimens include 11 States but no Provinces.

As part of a trapping study for the recently discovered European oak borer,

Agrilus sulcicollis Lacordaire (Coleoptera: Buprestidae) in North America (Jendek

and Grebennikov 2009, Haack et al. 2009), we collected numerous P. walshii adults in

southwestern Ontario. These collections represent the first verified records of the species

Published October 2014

* Author to whom all correspondence should be addressed.

2 School of Environmental Sciences, University of Guelph, Guelph, Ontario, Canada

N1G2W1

3 Invasive Species Centre, Sault Ste. Marie, Ontario P6A 2E5

Lyons et al.

JESO Volume 145, 2014

in Canada. Here we describe the circumstances of these collections.

Three woodlots (Table 1) selected for trapping in 2011, produced P. walshii

adults. In each woodlot, three trap/lure combinations were deployed on oak trees: 1)

unbaited sticky-band trap; 2) green prism trap baited with the green leaf volatile, 3-(Z)-

hexenol (Synergy Semiochemicals, Inc., Burnaby, BC); and 3) green prism trap baited

with a manuka oil/phoebe oil lure (Synergy Semiochemicals, Inc.). Three replicates of

each trap type were established per site, for a total of nine traps per location. Traps were

set up 19 May 2011 and monitored approximately every 6 or 7 days throughout the season

until 18 August 2011. The Bickford Line and Courtright Line sites were sampled again in

2012 using unbaited green prism traps and sticky-band traps. Lour replicates of each trap

type were established per site for a total of 8 traps per location. Traps were set up on 10 to

12 May 2012 and monitored weekly until 19 July 2012. Three additional woodlots (Table

2), each with three sticky-band traps, also produced P. walshii adults. These additional

traps were set up on 10 to 14 May 2012 and sampling was conducted once in mid-season

(26 to 28 June) and again later in the season (18 to 19 July). The numbers of specimens of

each sex collected at each location in each year are tabulated (Tables 1 and 2).

Males and females were collected from all three trap/lure types. Significantly

more beetles (Tig. 2) of both sexes were collected on the green prism traps baited with the

LIGURE 1 . Dorsal view of female of Ptosima walshii from Bickford Line site, Lambton,

Co., Ontario (photograph by G.C. Jones).

TABLE 1 . Number of adults of Ptosima walshii captured with green prism traps and sticky-

band traps in woodlots in southwestern Ontario in 201 1 and 2012.

2011 2012

Site

Latitude

Longitude

Males

Females

Males

Females

Bickford Line

42.7635°N

82.3096°W

Courtright Line

42.7987°N

82.2388°W

Thames Road

42.85 18°N

81.7256°W

Total

First records of Ptosima walshii in Canada JESO Volume 145, 2014

TABLE 2. Number of adults of Ptosima walshii captured with sticky-band traps in additional

woodlots sampled in southwestern Ontario in 2012.

Site

Latitude

Longitude

Males

Females

Hillsboro Road

43.005 1°N

82.0946°W

Coldstream C.A. 1

43.021 0°N

81.4981°W

Ladysmith Road

42.8164°N

82.3946°W

Waterworks Road

42.8954°N

82.2575°W

Total

‘C.A. = Conservation Authority

c n

3 12

® 10

o3 8

E o

=3 6

FIGURE 2. Number of males and females of Ptosima walshii captured in the three different

trap/lure types (S= unbaited sticky-band traps, MP = green prism traps baited with manuka

oil/phoebe oil; and GLV = green prism traps baited with 3-(Z)-hexenol) in four sites

sampled in 201 1 in southwestern Ontario. Different letters over the bars represent significant

differences (G-test) between the total number of beetles (males + females) captured in each

trap lure type.

i males

i females

S MP GLV

Trap/Lure type

Lyons et al.

JESO Volume 145, 2014

3-(Z)-hexenol compared to the other trap/lure types (G-test; a = 0.05, G 2 = 24.247) which

suggests that this green leaf volatile is attractive to P. walshii. All adults of P. walshii

were captured on either Quercus macrocarpa Michx. or Q. alba.L. Nelson (1978) listed

the host of P. walshii as unknown. In a subsequent paper Nelson et al. (1981) reported

that two adults had been collected by beating Q, macrocarpa. MacRae (2006) reported

the first rearing of adults from Q. macrocarpa. The extensive use of this host tree for our

trap placement serendipitously resulted in the capture of large numbers of P. walshii. The

flight periods of the collected adults are shown in Fig. 3 for 2011 and 2012 excluding the

additional sites. The early flight period of P. walshii observed in our study, especially in

2012, supports the assumption that the members of Ptosima overwinter as adults within

pupal cells in the host tree (Nelson 1978).

Voucher specimens of P. walshii are deposited in the Great Lakes Forestry Centre

Insect Collection (GLFC), the University of Guelph Insect Collection (DEBU) and the

Canadian National Collection of Insects and Arachnids (CNCI).

10 May 30 May 19 June 9 July 29 July 18 August

Day of year

FIGURE 3. Total number of adults of Ptosima walshii (males + females) captured in each

sample interval on all trap types (excluding additional woodlots in 2012), in 2011 and 2012

in southwestern Ontario, plotted in the middle of each sample interval.

First records of Ptosima walshii in Canada

JESO Volume 145, 2014

Acknowledgements

We are grateful to H. Evans, L. Breton, M. Campbell, S. Crispell, D. Nisbet, I.

Ochoa, R. Pinkham, L. Roscoe and K. Wainio for assistance in the field. We would also like

to thank the following landowners for pennission to access and trap on their properties: S.

Bunker, D. Chowen, J. Mardy, G. Wilhelm, M. Wolff, D. Young, and the St. Clair Region

Conservation Authority. Funding for the research was provided by the Canadian Forest

Service, the Ontario Ministry of Natural Resources and the Invasive Species Centre.

References

Bellamy, C.F. 2008. A world catalogue and bibliography of the jewel beetles (Coleoptera:

Buprestoidea) , Volume 1: Introduction; fossil taxa; Schizopodidae; Bnprestidae:

Julodinae - Chrysochroinae: Poecilonotini. Pensoft Series FaunisticaNo. 76, pp.

1-625.

Bright, D.E. 1987. The metallic wood-boring beetles of Canada and Alaska (Coleoptera:

Buprestidae). The Insects and Arachnids of Canada, Part 15, Biosystematics Research

Centre, Research Branch, Agriculture Canada, Publication 1810, Ottawa, Canada. 335

pp.

Haack, R.A., Petrice, T.R. and Zablotny, J.E. 2009. First report of the European oak

borer, Agrilus sulcicollis (Coleoptera: Buprestidae), in the United States. Great Lakes

Entomologist 42: 1-7.

Jendek, E. and Grebennikov, V.V. 2009. Agrilus sulcicollis (Coleoptera: Buprestidae), a

new alien species in North America. The Canadian Entomologist 141 : 236-245.

MacRae, T.C. 2006. Distributional and biological notes on North American Buprestidae

(Coleoptera), with comments on variation in Anthaxia (Haplanthaxia) cyanella Gory

and A. (H.) viridifrons Gory. Pan-Pacific Entomologist 82: 166-199.

Nelson, G.H., Verity D.S. and Westcott, R.L. 1981. Additional notes on the biology and

distribution of Buprestidae (Coleoptera) of North America. Coleopterists Bulletin 35 :

129-151.

Nelson, G.H., Walters Jr., G.C., Haines, R.D. and Bellamy. C.L. 2008. A catalog and

bibliography of the Buprestoidea of America North of Mexico. Coleopterists Society

Special Publication No. 4, North Potomac, Maryland. 274 pp.

Paiero, S.M., Jackson, M.D, Jewiss-Gaines, A., Kimoto, T., Gill B.D. and Marshall, S.A.

2012. Field guide to the jewel beetles (Coleoptera: Buprestidae) of northeastern North

America. Canadian Food Inspection Agency, Ontario, Canada. 411 pp.

Westcott, R. L. 1991. Distributional, biological, and taxonomic notes on North American

Buprestidae). Insecta Mundi 4 : 73-89.

Eclosion of Physocephala tibialis from a Bombus host

JESO Volume 145, 2014

ECLOSION OF PHYSOCEPHALA TIBIALIS (SAY) (DIPTERA:

CONOPIDAE) FROM A BOMBUS (APIDAE: HYMENOPTERA)

HOST: A VIDEO RECORD

J. F. GIBSON 1 *, A. D. SLATOSKY 2 , R. L. MALFI 2 , T. ROULSTON 2 , S. E. DAVIS 2

'Biodiversity Institute of Ontario and Department of Integrative Biology,

University of Guelph, 50 Stone Rd. E., Guelph ON NIG 2W1

email, jfgibson@uoguelph.ca

Abstract J. ent. Soc. Ont. 145: 51-60

Some members of Conopidae and other families of flies require development

within hymenopteran hosts. Rearing of parasitized Apoidea provides valuable

life history and ecological data but is rarely documented. Greater emphasis

on gathering and analyzing rearing data is required. Analysis of a new video

record of Physocephala tibialis (Say) reared from Bombus impatiens Cresson

provides detailed evidence of the use of the ptilinum, mouthparts, and legs for

eclosion within Conopidae. The previous literature on Conopidae/Apoidea

rearing is reviewed.

Published October 2014

Introduction

Some species of Conopidae (Diptera) develop exclusively within hymenopteran

hosts. Eggs are deposited inside adult bees or wasps. Following emergence from the egg,

the larva grows, develops, and pupates entirely within the body of the host. Following death

of the host and often after an overwintering period, the adult conopid ecloses from the host’s

corpse (Freeman 1966). While this behaviour is well noted within the conopid literature,

careful rearing of parasitized hymenopteran hosts is only rarely documented.

Meijere (1904), Freeman (1966), and Smith (1959, 1966) summarized known host

records and life histories for Conopidae, including many records of development within bee

hosts. However, they did not distinguish between studies that relied on confirmed rearing

records and studies that did not. Several studies tried to establish host records for Conopidae

based exclusively on associations with host species (Rasmussen and Cameron 2004; Rocha-

Filho et al. 2008) or on the discovery of eggs on pinned specimens (Stuckenberg 1963; Couri

and Pont 2006; Couri and Barros 2010; Couri et al. 2013). These records must be considered

* Author to whom all correspondence should be addressed.

2 Department of Environmental Sciences, University of Virginia, Charlottesville, VA

Gibson et al.

JESO Volume 145, 2014

tentative, as they do not confirm the successful development of Conopidae within a given

host. While most confirmed rearing records are based on dead or obviously parasitized bees

being held until parasitoids emerge, a few (Paxton et al. 1996; Polidori et al. 2005) were

based on careful observation of host nests until parasitoids were seen to emerge.

Past rearing efforts have also produced other important observations on conopid

life history. Schmid-Hempel et al. (1990) and Otterstatter et al. (2002) compared the relative

parasitism rates and overlapping phenologies of competing parasitoid species. Knerer and

Atwood (1967) reared one species of Conopidae, Thecophora occidensis (Walker), from six

different species of halictine bees. They proposed a complex life history for T. occidensis ,

including phenological host switching. Muller (1994) observed self-burying behaviour in

parasitized specimens of Bombns terrestris. Unfortunately, the conopid species reared out

was never identified. Malfi et al. (2014) reared 39 specimens of Physocephala tibialis from

three species of Bombns. They provided evidence of differential successful parasitism rates

among the host species. Self-burying behaviour was also observed in parasitized bees, with

a differential frequency of this behaviour depending on host species.

Several authors used rearing experiments to observe directly the process of

eclosion from the host by Conopidae. Cumber (1949) reared Physocephala rufipes from

Bombns agrorum, and noted that “the [conopid] adult emerges by pushing aside the anterior

segments with its ptilinum.” Polidori et al. (2005) observed an adult Zodion cinereum

emerging from a ground nest of Andrena agilissima (Andrenidae) with the ptilinum “still

being inflated rhythmically, indicating that they were freshly emerged adults.” Koeniger et

al. (2010) reared Physocephala paralleliventris from two different species of honey bees

{Apis cerana, A. koschevnikovi) in Borneo. They observed an active period of walking for

about fifteen minutes prior to inflation of the wings. They theorized that this active stage

was necessary for conopids to emerge from the leaf litter in which the host bee was buried.

For other families of Diptera, video recordings of parasitoid emergence have been

informative. Downing (1995) observed ptilinal expansion in Amobia (Sarcophagidae),

which cleptoparasitizes mud-tube nesting wasps of Trypoxylon (Sphecidae). Strohm (2011)

recorded cleptoparasitic members of Drosophilidae using ptilinal expansion to break out of

the closed brood cells of their hymenopteran hosts.

The video recording presented here produced both life history data and behavioural

observations for one species of Conopidae.

Materials and Methods

As part of ongoing research examining the impact of parasitism on bumble bees

(Hymenoptera: Apidae: Bombns) in northern Virginia (Malfi and Roulston, 2014; Malfi et

al. 2014), 445 foraging bumble bees were collected at Blandy Experimental Farm (Boyce,

VA, USA, 36.09°N, 78.06°W) in June and July of 2012. These bees were maintained until

death in the lab by housing bees in aquaria partially filled with soil and leaf litter and

providing them sugar water ad libitum (for details, see Malfi et al. 2014). After death, bees

were examined for the presence of a conopid parasite. Often this could be determined with

minimal disturbance of the bee corpse as the conopid pupa frequently occupies the entire

abdominal cavity of its host and a large, last-instar larva can be seen externally to move

Eclosion of Physocephala tibialis from a Bornbus host

JESO Volume 145, 2014

within the bee. A total of 120 bumble bee workers belonging to three different species

(B. bimaculatus , B. griseocollis, B. impatiens ) were parasitized with conopid larvae. In

September 2012, 94 conopid pupae harvested from these parasitized bees were placed in

individual vials in a household refrigerator (~4°C) and left there until April, 20 13, when they

were placed on a lab bench in a room at 2 1 .5° C and monitored for emergence. The emergence

of one parasitoid was observed in progress. After the corpse of its host, a Bombus impatiens

worker, began to move, one author (ADS) recorded the emergence process with a cell phone

camera held up to the eyepiece of a dissecting microscope. Recording of emergence began

at 8:50 AM on June 2 nd , 2013, and continued for eight minutes, at which point the fly had

completed its exit from the host.

Results

Thirty-nine of the conopid pupae emerged as adults and were identified as

Physocephala tibialis. A 30-second edited version as well as the full eight-minute video

may be viewed at (VIDEO). The edited video begins with the head of the conopid already

visible, emerging from the ventral anterior of the bee abdomen. The ptilinum is seen fully

inflated, with the antennae deflected to the ventral surface of the head (Figs. 1A, ID). The

mouthparts are displaced posteriorly, between the forelegs. At full inflation, the ptilinal sac

appears to be equal in volume to the rest of the head. Following full inflation, the ptilinum

deflates, but is still extruded (Figs. IB, IE). As the conopid pulls the rest of its body from

the host, the ptilinum continues rhythmically inflating and deflating, though not to a volume

equal to that visible as the head is first emerging. The conopid uses its legs and mouthparts

as levers to pry itself from the host’s body. When the conopid body is fully emerged from

the host, it begins to walk around with the deflated ptilinal sac still visible. Throughout the

eclosion process, it appears that the antennae, mouthparts, and legs are already sclerotized

and dark. The sclerites of the head, thorax, and abdomen appear pale and unsclerotized. The

wings are not inflated to any extent throughout the video. The adult P. tibialis is included

with other voucher specimens that have been deposited in the University of Guelph Insect

Collection, Guelph, ON.

Discussion

Schizophora (Diptera) have a membranous invagination in the head that is visible

in adults only as a ptilinal fissure (Reaumur 1738; Becker 1882; Cumming and Wood

2009). The first description of the ptilinum in Calliphoridae (Reaumur 1738) included the

suggestion that it was used to burst the puparium wall through expansion. Early research on

the structure and function of the ptilinum (reviewed in Laing 1935; Atkins 1949) was limited

to Calliphoridae and Drosophilidae. Strickland (1953) examined 150 species from over 40

schizophoran families and found that, in all cases, the ptilinum is lined with microscopic

scales that are used to improve the puparium- bursting capabilities of the ptilinum. Zdarek et

al. (1986) and Zdarek and Denlinger (1992) revealed pressure changes within the ptilinum

and associated structures during eclosion and bodily inflation of the adult. Reid et al. (1987)

Gibson et al.

JESO Volume 145, 2014

defined a series of phases, including ptilinal expansion in the ‘extrication behaviour’ of

Sarcophagidae. Our observations suggest that Conopidae demonstrate the same patterns

and behaviours of eclosion as those observed in Calliphoridae, Drosophilidae, and

Sarcophagidae.

Strickland (1953) noted that the ptilinum of Conopidae is larger, thicker, and

covered with more varied types of scales than that of any other fly family examined.

His detailed drawings of Physocephala furcillata indicate that the base and length of the

mouthparts as well as the ptilinum are covered with sclerotized scales. He contended that

these scales assist with eclosion of the fly and subsequent digging from a subterranean

location. The species we observed, P. tibialis , is morphologically similar to P. furcillata

(Camras 1957). Malfi et al. (2014) demonstrated an induced digging behaviour in bees

parasitized by P. tibialis. Our video demonstrates the use of the mouthparts and ptilinum as

part of both eclosion and subsequent digging.

For most species of Conopidae, there are no host records confirmed through

rearing. The known records for Conopidae reared from Apoidea are summarized here (Table

1 ). The seven genera listed represent only 1 1 .9% of the approximately 59 extant genera and

subgenera (Gibson and Skevington 2013; Gibson et al. 2013).

Using DNA barcoding approaches to associate larvae, pupae, or adults of parasitoids

with each other and their host species (e.g., Smith et al. 2006, 2007) may add ecological and

evolutionary data to rearing experiments. A search of the cytochrome oxidase c subunit I

(COI) sequences available on GenBank (June 10, 2014) revealed 45 species of Conopidae,

FIGURE 1 . (A) Physocephala tibialis. Fateral view with ptilinum (PT) inflated. (B) Fateral

view with ptilinum deflated. (C) Fateral view after sclerotization. (D) Dorsal view with

ptilinum (PT) inflated. (E) Dorsal view with ptilinum deflated. (F) Fateral view after

sclerotization (different specimen of same species). Photo supplied by T. Burt, Canadian

National Collection of Insects. (D) and (E) are still images extracted from the VIDEO.

Sclerotized adult head width for (D) and (E) = 3.69 mm. Line diagrams were created with

Adobe Illustrator, based upon the video.

Eclosion of Physocephala tibialis from a Bombus host

JESO Volume 145, 2014

TABLE 1. Tabanidae species and number of specimens collected in 2011 and 2012 using

Malaise traps and sweep netting, with abundance records.

Parasitoid

Host

Region

Reference

Dahnannia

Lasioglossum scitulum

Japan

Maeta and

signata Chen

Myopa buccata

(Smith)

MacFarlane 1993

Andrena japonica (Smith)

Japan

Maeta and

MacFarlane 1993

(Linnaeus)

A. scotica Perkins

Sweden

Paxton et al. 1996

M. rubida (Bigot)

A. vierecki Cockerell

California

Mac Swain and

Bohart 1947

M. testacea

A. scotica Perkins

Sweden

Paxton et al. 1996

(Linnaeus)

Physocephala

Centris analis (Fabricius)

Brazil

Santos et al. 2008

aurifrons Walker

P. bennetti

C. analis

Brazil

Santos et al. 2008

Camras

Xylocopa frontalis

(Olivier), X submordax

Cockerell

Trinidad

Camras 1996

X. carinata Smith, X.

Kenya,

Smith and

bimarginipennis

flavorufa Degeer

Uganda

Cunningham- Van

Karsch

Someren 1970

P. bipunctata

(Macquart)

Euglossa anodorhynchi

Nemes io

Brazil

Melo et al. 2008

Centris analis (Fabricius)

Brazil

Santos et al. 2008

P. cayennensis

Macquart

P. inhabilis

C. analis

Brazil

Santos et al. 2008

Megachile maculata Smith

Brazil

Stuke and Cardoso

2013

(Walker)

Centris analis

Brazil

Santos et al. 2008

P. furcillata

Bombus vagans Smith

Ontario

MacFarlane and

(Williston)

P. marginata

(Say)

Apis mellifera Linnaeus

Washington

Pengelly 1975

VanTJuzee 1934

Bombus fervidus

Ontario

MacFarlane and

(Fabricius)

Megachile mendica

North

Pengelly 1975

Krombein 1967

Cresson

Carolina

P. obscura

Megachile willughbiella

Japan

Maeta 1997

Matsumura

(Kirby)

Bombus ardens Smith,

Japan

Maeta and

B. diversus Smith

MacFarlane 1993

P. paralleliventris

Apis cerana Fabricius,

Borneo

Koeniger et al.

Krober

A. koschevnikovi Enderlein

2010

P. pusilla

Megachile rotundata

Mongolia,

Seidelmann 2005,

(Meigen)

P ruftpes

Fabricius

France

Tasei 1975

Bombus agrorum

England

Cumber 1949

(Fabricius)

Fabricius

B. terrestris Linnaeus

???

Meijere 1904

B. lapidarius (Linnaeus),

Switzerland

Schmid-Hempel

B. lucorum (Linnaeus),

and Schmid-

B. pascuorum (Scopoli),

B. terrestris

Hempel 1988

P. rufithorax

Krober

Centris analis

Brazil

Santos et al. 2008

P. sagittaria

(Say)

Apis mellifera

Washington

Van Duzee 1934

Bremus auricomus

Robertson

Illinois

Frison 1926

Gibson et al.

JESO Volume 145, 2014

TABLE 1 continued...

P spheniformis Centris analis

Camras

P. soror Krober Centris analis

P. texana Apis mellifera

(Williston)

P. tibialis (Say)

P. vittata

(Fabricius)

P. wulpi Camras

Physocephala sp.

Nomia melanderi

Cockerell

Bombns bifarius Cresson,

B. californicus Smith,

B. flavifrons Cresson,

B. occidentalis Greene

B. bimaculatus Cresson,

B. griseocollis DeGeer,

B. impatiens Cresson

Megachile maritima

(Kirby)

Xylocopa artifex Smith,

X. augusti Lepeletier,

X. splendidula Lepeletier

Bombns (9 spp.)

Physoconops

fronto (Williston)

Sicus ferrugineus

(Linnaeus)

Megachile perihirta

Cockerell

Bombns lucorum , B.

pascuorum, B. terrestris

Thecophora

occidensis

(Walker)

Zodion cinereum

(Fabricius)

Lasioglossnm forbesii

(Roberts), L. laevissimns

(Smith), L. lineatnlnm

(Crawford), Halictns

ligatns Say, H. rnbicnndus

(Christ), Evylaens

cinctipes (Provancher)

Halictns confnsns Smith

H. confnsns, H.

rnbicnndns, Lasioglossnm

cinctipes (Provancher),

L. imitatns (Smith), L.

lineatnlnm , L. forbesii

Andrena agilissima

(Scopoli)

Andrena pro st omias Perez

Z. fulvifrons Say

Zj,

obliqnefasciatnm

(Macquart)

z. vsevolodi

Zimina

A pis mellifera

Nomia melanderi

Ceratina flavipes Smith,

C. japonica Cockerell,

C. megastigmata

Yasumatsu and Llirashima,

Chalicodoma spissnla

(Cockerell), Hylaens

thoracicns Fabricius

Brazil

Washington,

Wyoming

Idaho

Alberta

Virginia

Netherlands

Argentina

Massa-

chusetts

California

Switzerland

Ontario

Indiana

Ontario

Italy

Japan

South Dakota

???

Japan

Santos et al. 2008

Van Duzee

1934, Riedel and

Shimanuki 1966

Foote and Gittins

1961

Otterstatter et al.

2002

Malfi et al. 2014

Meijere 1904

Stuke et al. 2011

Gillespie 2010

Bohart and

MacSwain 1940

Schmid-FIempel

and Schmid-

FIempel 1988

Knerer and Atwood

1967

Dolphin 1979

Smiih 1966

Polidori et al. 2005

Maeta and

MacFarlane 1993

Severin 1937

Howell 1967

Maeta and

MacFarlane 1993

Eclosion of Physocephala tibialis from a Bombus host

JESO Volume 145, 2014

including three species of Physocephala. The use of DNA methods of identification, when

target taxa have already been sequenced, greatly increases the value of studies that record

immature parasites in hosts but, for methodological reasons, are unable to rear out adults

(Gillespie 2010; Malfi and Roulston 2014).

Our study is an example of the added information about parasitoids that can be

gained through careful rearing. Previous theories about the function of the ptilinum and the

process of eclosion from the host have been strengthened with video evidence.

Acknowledgements

Helpful editorial input was provided by G. Capretta, J. Skevington, T. Burt and

an anonymous reviewer. JFG is supported by an NSERC PDF. ADS was supported by the

National Science Foundation’s REU Program (DBI1 156796).

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Checklist of Ontario Orthoptera (cont.)

JESO Volume 145, 2014

NEW CANADIAN AND ONTARIO ORTHOPTEROID RECORDS,

AND AN UPDATED CHECKLIST OF THE ORTHOPTERA OF

ONTARIO

S. M. PAIERO 1 * AND S. A. MARSHALL 1

1 School of Environmental Sciences, University of Guelph,

Guelph, Ontario, Canada NIG 2W1

email, steve.paiero@gmail.com

Abstract J. ent. Soc. Ont. 145: 61-76

The following seven orthopteroid taxa are recorded from Canada for the first

time: Anaxipha species 1, Cyrtoxipha gundlachi Saussure, Chloroscirtus

forcipatus (Brunner von Wattenwyl), Neoconocephalus exiliscanorns

(Davis), Camptonotus carolinensis (Gerstaeker), Scapteriscus borellii

Linnaeus, and Melanoplus punctulatus griseus (Thomas). One further

species, Neoconocephalus retusus (Scudder) is recorded from Ontario for the

first time. An updated checklist of the orthopteroids of Ontario is provided,

along with notes on changes in nomenclature.

Published December 2014

Introduction

Vickery and Kevan (1985) and Vickery and Scudder (1987) reviewed and listed

the orthopteroid species known from Canada and Alaska, including 141 species from

Ontario. A further 15 species have been recorded from Ontario since then (Skevington et

al. 2001, Marshall et al. 2004, Paiero et al. 2010) and we here add another eight species

or subspecies, of which seven are also new Canadian records. Notes on several significant

provincial range extensions also are given, including two species originally recorded from

Ontario on bugguide.net. Voucher specimens examined here are deposited in the University

of Guelph Insect Collection (DEBU), unless otherwise noted.

New Canadian records

Anaxipha species 1 (Figs 1, 2) (Gryllidae: Trigidoniinae)

This species, similar in appearance to the Florida endemic Anaxipha calusa

* Author to whom all correspondence should be addressed.

Paiero and Marshall

JESO Volume 145, 2014

Walker & Funk, is here recorded as new to Canada based on specimens found in 2013 in

the Wings of Paradise Butterfly Conservatory, Cambridge, Ontario. Numerous individuals

(including nymphs, adult males and females) were observed at that time, and discussions

with the conservatory staff indicate that this Anaxipha species has been established there for

some time. Two other Ontario butterfly houses were contacted to determine if this species

had been introduced elsewhere in the province, but no further populations were reported.

Anaxipha “species 1” is an undescribed species, probably the same as an unnamed species

known to occur in Central America (Funk, pers. comm.; see also http://entnem.ifas.ufl.edu/

walker/buzz/SM AcalusaRelatives.pdf ) and was probably accidentally introduced with

shipments of butterfly pupae from Central America. If it is indeed this tropical species, it

is unlikely to become established outdoors in natural habitats in Canada. Distinctive dark

markings on the fore wings distinguish males (Fig. 1) of Anaxipha “species 1” from other

FIGURES 1-6. 1, Anaxipha species 1, S- 2, Anaxipha species 1, $. 3, Cyrtoxipha gnndlachi.

4, Chloroscirtus forcipatus , $ . 5, Chloroscirtus forcipatus , male terminalia (dorsal view). 6,

Camptonotus carolinensis, $ .

Checklist of Ontario Orthoptera (cont.)

JESO Volume 145, 2014

Ontario cricket species. The pale veins bordering infuscate cells make the females (Fig. 2)

easy to separate from A. exigna (Say).

Specimen Records: ONTARIO: Wellington Co., Cambridge, Wings of Paradise Butterfly

Conservatory, 43 0 27’7”N 80°22’2”W, in butterfly greenhouse, 20 April 2013, Paiero &

Zinger (2 SS)\ 3 July 2013, Paiero & Jackson (1 $, debu00367134); 22 May 2014, Paiero

& Zinger (2 additional males and females from this collection were sent to Funk for

further examination).

Cyrtoxipha gundlachi Saussure (Gryllidae: Trigidoniinae)

Cyrtoxipha gundlachi (Fig. 3), a species native to Florida and parts of the Caribbean

(Eades et al. 2013), was found at the Cambridge Wings of Paradise Butterfly Conservatory

and the Niagara Butterfly Conservatory. During visits to both sites, nymphs and adults

were observed on foliage No native Cyrtoxipha species occur in Canada but Cyrtoxipha

columbiana , which is similar in appearance to C. gundlachi , is found as far north as New

Jersey (Walker 1969) and southern Ohio (Walker and Moore 2012). Cyrtoxipha columbiana ,

like Anaxipha species 1 , may have been introduced with shipments of butterfly pupae, but it

may have also been brought in with plants used within the greenhouses as both greenhouses

have previously received plants from Florida.

Specimen Records: ONTARIO: Wellington Co., Cambridge, Wings of Paradise Butterfly

Conservatory, 43 0 27’7”N 80°22’2”W, in butterfly greenhouse, August 2013, S.M.

Paiero (3 SS, 3 $ $, debu00367 122-27); Niagara Reg., Niagara Falls, Niagara Butterfly

Conservatory, 43°10 , 37”N 70 o 3’20”W, 22 June 2013, S.M. Paiero (2 ??, debu00367135-

36); same as previous except 8 August 2013 (1 $,3 $$, debu00367 150-53); same as

previous except 7 October 2013 (4 SS, 2 $$, debu00367128-133)

Neoconocephalus exiliscanorus (Davis ) (Tettigoniidae: Conocephalinae)

Neoconocephalus exiliscanorus (the Slightly Musical Conehead) is here recorded

from Canada for the first time, from a marsh adjacent to the Wheatley Provincial Park

campgrounds. Although Vickery and Kevan (1985) indicate that N. exiliscanorus was

expected to occur in Canada, the new Wheatley record is about 250 km north of the most

northerly previous record. Its apparent preference for marsh habitat may be why this species

had not previously been found. If N. exiliscanorus is established in extreme southern

Ontario, additional populations may occur in similar habitats or along the shores of Fake

Erie (e.g., Point Pelee).

Specimens examined: ONTARIO: Kent Co., Wheatley Provincial Park, 42°5’25”N

82°26’50”W, 10-11 August 2007, S.M. Paiero (2 SS, debu00286872-73); same as

previous except 7 September2007, S.M. Paiero (2 SS, debu0029 11 56-57).

Neoconocephalus retusus (Scudder) (Tettigoniidae: Conocephalinae)

Neoconocephalus retusus (the Round-tipped Conehead) is an eastern North

American species here newly recorded from Ontario on the basis of observations and

collections from the Haldimand -Norfolk region. Gartshore and Carson (pers. comm.) have

heard this species singing at the same site in successive years, suggesting that the species is

established in Ontario. If N. retusus was previously overlooked in Ontario it may be either

due to its later emergence (Rehn & Hebard 1 9 1 5) or because it has a restricted distribution in

Paiero and Marshall

JESO Volume 145, 2014

the Norfolk area or because it was only recently established. Catling et al. (2009) previously

recorded this species as new to Canada on the basis of specimens from Nova Scotia, but

suggested that the species was adventitious rather than established.

Specimens examined: ONTARIO: Hald. -Norfolk Reg., Walshingham, ~5km

SW, Pterophyla, 42°38 , 27”N 80 o 34’29”W, 31 August 2010, M. Gartshore, (1 S,

debu00340774, one of several males heard singing by M. Gartshore and P. Carson), 27

September 2014, Gartshore & Carson (2 SS).

Chloroscirtus forcipatus (Brunner von Wattenwyl) (Tettigoniidae: Phaneopterinae) (Fig. 4)

Chloroscirtus forcipatus is a Central American species that has established a

population at the Niagara Butterfly Conservatory, likely being transported there in shipping

materials. Feeding and ovipositionby this exotic phytophagous species has caused significant

damage to a wide variety of cultivated plants at the conservatory and discussions with staff

indicate that it has been present there for many years.

Chloroscirtus forcipatus (Fig. 4) is superficially similar to the common bush

katydids in the genus Scudderia but the hind femur has no spines on the lateral genicular

lobe (Nickle 1992a), the eyes are more pronounced with the hind margin somewhat angled,

the fore wings are more strongly microreticulate, the femora are shorter (only extending

2/3 length of fore wings; Scudderia extend to 4/5 or more), and the male cerci are distinct

(Fig. 5). It will key out in Walker and Moore (2012) as Turpilia rostrata (Rehn & Hebard)

although T. rostrata has spines on the lateral genicular lobe of the hind femur.

Specimens examined: ONTARIO: Niagara Reg., Niagara Falls, Niagara Butterfly

Conservatory, 43°10 , 37”N 70 o 3’20”W, 22 June 2013, S.M. Paiero (2 ??, debu00367137-

38); 8 August 2013 (1 S, 2 ??, debu00367 154-56); 7 October 20 13 (5 SS, 3

debu00367 160-67).

Camptonotus carolinensis (Gerstaeker) (Gryllacrididae)

Also known as the Carolina Feaf-rolling Cricket, C. carolinensis (Fig. 6) is here

newly recorded from Canada on the basis of a specimen from Point Pelee National Park.

Since C. carolinensis is the only representative of this group in North America, it represents a

new family record for Canada. This predaceous species was previously known from Indiana

to Florida (Walker and Moore 2012). Camptonotus carolinensis is easily overlooked as it

is nocturnal and creates a leaf shelter in which to hide during the day, which might explain

why it was not found earlier in Ontario.

Specimens examined: ONTARIO: Essex Co., Point Pelee Natl. Pk., West Beach,

41°59’0”N 82°32 , 50”W, wooded area, Malaise & pan traps, 10-23 September 1999, O.

Fonsdale (1 S, debuOOO 13739).

Melanoplus punctulatus griseus (Thomas) (Acrididae)

New records of M. punctulatus griseus from southern Ontario represent the first

Canadian records of this subspecies, previously known from nearby Michigan (Vickery

and Kevan 1985, Bland 2003). Melanoplus punctulatus punctulatus (Scudder), the other

subspecies in Ontario, is widespread and extends east into Quebec. Bland (2003) reviewed

the differences between the two subspecies.

Specimens examined: ONTARIO: Kent Co., Rondeau Prov. Pk., South Point Trail, nr. East

Checklist of Ontario Orthoptera (cont.)

JESO Volume 145, 2014

parking lot, oak savannah, 42°15 , 42 ,, N 81°50’49”W, 10 October 2003, S.A. Marshall (1

S, 1 ?, debuO 11 34545-46); Essex Co., Point Pelee National Park, Visitor Centre, malaise

& pans, O. Lonsdale, 11-18 August 2000 (2 SS, debuO 1004046-47); same as previous

except 5-26 September 2000 (2 SS, debuO 1006370, debuO 1006376); same as previous

except 27 August-5 September 2000 (1 S, debuO 1006403).

Scapteriscus borellii Linnaeus (Gryllotalpidae)

We identified one specimen of S. borellii from an Oakville bakery and it almost

certainly represents an adventitious individual. Scapteriscus borellii is a Neotropical species

(Nickel 1992b) that has become established in the southeastern U.S with its northernmost

limit in North Carolina.

Specimen Examined: ONTARIO: Halton Reg., Oakville, found in a bakery, 13 October

1983, R. Ostrow (1 S, debuOl 076772).

Significant provincial range extensions

Meconema thalssinum (DeGeer) (Tettigoniidae: Meconematinae)

Meconema thalssinum (the Drumming Katydid) was originally recorded in Canada

in 2004 (Marshall et ah 2004) on the basis of specimens from Harrow, and it has since been

recorded in British Columbia as well (Cannings et al. 2007). This European species appears

to have become widespread and established in Ontario from Windsor to Toronto, in both

rural and urban environments.

Specimens examined: ONTARIO: York Reg., Toronto, 43°42’N 79°24’W, 8

July 2007, T. Careless, (1$ 1 $, debu00296956-57); Toronto, prey of Isodontia

mexicana (Hymenoptera: Sphecidae), July 2013, P.D. Careless (nymphs and adults;

photograph); Toronto, 43°42’N 79°25’W, grass, sweep, 10 August 2007, A. Turlco, (1$,

debuO 103 1467); Essex Co., Harrow, 42°2’N 82°55’W, hand collection, 11 August 1997,

M. Beaudoin (1<$ debuO 103 1465); Durham Co., Darlington Prov. Pk., 43 0 52’17”N”

78°47’02”W”, 11 September 2007, G. Vogg, {\$, debuO 103 1466); Kent Co., Wheatley

Prov. Pk., 42°5’25”N 82°26 , 50”W, 22 July 2011, S.M. Paiero, (\S 1$, debu00340211-

12); Hald. -Norfolk Reg., Turkey Point Prov. Pk., site 2, 42°42 , 28”N” 80°20’29”W,

savannah, at night , 4 August 2011, S.M. Paiero (1$, debuOl 148938); Wellington Co.,

Guelph, Wellington Woods, 43°31’12”N 80°13 , 53”W, 11 August 2011, D.K.B. Cheung

(1 $, debu00340228); Halton Reg. Oakville, nr. Hwy 25 & Burnhamthorpe Rd., August

2012, S.M. Paiero (1& debu00361466).

Neoxabea bipunctata (DeGeer) (Gryllidae)

This Carolinian species was originally recorded from Ontario on the basis of

specimens from Essex County (Marshall et al. 2004). The more recent records presented

here suggest that N. bipunctata now has a much more extensive range in southern Ontario.

Specimens examined: ONTARIO: Brant Co., Newport Forest, 30 July 2009, S.A.

Marshall (1 nymph, photographed); Elgin Co., Springwater Forest, 3 October 2013, J.

Allair (1$, Allair 2013); Essex Co., Wheatley Prov. Pk., 1 Sep 2007, S.M. Paiero (1 $,

debu00291096); Hald. -Norfolk Reg., Walshingham, ~5km SW, Pterophyla, 42 0 38’27”N

80 o 34’29”W,31 August 2013, at light sheet, Carson & Gartshore 2$ $; observed);

same data as previous except 1 September 2013 2$ $; photograph/observed); same

Paiero and Marshall

JESO Volume 145, 2014

data as previous except 3 September 2013 (1 observed); Halton Reg., Oakville, nr.

Hwy. 25 & Burnhamthorpe Rd., 29 August 2014, S.M. Paiero (1$); Kent Co., Rondeau

Prov. Pk., campground, 42°19’4”N 81°50’41”W, 25-26 September 2009, S.M. Paiero (1$,

debu00318141); Middlesex Co., London, Environmental Sciences Western Field Station,

43°4’29”N 81°20’13”W 13 September 2013, L. Des Marteaux (1$, photograph).

Myrmecophila pergandei (Bruner) (Myrmecophilidae)

Although this species was first formally recorded from Canada during an insect

survey of Ojibway Prairie, Windsor, Ontario (Paiero et al. 2010), the earliest Canadian

collection of M. pergandei (the Eastern Ant Cricket) was from Ancaster, Ontario (Borer’s

Falls Conservation Area) in 2006, and we have also found it at Wheatley Provincial Park,

Ontario. Myrmecophila pergandei is rarely encountered outside of ant colonies and most of

the specimens we have observed were found in slave-maker ant colonies {Formica species)

in Ancaster. Specimens from Wheatley Provincial Park and Ojibway Prairie Provincial

Nature Reserve were found at night on the bark of trees, walking with foraging carpenter

ants {Camponotus species).

Specimens examined: ONTARIO: Hamilton- Wentworth Reg., Dundas, Borer’s Falls

Conservation Area, in slave maker ant colony, in rotten fallen log, 24 May 2006,

Umphrey, Marshall & Paiero (10 nymphs, debu00264402-264411); Kent Co., Wheatley

Provincial Park, 42°5’25”N 82°26’50”W, on tree with carpenter ants at night, 22 July

2011, S.M. Paiero (1 S 1 ?, debuO 11 54454-55); Essex Co., Ojibway Prairie, 42°15’51”N

83°4’30”W, 6 September 2007, S.M. Paiero (2 ??, debu0029 1089-90)

Ectobius lapponicus (Linnaeus) (Ectobiidae)

This species, previously recorded in Canada from the Maritimes (Chandler 1992,

Clements et al. 2013), is here recorded from Ontario for the first time although it appears to

have been established in the province at least since 2006. Several specimens were collected

during a 2014 “Bioblitz” in Toronto’s Humber Valley and the species appears to be well

established in the area. Several Ectobius nymphs collected during a 2013 “Bioblitz” in the

nearby Rouge Valley are also likely to be E. lapponicus. Hoebeke and Carter (2010) gave

features to separate this species from other introduced Ectobius in the northeast.

Specimen data: ONTARIO: Muskoka Distr., Gravenhurst, Muskoka Lake, 1 July 2006,

on tree leaf, J.S. Maclvor, (1 S debuO 1040861); same as previous except 2 July 2006 (1

$ debuO 1040862); Peel Reg., Mississauga, nr. Mississauga Rd. & Dundas Rd., 18 July

2008, S.M. Paiero, (1 S debu00302523); York Reg., McMichael Canadian Art Gallery,

43°50’30”N 79°37’2”W, nymphs collected on gravel nr. lights, 24 May 2014, S.M. Paiero

(5 SS 3 $ $ 3 nymphs); Halton Reg., Oakville, nr. Hwy.25 & Burnhamthorpe Rd., 25

June 2014, S.M. Paiero (1 <$); same as previous except 1 July 2014 (1 <$).

Ectobius lucidus (Hagenbach) (Ectobiidae)

Hoebeke and Carter (2010) reviewed the distribution of Ectobius in northeastern

North America and gave characteristics to separate this species from other northeastern

Ectobius , but did not record E. lucidus from Ontario. Ectobius lucidus , like E. lapponicus ,

was first recorded from Ontario (Orillia, 20 June 2008, “helmetinthebush”, 1 $; Barrie,

Ontario, 25 June 2005, “shemiles”, 1 S) on the basis of images posted to and identified

Checklist of Ontario Orthoptera (cont.)

JESO Volume 145, 2014

on BugGuide.net. The additional specimen records below confirm the establishment of E.

lucidus in Ontario, and further suggest that this species has been here since at least 1973.

Specimen data: ONTARIO: Simcoe Co., Barrie, 2 August 1973, R.J. Hellewell, (1 S,

debuO 1035451); Barrie, 11 July 2010, “vireo” (1 S, photo posted on bugguide.net);

Midhurst, Neretra St., 2 July 2007, A. Brunlce, (1 S, debuO 1035452); Simcoe Co.,

Georgian College, u.v. light, 20 July 1977, E.R. Fuller (1 S, ROM); Simcoe Co., S of

Washago, mixed forest, 23 July 1992, L.D. Coote (1 S, ROM); Middlesex Co., London,

Public Submission, 12 June 2001, T.A. Zowinski (1 S 1 ?).

Dubious record

Orocharis saltator Uhler (Gryllidae: Eneopterinae)

The University of Guelph Insect Collection has a specimen of O. saltator (the

Jumping Bush Cricket) labelled as “Brant Co.?” without a collector or date. As no other

Brant County exists in Canada or the United States, it is presumed that this locality refers

to Ontario where it would be both a new species and subfamily record for Canada. The

northern limit of the range of this species is close to southern Ontario but we have not

included it in the checklist because the single record is doubtful.

Complete checklist of Ontario Orthoptera

Table 1 is a list of all 134 species and two additional subspecies of Orthoptera

recorded from Ontario, including native species (125, including multiple subspecies),

introduced species occurring outdoors (2), introduced species only found indoors (6), and

adventitious species (7, including intercepted material). Table 2 is a list of the 36 other

orthopteroids recorded from Ontario, including native species (6), introduced species

occurring outdoors (8), introduced species only found indoors (8), adventitious species (9),

and cultured species (5). Cultured species used in the pet trade were not treated as provincial

records, as they are not known to occur in Ontario outside of captivity. Nomenclature follows

Eades et al. (2013) for the Orthoptera, Beccaloni (2014) for the Blattodea, Deem (2014) for

the Dermaptera, Otte et ah (2014) for the Mantodea, and Brock (2014) for the Phasmida.

Changes in status

Melanoplus differentialis differentialis (Thomas) was considered by Vickery

and Scudder (1987) to be an introduced species. We consider it to be a native species

as its occurrence in Ontario is close to the northern limit of its historical range. Syrbula

admirabilis (Uhler) was also treated as an invasive by Vickery and Scudder (1987), but this

species too has a historical range with Ontario being its northern limit. Although there are

no recent Ontario records of S. admirabilis , historical records from southwestern Ontario

are consistent with the overall range of the species, and it is here considered as native to the

province. Similarly, Parcoblatta caudelli Hebard was treated by Vickery and Kevan (1987)

as an adventitious species in Ontario but, based on the overall distribution of this species

and its occurrence in Rondeau Provincial Park, we treat it as a native species. Psinidia

fenestralis fenestralis (Audinet-Serville) was treated by Vickery and Scudder (1987) as

“expected to occur in Ontario”, but we have not included it in the checklist because we have

yet to see a verifiable Ontario record of this species. Schistocera americana (Drury) was

treated by Vickery and Scudder (1987) as an adventitious species not established in Ontario.

Paiero and Marshall

JESO Volume 145, 2014

TABLE 1. Checklist of Ontario Orthoptera. (* denotes an introduced species occurring

outdoors; $ denotes an introduced species only found indoors; £ denotes an intercepted or

adventitious species).

ACRIDIDAE - Grasshoppers (52 species

+ 1 subspecies)

Acridinae (1 species)

Metaleptea brevicornis (Johansson)

Cyrtacanthacridinae (3 species)

Schistocerca alutacea (Harris)

Schistocerca americana (Drury)

£ Schistocerca lineata Scudder

Gomphocerinae (8 species)

Chloealtis abdominalis (Thomas)

Chloealtis conspersa Harris

Dichromorpha viridis (Scudder)

Orphulella pelidna (Burmeister)

Orphulella speciosa (Scudder)

Pseudochorthippus curtipennis curtipennis

(Harris)

Pseudopomala brachyptera (Scudder)

Syrbida admirabilis (Uhler)

Oedipodinae (16 species)

Arphia conspersa Scudder

Arphia pseudonietana (Thomas)

Arphia snlphnrea (Fabricius)

Camnula pellucida (Scudder)

Chortophaga viridifasciata (DeGeer)

Dissosteria Carolina (Linnaeus)

Encoptolophus sordidus (Burmeister)

Pardalophora apiculata (Harris)

Spharagemon bolli Scudder

Spharagemon coll are (Scudder)

Spharagemon marmorata marmorata

(Harris)

Stethophyma gracile (Scudder)

Stethophyma lineata (Scudder)

Trimerotropis hnronia E.M. Walker

Trimerotropis maritima (Harris)

Trimerotropis verruculata verruculata

(Kirby)

Melanoplinae (23 species + 1 subspecies)

Booneacris glacialis canadensis (Walker)

Booneacris variegata (Scudder)

Dendrotettix quercus Packard

Melanoplus angnstipennis (Dodge)

Melanoplus bivitattus (Say)

Melanoplus borealis borealis (Fieber)

Melanoplus bruneri Scudder

Melanoplus confusus (Scudder)

Melanoplus dawsoni (Scudder)

Melanoplus differentialis differentialis

(Thomas)

Melanoplus eurycerus Hebard

Melanoplus fasciatus (F. Walker)

Melanoplus femurrubrum (DeGeer)

Melanoplus huroni Blatchley

Melanoplus islandicus Blatchley

Melanoplus keeleri luridus (Dodge)

Melanoplus mancus Smith

Melanoplus punctulatus griseus (Thomas)

Melanoplus punctulatus punctulatus

(Scudder)

Melanoplus sanguinipes (Fabricius)

Melanoplus scudderi scudderi (Uhler)

Melanoplus stonei Rehn

Melanoplus walshii Scudder

Paroxya hoosieri (Blatchley)

Oxyinae (1 species)

£ Oxya hyla intricata (Stal)

ROMALEIDAE - Lubber Grasshoppers

(1 species)

£ Romalea microptera (Beauvois)

TETRIGIDAE - Pygmy Grasshoppers (7

species)

Batrachideinae (1 species)

Tettigidea lateralis lateralis (Say)

Tetriginae (6 species)

Nomotettix cri status cristatus (Scudder)

Baratettix cucullatus (Burmeister)

Tetrix arenosa angusta (Hancock)

Tetrix brunneri (Bolivar)

Tetrix ornata ornata (Say)

Tetrix subulata (Linnaeus)

TRIDACTYLIDAE - Pygmy Mole

Crickets (3 species)

Ellipes gurneyi Gunther

Ellipes minuta (Scudder)

Neotridactylus apicalis (Say)

MYRMECOPHILIDAE - Ant Crickets (1

species)

Myrmecophilus pergandei Bruner

Checklist of Ontario Orthoptera (cont.)

JESO Volume 145, 2014

TABLE 1 continued...

GRYLLIDAE - Crickets (23 species)

Gryllinae (4 species)

SAcheta domesticus (Linnaeus)

SGryllodes sigillatus (F. Walker)

Gryllus pennsylv aniens Burmeister

Gryllus veletis (Alexander & Bigelow)

Nemobiinae (6 species)

Allonemobius allardi (Alexander &

Thomas)

Allonemobius fasciatus fasciatus (DeGeer)

Allonemobius griseus griseus (E.M.

Walker)

Allonemobius maculatus (Blatchley)

Eunemobius carolinus carolinus (Scudder)

Neonemobius palustris (Blatchley)

Oecanthinae (10 species)

Neoxabea bipunctata (DeGeer)

Oecanthus argentinus Saussure

Oecanthus exclamationis Davis

Oecanthus fultoni T.J. Walker

Oecanthus laricix T.J. Walker

Oecanthus latipennis Riley

Oecanthus nigricornis F. Walker

Oecanthus niveus (DeGeer)

Oecanthus pini Beutenmtiller

Oecanthus quadripunctatus Beutenmtiller

Trigonidiinae (3 species)

%Anaxipha species 1

Anaxipha exigua (Say)

%Cyrtoxipha gundlachi Saussure

GRYLLOTALPIDAE - Mole Crickets (2

species)

Neocurtilla hexadactyla (Perty)

| Scapteriscus borellii Linnaeus

RHAPHIDIPHORIDAE - Camel Crickets

(9 species + 1 subspecies)

Aemodogryllinae (1 species)

SDiestrammena asynamora (Adelung)

Ceuthophilinae (8 species + 1 subspecies)

Ceuthophilus brevipes Scudder

Ceuthophilus divergens Scudder

Ceuthophilus guttulosus guttulosus F.

Walker

Ceuthophilus guttulosus thomasi Hubbell

Ceuthophilus latens Scudder

Ceuthophilus maculatus (Harris)

Ceuthophilus meridionalis Scudder

Ceuthophilus pallidipes E.M. Walker

Ceuthophilus uhleri Scudder

GRYLLACRIDIDAE - Leaf-rolling

Crickets (1 species)

Camptonotus carolinensis (Gerstaeker)

TETTIGONIIDAE - Katydids (35

species)

Conocephalinae (20 species)

Conocephalus attenuatus (Scudder)

Conocephalus brevipennis (Scudder)

Conocephalus fasciatus (DeGeer)

Conocephalus nigropleurum (Bruner)

Conocephalus saltans (Scudder)

Conocephalus strictus (Scudder)

Neoconocephalus ensiger (Harris)

Neoconocephalus exiliscanorus (Davis)

Neoconocephalus lyristes (Rehn & Hebard)

Neoconocephalus retusus (Scudder)

Neoconocephalus robustus (Scudder)

% Neoconocephalus triops (Linnaeus)

Orchelimum campestre Blatchley

Orchelimum concinnum Scudder

Orchelimum delicatum Bruner

Orchelimum gladiator Bruner

Orchelimum nigripes Scudder

Orchelimum silvaticum McNeill

Orchelimum voltanum McNeill

Orchelimum vulgare Harris

Meconematinae (1 species)

*Meconema thalassinum (DeGeer)

Phaneopterinae (9 species)

Amblycorypha oblongifolia (DeGeer)

SChloroscirtus forcipatus (Brunner von

Wattenwyl, 1878)

Microcentrum rhombifolium (Saussure)

Scudderia curvicauda (DeGeer)

Scudderia fasciata Beutenmtiller

Scudderia furcata furcata Brunner von

Wattenwyl

Scudderia pistillata Brunner von Wattenwyl

Scudderia septentrionalis (Audinet-

Serville)

Scudderia texensis Saussure & Pictet

Pseudophyllinae (1 species)

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JESO Volume 145, 2014

TABLE 1 continued...

Pterophylla camellifolia camellifolia * Roeseliana roeselii roeselii (Hagenbach)

(Fabricius) Sphagniana sphagnorum (F. Walker)

Tettigoniinae (4 species)

Atlanticus davisi Rehn & Hebard

Atlanticus testaceus (Scudder)

TABLE 2. Checklist of Ontario’s other orthopteroids (Blattodea, Dermaptera, Mantodea,

Phasmatodea). (* denotes an introduced species occurring outdoors; $ denotes introduced

species only found indoors; f denotes an intercepted or adventitious species, % denotes a

species that is reared in cultures in Ontario but not known to be established outside of these

cultures).

BLATTODEA - Cockroaches (22 species)

and Termites (3 species)

Blattidae (4 species)

$Blatta orientalis Linnaeus

$Periplaneta americana (Linnaeus)

%Periplaneta australasiae (Fabricius)

%Shelfordella lateralis (Walker)

Ectobiidae (5 species)

*Ectobins lapponicus (Linnaeus)

*Ectobins lucidus (Hagenbach)

%Nyctibora laevigata (Palisot de Beauvois)

%Nyctibora noctivaga Rehn

%%Symploce pallens (Stephens)

Blatellidae (7 species)

%Blatella germanica Linnaeus

%Cariblatta sp. A

Parcoblatta caudelli Hebard

Parcoblatta pennsylvanica (DeGeer)

Parcoblatta uhleriana (Saussure)

Parcoblatta virginica (Brunner von

Wattenwyl)

SSupe/la longipalpa (Fabricius)

Blaberidae (6 species)

%Blaberus discoidalis Serville

%Blaberns giganteus (Linnaeus)

%Blaptica dubia (Serville)

%Gromphadorhina portentosa (Schaum)

| Panchlora nivea (Linnaeus)

$Pycnoscelis surinamensis (Linnaeus)

(Infraorder ISOPTERA)

Kalotermitidae (1 species)

| Crypt otermes brevis (F. Walker)

Rhinotermitidae (2 species)

*Reticulitermes flavipes (Kollar)

| Reticulitermes virginicus (Banks)

DERMAPTERA - Earwigs (8 species)

Anisolabididae (2 species)

*Anisolabis maritima (Bonelli)

%EuborieUa armnlipes (Lucas)

Forficulidae (4 species)

| Chelidurella acanthopygina (Gene)

Dorn aculeatum (Scudder)

| Doru taeniatum (Dohrn)

* Forficula auricularia Linnaeus

Spongiphoridae (2 species)

*Labia minor (Linnaeus)

%Marava arachidis (Yersin)

MANTODEA - Praying Mantids (2

species)

Mantidae (2 species)

*Mantis religiosa Linnaeus

*Tenodera sinensis Saussure

PHASMATODEA- Walkingsticks (1

species)

Heteronemiidae (1 species)

Diapheromera femorata (Say)

Checklist of Ontario Orthoptera (cont.)

JESO Volume 145, 2014

Since S. americana has a historical eastern North American range within flying distance of

Canada, we consider it to be a native, albeit migrant, species. Pycnoscelis surinamensis was

also recorded by Vickery and Kevan (1987) as an adventitious species in Ontario but this

species is now well established in many greenhouses.

Nomenclatural changes

Nomenclatural changes since Vickery and Scudder (1987) are summarized below.

Acrididae

Schistocerca emarginata (Scudder) is now treated as a synonym of S. lineata Scudder

(Song 2004).

Stethophyma was previously treated within the Acrididinae (Vickery and Kevan 1985)

and Oedopodinae (=Locustinae, Vickery and Scudder 1987); it is now treated in the

Oedopodinae (Chapco and Contreras 2007).

Chorthippus curtipennis curtipennis (Harris) is now treated as Pseudochorthippus

curtipennis curtipennis (Harris) (Defaut 2012).

Arphia pseudonietana pseudonietana (Thomas) is now treated as A. pseudonietana ; no

subspecies are recognized (Otte 1984).

Orphulella pelidna pelidna (Burmeister) is now treated as O. pelidna; subspecies are no

longer recognized (Eades et al. 2013).

Spharagemon bold bolli Scudder is now treated as S. bolli Scudder; subspecies are no

longer recognized (Otte 1984).

Trimerotropis verruculata (Kirby) is now treated as Trimerotropis verruculata verruculata

(Kirby) because an additional subspecies is now recognized ( T. verruculata suffusa\ Eades

et al. 2013).

Trimerotropis maritima interior Walker is now treated as T. maritima (Harris); subspecies

are no longer recognized (Otte 1984).

Melanoplus viridipes eurycercus Hebard is now treated as a separate species, M.

eurycercus Hebard (Otte 2002).

Melanoplus femurrubrum femurrumbrum (DeGeer) is now treated as M. femurrubrum\

subspecies are no longer recognized (Eades et al. 2013).

Romaleidae

Romalea guttata (Houttuyn) is now treated as R. microptera (Beauvois) (the Eastern

Lubber Grasshopper); it is apparently occasionally carried north from the southeastern

USA in plant shipments or as bait by fishermen, but is not established in Ontario.

Tridactylidae

Ellipes minutus minutus (Scudder) is now treated as E. minuta (Scudder) (spelling

corrected); subspecies are no longer recognized (Eades et al. 2013).

Tetrigidae

Tettigidea lateralis (Say) is now treated as T. lateralis lateralis (Say) as another subspecies

is recognized (Rehn and Grant 1958). Vickery and Kevan (1985) recognized the

Paiero and Marshall

JESO Volume 145, 2014

subspecies but it was omitted in Vickery and Scudder (1987).

Gryllidae

The Tropical House Cricket ( Gryllodes sigillatus (F. Walker)) was previously confused

with G. snpplicans (F. Walker) (Otte 2006). G. sigillatus is occasionally brought into

Canada as cultures and is commercially available as reptile food, but is not known to be

established outside of cultures.

Rhaphidophoridae

Tachycines asynamorus Adelung is now treated as Diestrammena ( Tachycines )

asynamorus (Adelung); Tachycines has been lowered to subgeneric rank (Sugimoto 2002).

Tettigoniidae

Atlanticus davisi Rehn & Hebard was previously treated by some authors as a synonym of

A. monticola Davis but its species status is here retained, following Eades et al. (2013).

Roesefiana roeselii (Hagenbach) was placed in Roesefiana by Zeuner (1941) but has since

been treated by most authors as belonging to Metrioptera or Sphagiana. Massa & Fontana

(2011) reinstated Roeseliana as a valid genus. Multiple subspecies are also recognized and

R. roeselii roeselii is the subspecies present in North America (Eades et al. 2013).

Pterophylla camellifolia (Fabricius) is now treated as P. camellifolia camellifolia as

several subspecies are recognized (Alexander 1968, Walker and Moore 2012).

Mantidae

Tenodera sinensis Saussure was originally treated as a subspecies of T. aridifolia Stoll;

Ehrmann (2002) treated it as a separate species.

Discussion

Although the orthopteroids are relatively well studied in North America, further

Canadian records are to be expected. Several species so far unknown from Canada have

known ranges that almost reach the border, especially near southern Ontario. For example,

Oecanthus forbesi Titus, morphologically indistinguishable from O. nigricornis F. Walker,

likely occurs in southwestern Ontario based on range maps (Walker and Moore 2012) but

has not yet been formally recorded from Canada. This species would be most effectively

sought out on the basis of its song. Other species currently known from nearby localities

just south of the border, such as Phyllopaplus pulchellus Uhler or Trachyrhachys kiowa

(Thomas) are likely candidate additions to our fauna in response to a warming climate, and

might already occur here as undocumented populations. There also remains some possibility

of adding previously undescribed species to the provincial fauna. Several Anaxipha, for

example, have only recently been described from the eastern USA (Walker and Funk 2014)

and some of these may yet be discovered in southwestern Ontario. There is also a high

probability of continued discoveries of newly introduced exotic species. Although most

of the recently discovered introductions are of tropical species that are unable to survive

Checklist of Ontario Orthoptera (cont.)

JESO Volume 145, 2014

outside during the winter in Canada, several exotic orthopteroids ( Meconema thalassinum ,

Roeseliana roeselii roeselii, Ectobius lapponicus and E. lucidus) have become successfully

established in Ontario. And while the presence of Anaxipha species 1 is surprising, finding

an undescribed tropical species in temperate climates is not unprecedented. Weissman et

al. (2012) found a previously undescribed Gryllus species (G. locorojo Weissman & Gray,

also known as the “crazy red”), apparently native to South America, being used as a feeder

cricket in parts of Denmark, England and the United States.

The ongoing task of documenting the presence and ranges of orthopteroids (and

other arthropods) in Canada is expedited by keys and photographic guides that enable

amateurs and biologists without entomological training to recognize species. Vickery and

Kevan (1985; available through the Entomological Society of Canada’s website) remains

an important resource although can be difficult to use without access to an extensive

reference collection. Bland (2003), Walker and Moore (2012) and Kirk and Bomar (2005)

are more extensively illustrated guides that cover parts of Ontario’s fauna and supplement

the keys given in Vickery and Kevan (1985). Correctly identified photos of most Canadian

orthopteroid species can be found on bugguide.net ( http://bugguide.net/node/view/15740 )

and Walker (2014) provided keys and songs to the Gryllidae and Tettigoniidae of North

America.

Acknowledgements

The authors would like to thank M. Tomlinson, D. Constantinides and other

personnel at the Niagara Butterfly Conservatory for access to specimens and the facility;

A. Stapley and other personnel at the Cambridge Butterfly Conservatory for access to

specimens and the facility; Drs. T. J. Walker and D.H. Funk for confirming the identity of the

Anaxipha species ; Dr. P. Naskreki, for confirming the identity of Chloroscirtus forcipatus ;

Dr. G. Umphrey for drawing the authors’ attention to Myrmecophila in Ontario; P. Carson

and M. Gartshore for sharing their observations and records of Neoxabea bipunctata and

Neoconocelphalus retusus with the authors; Dr. R. Cannings for sharing specimen data

for Ectobius lapponicus ; and P.D. Careless for sharing his observations on Meconema

thalassinum.

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JESO Volume 145, 2014

THE ENTOMOLOGICAL SOCIETY OF ONTARIO

OFFICERS AND GOVERNORS

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President: I. SCOTT

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ISSN 1713-7845

JOURNAL

of the

ENTOMOLOGICAL SOCIETY

ONTARIO

Volume One Hundred and Forty Five

2014

THE ENTOMOLOGICAL SOCIETY OF ONTARIO

OFFICERS AND GOVERNORS

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