Syrphidae (Diptera) of Northern Ontario and Akimiski Island, Nunavut: New Diversity Records, Trap Analysis, and DNA Barcoding

Syrphidae (Diptera) of northern Ontario and Akimiski Island, Nunavut: new diversity records, trap analysis, and DNA barcoding

A Thesis Submitted to the Committee of Graduate Studies in Partial Fulfillment for the Degree of Master of Science in the Faculty of Arts and Science

TRENT UNIVERSITY Peterborough, Ontario, Canada © Copyright by Kathryn A. Vezsenyi 2019 Environmental and Life Sciences M.Sc Graduate Program May 2019

ABSTRACT

Syrphidae (Diptera) of northern Ontario and Akimiski Island, Nunavut: new diversity records, trap analysis, and DNA barcoding Kathryn A. Vezsenyi

Syrphids, also known as hover flies (Diptera: Syrphidae) are a diverse and widespread family of flies. Here, we report on their distributions from a previously understudied region, the far north of Ontario, as well as Akimiski Island, Nunavut. I used samples collected through a variety of projects to update known range and provincial records for over a hundred species, bringing into clearer focus the distribution of syrphids throughout this region. I also analysed a previously un-tested trap type for collecting syrphids (Nzi trap), and report on results of DNA analysis for a handful of individuals, which yielded a potential new species.

KEYWORDS

Syrphidae, Diptera, insects, northern Ontario, Akimiski Island, diversity, insect traps,

long term study, range extension, new species, DNA barcoding

ACKNOWLEDGMENTS

This thesis is written in dedication to my co-supervisor David Beresford, without whom none of this would have been possible. You have spent many long hours helping me with my data, my writing, and overall, my life. Your tireless, unwavering belief in me is one of the things that got me to where I am today, and has helped me grow as a person.

Your collection of insects from Akimiski is incredible, and I hope these collections will continue for years to come. Words cannot express my thanks.

Thank you to my co-supervisor Jim Schaefer; your funding, expertise, time, and edits were invaluable, and you raised my work to a higher level.

Huge thanks to my committee members, Jeff Skevington and Bill Crins. Your expertise of syrphids has taught me so much and your phenomenal syrphid field guide was an integral part of my thesis; I couldn’t have done it without you. Jeff, thank you so much for giving me the opportunity to work with you and your team at the Canadian

National Collections of Insects, Arachnids and Nematodes (CNC); it was an incredible opportunity for which I am grateful.

Thanks to my friends and co-authors from the CNC: Andrew Young, Michelle

Locke, and Kevin Moran. Your knowledge and enthusiasm for syrphids has taught me so much, and I am eternally grateful. Also thanks to Scott Kelso and Victoria Nowell, for teaching me about insect prep, and DNA barcoding.

This work would not have been possible without the Far North Biodiversity

Project; thanks to Dean Phoenix, John Ringrose, Ken Abraham, and everyone else who had a hand in the FNBP collection. Your work is invaluable, and I am grateful.

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My labmates in the Beresford lab have spent many long hours tirelessly sorting samples, slogging through spreadsheets, and supporting my thesis. Kaitlyn Fleming,

Sarah Langer, Kayla Vizza, Sherri DeGasparro, Ayden Ricker-held, and all the other volunteers I’ve had the pleasure of working with: thank you. You’ve made being trapped in a freezing cold windowless room an enjoyable experience.

Finally, thanks to my family and friends, who have motivated me to keep going.

My Mom, Michelle George, was particularly motivating and always around to talk when

I needed her the most. My Aunt, Jennifer Irwin, for your excellent review of my thesis; my unnecessary commas didn’t stand a chance. Also, thanks to Austin George for support, and Hana Vezsenyi for your institutional access getting me around paywalls.

Matt Yee, Kaitlyn Fleming, Sarah Langer, Vaughn Mangal, Nat Cummings, and Charlie

Pilgrim made my life enjoyable in the meantime.

TABLE OF CONTENTS ABSTRACT ...... ii KEYWORDS ...... ii ACKNOWLEDGMENTS ...... iii LIST OF FIGURES ...... vii LIST OF TABLES ...... ix Preface ...... xi REFERENCES ...... xvi Chapter 1: Sampling Syrphidae using Malaise and Nzi traps on Akimiski Island, Nunavut...... 1 ABSTRACT ...... 1 INTRODUCTION ...... 2 METHODS AND MATERIALS ...... 3 RESULTS ...... 7 DISCUSSION ...... 9 REFERENCES ...... 16 TABLES AND FIGURES ...... 22 Chapter 2: Distribution of Syrphidae (Diptera) across the far north of Ontario ...... 31 INTRODUCTION ...... 31 METHODS AND MATERIALS ...... 33 RESULTS ...... 36 DISCUSSION ...... 38 ACKNOWLEDGMENTS ...... 42 REFERENCES ...... 43 TABLES AND FIGURES ...... 48 APPENDIX ...... 58 Chapter 3: Barcode Syrphidae data from the far north of Ontario, and Akimiski Island, Nunavut...... 60 INTRODUCTION ...... 60 METHODS ...... 60 RESULTS ...... 62 DISCUSSION ...... 63 ACKNOWLEDGEMENTSREFERENCES ...... 65 TABLES AND FIGURES ...... 68 Chapter 4: General Discussion ...... 71

REFERENCES ...... 75

LIST OF FIGURES

Chapter 1

Figure 1: Map of study site on Akimiski Island, Nunavut. Page 28

Figure 2: Malaise trap (left) and cloth Nzi trap (right) deployed on Akimiski Island,

Nunavut, 2017. Page 29

Figure 3: Rarefaction curve of Malaise and Nzi syrphid sampling data from 2012–2017 on Akimiski Island, Nunavut. The bars represent standard deviation. Page 30

Chapter 2

Figure 1: All sites in the far north of Ontario from which syrphids were collected 2009-

2015 (black circles) and FNBP sites sampled from which no syrphids were collected

(white circles). Page 55

Figure 2 A-D: Species maps of individuals collected from the far north of Ontario that extended their overall range distributions: (A) Eristalis stipator, (B) Helophilus bottnicus,

(C) Platycheirus orarius, (D) Platycheirus thompsoni. Original ranges are in light grey, collected specimen in black. Page 56

Figure 3 A-F: New Ontario species records of Syrphidae, collected in the far north of

Ontario. (A) Helophilus bottnicus, (B) Melangyna labiatarum, (C) Platycheirus pictipes,

(D) Orthonevra robusta, (E) Platycheirus orarius, (F) Eupeodes curtus. Photos done by

Kaitlyn Fleming and KV. Page 57

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Chapter 3

Figure 1: Neighbour-joining tree using COI data featuring Platycheirus kelloggi, with the specimen previously identified as P. kelloggi outlined in black. Page 68

Figure 2: Putative new species, previously thought to be Platycheirus kelloggi, found to have almost a 1% difference in COI from west coast P. kelloggi individuals. Collected from Akimiski Island, Nunavut. Photo by KV. Page 69

Chapter 4

Figure 1: Rarefaction analysis of collected syrphids from northern Ontario and Akimiski

Island, NU, across all sampled years. Dotted lines represent 95% confidence intervals.

Page 77

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LIST OF TABLES

Chapter 1

Table 1: All syrphid species records of males (M) and females (F) for Akimiski Island,

Nunavut, Canada; (A) species found <400 km away from nearest record, (B) species found 400–800 km away from nearest record, (C) species found >800 km away from nearest record. Sex, counts, trap type, and years caught are included. *= New species record for Nunavut. Page 22-24

Table 2: Syrphid individuals collected by each trap type for each year of sampling on

Akimiski Island, Nunavut, Canada. Blanks indicate the years when traps were not deployed. Page 25

Appendix Table 1: Total syrphid catch from Malaise and Nzi traps, 2012–2017,

Akimiski Island, Nunavut. Blanks represent zeros. Page 26-27

Chapter 2

Table 1: All species of Syrphidae collected from the far north of Ontario, 2009-2014, with male (M) and female (F) counts, as well as traps they were collected in. Species with

* are new Ontario records. Record types for syrphids found >400km away from nearest collected record are as follows: gap infill (inf), edge of range (edg (direction)), and range extension (r.e (direction)). Records found more than 800 km from the nearest record are indicated by (>800). Page 48-51

Table 2: New provincial records of Syrphidae, as well as the dates, sex (male (M) and female (F)), trap type, and locations. Page 52

Table 3: Number of syrphids caught in each trap type per year. Page 53

Table 4: Number of Syrphidae species shared between three projects: the far north of

Ontario, Abitibi Model Forest (Dean et al., 2007), and Algonquin Park (Proctor et al.,

2011). Page 54

Appendix Table 1: All syrphid species collected from the far north of Ontario, categorized by the distance to their nearest collected record of that species (<400 km,

400-800 km, >800 km), and whether these new records are a gap infill, edge of range, or range extension. Page 58-59

Chapter 3

Table 1: Syrphidae specimens barcoded using COI. Data have been uploaded to the

Barcode of Life Datasystems (BOLD) website. Page 70

Preface

When I first went looking for a project for my Master’s thesis, I wasn’t exactly sure what I was looking for. I knew I was interested in insects, anything scientific that involved diversity, and learning about the natural world. Syrphids, while unknown to me at the time, were the perfect candidate group for me to study. They are not as flashy as butterflies and bumblebees, and are below the radar in many ways, underappreciated pollinators, mimics of their more well-known hymenopteran cousins. Nevertheless, this understated group has much to recommend it.

Insects all over the world have been shown to be in decline, either directly

(Biesmeijer et al, 2006; Conrad et al., 2006; Shortall et al., 2009; Hallmann et al., 2017) or indirectly (Nebel et al., 2010; Hallmann et al., 2014). This decline is not limited to any one group. In this regard, bees get almost all the attention in the media, which gives some the mistaken impression that bees are the penultimate, most important pollinator. While bees may be good at what they do, there are many other taxa working just as hard to accomplish plant pollination alongside them. Flies, and in particular syrphids (Diptera:

Syrphidae) are excellent pollinators that work tirelessly behind the scenes, in plain sight.

As they are Batesian mimics of Hymenopterans, such as bees and wasps, they often get away with flying under the radar of the common observer- both humans and predators alike. Once you know what to look for, it’s as if a whole secret world opens up – the secret world of Syrphidae.

Syrphids are a large and diverse family of flies that are found nearly world-wide.

The adults, charismatic and variably coloured, and function as pollinators for many plants. As larvae, they cover a huge array of ecological niches; within Diptera, they are

perhaps second only to Phoridae in their diversity of roles (Kirk-Spriggs and Sinclair,

2017). Larval roles range from specialized to generalist predators, parasitoids, decomposers, plant feeders using many different modalities (leaf miners, stem borers, root feeders, etc.), to prey (Rotheray and Gilbert, 2011). Generalist larvae (Syrphinae) are those that prey on soft-bodied insects such as aphids (Sommaggio, 1999). These tend to be more common and frequent crop fields to feed on these pests (Chambers and Adams,

1986; Sommaggio, 1999; Nelson et al., 2012). Specialist larvae (Eristalinae,

Microdontinae) require specific habitat features and can be more difficult to find.

Microdontines mostly prey on ant larvae, and as such are heavily associated with ants, while Eristalines are more variable in larval habitat requirements but tend to center around decaying plants and tree sap (Sommaggio, 1999). Syrphid communities within an area are a mosaic that reflect the habitat from which they come, and are exciting insects to work with as you can always find something that surprises you!

We know a lot about where different Syrphidae species are found, but there is always more to learn. Indeed, one of the most basic and important questions in entomology is, where is this species found (Beresford, 2018)? There are 313 species and

35 species known in Ontario and Nunavut, respectively. These species known in Ontario are largely from areas easily accessed, though almost half of Ontario makes up an area that is not so easily sampled. The far north of Ontario (MNRF, 2014) is made up of two different ecozones across a large expanse of wilderness. As one of the largest contiguous habitats in the world (MNRF, 2015), it is a rare and valuable area of study.

The far north of Ontario is made up of the Hudson Bay Lowlands and Ontario

Shield ecozones (Crins et al., 2009). It is best experienced through the window of a small

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twin otter plane, climbing to an altitude wherein all you can see for miles in every direction is thick, seemingly unending coniferous forests, interspersed with large swaths of wetland. Deceivingly small while looking at a map, it isn’t until you are flying over this area for over an hour that you understand the scale at which this ecosystem dominates. The Ontario Shield ecozone is the Ontario portion of the national Boreal

Shield ecozone, with its upper half within the far north boundary lines (Crins et al.,

2009). It is composed of large expanses of boreal forest, interspersed by many smaller wetlands. The Hudson Bay Lowlands ecozone is comprised of forests, large rivers, and coastal marshes and mudflats, making up the better part of the third largest wetland in the world (Crins et al., 2009).

Akimiski Island, Nunavut, is a large island found just off the coast of northern

Ontario, in James Bay. It is roughly 3,800 km2 and is largely uninhabited by humans. The only human settlement on the island is a small Ontario Ministry of Natural Resources and

Forestry (OMNRF) research camp on the north-east side, from which many samples originated. It is a post-glacial isostatic rebound island, formed roughly 4,000 years ago

(Degasparro et al., 2019). It has expansive tidal flats on all sides, with gravel ridges in the median, and with forested wetlands in the interior (Martini and Glooschenko, 1984).

Though geographically wide-spread collection efforts of syrphids tend to be few, there have been collaborative work such as that by Skevington et al. (2019) to pull their distributions into a larger picture in North America. The theme of this thesis is that of discovery; syrphids are still being discovered, distributions maps still getting filled in.

Biogeography is an on-going process, and all begins with describing species in an area. It has never been more important to monitor insect populations, and that starts with a good

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baseline bioinventory. Here we give the first extensive report on the syrphids from the far north of Ontario and Akimiski Island, Nunavut.

Chapter 1 “Sampling Syrphidae using Malaise and Nzi traps on Akimiski Island,

Nunavut” (Vezsenyi et al. in press), examines syrphids from Akimiski Island, Nunavut, and discusses the species found there relative to their previously known ranges and collected records. It also features some trap analyses comparing a previously untested trap (for collecting syrphids), the Nzi trap, to the well-known Malaise trap. We also report on 55 new species records to the territory of Nunavut, more than double the number of species currently known from the region.

Chapter 2, “Distribution of Syrphidae across the far north of Ontario” keeps with the theme of examining syrphids from remote locations, this time in the far north of

Ontario. Though not often realized, about 42% of Ontario is inaccessible by road, and as such, insect collections from the region are few and far between. This chapter examines syrphids collected from across this region, from a number of different projects. These syrphids are also compared to their overall distributions and previously collected records; we also compare the different syrphid sub-families sampled to other relatively nearby studies, in terms of larval life history traits.

Chapter 3, “Barcoding Syrphidae from the far north of Ontario and Akimiski

Island” takes a brief look at COI data taken from some of the more geographically and morphologically unusual or rare syrphids from our collections. Though most sequences were as expected, we sequenced one species for the first time, and one individual from

Akimiski Island, Nunavut, has barcode data suggesting it is a new species that has yet to be described.

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Chapter 4 “General Discussion” briefly summarizes and compares Chapters 1-3, and gives some closing remarks on my thesis.

REFERENCES

Beresford, D. V. 2018. President's Address. Entomological Society of Ontario Fall

Newsletter 23(2): 3.

Biesmeijer, J. C., S. P. M. Roberts, M. Reemer, R. Ohlemüller, M. Edwards, T. Peeters,

A. P. Schaffers, S. G. Potts, R. Kleukers, C. D. Thomas, J. Settele, W. E. Kunin.

2006. Parallel declines in pollinators and insect-pollinated plants in Britain and

the Netherlands. Science 313(5785): 351-354.

Chambers, R. J., T. H. L. Adams. 1986. Quantification of the impact of hoverflies

(Diptera: Syrphidae) on cereal aphids in winter wheat: an analysis of field

populations. Journal of Applied Ecology 23(3): 895-904.

Conrad, K. F., M. S. Warren, R. Fox, M. S. Parsons, I. P. Woiwod. 2006. Rapid declines

of common, widespread British moths provide evidence of an insect biodiversity

crisis. Biological Conservation 132: 279-291.

Crins, W.J., P.A. Gray, P.W.C. Uhlig, M.C. Wester. 2009. The Ecosystems of Ontario,

Part 1: Ecozones and Ecoregions. Science and Inventory Branch, Inventory,

Monitoring, and Assessment, Ontario Ministry of Natural Resources,

Peterborough, Ontario.

Degasparro, S. L., G. S. Brown, Y. Alarie, D. V. Beresford. 2019. Records of predaceous

diving beetles (Coleoptera: Dytiscidae) from Akimiski Island, Nunavut, Canada.

The Coleopterists Bulletin 72(4): 866-869.

Hallmann, C. A., R. P. B. Foppen, C. A. M. van Turnhout, H. de Kroon, E. Jongejans.

2014. Declines in insectivorous birds are associated with high neonicotinoid

concentrations. Nature 511(7509): 341.

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Hallmann, C. A., M. Sorg, E. Jongejans, H, .Siepel, N. Hofland, H. Schwan, W.

Stenmans, A. Müller, H. Sumser, T. Hörren, D. Goulson, H. de Kroon. 2017.

More than 75 percent decline over 27 years in total flying insect biomass in

protected areas. PLOS One https://doi.org/10.1371/journal.pone.0185809.

Kirk-Spriggs, A. H., and B. J. Sinclair. 2017. Manual of Afrotropical Diptera Volume 1

& 2. SANBI Graphics & Editing, Pretoria, South Africa.

Martini, I. P., and W. A. Glooschenko. 1984. Emergent coasts of Akimiski Island, James

Bay, Northwestern Territories, Canada: geology, geomorphology, and vegetation.

Sedimentary Geology 37(4): 229-250.

MNRF. 2014. Far North land cover data specifications version 1.14. Ministry of Natural

Resources and Forestry, Ontario, Canada.

MNRF. 2015. Far North land use strategy: a draft. Ministry of Natural Resources,

Ontario, Canada.

Nebel, S., A. Mills, J. D. McCracken, P. D. Taylor. 2010. Declines of aerial insectivores

in North America follow a geographic gradient. Avian Conservation and Ecology

5(2): 1. [online] http://dx.doi.org/10.5751/ACE-00391-050201.

Nelsen, E. H., B. N. Hogg, N. J. Mills, K. M. Daane. 2012. Syrphid flies suppress lettuce

aphids. BioControl 57(6): 819-826.

Rotheray, G. E., and F. Gilbert. 2011. The natural history of Hover Flies. Tresaith, Wales,

United Kingdom: Forrest Text 333 pp.

Shortall, C. R., A. Moore, E. Smith, M. J. Hall, I. P. Woiwod, R. Harrington. 2009. Long-

term changes in the abundance of flying insects. Insect Conservation and

Diversity 2: 251-260.

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Skevington, J. H., Locke, M. M., Young, A. D., Moran, K., Crins, W. J., Marshall, S. A.

2019. Field Guide to the Flower Flies (Hover Flies) of Northeastern North

America. Princeton University Press, Princeton, NJ 512 pp.

Sommaggio, D. 1999. Syrphidae: can they be used as environmental bioindicators?

Agriculture, Ecosystems and Environment 74: 343-356.

Vezsenyi, K. A., J. H. Skevington, K. Moran, A. D. Young, M. M. Locke, J. A. Schaefer,

D. V. Beresford. In press. Sampling Syrphidae using Malaise and Nzi traps on

Akimiski Island, Nunavut. Journal of the Entomological Society of Ontario

(accepted January 14, 2019).

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Chapter 1: Sampling Syrphidae using Malaise and Nzi traps on Akimiski Island, Nunavut.

(paper accepted and in press for the Journal of the Entomological Society of Ontario, January 14, 2019) ABSTRACT

Flower flies (Diptera: Syrphidae) are a diverse group of pollinators found almost worldwide. Species surveys of these flies provide unique challenges, as they can be difficult to collect due to different trapping biases. Here, we test the efficacy of the Nzi trap for use in the collection of syrphids by comparing the richness and abundance of syrphids caught in a Malaise and Nzi trap, July, 2012-2017, on Akimiski Island, Nunavut.

We found that the Nzi trap caught many of the same species in similar abundances as the

Malaise trap, except for Platycheirus kelloggi (Snow), which was caught more in the Nzi than the Malaise. The high capture rate of P. kelloggi using Nzi traps could be due to unique behaviours of the flies’ shelter or mate seeking related to structure or colour.

Using collections from 2008-2017, we also provide new territory records for 55 species and range extensions for 19 species. Two of these, Platycheirus kelloggi and Platycheirus latitarsis Vockeroth had previously been reported only west of the Rocky Mountains.

KEYWORDS Akimiski Island, Diptera, insect pollinators, Nunavut, Syrphidae, range extensions

INTRODUCTION

Flower flies (Diptera: Syrphidae), also known as hover flies or syrphids, are a common and diverse family of dipteran pollinators (Miranda et al 2013). To elucidate the impact of individual flower fly species on wild pollination, we first need to understand those species’ geographic distributions. Unfortunately, our knowledge of species ranges is often hampered by a lack of species surveys in remote locations. In this paper, we report on the syrphid species caught on Akimiski Island, Nunavut—a northern island in

James Bay, Canada. Few studies of insect diversity have been conducted on Akimiski

Island, with previous studies focusing on biting flies (Beresford et al 2010) and beetles

(Fleming and Beresford 2016; DeGasparro et al 2018).

Species surveys of syrphids are often challenging due to the specialized niches many species inhabit. Hand netting is the most common sampling method, although results may be influenced by time of day, season, habitat, and collector expertise

(D’Amen et al 2008; Gill and O’Neal 2015). The use of passive intercept traps to capture flying adults, such as Malaise traps, removes many of these problems as they can be placed for longer periods of time. While Malaise traps capture a wide variety of insect species with little effort (Brown 2005), they may miss species that are trap-shy or occur only in specialized niches. Traps that target pollinators, such as pan traps, have produced mixed results for sampling syrphids (Campbell and Hanula 2007; Sadeghi Namaghi and

Husseini 2009; Proctor et al 2012).

Here, we report on syrphid species collected using five different methods, along with range extensions and new species records for Nunavut. Some syrphid species are attracted to blue (Chen et al 2004; Campbell and Hanula 2007), so it is likely that blue

and black cloth traps, or Nzi traps (Mihok 2002), may provide an additional tool for sampling previously under-collected species. To test this hypothesis, we compared the

Nzi trap catches to Malaise trap catches deployed at the same time, at the same location.

Malaise traps are similar to mist nets, intercepting flying insects, and are non-attracting

(more or less). As a starting point, we surmised that the effectiveness of an intercept trap might be based on its interception surface area. Our null hypothesis was that the Nzi trap works as a smaller non-attracting intercept trap along the lines of the Malaise trap, its effectiveness based on interception surface area. Other trapping methods used for the general collection of insects from Akimiski Island (International Polar Year trap, sticky trap, hand netting) were not included in this comparison as they were not standardized over time and were used on an ad hoc basis.

METHODS AND MATERIALS

Study site: All collecting took place on Akimiski Island, Nunavut, Canada, at the biological research station (53°6'18"N, 80°57'25"W) operated by the Ontario Ministry of

Natural Resources and Forestry. This large island (3,800 km2) is located in James Bay, near the coast of northern Ontario (Fig. 1). It is part of the Hudson Plains ecozone, with an extensive sand and gravel shoreline, and many bogs, fens, and spruce stands in the interior (Martini and Glooschenko 1984). The mean annual temperature is -3.5ºC, and mean annual precipitation is around 700 mm (Ecological Stratification Working Group

1995). Uninhabited by humans, two thirds of the island is a migratory bird sanctuary, serving as an important stopover site for many migrating shorebirds and geese (Abraham et al 1999; Jefferies et al 2006; Pollock et al 2012).

Collection methods: Collecting occurred in the last two weeks of July 2009–

2017, and in June and August, 2008. Trapping methods included Malaise, Nzi, sticky traps, hand netting, and IPY (International Polar Year) traps (McKinnon et al 2008).

We used a single Malaise trap (Fig. 2; Lightweight Malaise Trap, Townes Style;

176 cm high, 165 cm long; model no. 2868, BioQuip Products, Rancho Dominguez, CA) from 2012 to 2017. This was placed within the research station fenced-in area to prevent polar bears from damaging the trap, 10 m from the Nzi trap.

Nzi traps (Fig. 2) are blue and black cloth traps designed for biting flies (Mihok et al 2006). A single cloth Nzi trap was deployed each year, 2008–2017 (Fig. 2); in 2010 and 2011 a second Nzi trap made from Coroplast® (Coroplast, a division of Great Pacific

Enterprises Inc., Granby, QC, and Dallas, TX) plastic panels was also used in addition to the cloth Nzi trap. Nzi and Malaise collecting heads were charged with propylene glycol

(non-toxic RV antifreeze) as a drowning agent and preservative; these were emptied every 12 hours (at dawn and dusk), and specimens were stored in 80% ethanol for preparation later.

Different coloured sticky card traps (Beresford and Sutcliffe 2006) were set in

August 2008. Coloured Coroplast (black, white, yellow, blue, red) cards, 20 cm x 30 cm, were coated with Tangle Trap® (Tangle Foot Co., Chicago, IL) and screwed to a wooden stake, 1 m apart. Acetone was applied to the cards with a dropper so we could remove the specimens. Flies were then removed and pinned, then soaked in acetone to completely remove the Tangle Trap.

Hand netting took place on an ad hoc basis during the day, throughout the sampling period, sweeping over blossoms. Specimens were then pinned.

IPY traps (Gan et al 2009) consisted of a 40-cm wide black screen stretched between two 50-cm tall wooden stakes, with the screen overtop of a plastic tray, and the entire structure surmounted by a white plastic funnel leading upward to a collecting bottle. This trap intercepted low flying insects, capturing those that either crawled up or down to escape as well as insects that crawled into the bottom tray (Gan et al 2009). The bottom tray and the upper bottle were charged with soapy water. They were only placed during the sampling period from 2008 to 2009. Fifteen IPY traps were placed near the shoreline between the high tide mark and the beach area along the coast and set about 50 m apart (Gan et al 2009; Beresford et al 2010).

Specimen preparation: All specimens were pinned; most were first preserved by critical point drying using a Leica critical point dryer (Leica EM CPD300, Leica

Microsystems Inc., Concord, ON), then pointed. Identifications were performed using

Skevington et al (2019), supplemented by keys from Vockeroth (1992), Miranda et al

(2013), and Young et al (2016). The collection is currently housed in the Trent University

Entomology lab and will be deposited in the Canadian National Collections of Insects,

Arachnids and Nematodes (Ottawa, ON) in the future.

Analyses: Nzi and Malaise comparison We compared the proportion of singleton species, and the number of unique species, caught by Malaise and cloth Nzi traps for the years 2012-2017 with 2 x 2 contingency table tests for independence (chi square test, d.f. = 1).

We also compared the abundance of each species caught by Malaise and cloth Nzi traps for the years 2012-2017. Because the Malaise trap is larger than the Nzi trap, and if

Nzi traps do not attract but only intercept insects, we expected to catch fewer individuals

in the Nzi trap, without making any assumption of how the effective surface area of each trap might change with trap height, direction or placement. The Malaise trap had a surface area of 2.9 m2 and the Nzi trap had an effective surface area of 1.6 m2. Based on the simple premise that interception area governs trap rate, our null hypothesis was that trap catches were independent of trap type. We tested this for species with total catches of ten or more individuals using a Fisher's exact test of independence, set up as the total number of a species in each trap type versus the number of other species in each trap type

(all years combined). We then used a sequential Bonferroni correction (Sokal and Rohlf

1997) for these tests. These tests were performed using STATISTICA 7 (StatSoft Inc.

Tulsa, OK).

In order to estimate how many additional species had yet to be caught from this community, we fitted the abundance data, using the total combined Nzi and Malaise catches from 2012-2017, to a log normal distribution following the method used by

Preston (Preston 1948; Ludwig and Reynolds 1988). In addition, we estimated species richness using several other methods: Chao 1, Chao 2, Jackknife 1, Jackknife 2, and the bootstrap method (Gotelli and Colwell 2011). These were performed using PAST 3.20

(Hammer et al 2001).

Range Extensions: The known ranges were determined for each species caught using data from the Field Guide to the Flower Flies (Hover Flies) of Northeastern North

America (Skevington et al 2019). We defined a range extension as a new record for a species more than 400 km away from the nearest previous record or from a line between two previous records that extended the known range in any direction. We defined a gap infill as a new record in an area between previous records that were at least 400 km away,

but did not extend the known range in any one direction. This range extension distance was chosen arbitrarily; however, a distance of 400 km crosses a number of plant hardiness zones from north to south, indicative of changing plant communities. A distance of 800 km almost always crosses into a different ecozone outside of the Hudson

Plains, meaning these records originated from fundamentally different habitats.

RESULTS

We caught 598 individual syrphids, and were able to identify 553 specimens to 73 species; the remaining 45 individuals could be identified only to genus. These unknowns were either damaged (two specimens) or were females (43 specimens) from genera in which males are required for identification. Although we could not identify these directly to species, we do know that at least two of these groups were additional species as we had no other individuals from these particular groups: Eupeodes americanus

(Wiedemann)/pomus (Curran) (4 individuals), and Paragus (Paragus) sp. (3 individuals).

Of the 73 species collected, 55 were new species records for Nunavut (Table 1 a–c). The most abundant species were Cheilosia latrans (Walker), making up 20% (n= 103) of all syrphid catches, followed by Melanostoma mellinum (Linnaeus) at 12% (n= 36). There were 29 singleton species, 39.7% of all species identified, but only 4.8% of the total catch

(29/598 specimens).

The catches by trap type are summarized in Table 2. Most specimens were obtained using the Malaise traps and the Nzi traps (by species Table 1; summary Table

2).

The estimates of species richness were (in descending order): Jackknife 2 =

131.76, Chao 2 = 123.14 (SD = 21.79), Chao 1 = 116.2 (range 79.92 to 118.1), Jackknife

1 = 109 (SD = 10.39), Bootstrap = 88.5, Preston’s log-normal = 76.1.

Range extensions: We found 54 species that were expected in this area, as they were found less than 400 km away from the nearest record (Table 1a). There were nine species found 400–800 km away from the nearest record (Table 1b), and 10 species records that were more than 800 km away from the nearest collection record (Table 1c).

Of the species from Table 1c, three of these (Cheilosia laevis (Bigot), Eristalis hirta

Loew, and Platycheirus neoperpallidus (Young) represent large gap infills, with the others being large range extensions.

Trap Analysis: Overall, we caught 51 species (271 individuals) in the Malaise traps and 29 species (122 individuals) in the Nzi traps, 2012–2017. Of these, Malaise and

Nzi traps had more-or-less the same proportion of singletons (21 and 8 singleton species respectively; 2 x 2 contingency test: χ2 = 0.71, p = 0.40). For 2012–2017, Malaise and

Nzi traps each caught 33 and 11 unique species respectively, which was not significantly different (2 x 2 contingency test: χ2 = 1.64, p = 0.20).

When we compared catches by species, three species appeared to have higher Nzi trap numbers than expected if the Nzi trap was simply acting as a flight intercept trap:

Lapposyrphus lapponicus (Zetterstedt) (Malaise: Nzi, 17:17), Platycheirus pictipes

(Bigot) (7:10), and Platycheirus kelloggi (Snow) (3:10). After we applied the sequential

Bonferroni correction, only P. kelloggi showed a significant trap bias (Fisher exact test, p

= 0.0011, sequential Bonferroni corrected α= 0.05/8 = 0.00625), with substantially higher numbers in the Nzi trap. From this, we tested whether these traps differed in how many

Platycheirus spp. (excluding P. kelloggi) were caught and found significantly more

Platycheirus spp. caught by the Nzi trap (26) than by the Malaise trap (24) (χ2 = 11.99, p

= 0.0005).

The rarefaction curve of the Malaise and Nzi trap catches (Fig. 3) shows that, when corrected for sample size, the number of species caught in each trap was not substantially different, although the curve for the Nzi trap was somewhat lower.

DISCUSSION

Our Nzi and Malaise comparison shows that, aside from Platycheirus kelloggi, the Nzi trap likely functions as a flight intercept trap for syrphids. Although the Nzi did collect a number of species that the Malaise did not, most were composed of single individuals, and from our analysis, these individuals were most likely an aimless occurrence in one of the two traps.

Like many syrphids, Platycheirus Lepeletier & Serville often congregate around specific landscape features to find mates (Barkalov and Nielsen, 2007). However, unlike most syrphids, Platycheirus are often active on overcast or light rain days, and in cool, shaded areas (Barkalov and Nielsen 2007; Skevington et al 2019). All individuals of P. kelloggi collected in Nzi traps were female, which indicates that this species likely seeks out a different microhabitat when looking for mates. It is possible that P. kelloggi and other Platycheirus were collected more frequently in the Nzi traps because other syrphids avoid the dark colours and shaded areas created by the trap. Conversely, it is possible that female P. kelloggi were specifically attracted to the Nzi trap due to the colour, mistaking it either for an oviposition site or for a male conspecific. The latter scenario could be taking place because male P. kelloggi, unlike most Platycheirus species, have an overall

blue colouration. This would explain why only P. kelloggi was significantly associated with the traps, and not other shade-loving species of Platycheirus.

Studies with blue traps from warmer regions have reported catches of different species in high abundance: Allograpta obliqua (Say) (Chen et al 2004) and Melangyna viridiceps (Macquart) (Broughton and Harrison 2012), so it is likely that using an Nzi trap south of our study area would yield different proportions of species. The total number of individuals caught by the Nzi trap was still more than expected given the smaller trap surface. Even so, for those interested in general syrphid surveys, the Malaise remains the better choice, with higher catch rates and more species (Table 1 a–c, Table

2).

Preston’s log normal estimate (76 species) came the closest to the total number of species that we captured (73 to species, and two additional species that could not be identified). The other methods, however, suggested as many as 57 additional species. It is quite likely that new species could be found by sampling different times of year, as well as different microhabitats across the island. Generally, the Chao 1 and Jackknife 1 methods are appropriate tests for estimating species richness (Gotelli and Colwell 2011;

Samarasin et al 2017) — in our case, 116 and 109 species respectively. These methods are based on the number of rare species, those with one or two individuals, and are not based on assuming an underlying distribution, whereas the log-normal method does assume that species abundance has a log-normal distribution, and that sampling is more or less unbiased. Any trap biases would have a larger effect on log-normal estimates. In our study, the estimates least affected by sampling bias were Jackknife 2 (132 species;

Hortal et al 2006) and Chao 2 (123 species; Colwell and Coddington 1994).

We found 55 new Nunavut species records, which more than doubles the current known number of species for the territory (n=35). Most of these were expected based on their current distributions in nearby jurisdictions (Manitoba, Ontario, Quebec). Akimiski

Island is considerably south of the Nunavut mainland, so while these records may be expected from the area ecologically, they were not expected from Nunavut. Jurisdictional boundaries can be important in conservation practice; in that regard, Akimiski Island is significant for Nunavut. It is the only island in Hudson Bay of that size without permanent residents and has protections through the bird sanctuary on the eastern half of the island.

Table 1a represents expected species, found less than 400 km away from the nearest previous record. Most of these represent gap infills in their distributions. Quite a few of these species, though not far enough to constitute range extensions by our criterion, were found at the edge of their species range. Scaeva affinis (Say) is quite common in western North America, with our record and a nearby record from the far north of Ontario representing the eastern-most points of its distribution. Though there is one southern Ontario record, it is presumed to be a vagrant (the species appears to be migratory in the west; Skevington, unpublished data).

Table 1b represents species with intermediate-sized gaps in distributions; this could be due to lack of sampling from the nearby regions. Orthonevra robusta (Shannon) is found largely in the western United States and has a small disjunct population along the southern shores of Hudson Bay, which includes our record from Akimiski Island, as well as one record from Churchill, Manitoba, and one from Fort Severn, Ontario.

Ten species were found more than 800 km away from previous records (Table

1c). Cheilosia laevis is found primarily in western North America towards the Pacific coast, with an outlying record on Anticosti Island towards the southeast of Quebec.

Eristalis hirta is found primarily west of the Rocky Mountains, although its distribution also extends into the northern territories and eastward. The nearest records are in northern

Manitoba and northern Quebec; the records from northern Manitoba are also in the

Hudson Plains ecozone. Platycheirus neoperpallidus is a boreo-montane species with the nearest records from southern Manitoba, to the west, and eastern Quebec, to the east. Our record falls midway between these two closest known localities. Platycheirus nielseni

Vockeroth is found primarily in the Yukon, and more sparsely distributed to the east, in

Nunavut, Newfoundland, Manitoba, and further south in Colorado. Each of these records represent gap infills for their respective populations.

The remaining six species from Table 1c are all large range extensions.

Platycheirus kelloggi and Platycheirus latitarsis Vockeroth represent eastern extensions of their populations. Both are known strictly west of the Rocky Mountains, which possibly acts as an ecological barrier to their distribution. Eumerus strigatus (Fallén) and

Neocnemodon elongata (Curran) represent northern range extensions. Both populations extend from the Pacific coast to the Atlantic coast, along the border between the United

States and Canada. Eumerus strigatus larvae tend to be associated with the bulbs of flowering plants, so this new record may have been associated with native bulb plants on

Akimiski Island, although finding this species in an area devoid of horticulture might also suggest that it is a vagrant (Blaney and Kotanen 2001; Kizil et al 2008). Pipiza atrata

Curran is a northeastern range extension. It is a rare species, mostly found west of the

Rocky Mountains, though it has been recorded from Minnesota, and southern Ontario.

Platycheirus jaerensis (Nielsen) is the only western range extension; it is a species found strictly along the east coast, ranging from Labrador to Maine.

The IPY traps caught a fairly high number of syrphids given the small trap size; they were composed mainly of two species: Eristalis brousii Williston (n= 34) and

Orthonevra robusta (n= 32). They were caught in low numbers when IPY traps were not in use (2009–2017), though this difference in abundance is likely due to the difference in habitats where trapping took place. IPY traps were deployed along the shoreline, while all other trapping was done in camp, more inland towards the tree line. Both are species of interest: O. robusta was previously collected this far north only in Churchill, which is considerably disjunct from its western population in the United States. Eristalis brousii was once a common and widespread species whose range has since diminished and become restricted to northern latitudes, making it one of the most at-risk species of syrphid in North America (Skevington et al 2019). Since recognizing E. brousii in the

IPY trap samples, we have collected two more specimens by hand netting, one in 2010 and one in 2013.

The other notable catches from the IPY were Cheilosia lasiophthalmus Williston, an uncommon species, our record being on the western edge of the eastern-most population, as well as Platycheirus jaerensis, an eastern, coastal species. The latter’s appearance on Akimiski was unexpected, though the island is considered coastal and likely has the required habitat.

Akimiski Island includes a mix of unique habitats, on a low arctic island, that likely provides specialized microhabitats for many syrphid species. The landscape of

Akimiski Island is a combination of arctic-coastal and boreal forest, providing habitats for northern species and southern species. The placement and size of the island in James

Bay could also be responsible for many large range extensions – insects transported by prevailing westerly winds could be deposited on the landscape when they encountered cold air over James Bay. As an island in James Bay, Akimiski has a maritime climate, with extreme temperatures moderated in comparison to continental landscapes on the adjacent mainland; this may provide a climate refuge for species normally found in more southern regions. Another feature unique to this island is the seasonal residence of abundant geese and shorebird populations. These have profound effects, denuding many areas of plants and changing plant community composition (Jefferies et al 2006), with

(we expect) similar effects on pollinator species such as syrphids. Insectivorous shorebirds rely on insects for food (Tulp et al 2008; Bolduc et al 2013), so the high number of shorebirds on Akimiski could also conceivably affect these insect populations.

In this paper, we reported on an interesting response to Nzi traps by species of the genus Platycheirus, P. kelloggi in particular. This study shows the continued importance and need of faunistic studies in remote regions. We have increased the number of known syrphid species from Nunavut, and reported several large range extensions, filling in much needed distributional information for flower flies in our northern ecosystems.

Perhaps most important in the long term, projects like these create a rich comparative dataset that will allow us to track distribution changes in response to changing environmental pressures and help us to discover and highlight habitats and species at risk.

ACKNOWLEDGMENTS

We thank the MNRF Wildlife Division for support for this work, in particular

Ken Abraham, Glen Brown, Rod Brook, Kim Bennett, Bill Crins, and Sarah Hagey; Dan

Steckly, Michelle Carlisle, and Karen Shearer for help netting specimens; Danica Hogan

and Lisa Pollock for collecting the IPY specimens; Dean Phoenix and Dave Etheridge

(MNRF Far North) for funding KV’s contribution to this work; Kaitlyn Fleming, Sarah

Langer, Sherri DeGasparro, Kayla Vizza, and everyone else from the Trent Entomology

lab for their kindness and support.

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TABLES AND FIGURES

Table 1. All syrphid species records of males (M) and females (F) for Akimiski Island,

Species Count Collection method Years M F

Anasimyia anausis* 3 Malaise 2014, 2017

Chalcosyrphus vecors 1 sticky trap 2008 Cheilosia latrans 64 39 Malaise, Nzi, net 2010, 2012-17 Cheilosia orilliaensis* 3 Nzi, net 2010, 2016

Cheilosia rita* 1 Nzi 2016

Cheilosia shannoni* 1 Malaise 2016 Chrysotoxum derivatum* 6 1 Malaise, Nzi 20012, 2014, 2016-17 Chrysotoxum flavifrons* 2 Malaise 2017 Chrysotoxum plumeum* 7 13 Malaise, Nzi 20012, 2016-17 Dasysyrphus amalopis* 1 Malaise 2012

Dasysyrphus limatus* 1 Nzi 2016

Dasysyrphus venustus 1 Nzi 2016

Doros aequalis* 1 Malaise 2014

Epistrophe grossularie* 1 sticky trap 2008

Epistrophe nitidicollis* 1 Malaise 2013

A: <400 km A: Epistrophella emarginata* 1 Malaise 2009 Eristalis brousii 18 18 IPY, net 2008-10, 2013 Eristalis dimidiata* 10 14 IPY, Nzi, sticky trap, net 2008-11

Eristalis flavipes* 2 Malaise, Nzi 2011, 2014

Eupeodes curtus 1 Malaise 2016 Eupeodes flukei* 1 5 Malaise, Nzi 2015, 2016-17

Eupeodes luniger 1 Malaise 2017

Eupeodes perplexus* 1 Nzi 2015

Ferdinandea buccata* 3 Malaise 2017

Helophilus groenlandicus 7 IPY 2008

Helophilus hybridus* 2 Malaise, Nzi 2015-16

Helophilus lapponicus 2 Malaise 2014, 2017 Helophilus obscurus* 3 7 Malaise, net 2008, 2013-16

Lapposyrphus lapponicus* 28 25 IPY, Malaise, Nzi, sticky 2008, 2010, 2012-17 trap, net Melanostoma mellinum 32 40 Malaise, Nzi, net 2012-17

Meligramma guttata* 1 Malaise 2017 Meliscaeva cinctella* 4 11 Malaise, Nzi 2013-17

Parasyrphus genualis* 1 Malaise 2012 Parhelophilus porcus* 1 1 Malaise, Nzi 2013, 2015

Platycheirus albimanus* 7 Malaise, Nzi 2010-12, 2016-17 Platycheirus amplus* 2 1 Malaise, Nzi 2015-16

Platycheirus clypeatus 1 Nzi 2016 Platycheirus granditarsis* 1 5 Malaise, Nzi, pitfall, net 2010-12, 2015-16

Platycheirus hyperboreus 1 Malaise 2014 Platycheirus inversus* 1 1 Malaise 2014-15

Platycheirus naso* 3 Malaise, Nzi 2012, 2015-16

Platycheirus obscurus 2 Malaise 2013, 2015 Platycheirus pictipes* 2 15 Malaise, Nzi 2012, 2015-17

Platycheirus podagratus* 2 Malaise 2014, 2016

Platycheirus varipes* 4 Malaise, Nzi 2012, 2014, 2016-17

Polydontomyia curvipes* 1 Malaise 2013

Scaeva affinis* 1 Malaise+D175 2016

Sphaerophoria 2 Nzi, net 2010, 2015 abbreviata* Sphaerophoria philanthus 15 Malaise, Nzi, net 2008, 2010, 2012, 2014- 17 Syrphus attenuatus* 1 Malaise 2014

Syrphus ribesii 11 Malaise, Nzi 2010, 2012-16 Syrphus vitripennis 1 1 Malaise 2015

Toxomerus marginatus* 1 IPY 2008

Volucella facialis* 1 Nzi 2017 Cheilosia lasiophthalmus* 1 IPY 2009 Eupeodes confertus* 1 2 Malaise, Nzi 2013, 2016-17

Megasyrphus laxus* 1 Nzi 2010

Meligramma 1 Malaise 2016 triangulifera*

800 km - Orthonevra robusta* 17 24 IPY, Malaise 2008-09, 2016-17

Paragus haemorrhous* 1 Malaise 2015

B: 400 Pipiza quadrimaculata* 1 Malaise 2014

Platycheirus luteipennis* 2 Nzi 2017

Syrphus sexmaculatus* 2 Malaise 2014, 2016

Cheilosia laevis 1 Malaise 2016 Eristalis hirta 4 net 2010, 2013 Eumerus strigatus* 1 net 2009 Neocnemodon elongata* 1 Malaise 2016

C: > 800 km C: > 800

Pipiza atrata* 1 Malaise 2017 Platycheirus jaerensis* 1 IPY 2009 Platycheirus kelloggi* 16 Malaise, Nzi 2009-10, 2012, 2016-17 Platycheirus latitarsis* 1 Nzi 2016 Platycheirus 3 Nzi 2017 neoperpallidus* Platycheirus nielseni 1 Malaise 2017

Table 2: Syrphid individuals collected by each trap type for each year of sampling on

Akimiski Island, Nunavut, Canada. Blanks indicate the years when traps were not deployed.

Year Malaise NZI Sticky Netted IPY trap trap trap 2008 1 12 9 77 2009 1 1 8 2010 23 17 2011 3 2 2012 22 13 0 2013 24 2 7 2014 70 11 0 2015 29 15 0 2016 38 55 10 2017 109 39 0 Total 292 163 12 46 85

Appendix Table 1: Total syrphid catch from Malaise and Nzi traps, 2012–2017, Akimiski

Island, Nunavut. Blanks represent zeros.

Species Malaise Nzi Total Anasimyia anausis 3 3 Cheilosia laevis 1 1 Cheilosia latrans 64 30 94 Cheilosia orilliensis 2 2 Cheilosia rita 1 1 Cheilosia shannoni 1 1 Chrysotoxum derivatum 5 2 7 Chrysotoxum flavifrons 2 2 Chrysotoxum plumeum 17 3 20 Dasysyrphus amalopis 1 1 Dasysyrphus limatus 1 1 Dasysyrphus venustus 1 1 Doros aequalis 1 1 Epistrophe nitidicollis 1 1 Epistrophella emarginata 1 1 Eristalis flavipes 1 1 Eupeodes confertus 2 1 3 Eupeodes curtus 1 1 Eupeodes flukei 3 3 6 Eupeodes luniger 1 1 Eupeodes perplexus 1 1 Ferdinandea buccata 3 3 Helophilus hybridus 1 1 2 Helophilus lapponicus 2 2 Helophilus obscurus 5 5 Lapposyrphus lapponicus 17 17 34 Melanostoma mellinum 56 14 70 Meligramma guttata 1 1 Meligramma triangulifera 1 1 Meliscaeva cinctella 10 5 15 Neocnemodon elongata 1 1 Orthonevra robusta 9 9 Paragus haemorrhous 1 1 Parasyrphus genualis 1 1 Parhelophilus porcus 1 1 2 Pipiza atrata 1 1 Pipiza quadrimaculata 1 1

Platycheirus albimanus 4 1 5 Platycheirus amplus 1 2 3 Platycheirus clypeatus 1 1 Platycheirus granditarsis 1 2 3 Platycheirus hyperboreus 1 1 Platycheirus inversus 2 2 Platycheirus kelloggi 3 10 13 Platycheirus latitarsis 1 1 Platycheirus luteipennis 2 2 Platycheirus naso 2 1 3 Platycheirus neoperpallidus 3 3 Platycheirus nielseni 1 1 Platycheirus obscurus 2 2 Platycheirus pictipes 7 10 17 Platycheirus podagratus 2 2 Platycheirus varipes 1 3 4 Polydontomyia curvipes 1 1 Scaeva affinis 1 1 Sphaerophoria abbreviata 1 1 Sphaerophoria philanthus 12 1 13 Syrphus attenuatus 1 1 Syrphus ribesii 9 9 Syrphus sexmaculatus 2 2 Syrphus vitripennis 2 2 Volucella fascialis 1 1

collection site

bird sanctuary

Akimiski Island

Figure 1: Map of the study site on Akimiski Island, Nunavut.

Figure 2: Malaise trap (left) and cloth Nzi trap (right) deployed on Akimiski Island,

Nunavut, 2017.

Malaise traps 50

20 Nzi traps Numberof species

0 0 50 100 150 200 250 300 Specimens

Figure 3: Rarefaction curve of Malaise and Nzi syrphid sampling data from 2012–2017 on Akimiski Island, Nunavut. The bars represent standard deviation.

Chapter 2: Distribution of Syrphidae (Diptera) across the far north of Ontario

INTRODUCTION

Ontario is one of the most populated provinces in Canada and is the third largest jurisdiction in Canada overall. Despite this, the far north of Ontario, almost 42% of

Ontario’s land mass, an area larger than Germany, is largely inaccessible and unpopulated. It has low human population density and very few roads (MNRF, 2014); access during the warmer months often is limited to airplanes or helicopters. At approximately 441,000 km2, it is globally one of the largest intact ecosystems in the world (MNRF 2015). It is an interesting area of study because it is still relatively untouched by anthropogenic changes in comparison to the extensive urban and agricultural development in the rest of the province. The area is made up of two ecozones: the Hudson Bay Lowlands and the Ontario Shield (Crins et al., 2009). The

Hudson Bay Lowlands ecozone is a coastal region adjacent to Hudson and James Bays, influenced by salt water tides, and makes up the core of the third largest wetland in the world (Crins et al., 2009, MNRF, 2015). The far north portion of the Ontario Shield ecozone is made up of boreal forest interspersed by many wetlands (Crins et al., 2009).

The far north of Ontario is rich in resources, and there are currently plans drafted for future development in the area. Resource extraction under consideration includes, but is not limited to mining, forestry, oil, aggregates, peat extraction, trapping, and fishing

(MNRF 2015). While economic development would likely aid local communities, it would also have fundamental impacts on the environment, and the species in it.

Syrphids (Diptera: Syrphidae) are a diverse and widely distributed family of flies, known for their Batesian mimicry of Hymenoptera, and for their pollination services

(Howarth and Edmunds, 2000). Adults sport striking colours, and frequent flowers, pollinating them in their search for nectar and pollen. Their larvae have highly diverse life history traits, and can be zoophagous, saprophagous, phytophagous, or mycophagous

(Sommaggio, 1999). Often, the distribution of syrphid species is dictated by larval habitat, especially for low-dispersing specialist species (Schweiger et al., 2007; Aguirre-

Gutiérrez et al., 2016).

Our knowledge of Syrphidae species’ distribution is becoming more well-known across the more accessible parts of North America, with extensive collections housed in institutions such as the Canadian National Collections of Insects, Arachnids and

Nematodes. Nevertheless, vast regions are still unstudied due to a lack of sampling from the difficulty of access, in spite of the ecological importance of these pollinators

(Vezsenyi et al., 2019). One such region is Ontario’s far north. Insect collections from the far north of Ontario tend to be from larger communities such as Moosonee (Freeman and

Twinn, 1954; Huckett, 1965; Cordero et al. 2017) and Fort Severn (Shorthouse et al.,

2003; Jones et al., 2014) as they are more easily accessed by plane or rail. Until recently, collections from the interior and away from the communities were few to nonexistent.

The most recent studies have been based on collections made in the interior of the far north, through the Far North Biodiversity Project, by the Ontario Ministry of Natural

Resources and Forestry. Of these, the dipteran studies were conducted by Ringrose et al.

(2013), and Ringrose et al. (2014) who examined tabanids and mosquitoes from the area.

Beetle studies were carried out by Jumean et al. (2017), Degasparro et al. (2018),

Fleming et al. (2019), and Ringrose et al. (2019) who studied tiger beetles, dytiscids,

Elaphrus beetles, and burying beetles, respectively. The only hymenopteran study so far from this project has been that of Gibson et al. (2018), looking at bumblebees. Non-insect invertebrate studies from this collection project include a mollusc paper by Forsyth and

Oldham (2016), and a leech paper by Langer et al. (2018).

Syrphids have potential as bioindicators, particularly for changes to forests

(Sommaggio, 1999; Deans et al., 2007; Sommaggio and Burgio, 2014). Shifts in species ranges or interspecific ratios could be indicative of environmental changes (Danks,

1992). To understand how a changing landscape affects the species in an area, we need baseline distributional data. Kearns (2001) highlights the particular importance of this for dipteran pollinator populations, as baseline data of abundances and species compositions are needed in order to understand the impacts that anthropogenic changes can have on these populations.

In this paper we present the first extensive inventory of syrphids across the vast expanse of the far north of Ontario, with nearly a decade’s worth of collected specimens, and summarize the most significant discoveries from this area.

METHODS AND MATERIALS

Collection: Specimens were gathered from three different projects conducted in the far north of Ontario. All projects were general surveys, and collections were non- specific to syrphids. Dates, locations, and collecting methods were different between each project, and are as follows:

The Far North Biodiversity Project (FNBP) was conducted by the OMNRF to identify and catalogue biodiversity in the far north of Ontario. This project covered more than 400 sites over six years (see: Ringrose et al., 2013; Ontario Biodiversity Council,

2015). Starting in 2009, a number of areas were sampled across Ontario’s far north. From

2010-2014 sample sites were randomly selected within a 150 km radius of a first nations’ community each year: Webequie in 2010 and 2013, Big Trout Lake and Sandy Lake in

2011, Fort Albany in 2012, and Fort Severn in 2014. Ringrose et al. (2013) details the site selection process, as well as site descriptions from 2011 and 2012. Sites were sampled intensively for one week. Insect sampling methods included Malaise traps, Nzi traps, bottle traps, pan traps, vegetation sweeps, net sweeps, pitfall traps, and aquatic sampling. All specimens were stored in 70% ethanol. Simultaneous to this, additional surveys were conducted by ad hoc hand netting, using helicopters to access remote areas.

These were part of a larger survey of plants and animals.

Sampling was also conducted in Moosonee, Ontario, 19-28 July, 2015. Two

Malaise traps (Lightweight Malaise Trap, Townes Style, 176 cm high, 165 cm long, model no. 2868, BioQuip Products, Rancho Dominguez, CA) were set up with non-toxic antifreeze and were emptied daily. Coloured pan traps (yellow (RAL 1026), light blue

(RAL 5012), dark blue (RAL 5017), white (RAL 9010), and pink (RAL 4003)) were also deployed using soapy water, and were emptied every other day. Pan traps were set up in similar fashion to those from the FNBP, and used three similar colours (yellow, light blue, and white). They were put out in an ‘x’ formation, randomly with either all bowls of the same colour, or a mix of colours. All specimens were placed in 70% ethanol after collection.

A single Malaise trap (Lightweight Malaise Trap, Townes Style, 176 cm high,

165 cm long, model no. 2868, BioQuip Products, Rancho Dominguez, CA) was deployed at the MNRF research station in Polar Bear Provincial Park near Burnt Point, Ontario

(Beresford, 2011; by Kim Bennett) 18 June to 30 June, 2016. Specimens were placed into

70% ethanol.

Processing: Syrphids were removed from all collected samples. Excluding previously pinned individuals, all specimens were then critically point dried using a Leica critical point dryer (Leica EM CPD300, Leica Microsystems Inc., Concord, ON), and then pointed onto pins. Identifications were conducted using Skevington et al. (2019), supplemented by keys from Vockeroth (1992), Miranda et al. (2013), and Young et al.

(2016).

Voucher specimens are currently housed at Trent University, with plans to move them to the Canadian National Collection of Insects, Arachnids and Nematodes for curation and long term storage.

Analysis: Range records: Range records were determined using Skevington et al.

(2019), using the same methods as Vezsenyi et al. (2019).

Specimens were divided into three categories based on their distance to the nearest collected specimen of that species: near (<400 km), intermediate (400-800 km), and far (>800 km). Though these distances are arbitrary, they were chosen based on factors such as plant hardiness zones and ecozones. Near records pass through few plant hardiness zones; intermediate records pass through a number of plant hardiness zones but tend to stay only in the two ecozones found within northern Ontario, while far records

pass through numerous plant hardiness zones, as well as different ecozones from those of the far north of Ontario.

Records were also classified as the following: gap infill, edge of range, and range extension. Gap infills were defined as records which fall between multiple points within the species’ range, regardless of the distance between these points. Edge of range records were records found on the edge of the species’ range, but do not extend the overall range in any particular direction. Range extensions were records that extend the overall distribution in a particular direction.

Range maps were produced using ArcMap 10.6.1 (ESRI, 2011). Previous ranges were approximated using Skevington et al. (2019).

Richness estimates: We used Preston’s lognormal as well as Chao 1 (Hammer et al., 2001) to estimate how many remaining species could be found in the far north of

Ontario (Preston 1948; Ludwig and Reynolds 1988; Gotelli and Colwell 2011). Chao 2 and Jackknife 1 and 2 could not be used, as we did not have consistent presence/absence data for each location sampled.

Larval life history traits: We used work by Vockeroth and Thompson (1987),

Sommaggio (1999), and Skevington et al. (2019) to categorize specimens collected into mycophagous, phytophagous, saprophagous, and zoophagous categories.

RESULTS

A total of 1514 individuals of 120 species were collected (Table 1). Of these species, six are new provincial records for Ontario (Table 2; Figure 3 A-F). The most abundant species caught were Melanostoma mellinum (n=124), Eristalis dimidiata

(n=98), and Syrphus vitripennis (n=88), which made up 8.2%, 6.5%, and 5.8% of the total catch, respectively. Species with only a single individual made up 30.8% of all species found (n=36); 66.7% of species had five or fewer individuals caught. We were unable to identify 219 individuals to species level using current keys, mainly because females of several genera are not yet distinguishable (e.g., Eupeodes, Sphaerophoria, some species of Platycheirus). In terms of life history traits, four species were phytophagous/mycophagous (n=34 individuals), 47 species were saprophagous (n=384 individuals), and 69 species were zoophagous (n=877 individuals).

Using Preston’s lognormal, it was estimated that there are at least 17 more species to be found (n= 137), while Chao 1 estimates a further 53 species (n= 173).

The largest proportion of syrphids (69.2%) were caught with Malaise traps, followed by net sweeps (21.3%), Nzi traps (3.7%), and vegetation sweeps (2.91%); all other trap types accounted for 2.9% of individual catches (Table 3).

Five syrphid species were found a significant distance away (>800 km) from their nearest known record; one of these was a range extension (Figure 2C), the other four were gap infills (Table 1). Thirty-one species were found an intermediate distance away

(400-800 km): three range extensions (Figure 2A-B, D), three edge of range records, and

25 gap infills (Table 1). Eighty-four species were found relatively close to their nearest record (<400 km): 11 edge of range records, and 73 gap infills (Appendix).

Species richness was compared to studies by Deans et al. (2007) and Proctor et al.

(2012) (Table 4).

DISCUSSION

This study provides the first extensive look at the Syrphidae residing in the far north of Ontario. We have filled in distributional gaps for many different species and extended the ranges of several species. With 120 species collected from the Far North of

Ontario, we collected over one-third of all syrphid species known provincially. The total number of species found in the far north of Ontario is likely closer to at least half of the species richness of the provincial total, given our estimates using Preston’s lognormal (n=

137) and Chao 1 (n= 173). Species were almost certainly missed given that we collected only during the summer months, which would have missed early and late season emergent species. More highly specialized species, such as those of the subfamily

Microdontinae, were also likely missed due to a lack of targeted collecting. The highest proportion of individuals caught were zoophagous (58%) which tend to be more generalists, feeding on soft-bodied insects, while saprophagous species accounted for the next highest proportion of individuals (25%). These proportions, in terms of life history traits, seem to be in line with other “nearby” studies with large collections of syrphids

(Deans et al., 2007; Proctor et al., 2012).

The study by Deans et al. (2007) took place in the Lake Abitibi Model Forest, which is just south of the far north boundary line in Ontario; they found richness results similar to ours, with 105 species of 3,209 individuals. Proctor et al. (2012) sampled syrphids in Algonquin Provincial Park, and found 141 species of 7,992 individuals.

Though these studies had vastly different collection efforts and sampling areas than ours, the number and identity of species were comparable. Between the Far North of Ontario,

Lake Abitibi Model Forest, and Algonquin Park, 60 species were shared across all three

locations (Table 4). The far north also shared nine species with the Abitibi site, and shared 13 species with the Algonquin site. Our collections also included 38 species that were not seen at either of these locations. Another nearby collection is that of Beresford

(2011), who collected insects in Polar Bear Provincial Park in the far north for a week during the summer of 2009, and caught five species, three of which were also found in our collection. As with any taxa, common species can be found across large areas, while varying habitats between sites result in differing arrays of species. Though we had lower sample effort per site, with fewer traps set over a shorter timeframe, we had the advantage of sampling over a greater number of sites set across a huge expanse of area.

It is unsurprising that Malaise traps collected the most individuals, as they are well known as one of the better tools for collecting syrphids (Skevington et al., 2019). On the other hand, our pan traps yielded few syrphids; only four individuals in the three years that FNBP used these traps (2012-2014), and another 19 from the 2015 Moosonee collection. This seems to be consistent with other studies, such as Cane et al. (2000) and

Proctor et al. (2012), who also had poor yields using these traps. Though many think of pan traps as an essential tool for collecting syrphids, in the case of faunistic studies it is best to use a wide array of sampling techniques in attempts to maximize the species found. This is true for syrphids, as there is no targeted trap to catch them.

Most of the species collected more than 800 km away from their nearest record were gap infills, and as such could be expected from the area. Of these, Anasimyia bilinearis was the most common, as it has evenly distributed populations from east to west in both Canada and the United States. Eupeodes volucris is very common in western

Canada, and less commonly extends to the east coast. Conversely, Platycheirus

quadratus and Sphegina brachygaster are commonly found in the east, with more sparse populations extending to the west. The only range extension of this group was

Platycheirus orarius (Figure 2E), whose population is very strictly on the east coast; the specimen collected nearest to ours was from eastern Quebec, near New Brunswick. This jump from the Atlantic Maritime ecozone to the Hudson Plains ecozone was unexpected, though the Hudson Plains ecozone does provide salt marshes and tidal flats from which this species is known (Skevington et al., 2019).

All other range extensions were between 400 km and 800 km from their nearest collected specimen. These include Eristalis stipator (Figure 2A), which is found quite commonly in western North America, with populations extending east to a lesser extent.

Our record of this species represents the northern-most point from which this species has been collected. Helophilus bottnicus (Figure 2B) is another species more commonly found in north-west North America, ranging around northern Yukon and Northwest

Territories; our specimen is the eastern-most point that this species has been collected in

North America. The closest specimen to ours was also from the Hudson Plains ecozone, in Churchill, Manitoba. Lastly, there is Platycheirus thompsoni (Figure 2D), whose range is quite similar to that of Platycheirus orarius being strictly found eastern, with the exception of one record from northern Minnesota. Our collected specimen represents the northern-most point for this species; it extends the overall distribution to the northeast.

Ecologically, these range extensions were not unexpected as this species is commonly associated with marshes and bogs (Skevington et al., 2019).

As northern Ontario has been vastly under-sampled, it was expected that there would be a large number of gap infill records and edge of range records. Among these

were several new Ontario provincial records. Helophilus bottnicus (Figure 3A) and

Platycheirus orarius (Figure 3E) were already discussed as range extensions. Eupeodes curtus (Figure 3F), Melangyna labiatarum (Figure 3B), Orthonevra robusta (Figure 3D), and Platycheirus pictipes (Figure 3C) are all new provincial records, and gap infills for their overall distributions. Each of these has been collected in surrounding areas, such as

Churchill (MB), Quebec, Akimiski Island (NU), and southern Nunavut. Orthonevra robusta was the only one of these not found in Quebec.

With increased pressure for development, changes to the landscape and overall habitat will likely be reflected in these syrphid communities. We have provided a solid baseline of the syrphids found in the far north of Ontario. Despite the practical challenges of sampling in the North, repeated sampling is crucial. It remains the basis for mapping species ranges, describing biological communities, and assessing biological change.

ACKNOWLEDGMENTS

We would like to thank Ontario Ministry of Natural Resources and Forestry for resources and funding; Dean Phoenix, John Ringrose, Dave Etheridge, Ken Abraham,

Glenn Brown, Rod Brook, Kim Bennett, and all other field volunteers for all their hard work collecting insects and data from these remote regions.

Thank you to all those at the Canadian National Collections of Insects, Arachnids and Nematodes, for allowing me to work in their facilities, and passing on their syrphid expertise; particularly Jeff Skevington, Andrew Young, Michelle Locke, Kevin Moran, and Victoria Nowell.

Thanks to those in the Beresford lab, Sarah Langer, Kaitlyn Fleming, Kayla

Vizza, Ayden Ricker-held for all their hard work sorting samples, and support.

Finally, thank you to my supervisors Dave Beresford and James Schaefer, as well as my committee members Jeff Skevington and Bill Crins for all of their edits and work on my thesis.

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TABLES AND FIGURES

Table 1: All species of Syrphidae collected from the far north of Ontario, 2009-2014, with male (M) and female (F) counts, as well as traps they were collected in. Species with

Species Count Trap Years Record M F type

Anasimyia anausis 3 2 Malaise 2010-2012

Anasimyia bilinearis 13 Malaise, Nzi 2012 inf (>800) Baccha cognata 1 Malaise 2011 inf

Blera confusa 1 net 2011

Blera nigra 2 1 Malaise, Nzi 2011, 2013

Brachyopa notata 1 Malaise 2010 inf

Chalcosyrphus anomalus 1 Malaise 2012 inf

Chalcosyrphus inarmatus 2 1 Malaise 2011, 2013

Chalcosyrphus libo 1 Malaise 2013

Chalcosyrphus nemorum 7 Malaise 2010-2012, 2015 Cheilosia hunteri 2 2 Malaise, net, Nzi 2012

Cheilosia latrans 12 11 Malaise, net, Nzi 2010-2013, 2015 Cheilosia rita 6 Malaise 2011-2012

Cheilosia shannoni 1 net 2011

Chrysotoxum derivatum 3 Malaise, net 2009-2010

Chrysotoxum flavifrons 11 24 Malaise, net 2009-2013

Chrysotoxum plumeum 1 2 Malaise, net, Nzi 2011-2012, 2014 Dasysyrphus venustus 2 net, Nzi 2011, 2013

Epistrophe nitidicollis 1 2 Malaise, net, Nzi 2011-2012, 2015 Epistrophe terminalis 2 2 Malaise 2011, 2013, 2016 Epistrophella emarginata 1 net 2009

Eristalis cryptarum 1 Malaise 2011 inf

Eristalis dimidiata 5 93 Malaise, net, Nzi 2011-2014, 2016 Eristalis flavipes 2 9 Malaise, net 2009-2013

Eristalis obscura 1 net 2009

Eristalis stipator 1 Malaise 2016 r.e (N)

Eristalis tenax 1 pan trap 2015

Eristalis transversa 4 Malaise, net 2012-2013

Eupeodes americanus 76 Malaise, net, Nzi, veg sweep 2010, 2012- 2013 Eupeodes confertus 3 Malaise, Nzi 2011, 2013 edg (W)

Eupeodes curtus* 1 Malaise 2010

Eupeodes flukei 1 1 Malaise 2016

Eupeodes latifasciatus 1 7 Malaise 2010-2013

Eupeodes luniger 8 13 Malaise, net, veg sweep 2009-2010, 2013 Eupeodes perplexus 6 Malaise, net, Nzi 2009-2013

Eupeodes pomus 3 Malaise 2010-2011 inf

Eupeodes volucris 1 Malaise 2012 inf (>800) Ferdinandea buccata 1 1 Malaise 2010, 2013

Hammerschmidtia rufa 1 Malaise 2012

Helophilus bottnicus* 1 Malaise 2011 r.e (E)

Helophilus fasciatus 2 2 Malaise, net 2012

Helophilus lapponicus 5 Malaise, net 2011-2013

Helophilus obscurus 3 5 Malaise, net, pan trap 2011, 2015

Lapposyrphus lapponicus 35 18 bottle trap, Malaise, mosquito 2010-2016 trap, net, Nzi Melangyna fisherii 2 Malaise 2010 inf

Melangyna labiatarum* 1 Malaise 2010 inf

Melangyna lasiophthalma 2 Malaise 2011, 2013

Melangyna umbellatarum 1 Malaise 2015

Melanostoma mellinum 24 10 aquatic net, Malaise, net, Nzi, 2009-2016 0 veg sweep Meligramma guttata 1 1 Malaise 2010

Meligramma triangulifera 2 Malaise 2011 inf

Meliscaeva cinctella 1 3 Malaise, net 2010-2012

Microdon tristis 2 Malaise, Nzi 2011, 2013 inf

Neoascia metallica 3 net, pitfall 2009

Neoascia sandsii/ 1 net 2009 undescribed species 1

Neoascia tenur 9 Malaise, pan trap, veg sweep 2011-2012, inf 2014 Neocnemodon coxalis 1 Malaise 2010 inf

Ocyptamus fascipennis 1 net 2013

Orthonevra pulchella 2 12 Malaise, net 2012-2013

Orthonevra robusta* 1 veg sweep 2014 inf

Parasyrphus genualis 6 Malaise 2009-2011

Parasyrphus nigritarsis 1 6 Malaise, Nzi 2010-2012

Parasyrphus vockerothi 3 Malaise 2011

Parhelophilus porcus 4 8 Malaise, net, Nzi 2010-2011, 2014-2015 Parhelophilus rex 10 4 Malaise 2010-2012 inf

Pipiza macrofemoralis 1 Nzi 2012 inf

Platycheirus albimanus 4 Malaise, net 2010-2011

Platycheirus amplus 1 3 Malaise, pan trap 2010, 2015

Platycheirus clypeatus 1 1 Malaise, net 2010, 2014

Platycheirus confusus 1 Malaise 2010 inf

Platycheirus granditarsis 13 22 Malaise, net, Nzi, veg sweep 2009-2012, 2015 Platycheirus hyperboreus 6 14 Malaise, net 2009-2012, 2015 Platycheirus immarginatus 6 12 aquatic net, Malaise, net 2009-2011 Platycheirus inversus 1 31 Malaise, veg sweep 2010-2011, edg (W) 2013, 2015 Platycheirus modestus 7 1 Malaise, mosquito trap 2010 inf

Platycheirus naso 3 6 Malaise, net, veg sweep 2010-2011, 2014-2015 Platycheirus nearcticus 4 6 Malaise, net 2010-2012, 2015 Platycheirus nodosus 3 3 Malaise, net 2010-2012 inf

Platycheirus obscurus 3 3 Malaise, net 2009-2011

Platycheirus orarius* 1 net 2009 r.e (W) (>800) Platycheirus pictipes* 1 Malaise 2010

Platycheirus podagratus 1 2 Malaise 2010, 2014

Platycheirus quadratus 1 Malaise 2013 inf (>800) Platycheirus rosarum 3 40 Malaise, net, Nzi, veg sweep 2010-2013, 2015 Platycheirus scambus 4 6 Malaise, net 2009-2012, 2015 Platycheirus scutatus 1 Malaise 2010

Platycheirus thompsoni 1 Malaise 2011 r.e (NE)

Platycheirus thylax 1 Malaise 2011 inf

Platycheirus varipes 1 Malaise 2014

Polydontomyia curvipes 1 pan trap 2012

Scaeva affinis 1 Malaise 2016

Sericomyia lata 1 Malaise 2011 inf

Sericomyia militaris 2 8 Malaise, net 2011

Sericomyia sexfasciata 1 2 Malaise, net 2012-2014

Sericomyia transversa 4 7 Malaise, net, Nzi 2011, 2013

Sphaerophoria abbreviata 12 Malaise, net 2010-2012, 2014-2015 Sphaerophoria 2 Malaise, net 2010, 2015 asymmetrica Sphaerophoria 1 2 Malaise 2011, 2015 inf novaeangliae Sphaerophoria philanthus 19 Malaise, net, Nzi 2009-2010, 2012, 2015 Sphegina brachygaster 1 Nzi 2013 inf (>800) Sphegina flavimana 1 Malaise 2012 edg (N)

Sphegina rufiventris 1 1 net 2009 Syrphus attenuatus 4 1 Malaise, net 2011, 2014 inf Syrphus rectus 14 10 Malaise, net 2010, 2012, inf 2015 Syrphus ribesii 21 41 bottle trap, Malaise, net 2009-2015

Syrphus torvus 5 4 Malaise, net 2009-2013, 2015 Syrphus vitripennis 10 78 Malaise, net 2009-2012, 2014, 2016 Temnostoma excentrica 2 1 Malaise 2011

Temnostoma obscurum 3 Malaise, pan trap 2011-2012, 2015 Toxomerus geminatus 1 3 Malaise, net 2010-2012

Toxomerus marginatus 14 24 aquatic net, Malaise, net, Nzi, 2010-2013, pan trap, veg sweep 2015 Volucella facialis 2 5 net, Nzi, pan trap, veg sweep 2010, 2012- 2013, 2015 Xylota annulifera 3 12 Malaise, Nzi 2010-2013, 2015 Xylota confusa 3 Malaise, pan trap 2011, 2015

Xylota flavifrons 1 veg sweep 2014 Xylota flavitibia 1 3 Malaise 2010, 2012 inf

Xylota flukei 1 5 Malaise, pan trap 2013-2014 Xylota hinei 1 7 Malaise, pan trap, pitfall 2009-2013, inf 2015 Xylota quadrimaculata 18 60 Malaise, net, Nzi, pan trap, veg 2010-2014 inf sweep Xylota subfasciata 2 4 Malaise 2009-2012, 2014

Table 2: New provincial records of Syrphidae, as well as the dates, sex (male (M) and female (F)), trap type, and locations.

Species Sex Trap Date Lat Long Eupeodes curtus F Malaise trap 25 July, 2010 52.73655 -86.0824 Helophilus bottnicus F Malaise trap 13 July, 2011 53.75296 -88.9192 Melangyna labiatarum F Malaise trap 2 July, 2010 54.91296 -85.4801 Orthonevra robusta M low vegetation sweep 27 July, 2014 55.48618 -87.8807 Platycheirus orarius F incidental 5 July, 2009 51.40363 -79.6782 Platycheirus pictipes M Malaise trap 3 July, 2010 54.44208 -84.7681

Table 3: Number of syrphids caught in each trap type per year. Year Aquati Bottle Incidental Low Malais Mosquito NZI Pan Pitfall c net trap veg e trap trap trap trap swee p 2009 36 45 2

2010 4 67 11 387 3 7

2011 1 89 6 236 1

2012 1 91 5 190 28 2

2013 1 37 4 84 25 1

2014 2 18 43 3 1

2015 52 19 1

2016 11 Total 5 2 322 44 1048 3 56 23 11

Table 4: Number of Syrphidae species shared between three projects: the far north of

Ontario, Abitibi Model Forest (Dean et al., 2007), and Algonquin Park (Proctor et al.,

2011).

Abitibi Model Far North All Not Forest Ontario locations shared Algonquin Park 16 13 60 48 Abitibi Model 9 60 16 Forest Far North Ontario 60 38

Figure 1: All sites in the far north of Ontario from which syrphids were collected 2009-

2015 (black circles) and FNBP sites sampled from which no syrphids were collected

(white circles).

A B

C D

Figure 2 A-D: Species maps of individuals collected from the far north of Ontario that

extended their overall range distributions: (A) Eristalis stipator, (B) Helophilus bottnicus,

collected specimen in black.

Figure 3 A-F: New Ontario species records of Syrphidae, collected in the far north of

Ontario. (A) Helophilus bottnicus, (B) Melangyna labiatarum, (C) Platycheirus pictipes,

(D) Orthonevra robusta, (E) Platycheirus orarius, (F) Eupeodes curtus. Photos done by

Kaitlyn Fleming and KV.

APPENDIX

Appendix Table 1: All syrphid species collected from the far north of Ontario, categorized by the distance to their nearest collected record of that species (<400 km,

400-800 km, >800 km), and whether these new records are a gap infill, edge of range, or range extension.

Species Distance to nearest species record (km) Record type Anasimyia anausis <400 km gap infill Blera confusa <400 km edge of range (north) Blera nigra <400 km gap infill Chalcosyrphus inarmatus <400 km edge of range (north) Chalcosyrphus libo <400 km gap infill Chalcosyrphus nemorum <400 km gap infill Cheilosia hunteri <400 km gap infill Cheilosia latrans <400 km gap infill Cheilosia rita <400 km gap infill Cheilosia shannoni <400 km gap infill Chrysotoxum derivatum <400 km gap infill Chrysotoxum flavifrons <400 km gap infill Chrysotoxum plumeum <400 km gap infill Dasysyrphus venustus <400 km gap infill Epistrophe nitidicollis <400 km gap infill Epistrophe terminalis <400 km gap infill Epistrophella emarginata <400 km gap infill Eristalis dimidiata <400 km gap infill Eristalis flavipes <400 km gap infill Eristalis obscura <400 km gap infill Eristalis tenax <400 km gap infill Eristalis transversa <400 km edge of range (north) Eupeodes americanus <400 km edge of range (north) Eupeodes curtus <400 km edge of range (south) Eupeodes flukei <400 km gap infill Eupeodes latifasciatus <400 km gap infill Eupeodes luniger <400 km gap infill Eupeodes perplexus <400 km gap infill Ferdinandea buccata <400 km gap infill Hammerschmidtia rufa <400 km gap infill Helophilus fasciatus <400 km gap infill Helophilus lapponicus <400 km gap infill Helophilus obscurus <400 km edge of range (north) Lapposyrphus lapponicus <400 km gap infill Melangyna lasiophthalma <400 km gap infill Melangyna umbellatarum <400 km gap infill Melanostoma mellinum <400 km gap infill Meligramma guttata <400 km gap infill Meliscaeva cinctella <400 km gap infill Neoascia metallica <400 km gap infill Neoascia sandsii/ undescribed species 1 <400 km edge of range (north) Ocyptamus fascipennis <400 km gap infill Orthonevra pulchella <400 km gap infill Parasyrphus genualis <400 km gap infill Parasyrphus nigritarsis <400 km gap infill Parasyrphus vockerothi <400 km gap infill Parhelophilus porcus <400 km gap infill Platycheirus albimanus <400 km gap infill Platycheirus amplus <400 km gap infill Platycheirus clypeatus <400 km gap infill Platycheirus granditarsis <400 km gap infill Platycheirus hyperboreus <400 km gap infill Platycheirus immarginatus <400 km gap infill

Platycheirus naso <400 km gap infill Platycheirus nearcticus <400 km gap infill Platycheirus obscurus <400 km edge of range (north) Platycheirus pictipes <400 km gap infill Platycheirus podagratus <400 km gap infill Platycheirus rosarum <400 km gap infill Platycheirus scambus <400 km gap infill Platycheirus scutatus <400 km gap infill Platycheirus varipes <400 km gap infill Polydontomyia curvipes <400 km gap infill Scaeva affinis <400 km gap infill Sericomyia militaris <400 km gap infill Sericomyia sexfasciata <400 km gap infill Sericomyia transversa <400 km gap infill Sphaerophoria abbreviata <400 km gap infill Sphaerophoria asymmetrica <400 km gap infill Sphaerophoria philanthus <400 km gap infill Sphegina rufiventris <400 km edge of range (northwest) Syrphus ribesii <400 km gap infill Syrphus torvus <400 km gap infill Syrphus vitripennis <400 km gap infill Temnostoma excentrica <400 km gap infill Temnostoma obscurum <400 km gap infill Toxomerus geminatus <400 km gap infill Toxomerus marginatus <400 km gap infill Volucella facialis <400 km gap infill Xylota annulifera <400 km edge of range (northwest) Xylota confusa <400 km gap infill Xylota flavifrons <400 km gap infill Xylota flukei <400 km edge of range (west) Xylota subfasciata <400 km gap infill Baccha cognata 400-800 km gap infill Brachyopa notata 400-800 km gap infill Chalcosyrphus anomalus 400-800 km gap infill Eristalis cryptarum 400-800 km gap infill Eristalis stipator 400-800 km range extension (north) Eupeodes confertus 400-800 km edge of range (west) Eupeodes pomus 400-800 km gap infill Helophilus bottnicus 400-800 km range extension (east) Melangyna fisherii 400-800 km gap infill Melangyna labiatarum 400-800 km gap infill Meligramma triangulifera 400-800 km gap infill Microdon tristis 400-800 km gap infill Neoascia tenur 400-800 km gap infill Neocnemodon coxalis 400-800 km gap infill Orthonevra robusta 400-800 km gap infill Parhelophilus rex 400-800 km gap infill Pipiza macrofemoralis 400-800 km gap infill Platycheirus confusus 400-800 km gap infill Platycheirus inversus 400-800 km edge of range (west) Platycheirus modestus 400-800 km gap infill Platycheirus nodosus 400-800 km gap infill Platycheirus thompsoni 400-800 km range extension (northwest) Platycheirus thylax 400-800 km gap infill Sericomyia lata 400-800 km gap infill Sphaerophoria novaeangliae 400-800 km gap infill Sphegina flavimana 400-800 km edge of range (north) Syrphus attenuatus 400-800 km gap infill Syrphus rectus 400-800 km gap infill Xylota flavitibia 400-800 km gap infill Xylota hinei 400-800 km gap infill Xylota quadrimaculata 400-800 km gap infill Anasimyia bilinearis >800 km gap infill Eupeodes volucris >800 km gap infill Platycheirus orarius >800 km range extension (west) Platycheirus quadratus >800 km gap infill Sphegina brachygaster >800 km gap infill

Chapter 3: Barcode Syrphidae data from the far north of Ontario, and Akimiski Island, Nunavut.

INTRODUCTION

Using dichotomous keys, many insects can be identified to species level; however, morphological identifications are not without drawbacks. Identifications require some degree of training (Jinbo et al., 2011) and are only as good as the keys available to the user, with many groups in desperate need of revision. Additionally, identifications become difficult, if not impossible, if the specimen has been damaged. With the cost of

DNA barcoding decreasing over time it is becoming a more common tool in the field of entomology. Though more expensive than morphological identifications, it can be faster when dealing with large collections, and may sometimes be used to identify specimens that cannot be determined morphologically. Within the family Syrphidae (Diptera) in particular, many genera have females that are indistinguishable morphologically, and require DNA barcoding to get to species level (Skevington et al., 2019).

Here, we used the DNA barcoding region Cytochrome c Oxidase I (COI) on several syrphid individuals from the far north of Ontario, and Akimiski Island, Nunavut.

This was done to identify females from genera where morphological identifications were not possible, and to confirm morphological identifications for individuals of rare species or of species whose populations are normally found significant distances away from our study area.

METHODS

For collection methods, see Vezsenyi et al. (in press) and chapter 2 of this thesis.

Morphological identifications were made using Skevington et al. (2019).

Seven individuals were selected for species identification, as they are from groups where morphological identifications of females are not possible. These individuals were morphologically identified to three genera: Neocnemodon (n=1), Anasimyia (n=5), and

Platycheirus (n=1). The remaining 10 individuals were selected to confirm identifications. Cheilosia hunteri (n=1) and Cheilosia latrans (n=2) were selected as there were some difficulties with their morphological identifications. Helophilus bottnicus (n=1) and Platycheirus thylax (n=1) are both rare species and were found more than 400 km from previously collected specimens within their species. Platycheirus kelloggi (n=1) and Platycheirus latitarsis (n=1) were chosen as their populations are normally restricted to the west of the Rocky Mountains, almost 3,000 km away.

Chyrysotoxum derivatum (n=1), and Orthonevra robusta (n=2) were selected due to their perceived rarity.

All specimens were critically point dried and pointed onto pins. Specimens were sent to the Canadian National Collections of Insects, Arachnids, and Nematodes (CNC) for sequencing by Scott Kelso. The right hind leg was removed from each specimen; smaller specimens had the left mid and hind legs removed for DNA extraction. The

Cytochrome c Oxidase I (COI) mitochondrial region was sequenced using the HebF

(LCO1490) x 780R primer set (Folmer et al., 1994), using protocols outlined in

Hajibabaei et al. (2005). Sequences were scored and aligned using Sequencher 5.4.6

(2018) and Mesquite (Maddison and Maddison, 2010) respectively. Sequences were then uploaded to the Barcode of Life Datasystems (BOLD) website and compared to other specimens in the BOLD database to confirm species identifications. We compared our specimens with others available through the BOLD database using a neighbor-joining

62 tree created through BOLD using the Kimura 2 Parameter model. Voucher specimens are currently housed in the Trent University entomology lab, with plans to move them to the

CNC for curation and long-term storage.

RESULTS

In total, we barcoded 17 individuals of 11 different species (Table 1). For females from groups wherein morphological identifications are not possible, we identified

Anasimyia anausis (n=5), and Neocnemodon coxalis (n=1). The female Platycheirus was only identifiable to P. quadratus/neoperpallidus/perpallidus, as females of these species cannot be separated morphologically or with COI barcodes.

Molecular data supported the morphological identifications of Cheilosia hunteri

(n=1), Cheilosia latrans (n=2), Helophilus bottnicus (n=1), and Platycheirus thylax

(n=1). BOLD systems had no Platycheirus latitarsis (n=1) sequences to compare with; thus our sequence is currently the only one available in the database for that species. Our putative morphologically identified specimen of Chrysotoxum derivatum was found to be a dark morph of the more common Chrysotoxum plumeum, in this case identifiable only using DNA (n=1). Using a nearest-neighbor phylogram, Platycheirus kelloggi (n=1) was found to have about a 1% difference in COI from others of its species found on the west coast of North America (Figure 1, Figure 2). This degree of genetic differentiation is typical of different species within the genus Platycheirus, suggesting that this is an undescribed species. Description of this putative new species awaits the discovery of a male specimen (males have distinctive secondary sexual characters and are typically used in species descriptions).

DISCUSSION

We found COI analysis of our specimens to be useful for obtaining and supporting identifications, although no more of a panacea than morphology. Many female syrphid species are difficult to impossible to tell apart using morphological identification methods. Therefore, it is important to confirm their identifications genetically. Using only males for confirmation of the presence of species within an area will miss potential species in which only females were caught that could not be morphologically identified, as was the case with the Neocnemodon coxalis and Platycheirus kelloggi females we sequenced.

Our sequences of rare species, such as those of Helophilus bottnicus and

Platycheirus thylax serve as surrogate vouchers and important data, as they better lend to our understanding of these individuals and serve as strong datapoints to compare others to in the future. This is especially true for the specimen of Platycheirus latitarsis that was sequenced, as it is the first sequence of this species in BOLD. Young et al. (2016) examined this species during their revision of Platycheirus but were unable to obtain any sequences for the species.

Platycheirus latitarsis and P. kelloggi were sequenced as they were collected almost 3,000 km away from their nearest known populations. We were unsure if they represent disjunct populations from their west coast distributions, or if they were extensions of these west coast populations that have yet to be collected from the areas in- between. Sequence data for P. latitarsis was inconclusive, as no other sequence for this species exists. Platycheirus kelloggi appears to be geographically isolated (Figure 1) and is genetically distinct, suggesting a disjunction.

Most closely related Platycheirus species are very similar genetically; some groups of species are invariant in their COI sequences, but are considered different species as they are morphologically different (Young et al., 2016). The Platycheirus kelloggi individual that was sequenced was found to be significantly different from the P. kelloggi sequenced from the west coast of North America (1% different). As this individual is both genetically and geographically isolated, it would suggest that it is a new undescribed species. Although it could be described on these bases alone, (see Jinbo et al., 2011; Young et al., 2016 for examples), we prefer to wait until a male is captured.

Males have distinctive secondary sexual structures on their legs and we hypothesize that this putative new species should differ from its putative western sister taxon. Females have silver pollinosity on the dorsal abdominal spots, contrasting with the bright orange dorsal abdominal spots of west coast P. kelloggi that largely lack pollinosity (Young et al., 2016). We currently have an additional 15 other females of this species that have not yet been sequenced. They were caught across multiple years on Akimiski Island (2009 n=1, 2010 n=2, 2012 n=1, 2016 n=7, 2017 n=5). We clearly need to do additional field work to try to find some males of this presumed new species.

In summary, genetic identification using COI has proven useful for my project.

We have identified species that would have otherwise been missed, supported identifications for some of our morphological identifications, sequenced Platycheirus latitarsis for the first time, and found a probable new species at this point only known from a remote island in James Bay.

ACKNOWLEDGEMENTS

I would like to thank Jeff Skevington for giving me the opportunity to barcode my

specimens and for teaching me about syrphid taxonomy; your work on this chapter was much needed, and greatly appreciated! I would also like to thank Scott Kelso for walking me through the DNA extraction process from start to finish, and for answering all of my

questions.

REFERENCES

Folmer, O., M. Black, W. Hoeh, R. Lutz, R. Vrijenhoek. 1994. DNA primers for

amplification of mitochondrial cytochrome c oxidase subunit I from diverse

metazoan invertebrates. Molecular Marine Biology and Biotechnology 3(5): 294-

299.

Hajibabaei, M., J.R. deWaard, N.V. Ivanova, S. Ratnasingham, R.T. Dooh, S.L. Kirk,

P.M. Mackie and P.D.N. Hebert. 2005. Critical factors for assembling a high

volume of DNA barcodes. Philosophical Transactions of the Royal Society B:

Biological Sciences 360(1462): 1959–1967.

Jinbo, U., T. Kato, M. Ito. 2011. Current progess in DNA barcoding and future

implications for entomology. Entomological Science 14: 107-124.

Maddison, W.P. and D.R. Maddison 2010. Mesquite: a modular system for evolutionary

analysis. Available from: https://www.mesquiteproject.org/.

Sequencer. 2018. Sequencher DNA sequence analysis software. Ann Arbor, MI, USA,

Gene Codes Corporation. Available from: https://www.genecodes.com/.

Skevington, J.H., M.M. Locke, A.D. Young, K. Moran, W.J. Crins, S.A. Marshall. 2019.

Field Guide to the Flower Flies (Hover Flies) of Northeastern North America.

Princeton University Press, Princeton, NJ 512 pp.

Vezsenyi, K.A., J.H. Skevington, K. Moran, A.D. Young, M.M. Locke, J.A. Schaefer,

D.V. Beresford. In press. Sampling Syrphidae using Malaise and Nzi traps on

Akimiski Island, Nunavut. Journal of the Entomological Society of Ontario

(accepted January 14, 2019).

Young, A.D., S.A. Marshall, J.H. Skevington. 2016. Revision of Platycheirus Lepeletier

and Serville (Diptera: Syrphidae) in the Nearctic north of Mexico. Zootaxa

4082(1): 1-317.

TABLES AND FIGURES

Figure 1: Phylogenetic tree using COI data featuring Platycheirus kelloggi, with the specimen previously identified as P. kelloggi outlined in black.

Table 1: Syrphidae specimens barcoded using COI. Data has been uploaded to the

Barcode of Life Datasystems (BOLD) website.

Species BOLD ID Sample ID Collection Date Location Anasimyia anausis SRCNC635-17 CNC640314 01-Jul-2013 Akimiski Anasimyia anausis SRCNC637-17 CNC640352 21-Jun-2011 Ontario Anasimyia anausis SRCNC638-17 CNC640356 01-Jul-2013 Akimiski Anasimyia anausis SRCNC639-17 CNC640369 17-Jun-2012 Ontario Anasimyia anausis SRCNC641-17 CNC640425 21-Jun-2011 Ontario Cheilosia latrans SRCNC632-17 CNC640238 15-Jul-2010 Ontario Cheilosia latrans SRCNC636-17 CNC640321 26-Jul-2016 Akimiski Cheilosia hunteri SRCNC642-17 CNC640433 10-Jun-2012 Ontario Chrysotoxum plumeum SRCNC628-17 CNC639825 20-Jul-2012 Akimiski Helophilus bottnicus SRCNC643-17 CNC640434 13-Jun-2011 Ontario Heringia coxalis SRCNC629-17 CNC639932 22-Jun-2010 Ontario Orthonevra robusta SRCNC631-17 CNC640232 27-Jun-2014 Ontario Orthonevra robusta SRCNC633-17 CNC640246 20-Jul-2016 Akimiski Platycheirus kelloggi SRCNC634-17 CNC640285 17-Jul-2016 Akimiski Platycheirus latitarsis SRCNC630-17 CNC640036 21-Jul-2016 Akimiski Platycheirus sp. SRCNC645-17 CNC640464 01-Jul-2010 Ontario Platycheirus thylax SRCNC640-17 CNC640408 11-Jun-2011 Ontario

Chapter 4: General Discussion

Insects are a precious and important part of this world, and deserve to be monitored and protected just as carefully as fish, mammals, and birds. Long-term monitoring programs are of the utmost importance, as they allow us to see impacts to species richness, diversity, and abundance. These types of studies allow us to detect and document declines in populations (Biesmeijer et al., 2006; Conrad et al., 2006; Shortall et al., 2009; Potts et al., 2010). This is especially apparent in a recent study by Hallmann et al. (2017) who found more than a 75% decline in flying insect biomass using their 27 year-long dataset of Malaise traps deployed in protected areas. Without these long-term studies, even if just observing declines in seasonal traps, there would be no quantitative basis to emphasize the exact changes occurring to these populations. Despite their importance, studies of this nature tend to be few, with noted changes in insect populations often coming through studies of insectivorous birds (Benton et al., 2002; Nebel et al.,

2010; Hallmann et al., 2014).

Here, we have provided novel Syrphidae species and distributional information for both Akimiski Island, Nunavut, as well as the far north of Ontario. We have over a decade’s worth of insect collections from Akimiski Island, with plans to continue these collections into the future. Unfortunately, sampling for the Far North Biodiversity Project stopped in 2014. The collections from this project can still be used for studying other taxa within this area, but by stopping the collection we are left with only a brief snapshot of populations from this period. This is a lost opportunity to discover how their populations have been changing and will change in the future. Nevertheless, while one can always make the argument that we need to collect more, the huge investment in the Far North

Biodiviersy Project gave us new insights that would never have been possible for this remote region, bringing into clearer focus the insects, and in particular, syrphids from this area.

It was interesting having the opportunity to work with these two datasets. While superficially very similar, having insects from largely inaccessible remote areas, they are the two extremes when it comes to insect collecting: a very long dataset from a single place, and very short datasets from a manifold of sites spanning a vast area. Neither is better than the other, and both were done to suit the needs of the projects and researchers.

Figure 1 illustrates the difference in species richness between the two collections. It is unsurprising that there is a higher richness in the far north of Ontario, as a vastly greater area was sampled, despite being sampled for a shorter time. Sites sampled in Ontario were also part of a contiguous land mass, while Akimiski Island, as an island, is less connected to other areas, even for flying insects. It is nearest to mainland Ontario via the

Akimiski Strait, which is about 20 km wide. One would expect that most, if not all species found on Akimiski would be the same as those found along the coast of northern

Ontario. However, 34% of the species collected on Akimiski Island were not found in any of the northern Ontario collections. This may be an artifact of collecting; maybe those species are also found in Ontario, but we were unable to catch them. A more plausible explanation is that this island has unique species due to some island-specific habitat characteristics, such as a moderated temperature. One hypothesis that was previously suggested (Ringrose 2014) was that insects being carried on weather systems would be deposited on the edge of James Bay (Taylor, 1974; Chapman et al., 2011). The prevailing wind direction in Canada is from west to east, meaning that Akimiski may

73 have accumulated a suite of species by this mechanism over the island's 4,000 year history.

Both the far north of Ontario and Akimiski Island were found to have a number of range extensions and new provincial/terreorial records. While the high number of new territory records for Nunavut is an artifact of political boundaries, the new records for

Akimski Island itself are biologically significant. Akimiski Island has a much longer growing season and more moderated temperatures than the nearest mainland area.

Though our trap analysis in Chapter 1 had found that the Nzi trap was not as effective at collecting syrphids as we had expected, it was still an interesting avenue to explore. It would be worthwhile to design an Nzi trap with colours more attractive to pollinators, such as combinations of white and yellow, and see how that impacts syrphid collection. The search for the perfect syrphid trap continues; Malaise traps and hand netting are the best options in the meantime. For broad insect surverys, a variety of traps should always be used to maximize catch potential for any taxa.

Finally, we found using genetic analysis for select individuals useful in acquiring and confirming species identifications, as well as searching for novel species. Collections from remote areas such as these are rare, so it is always ideal to sequence individuals such as these, given the opportunity.

My thesis was one of discovery. We have greatly expanded on the known syrphid distributions for the regions of the far north of Ontario, and Akimiski Island, as well as recorded dozens of new provincial/terriorial species records for both Ontario and

Nunavut. We explored the use of the Nzi trap for the collection of syrphids, and though finding it unsatisfactory, had some interesting observations nonetheless. Finally, we

74 suspect that we found a new species that will likely be fully described in the future. All of this work serves as a fundamental baseline bioinventory for the region, which can be built upon and used in the future to examine the impacts of future development, or other ecological changes.

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125

100 Ontario

50 Akimiski

0 Number of species (95% (95% C.I.) species of Number 200 400 600 800 1000 1200 Specimens

Figure 1: Rarefaction analysis of collected syrphids from northern Ontario and Akimiski

Island, NU, across all sampled years. Dotted lines represent 95% confidence intervals.