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Mercury Concentrations in Aquatic Insects of Two Maine Coastal Streams: Temporal Patterns of Mercury in Aquatic Insect Biosentinels and Implications for Methylmercury Flux
Megan C. Hess University of Maine, [email protected]
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Recommended Citation Hess, Megan C., "Mercury Concentrations in Aquatic Insects of Two Maine Coastal Streams: Temporal Patterns of Mercury in Aquatic Insect Biosentinels and Implications for Methylmercury Flux" (2019). Electronic Theses and Dissertations. 3190. https://digitalcommons.library.umaine.edu/etd/3190
This Open-Access Thesis is brought to you for free and open access by DigitalCommons@UMaine. It has been accepted for inclusion in Electronic Theses and Dissertations by an authorized administrator of DigitalCommons@UMaine. For more information, please contact [email protected]. MERCURY CONCENTRATIONS IN AQUATIC INSECTS OF TWO MAINE COASTAL STREAMS:
TEMPORAL PATTERNS OF MERCURY IN AQUATIC INSECT BIOSENTINELS
AND IMPLICATIONS FOR METHYLMERCURY FLUX
By
Megan C. Hess
B.S. University of Wisconsin – La Crosse, 2016
A THESIS
Submitted in Partial Fulfillment of the
Requirements for the Degree of
Masters of Science
(in Ecology and Environmental Sciences)
The Graduate School
The University of Maine
December 2019
Advisory Committee:
Sarah J. Nelson, Professor of Biogeochemistry, University of Maine, Advisor
Director of Research, Appalachian Mountain Club, NH
Hamish S. Greig, Professor of Stream Ecology, University of Maine, Advisor
Collin A. Eagles-Smith, Supervisory Research Ecologist, U.S. Geological Survey
Amanda J. Klemmer, Professor of Food-web Ecology, University of Maine
MERCURY CONCENTRATIONS IN AQUATIC INSECTS OF TWO MAINE COASTAL STREAMS:
TEMPORAL PATTERNS OF MERCURY IN AQUATIC INSECT BIOSENTINELS
AND IMPLICATIONS FOR METHYLMERCURY FLUX
By Megan C. Hess
Thesis Advisors: Dr. Sarah J. Nelson & Dr. Hamish S. Greig
An Abstract of the Thesis Presented in Partial Fulfillment of the Requirements for the Degree of Master of Science (in Ecology and Environmental Sciences) December 2019
Mercury (Hg) is a widespread water quality concern because it is a potent neurotoxin that biomagnifies through food webs, posing risk to biota higher in the food chain. Aquatic insects are effective for characterizing relative Hg risk in freshwater food webs because they live and forage in aquatic habitats where Hg is present and converted to its more toxic bioavailable form, methylmercury
(MeHg). Specifically, dragonfly nymphs (Odonata) are increasingly being used as a Hg bioindicator.
Therefore, I (Chapter 1) conducted a systematic review of the literature to evaluate the utility of dragonflies as biosentinels for Hg. Dragonfly nymphs have been used throughout the world to provide baseline Hg concentrations within waterbodies, analyze factors affecting bioaccumulation in invertebrates, and determine Hg concentrations within food webs. Dragonfly adults have been used for monitoring Hg concentrations as vectors into and among terrestrial habitats and food webs. There has been an increase in the use of dragonflies for Hg monitoring in the last 10 years, with more than half of the studies in the review conducted after 2009.
However, research evaluating temporal Hg variability in dragonfly nymphs is limited, which can have important implications for monitoring programs and risk assessment because risk to wildlife may be under- or overestimated due to sample timing asynchrony with representative life-history events or
body burden variation. Thus, I (Chapter 2) assessed the temporal variation in total mercury (THg) concentrations in similar aged cordulegastrid dragonfly nymphs from two coastal Maine streams over the course of one year. When we accounted for stream, there was no overall temporal pattern in dragonfly THg concentrations throughout the year. We found that the two streams had similar annual mean THg concentrations; however, within stream variation of THg among months was high and differed among months. Therefore in temperate streams, multiple collections, ideally in different seasons, could be warranted to obtain a robust estimate of Hg concentrations within a lotic ecosystem.
Besides dragonfly nymphs, other aquatic insects also serve as an important link in the MeHg pathway in that they transfer MeHg from single-celled organisms to larger predators in the aquatic environment. Additionally, emergent aquatic insects make up a substantial proportion of the diet of many riparian terrestrial consumers and are a primary mechanism by which MeHg is incorporated into terrestrial systems. Therefore, I (Chapter 3) aimed to determine taxonomic composition, biomass, temporal trends, MeHg concentrations, and MeHg flux of emerging aquatic insects in two coastal streams in Maine using weekly emergence trap collections. Different stream reaches harbored different aquatic insect communities and thus, yielded differences in cross-ecosystem subsidies biomass and temporal flux with respect to aquatic insect emergence. These differences shed light on which organisms were vectors for MeHg and the timing of MeHg pulses to the terrestrial environment in natural systems.
My research indicate that remote protected natural streams, such as those in Acadia National
Park, can yield high MeHg concentrations in biota and thus, large amounts of MeHg flux to the terrestrial environment and predators.
DEDICATION
This is dedicated to my parents, Randy and Chris Hess. I would not have been here without your encouragement to study hard, willingness to allow me to spread my wings, and financial support throughout school. I feel incredibly blessed to have you both so present in my life and this could not have been completed without your support.
ii
ACKNOWLEDGEMENTS
My master’s degree could not have been completed without the support from multiple institutions and dedicated people. I thank my funding sources at the University of Maine, including the Research Reinvestment Fund and the Graduate Student Government. I am beyond grateful to the Dragonfly Mercury Project for welcoming me into the team, entrusting me with coordination duties, and allowing me to use their data to brainstorm thesis questions. I owe many thanks to Acadia
National Park for granting me research permits, allowing me to conduct research within the beautiful park, and being supportive of my research questions. Additionally, I would like to thank Abe Miller- Rushing and Bill Gawley for their willingness to answer questions and share Acadia National Park data with me to help answer my research hypotheses.
I would graciously like to thank Sarah Nelson for actively pursuing funding options to be able to take me on as a graduate student. I am eternally grateful that Sarah not only saw my passion and made the graduate position possible for me, but provided multiple opportunities to grow as a scientist and attend science conferences and meetings throughout the United States and internationally.
Additionally, I would like to extend my gratitude to Hamish Greig for welcoming me to participate in his lab from the beginning and the willingness to be my co-advisor later in my studies. Hamish’s class and enthusiasm reignited my passion for aquatic insects and I appreciate both of my advisors encouragement, inspiration, and laid-back personalities that allowed me to ask questions and explore multiple avenues of research. I thank my committee member, Collin Eagles-Smith, who provided tremendous research and statistical analysis expertise that helped me grow as an ecotoxicologist. I appreciate the guidance of my committee member, Amanda Klemmer, in R code analysis and her enthusiasm and support in collecting emergence traps data. I would also like to thank non-committee members Dr. Brian McGill and Dr. James Willacker for graciously taking time out of their day to answer my statistical questions, more than once. Lastly, I would like to thank my friends here in Maine for keeping me grounded and volunteering to help me with field work, especially Sarah Vogel.
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TABLE OF CONTENTS
DEDICATION ...... ii
ACKNOWLEDGEMENTS ...... iii
LIST OF TABLES ...... viii
LIST OF FIGURES ...... xi
CHAPTERS
1. DRAGONFLY (ODONATA:ANISOPTERA) USE IN MERCURY MONITORING:
A LITERATURE REVIEW ...... 1
Introduction ...... 1
Methods ...... 5
Results ...... 6
History of dragonfly Hg research in the literature ...... 7
Literature examining the usefulness of dragonflies as Hg biosentinels ...... 10
Mercury concentrations compared across dragonfly studies ...... 12
Discussion ...... 16
Dragonflies are used for a variety of purposes in mercury research and monitoring ...... 16
Field collection methods differ by dragonfly life stage and study goals ...... 21
Methodological considerations ...... 23
Research needs ...... 25
Conclusions ...... 26
iv
2. TEMPORAL VARIATION IN MERCURY CONCENTRATIONSIN DRAGONFLY NYMPHS FROM
TWO STREAMS IN ACADIA NATIONAL PARK, MAINE, USA ...... ……28
Introduction ...... 28
Methods ...... 30
Study streams ...... 30
Sample species ...... 32
Dragonfly collection and preparation ...... 32
Mercury analysis ...... 33
Statistical methods ...... 33
Results ...... 34
Discussion ...... 44
Dragonflies as bioindicators for temporal Hg in aquatic environments ...... 45
Temporal differences in THg within each stream provide evidence that temporal
THg variation depends on stream characteristics ...... 46
Monthly THg differences between streams suggests benefits of multiple
samples per year to compare streams ...... 48
Conclusion ...... 48
3. AQUATIC INSECT EMERGENCE AND METHYLMERCURY FLUX IN TWO COASTAL
MAINE STREAMS ...... 50
Introduction ...... 50
Methods ...... 52
Study sites ...... 52
Emergent aquatic insect collection and preparation ...... 52
v
Methylmercury analysis ...... 54
Statistical analyses ...... 54
Total biomass ...... 54
Temporal biomass ...... 55
Methylmercury ...... 55
Methylmercury flux ...... 55
Results ...... 56
Water temperature ...... 56
Stream discharge ...... 57
Taxonomic composition of insects ...... 58
Emergent aquatic insect biomass ...... 60
Temporal emergence of insect orders between months ...... 62
Methylmercury concentrations in insects ...... 70
Methylmercury flux via emergent aquatic insects ...... 72
Discussion ...... 75
Taxonomic composition and biomass differences between stream reaches ...... 75
Temporal variation of insect biomass within streams ...... 76
Methylmercury concentrations in insect orders ...... 78
Methylmercury flux from emergent aquatic insects ...... 81
Conclusion ...... 83
BIBLIOGRAPHY ...... 85
APPENDICES ...... 96
Appendix A. Additional Information for Chapter 1 ...... 96
Appendix B. Additional Information for Chapter 2 ...... 99
vi
Appendix C. Additional Information for Chapter 3 ...... 104
BIOGRAPHY OF THE AUTHOR ...... 109
vii
LIST OF TABLES
Table 1.1. Number of publications and locations of studies involving dragonfly Hg
data by decade ...... 7
Table 1.2. Dragonfly mercury concentrations in the literature ...... 14
Table 1.3. Utilization of dragonflies in mercury literature ...... 17
Table 2.1. Geometric mean total mercury (THg) concentrations in cordulegastrid
dragonfly nymphs for each month in two streams of Acadia National Park,
Maine from March 2018 to April 2019 ...... 36
Table 2.2. Statistical results (degrees of freedom (Df) and residuals, F statistics,
and p-values)from ANOVAs testing for main effects of stream and month
and a stream and month interaction effect on natural log total mercury
concentrations in Cordulegastridae dragonfly nymphs ...... 38
Table 2.3. Significant statistical results (degrees of freedom (Df), T ratio, and p-values)
from multiple comparisons testing for THg differences between streams in
the same month and THg differences among months within each stream ...... 39
Table 2.4. Statistical results (degrees of freedom (df), T ratio, and p-values) from
multiple comparisons testing for THg differences between streams in the
same season and THg differences among seasons within each stream ...... 41
Table 3.1. Taxonomic composition of emergent insects and Araneae collected from
two streams of Acadia National Park, Maine from May–September 2018 ...... 59
Table 3.2. Biomass (mg dry mass) for aquatic insect orders from each stream with all
trap data combined ...... 60
viii
Table 3.3. Statistical results (degrees of freedom (Df), F statistics, and p-values) from
ANOVAs testing for main effects of stream and order and a stream and
order interaction effect on log total biomass by insect order ...... 62
Table 3.4. Significant statistical results (degrees of freedom (df), T-ratio, and p-values)
from multiple comparisons testing for insect order biomasses differences
within months and insect order biomass differences among months from
Marshall Brook ...... 65
Table 3.5. Significant statistical results (degrees of freedom (df), T-ratio, and p-values)
from multiple comparisons testing for insect order biomasses differences
within months and insect order biomass differences among months from
Hunters Brook ...... 66
Table 3.6. Significant statistical results (degrees of freedom (df), T-ratio, and p-values)
from multiple comparisons testing for Araneae biomasses differences
among months from both streams ...... 70
Table 3.7. Summary statistics of MeHg concentration data for insect orders from
Hunters Brook and Marshall Brook, Acadia National Park, Maine ...... 72
Table 3.8. Tukey HSD output comparing MeHg concentrations means by insect order ...... 73
Table 3.9. Summary statistics of total MeHg flux data for insect orders from Hunters
Brook and Marshall Brook, Acadia National Park, Maine ...... 73
Table 3.10. Comparing MeHg concentrations with other adult insects from other
studies ...... 80
Table A.1. The environmental protection agency (EPA) hazardous waste test
methods (SW-846) that documents the protocol and standards for
ix
measuring inorganic mercury, total mercury, and organic methylmercury
used in biological tissues ...... 97
Table B.1. Cordulegastrid dragonfly nymph geometric mean total mercury (THg) for
each season in two streams of Acadia National Park, Maine from March
2018 to April 2019 ...... 103
Table C.1. Trichoptera that were analyzed for MeHg from Marshall Brook ...... 106
Table C.2. Plecoptera that were analyzed for MeHg from Hunters Brook ...... 108
x
LIST OF FIGURES
Figure 1.1. A conceptual model showing one possible route of mercury transfer through
an aquatic food web ...... 2
Figure 2.1. Sampling locations for dragonfly nymphs ...... 31
Figure 2.2. Total mercury (THg) concentrations among twelve months in cordulegastrid
dragonfly nymphs from two streams in Acadia National Park,
Maine, 2018-2019 ...... 37
Figure 2.3. Total mercury (THg) concentrations among seasons in cordulegastrid
dragonfly nymphs from two streams in Acadia National Park, Maine,
March 2018-April 2019 ...... 42
Figure 2.4. Percent difference in monthly geometric mean THg cordulegastrid dragonfly
nymph from the stream-specific annual THg geometric mean across twelve
months from two streams in Acadia National Park, Maine, 2018-2019 ...... 43
Figure 3.1. Water temperature (C) from the end of July to the middle of November in
two streams, Hunters Brook and Marshall Brook, from Acadia National Park,
Maine ...... 56
Figure 3.2. Stream discharge (cfs) from January to December 2018, from Otter Creek
in Acadia National Park, Maine ...... 57
Figure 3.3. The distributions of total biomass of each aquatic insect order collected in
each of six traps from Hunters Brook and Marshall Brook, Acadia National
Park, Maine from May-September 2018 ...... 61
Figure 3.4. The distributions of total biomass (mg/m2/week) of each aquatic insect
order (Diptera, Ephemeroptera, Plecoptera, Trichoptera) per week collected
xi
in emergence traps from Hunters Brook and Marshall Brook, Acadia
National Park, Maine from May–September 2018 ...... 63
Figure 3.5. Total biomass (mg/m2/month) of aquatic insect orders per month in Hunters
Brook and Marshall Brook ...... 64
Figure 3.6. The distributions of total biomass (mg/m2/week) of Plecoptera collected in
six traps weekly from Hunters Brook, Acadia National Park, Maine from
May–September 2018 ...... 67
Figure 3.7. The distributions of total biomass (mg/m2/week) of Trichoptera collected in
six traps weekly from Marshall Brook, Acadia National Park, Maine from
May–September 2018 ...... 68
Figure 3.8. The distributions of total biomass (mg/m2/week) of Tetragnathids per week
collected in six traps from Hunters Brook and Marshall Brook,
Acadia National Park, Maine from May–September 2018 ...... 69
Figure 3.9. A) Total insect MeHg concentrations between Hunters Brook and Marshall
Brook and B) MeHg concentrations among insect orders while accounting
for site ...... 71
Figure 3.10. Emergent aquatic insect total MeHg flux (ng/m2/month) from two streams,
Hunters Brook and Marshall Brook, in Acadia National Park, Maine ...... 74
Figure B.1. Total mercury among dragonfly nymph instars ...... 99
Figure B.2. Relationship between total mercury (THg) and methylmercury (MeHg)
concentrations in thirteen cordulegastrid dragonfly nymphs from Marshall
Brook in Acadia National Park, Maine ...... 100
Figure B.3. Relationship between environmental factors and monthly mean THg in
dragonflies ...... 101
xii
Figure B.4. Relationship between monthly mean THg concentrations in dragonfly
nymphs and water temperature in Hunters Brook ...... 102
Figure C.1. Marshall Brook sampling reach with emergence traps and substrate picture . .... 104
Figure C.2. Hunters Brook sampling reach with emergence traps and substrate picture ...... 104
Figure C.3. (A) Methylmercury concentrations within each Trichoptera vial, indicating
which Trichoptera families are within each vial and (B) The average
caddisfly biomass within each vial against the vials MeHg concentration ...... 106
Figure C.4. (A) Methylmercury concentrations within Plecoptera (Chloroperlidae and
Leuctridae) in Hunters Brook and (B) The average Plecoptera biomass
within each vial against the vials MeHg concentration ...... 107
xiii
CHAPTER 1
DRAGONFLY (ODONATA:ANISOPTERA) USE IN MERCURY MONITORING:
A LITERATURE REVIEW
Introduction
Dragonflies (Odonata:Anisoptera) have long served as effective research models for foundational ecological and biological inference rdoba-Aguilar, 2008; Silva, De Marco, & Chaves
Resende, 2010) and bio-sentinels of environmental quality (Barros Miguel, Max, Oliveira-Junior, Ligeiro,
& Juen, 2017; Bulankova, 1997). Additionally, they are increasingly being used as bio-sentinels for monitoring mercury (Hg) concentrations in aquatic habitats and potential export to terrestrial habitats
(Eagles-Smith, Nelson, Willacker Jr., Flanagan Pritz, & Krabbenhoft, 2016; Haro et al., 2013; Jeremiason,
Reiser, Weitz, Berndt, & Aiken, 2016; Nelson & Flanagan Pritz, 2014). A bio-sentinel is an organism with tissue contaminant concentrations that are indicative of environmental contaminant exposure and risk to ecosystem health (Beeby, 2001). Dragonflies are useful bio-sentinels for monitoring Hg contamination for multiple reasons; they reflect both the bioavailability of MeHg in aquatic ecosystems during their nymphal stage, as well as availability of MeHg to terrestrial communities via emergents (Figure 1.1).
1
Figure 1.1. A conceptual model showing one possible route of mercury transfer through an aquatic food web. (1) Inorganic mercury (Hg2+) is deposited from the atmosphere onto the watershed and transported via runoff to the aquatic system (blue dashed line) where it can be methylated by sulfate
+ reducing bacteria into its organic and more toxic form, methylmercury (CH3Hg or MeHg). (2) The MeHg is now in a bioavailable form that can bioaccumulate in single celled organisms and primary producers
(i.e. diatoms, algae). (3) MeHg can be efficiently assimilated by multicellular organisms (i.e.
Heptageniidae mayfly) and transferred up the food chain with increasing concentration (biomagnified).
(4) Once MeHg is assimilated into a dragonfly nymph’s tissues via diet, aquatic predators i.e. fish) can consume the nymph and the MeHg continues to persist in the predator, with little lost to depuration. (5)
Alternatively, the dragonfly nymph can emerge into a terrestrial flying adult leading to MeHg flux from the aquatic habitat to the terrestrial environment. (6) This dragonfly-mediated MeHg that originated in the aquatic environment is then assimilated by terrestrial predators (e.g. birds).
2
Dragonflies share many characteristics of other Hg bio-sentinels and several aspects of dragonfly ecology and natural history enhance their suitability for this use. First, they are found on every continent except for Antarctica and inhabit a wide range of lentic and lotic freshwater ecosystems (Corbet, 1999).
Additionally, dragonfly nymphs are constrained to their specific water body and have relatively sedentary behavior (Corbet, 1999), thus Hg concentrations within dragonfly tissue are highly site- specific. Third, they are easily captured using minimal equipment (i.e. dip net) and because they are invertebrates, do not require additional sampling permits or certifications as some vertebrate bio- sentinels require (i.e. IACUC and electrofishing certification). Fourth, unlike most other aquatic insects, dragonfly nymphs can take up to six years to develop in the aquatic habitat and thus can document Hg exposure integrated throughout their lifetime (Corbet, 1999).Lastly, individual dragonflies have large body size in comparison with other invertebrates, and represent a substantial portion of invertebrate biomass in aquatic ecosystems. Combined with their commonly elevated Hg concentration, they have relative high MeHg burdens and represent the highest proportion of MeHg flux to terrestrial environments (Chumchal et al., 2017).
Mercury concentrations in dragonfly nymphs are reflective of aqueous MeHg concentrations, and predictive of those in higher order predators (Haro et al., 2013; Jeremiason et al., 2016). Aqueous
MeHg concentrations are highly variable over time, making it difficult to understand potential environmental health risks from any single sampling event. This requires multiple sampling events, whereas dragonfly larvae may integrate over a broader time period and better reflect general Hg availability. Additionally, dragonfly nymphs may be effective biosentinels for waterbodies where more commonly sampled biosentinels, such as fish, are absent (Haro et al., 2013). However, dragonfly nymphs also are a key link in aquatic food webs between primary consumers and larger predators, so are equally informative in waterbodies with fish as well (Haro et al., 2013) (Figure 1.1). In fact, dragonfly nymph
MeHg concentrations correlate with those in fish from the same waterbodies, which provides a logistical
3 advantage as compared to fish sampling that can be time and resource intensive or require additional permitting.
Lastly, most dragonfly nymph Hg is in the form of MeHg (75%-100%) (Gorski, Cleckner, Hurley,
Sierszen, & Armstrong, 2003; Haro et al., 2013; Jones, Chumchal, Drenner, Timmins, & Nowlin, 2013;
Nelson, Webber, & Flanagan Pritz, 2015; Tremblay, Lucotte, Meili, Cloutier, & Pichet, 1996). Because
MeHg is roughly 3x more expensive to analyze in the laboratory than total mercury (THg = MeHg + inorganic Hg), most research projects analyze THg as a proxy for MeHg. Analysis of THg in dragonfly larvae offers researchers the advantage of larger sample sizes that can better characterize variability within and across sites, while still being cost-effective. Thus, researchers can measure THg in dragonfly nymphs with confidence that it is a good estimate of the organic and biotransferred MeHg (Haines, May,
Finlayson, & Mierzykowski, 2003; Mason, Laporte, & Andres, 2000; Nelson & Flanagan Pritz, 2014).
Dragonflies can also be useful for estimating potential Hg flux to terrestrial environments as well as Hg exposure to terrestrial predators, such as birds (E. B. Williams, Chumchal, Drenner, & Kennedy,
2017; Wolfe, Lane, Brigham, & Hall, 2018) because after the nymphs emerge from the water, they spend between a few weeks to a year as an adult in the terrestrial habitat (Chumchal & Drenner, 2015;
Chumchal et al., 2017; Corbet, 1999; Jones et al., 2013; Speir et al., 2014). In some environments, such as small seasonal cattle ponds, adult dragonflies transfer proportionally more MeHg to the terrestrial environment compared to other aquatic emergent insects (e.g. Diptera, Trichoptera, Ephemeroptera,
Zygoptera) both because their biomass is substantially larger than other emergent taxa, and because they accumulate higher MeHg concentrations than most other invertebrate taxa (Chumchal & Drenner,
2015; Chumchal et al., 2017; Tweedy, Drenner, Chumchal, & Kennedy, 2013). As a result, avian predators that feed on emerging and adult dragonflies could have elevated Hg exposure in comparison to those feeding on other prey types (Blancher & McNicol, 1991; Grof-Tisza, LoPresti, Heath, & Karban,
4
2017; Williams et al., 2017; Wolfe et al., 2018). Moreover, their wide home ranges and migration make adult dragonflies potential vectors of Hg well beyond riparian zones (Corbet, 1999; Wikelski et al., 2006).
Despite their potential utility, the implementation of dragonfly nymphs as bio-sentinels is still relatively limited, thus there are no syntheses in the literature documenting the strengths and weaknesses of this approach to research and monitoring. Here we synthesize the information in peer- reviewed and gray literature to inform the use and interpretation of dragonfly nymph and adult biosentinels. We searched for studies that determined Hg concentrations in any dragonfly life stage and included results that provided Hg body burdens (Hothem, 2008; Nelson & Flanagan Pritz, 2014) or measured Hg concentrations in dragonflies in order to evaluate food web Hg, which is typically achieved by extrapolating Hg concentrations to fish predator concentrations or terrestrial flux of emerging dragonflies to birds (Haro et al., 2013; Tsui et al., 2012; Vermeer, Armstrong, & Hatch, 1973). We focus our review on (1) documenting the history of dragonfly use for measuring Hg, (2) synthesizing the literature that has examined the effectiveness of dragonflies as Hg biosentinels, (3) discussing ways in which dragonflies are used for various Hg research and monitoring goals, and (4) synthesizing analytical and field methods used in dragonfly Hg research. A review of prior literature is relevant as dragonflies are increasingly used in the science of Hg research and in Hg monitoring networks (VanderMeulen et al.,
2018).
Methods
We conducted a systematic review of the literature that reported Hg data for Anisoptera to evaluate the utility of dragonflies as biosentinels for Hg. We searched during November 2017–January
2019 using the following standardized search terms: “mercury”, “methylmercury”, “dragonfly”, and
“odonata” within Web of Science and Google Scholar databases. We also examined the references cited in the papers found in the initial search and added those to the analysis where appropriate. Papers that
5 included the term “odonata” were examined to make sure the studies included dragonflies Odonata:
Anisoptera) and not only damselflies (Odonata: Zygoptera); a paper was not included in analysis if it did not taxonomically differentiate dragonflies vs. damselflies. Peer-reviewed papers that stated at least one measurement of Hg concentration (THg or MeHg) in a dragonfly nymph or adult, or discussed the role that dragonflies play in Hg research, were selected for the analysis.
Papers selected for further analysis were categorized by how the dragonfly Hg data were used in the main objective of the paper; however, most of the studies were relevant to multiple topics (Table
1.3). Main objective categories were (1) dragonflies as a Hg biosentinel, (2) determining baseline Hg concentrations, (3) the influence of anthropogenic changes to the landscape on Hg concentrations, (4) environmental factors affecting Hg concentrations, (5) food-web transfer of Hg, and (6) factors influencing bioaccumulation of Hg in biota (Table 1.3). THg, MeHg, and percent MeHg concentrations presented in studies were converted to ng/g dry weight to compare across studies. There were only two studies not reported in Table 1.3 because they spanned multiple areas across the United States, and contained data from all dragonfly families, and aquatic ecosystems, therefore these studies could not be represented in the table in the same manner as the other studies (Nelson et al., 2015; Wiener et al.,
2013)(Table 1.2). Additionally, studies that reported Hg concentrations as wet weight were not included in Table 1.2 (Hothem, 2008; Vermeer et al., 1973).
Results
A total of 53 peer-reviewed articles were found and analyzed in this review. Articles included 40 peer-reviewed primary research articles, six data reports, four articles, two research syntheses, and one
Master’s thesis Table 1.3). The majority of the studies were conducted in North America with 62% completed in the United States and 22% completed in Canada. The remaining 18% of studies were from
Sweden, Greece, Iran, China, South Africa, Bolivia, and Argentina (Table 1.1). Two studies contained data
6 from two countries, the United States and Canada (Chételat, Amyot, & Garcia, 2011) and Canada and
Sweden (Tremblay et al., 1996). We found literature reporting Hg data in dragonfly nymphs and adults spanning six of the seven dragonfly families (Aeshnidae, Cordulegastridae, Cordullidae, Gomphidae,
Libellulidae, and Macromiidae). There are only 11 species world-wide of the seventh family (Petaluridae) and no studies that we encountered utilized them for Hg research.
Table 1.1. Number of publications and locations of studies involving dragonfly Hg data by decade.
Years Number of Location (Number of studies) published publications 1970-1979 1 Canada 1980-1989 0 - 1990-1999 7* Canada (3), Sweden (2), USA (2), Greece 2000-2009 11 USA (8), Canada (2), China 2010+ 34* USA (23), Canada (6), Argentina (2), Bolivia, Iran, China, South Africa
*Two of the studies contained data from two countries, the United States and Canada (Chételat et al., 2011) and Canada and Sweden (Tremblay et al., 1996).
History of dragonfly Hg research in the literature
The use of dragonfly nymphs as Hg biosentinels, to our knowledge, was first documented in
1973 in Canada (Vermeer et al., 1973) and dragonfly Hg measurements were not mentioned again in the literature until twenty years later in Sweden (Parkman & Meili, 1993). Several papers during this interval provided “Odonata” Hg concentrations but did not specify the sub-order and thus were not included in our analyses. The first paper we found that specifically referenced dragonfly Hg concentrations examined the effects of Hg on aquatic birds in western Ontario in 1973 (Vermeer et al., 1973). Five dragonfly nymphs were analyzed in this study for Hg concentrations to compare with Hg concentrations in different invertebrate prey of aquatic birds including common goldeneye ducks (Bucephala clangula).
The goal of the research was to determine how much Hg was in the breast muscle of duck species and
7 examine a correlation between Hg concentrations in duck prey items including dragonfly nymphs.
Twenty years later, Parkman and Meili (1993) used macroinvertebrates to monitor Hg in eight diverse lakes in Sweden to assess the influence of ecological, chemical, regional, and climatic factors on the accumulation of Hg. It was not uncommon to study many lakes across a region to start to explore the factors that influence Hg accumulation in organisms; however, this was the first study we found that included predatory dragonfly nymphs and a range of other macroinvertebrate orders for broad scale Hg comparisons.
Throughout the 1990s and 2000s, studies we reviewed used macroinvertebrates, including dragonflies, for one of two purposes: (1) to report baseline Hg concentrations in biota to determine risk of wildlife within a region or waterbody (Bank, Burgess, Evers, & Loftin, 2007; George & Batzer, 2008;
Haines et al., 2003; Hall, Rosenberg, & Wiens, 1998; Hothem, 2008; Slotton, Ayers, Reuter, & Goldman,
1995; Tremblay & Lucotte, 1997; Wiener & Shields, 2000); and, (2) to determine factors controlling bioaccumulation in biota (Gorski et al., 2003; Mason et al., 2000; Parkman & Meili, 1993; Tremblay et al., 1996; Wyn, Kidd, Burgess, & Curry, 2009). This included comparing multiple aquatic habitats within a region to determine ecological, spatial, and temporal trends in Hg accumulation (George & Batzer, 2008;
Parkman & Meili, 1993; Tremblay et al., 1996), examining factors that control bioaccumulation in aquatic biota (Gorski et al., 2003; Mason et al., 2000; Wyn et al., 2009), and quantifying Hg in the prey of bird species (Goutner & Furness, 1997). Dragonfly Hg data were also used to assess the impacts of anthropogenic changes to ecosystems, including examining invertebrate Hg concentrations from reservoirs (Hall et al., 1998; Tremblay & Lucotte, 1997), assessing the impacts of gold mining on food web Hg (Slotton et al., 1995) and assessing the impacts of the Nyanza Chemical Waste Dump Superfund
Site on the Sudbury River in Massachusetts (Haines et al., 2003; Wiener & Shields, 2000). All of these studies sampled different groups of macroinvertebrates to assess Hg concentrations in the environment.
8
Dragonfly nymphs were one of the groups of macroinvertebrates captured but were not the primary focus of the study.
Studies further expanded the use of dragonfly nymph Hg biosentinels throughout the 2010s often with a broader focus on how nymphs can play a role in Hg research. For example, dragonfly nymphs were used to document relative Hg burdens for predatory aquatic insect taxa in Hg trophic transfer studies (Chételat et al., 2011; Jardine, Kidd, & Rasmussen, 2012; Molina et al., 2010; Riva-
Murray et al., 2011). Dragonflies were now recognized as an important biotic compartment that should be analyzed when discussing bioaccumulation and biomagnification of Hg in food webs (Arribére et al.,
2010; Bates & Hall, 2012; Beganyi & Batzer, 2011; Karimi, Fisher, & Meliker, 2014; Rizzo, Arcagni,
Arribére, Bubach, & Ribeiro Guevara, 2011). This decade also included the first studies to use recently- emerged dragonfly adults and exuviae to estimate flux of Hg to the terrestrial environment. Ten experimental ponds were created near Fort Worth, Texas to understand how fish predators (Tweedy et al., 2013), pond permanence and seasons (Williams et al., 2017), and drying disturbance (Chumchal et al., 2017) impacted insect-mediated Hg flux from the aquatic environment to the terrestrial environment and terrestrial predators (Speir et al., 2014; Williams et al., 2017). Another study used these ponds in conjunction with a mesocosm experiment to examine how nutrients and fish predators regulated MeHg flux to the terrestrial environment via aquatic emergent insects, including dragonflies
(Jones et al., 2013). Dragonfly adults were collected in 2014 for MeHg analysis (Chumchal et al., 2017;
Speir et al., 2014; Tweedy et al., 2013) and subsequent MeHg concentrations were used along with exuviae collections to estimate MeHg flux (Williams et al., 2017).
Including the study above, 10 of the 53 publications we found contained data on Hg in dragonfly adults, either as recently emerged nymphs (Chumchal et al., 2017; Jones et al., 2013; Speir et al., 2014;
Tweedy et al., 2013; Williams et al., 2017) or as flying adults (Lesch & Bouwman, 2018; Ortiz, Weiss-
9
Penzias, Fork, & Flegal, 2015; Tsui et al., 2012; Z.-S. Zhang, Lu, Wang, & Zheng, 2009; Z. Zhang, Song,
Wang, & Lu, 2012). Two broad themes were highlighted in these studies: using dragonflies to determine relative Hg baselines and burdens; or placing emerged dragonflies in the context of food webs and flux into the terrestrial environmental, including predators. Together, these studies provided evidence that dragonfly adults can play a large role in MeHg flux from aquatic systems because of their high MeHg concentrations and larger biomass compared to other aquatic insects.
Literature examining the usefulness of dragonflies as Hg biosentinels
The efficacy of using dragonfly nymphs as Hg biosentinels was explicitly tested in several studies in the 2010s. Haro et al. (2013) concluded that dragonfly nymphs of the family Gomphidae were useful
Hg biosentinels because they contain most of their Hg in the form of bio-transferrable MeHg (>60%
MeHg of total Hg in 761 gomphid nymphs), and their MeHg concentrations correlated with MeHg in unfiltered lake water. In fact, aqueous MeHg concentrations explained 63% of the variation in mean
MeHg levels in gomphids (Haro et al., 2013). Additionally, mean THg and MeHg concentrations in gomphid nymphs were positively correlated with THg in both prey fish and game fish in 12 lakes in the
Upper Midwest region of the US (Haro et al. 2013). Haro et al. (2013) further documented that a sample size of 10 nymphs or more was sufficient to determine an accurate MeHg concentration, as compared to a sample size of 40 prey fish species needed for similar accuracy, indicating that larval gomphids were as effective as, and more efficient than, small game fish as indicators of Hg exposure in game fish (Haro et al., 2013). Jeremiason et al. (2016) investigated another dragonfly family, Aeshnidae, as biosentinels of
MeHg concentrations for streams in northern Minnesota that were elevated in sulfate from acid mine drainage. There was a significant positive correlation in MeHg concentrations of aeshnid dragonfly nymphs and dissolved MeHg in water, thus Jeremiason et al. (2016) concluded that Aeshnid dragonfly nymphs were effective biosentinels in streams of that region.
10
Invertebrate life stages can play a role in how Hg moves through food webs (Chételat et al.,
2011). Thus, Buckland-Nicks et al. (2014) examined the use of dragonflies as Hg biosentinels in nymphs, exuviae, emerging adults, and adults for MeHg, Hg(II) and THg. Using general linear models from 66 nymphs, 32 exuviae, 10 emerging adults, and 20 mature adults, the authors concluded that dragonfly nymphs, emerging adults, and mature adults contained similar THg and MeHg concentrations (Buckland-
Nicks, Hillier, Avery, & O’Driscoll, 2014). However, Hg(II) has a high affinity for cutaneous tissues and thus a portion (~5%) of THg concentrations can be sequestered in the chitin and not available to terrestrial predators (Buckland-Nicks et al., 2014; Camci-Unal & Pohl, 2010; Nelson et al., 2015).
Nonetheless, there was not a significant difference in THg or MeHg concentrations between life stages
(e.g. nymphs vs adults) of dragonflies (Buckland-Nicks et al., 2014; Nelson et al., 2015).
Beginning in 2011, a spatially extensive survey, the Dragonfly Mercury Project (DMP), sampled dragonfly nymphs to obtain baseline Hg data for 22 national parks across the USA (Nelson et al., 2015).
The DMP data release in 2014 highlighted the spatial variation of Hg concentrations in dragonfly nymphs from collections that were completed in 2013 across national parks (Flanagan Pritz, Eagles-Smith, &
Krabbenhoft, 2014; Flanagan Pritz, Nelson, & Eagles-Smith, 2014; Nelson, Chen, Kahl, & Krabbenhoft,
2014; Nelson et al., 2015). These data provided baseline Hg concentrations in a variety of ecosystem types by using dragonfly nymphs as THg biosentinels. Like Haro et al. (2013) reported for MeHg, the
DMP found that water THg described some (~31% in stepwise regression) of the variability in dragonfly nymph THg and that MeHg was on average 88 ± 17% of THg in dragonfly nymphs. In addition, analyses indicated that THg concentrations were significantly different among three guilds of dragonflies
(sprawlers, climbers, and burrowers) when sites and parks were accounted for in statistical models.
Likewise, Buckland-Nicks et al. (2014) found that MeHg, THg, and Hg(II) differed among families;
Aeshnidae had significantly higher concentrations of MeHg and THg than Gomphidae and Libellulidae, and Gomphidae and Cordulidae had higher Hg(II) concentrations than in Aeshnidae and Libellulidae
11
(Buckland-Nicks et al., 2014). Evidence from these studies emphasizes the utility of dragonflies as biosentinels of Hg but also highlights a role for ecological or taxonomic context when interpreting dragonfly Hg data.
In 2018, the usefulness of adult dragonflies to monitor ten environmental metallic elements, including Hg, was examined in South Africa (Lesch & Bouwman, 2018). Adult dragonflies (n=105) were collected and identified to 15 different species. There was no significant difference among mean Hg concentrations in adult dragonflies from 21 different sites across South Africa. Elevated concentrations in dragonfly Hg were found in species located close to (1) a river that is heavily polluted from mining and industrial activities and (2) a wetland that is close to a wastewater treatment plant (Lesch & Bouwman,
2018). Furthermore, previous studies in the North-West Province of South Africa indicated elevated Hg concentrations in water, soil, and humans which coincided with elevated Hg concentrations in adult dragonflies of that region (Lesch & Bouwman, 2018; Lusilao-Makiese, Cukrowska, Tessier, Amouroux, &
Weiersbye, 2013; Oosthuizen, John, & Somerset, 2010). This was the first study that specifically tested adult dragonflies as a biomonitoring tool for metallic elements, but did not focus solely on Hg (Lesch &
Bouwman, 2018).
Mercury concentrations compared across dragonfly studies
THg, MeHg, and percent MeHg vary in dragonfly nymphs from lentic and lotic aquatic systems and dragonfly adults in terrestrial systems around the world (Table 1.2). Studies examined dragonfly nymphs in nearly every aquatic habitat from lakes and ponds to wetlands and streams (Table 1.2). Adult dragonflies were also analyzed for Hg in terrestrial habitats including adults captured in grasslands and riparian zones (Lesch & Bouwman, 2018; Ortiz et al., 2015; Tsui et al., 2012; Z.-S. Zhang et al., 2009). Two studies used experimental ponds in Texas to examined the influence of dragonfly MeHg flux to the
12 terrestrial environment (Chumchal et al., 2017; Williams et al., 2017). The majority of studies presented here studied Hg concentrations in dragonflies in either lake/pond or, stream/river habitats (Table 1.2).
Thirty percent of papers in this review identified dragonflies to sub-order (Anisoptera) whereas the rest (70%) identified them, at least, to family. Only six papers identified dragonflies to genus and seven papers to species but the supplemental information needed to be examined to determine species identification for some studies (Haro et al., 2013; Wiener et al., 2013). Unless the study focused heavily on dragonflies as their Hg biosentinel, Hg concentrations in all dragonfly families reported were lumped into “Anisoptera” (Arribére et al., 2010; Beganyi & Batzer, 2011; Chételat et al., 2011; Gorski et al., 2003;
Goutner & Furness, 1997; Rizzo et al., 2011; Tremblay & Lucotte, 1997; Wiener & Shields, 2000; Z.-S.
Zhang et al., 2009; Z. Zhang et al., 2012). Mean THg concentrations ranged from 34–560 ng/g dw in lakes or ponds, 20–15,654 ng/g dw in wetlands, 40–600 ng/g dw in streams or rivers, and 175–632 ng/g dw in terrestrial habitats. Mean MeHg concentrations ranged from 25–489 ng/g dw in lakes or ponds, 112–
145 ng/d dw in experimental ponds, 19–1200 ng/g dw in wetlands, 29–~450 ng/g dw in streams or rivers, and 180 ng/g dw in terrestrial habitats, where only one mean Hg concentration was available
(Table 1.2). The percent of THg as MeHg (MeHg %) was consistently high in wetlands (75.5-110%) and streams or rivers (68-97%) with increased variation in mean MeHg % in lakes and ponds (34-96.1%).
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Table 1.2. Dragonfly mercury concentrations in the literature. Mean ranges in mercury concentrations
(Total mercury (THg), Methylmercury (MeHg), and percent methylmercury (MeHg (%))) reported or converted into ng/g dry weight in select peer reviewed articles categorized by the aquatic or terrestrial system and the region in which the study took place. Estimates are indicated with a tilde (~) if concentrations were not directly stated and only shown graphically. Taxa indicates the family identification reported with the mercury concentration; Anisoptera did not distinguish Hg concentration within a specific family, A= Aeshnidae, L = Libellulidae, G= Gomphidae, C= Corduliidae, CR=
Cordulegastridae, M= Macromiidae.
THg MeHg MeHg Site Location Taxa Citation (ng/g dw) (ng/g dw) (%)
Lakes/Ponds (Nymphs) United States Anisoptera 155 Chételat et al. 2011 WI Anisoptera 77.9-105.9 48.5-57.2 54.1-62.2 Gorski et al. 2003 WI (ISRO) G 47.0-119.8 16.8-102.5 27.8-85.6 Haro et al. 2013 ME Anisoptera 208-560 130-319 53-83 Burgess 1997 TX A 96.1 Speir et al. 2014 L 74.3 Speir et al. 2014 NH, VT L, G 34-264 Karimi et al. 2016 MN 50.0-143.6 Knights et al. 2005 MN (VOYA) G 80.9-128.6 55.7-109.0 47.6-91.4 Haro et al. 2013 MI (PIRO) G 36.8-145.9 37.8-99.8 60.2-102.9 Haro et al. 2013 MI (SLBE) G 18.8-55.6 4.1-43.7 22.0-87.1 Haro et al. 2013 Sweden A 140 - 593 Parkman and Meili 1993 L 93 - 487 Parkman and Meili 1993 C 157-501 104-352 63-84 Tremblay et al. 1996 Canada C 80-217 64-167 34-92 Tremblay et al. 1996 Anisoptera 155 Chételat et al. 2011 Tremblay and Lucotte Quebec C ~150-400 ~200-400 1997 Tremblay and Lucotte C ~100-200 ~100-350 1997 Nova Scotia A 170-440 Wyn et al. 2009 A, C, G, L Buckland-Nicks et al. 2014 Alberta A 83-84 Allen et al. 2005 C 38-55 Allen et al. 2005 Bolivia A 459-533 319-395 Molina et al. 2010 L 56-555 25-489 Molina et al. 2010 Argentina Anisoptera 260 Arribere et al. 2010 Anisoptera 170 Rizzo et al. 2011
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Table 1.2 cont. Experimental Ponds (Nymphs) United States TX A, L 112 Chumchal et al. 2017 A, L 120-145 Williams et al. 2017 Wetlands (Nymphs) Canada 19.4 - Ontario A 191.3 Hall et. al 1998 59.4 - C 200.3 Hall et. al 1998 Saskatchewan A, L 215 187.6 75.7 Bates and Hall 2012 A 123 - 136 Wolfe et al. 2018 Toronto A 280-1250 90-110 Sinclair et al. 2012 L 76-1100 77-103 Sinclair et al. 2012 G 38 93 Sinclair et al. 2012 Iran A 70-15,654 Nasirian & Irvine 2017 United States GA C, L 20-40 Beganyi and Batzer 2011 River/Streams (Nymphs) United States MA Anisoptera ~200-450 Wiener and Shields 2000 A, C, G, L, M 272-514 185-407 68-95 Haines et al. 2003 CA G 70-600 Slotton et al. 1995 MD A ~30-50 70-95 Mason et al. 2000 177.33- NY, SC A 541.00 Riva-Murray et al. 2011 154.65- L 282.54 Riva-Murray et al. 2011 CA A, G 79.1 70.8 Tsui 2012 MN A 129-233 118-190 81-94 Rolfhus et al. 2015 CR 126-168 118-159 95 Rolfhus et al. 2015 C 140-194 136-185 91-97 Rolfhus et al. 2015 L 134-177 111-124 92-63 Rolfhus et al. 2015 G 70 54 77 Rolfhus et al. 2015 A 40-253 29-187 Jeremiason et al. 2016 Greece Anisoptera 150 Goutner and Furness 1997 Terrestrial (Adults) China Liaoning Anisoptera 151,700 Zhang et al. 2009 Providence Anisoptera 4,234.14 Zhang et al. 2012 South Africa L 0.6 - 2700 Lesch and Bouwman 2018 United States CA L, CR 220 - 632 Ortiz et al. 2015 CA A, G 175 180 Tsui 2012
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Discussion
Dragonflies have been studied in Hg research since 1973, for a variety of purposes. Our review highlights the continued development of dragonflies as biosentinels, from studies presenting dragonfly
Hg burdens to the emergence of research using dragonflies to quantify the spatial and temporal variation in environmental Hg. Unsurprisingly, these developments have been coupled with research critically evaluating the efficacy of dragonflies as biosentinels of THg and MeHg in both aquatic and terrestrial habitats.
Dragonflies are used for a variety of purposes in Hg research and monitoring
The 53 studies we reviewed were categorized based on how the dragonfly Hg data were used in terms of the main objective of the study (Table 1.3); however, most the studies were relevant to multiple other topics. Some studies were included within two categories. For example, Jeremiason et al.
(2016) used dragonfly nymphs to investigate two main objectives; (1) the impact of elevated sulfate loads on MeHg concentrations in water, indicating the study could be categorized as ‘Influence of anthropogenic changes to the landscape on Hg concentrations’, and 2) whether aeshnid dragonflies are useful Hg biosentinels of sulfate-impacted streams indicating that they could fall under the ‘Hg biosentinel’ category as well (Jeremiason et al., 2016).
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Table 1.3. Utilization of dragonflies in mercury literature. Mercury studies in the literature that report
Hg* concentrations in dragonflies were categorized based on how the dragonfly Hg data were utilized in the main objective of the study.
Dragonflies as Hg biosentinels Utility of dragonflies as a (Haro et al., 2013; Jeremiason et al., 2016; Lesch & Bouwman, 2018; Nasirian & Hg biosentinel Irvine, 2017; Nelson et al., 2015; Z. Zhang et al., 2012) Biological influences on (Buckland-Nicks et al., 2014; Fletcher, Lindell, Stillings, Blas, & McArthur, 2017) dragonflies as Hg indicators Baseline Hg concentrations in dragonflies California (Hothem, 2008; Ortiz et al., 2015) National Parks (Bank et al., 2007; Burgess, 1997; Flanagan Pritz et al., 2014b, 2014a; Flanagan Pritz and Nelson, 2017; Gorski et al., 2003; Haro, 2014; Haro et al., 2013; Knights et al., 2005; Nelson et al., 2015, 2014; Rizzo et al., 2011; Rolfhus et al., 2015; Wiener et al., 2013) Wetlands (George & Batzer, 2008; Nasirian & Irvine, 2017; Sinclair, Xie, & Mitchell, 2012) Sudbury River, MA (Haines et al., 2003; Wiener & Shields, 2000) Prairie Pothole Region (Bates & Hall, 2012; Wolfe et al., 2018) Influence of anthropogenic changes to the landscape on Hg concentrations in aquatic biota Gold mining (Slotton et al., 1995) Created wetlands (Chumchal & Drenner, 2015; Sinclair et al., 2012) Reservoirs (Hall et al., 1998; Tremblay & Lucotte, 1997) Superfund site (Haines et al., 2003; Wiener & Shields, 2000) Coal combustion waste (Fletcher et al., 2017) Sulfate loading (Jeremiason et al., 2016) Environmental factors affecting Hg concentrations in aquatic insects Wildfire (Allen, Prepas, Gabos, Strachan, & Zhang, 2005; Beganyi & Batzer, 2011) Pond permanence (Chumchal et al., 2017; Williams et al., 2017) Flooding (Hall et al., 1998) Food-web transfer of Hg Avian predators (Goutner & Furness, 1997; Vermeer et al., 1973; Williams et al., 2017; Wolfe et al., 2018) Fish predators (Gorski et al., 2003; Jones et al., 2013; Karimi, Chen, & Folt, 2016; Tweedy et al., 2013; Wyn et al., 2009) Terrestrial Subsidies (Chumchal & Drenner, 2015; Jones et al., 2013; Speir et al., 2014; Tsui et al., 2012) Biomagnification (Arribére et al., 2010; Haines et al., 2003; Molina et al., 2010; Rizzo et al., 2011; Wyn et al., 2009) Factors influencing bioaccumulation in aquatic insects In Lakes (Chételat et al., 2011; Gorski et al., 2003; Parkman & Meili, 1993; Tremblay & Lucotte, 1997; Tremblay et al., 1996; Wyn et al., 2009) In Streams (Mason et al., 2000; Riva-Murray et al., 2011) Diet (Jardine et al., 2012; Karimi et al., 2016) Biogeochemistry (Z.-S. Zhang et al., 2009; Z. Zhang et al., 2012) *Hg is used here as a broad term encompassing studies measuring total mercury and/or methylmercury
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Dragonflies have been used to obtain baseline Hg concentrations for locations all over the world. Several studies sampled a suite of macroinvertebrates, including dragonflies, to report baseline
Hg concentrations in biota to determine regional risk of Hg to wildlife (Bank et al., 2007; George &
Batzer, 2008; Haines et al., 2003; Hothem, 2008; Ortiz et al., 2015; Rolfhus et al., 2015; Wiener &
Shields, 2000), while others reported baseline dragonfly Hg concentrations to determine spatial patterns across the United States using dragonfly biosentinels (Nelson & Flanagan Pritz, 2014; Nelson et al., 2015)
(Table 1.3). A frequent use of dragonfly larvae in the literature is as a spatial biosentinel; that is, to indicate variation in Hg body burdens among different sites. Adult dragonflies have also been used to determine baseline concentrations in the environment including two studies that examined the biogeochemistry in soil in order to predict Hg concentrations in terrestrial insects in Huludao City, China
(Z.-S. Zhang et al., 2009; Z. Zhang et al., 2012) and a study in California that examined the baseline concentrations of Hg in terrestrial arthropods (Ortiz et al., 2015).
Dragonflies have been used to examine Hg concentrations in biota affected by anthropogenic changes to the landscape. Dragonfly nymphs, along with other macroinvertebrates, were sampled to understand the impacts of gold mining (Slotton et al., 1995), constructed wetlands and reservoirs
(Chumchal & Drenner, 2015; Hall et al., 1998; Sinclair et al., 2012; Tremblay & Lucotte, 1997), and the
Nyanza Chemical Waste Dump Superfund Site (Haines et al., 2003; Wiener & Shields, 2000) (Table 1.3).
Similar to the baseline and monitoring category above, these studies used aquatic invertebrates to understand how anthropogenic changes to hydrology or land use can influence Hg bioaccumulation and magnification. Two of these studies documented elevated Hg concentrations in dragonfly nymphs from the affected sites in comparison to control or unaffected sites (Haines et al., 2003; Slotton et al., 1995) while others did not find a significant difference in dragonfly nymphs of contaminated vs. reference sites
(Jeremiason et al., 2016). Other studies indicated that human-made ponds and wetlands introduce additional Hg flux to ecosystems (Chumchal & Drenner, 2015; Sinclair et al., 2012). The differences in Hg
18 in biota from experimental or hydroelectric reservoirs compared to reference wetlands or lakes were examined (Hall et al., 1998; Tremblay & Lucotte, 1997). One study showed that there was a significant difference in Hg biota concentrations between hydroelectric reservoirs and reference lakes (Tremblay &
Lucotte, 1997), while the other study did not find a significant difference (Hall et al., 1998). Therefore, dragonflies have the potential to monitor many different anthropogenic landscape and aquatic changes, not only the subjects listed here, and provide useful data to describe human impacts on Hg concentrations in the environment.
Dragonflies have been used to examine how natural environmental disturbances can impact Hg concentrations in biota. Two studies examined how wildfires can impact changes to Hg load to a waterbody and thus, subsequent Hg concentrations in biota (Allen et al., 2005; Beganyi & Batzer, 2011).
Seasonal drying disturbance in a waterbody can determine when aquatic insects need to emerge and thus, the timing of MeHg flux to the terrestrial environment (Chumchal & Drenner, 2015; Chumchal et al., 2017; Williams et al., 2017). Lastly, although they used an experimental reservoir, Hall et al. (1998) reported an increase in MeHg in most aquatic insects after flooding occurred. These types of studies provide important context for the natural environmental gradients or disturbances that warrant consideration when designing or interpreting dragonfly biosentinel studies.
Dragonflies have been used in determining Hg transfer through food webs. Dragonflies can transfer aquatically-derived MeHg to larger aquatic predators, like fish, or to terrestrial predators after emergence (Figure 1). Some studies included dragonflies in their Hg monitoring focused on detecting spatiotemporal patterns in concentration of MeHg in aquatic food webs (Rolfhus et al., 2015; Wyn et al.,
2009) while other studies examined the insect mediated Hg flux and trophic transfer to terrestrial predators (Speir et al., 2014; Tsui et al., 2012; Williams et al., 2017). Dragonfly nymphs can be a large source of Hg to waterfowl (Goutner & Furness, 1997; Vermeer et al., 1973) and fish (Jones et al., 2013;
19
Tweedy et al., 2013). Similarly, dragonfly emergents and adults can transfer Hg to avian predators like red-winged blackbirds (Agelaius phoeniceus) (Williams et al., 2017; Wolfe et al., 2018). Biomagnification of Hg has been examined via trophic position by categorizing aquatic biota into functional feeding groups or determining trophic level via stable isotope analysis (Arribére et al., 2010; Haines et al., 2003;
Molina et al., 2010; Rizzo et al., 2011; Wyn et al., 2009). Not surprisingly, all biomagnification studies showed that an increase of trophic position positively correlated with an increase of Hg concentration in biota (Arribére et al., 2010; Haines et al., 2003; Molina et al., 2010; Rizzo et al., 2011; Wyn et al., 2009).
Dragonfly Hg concentrations have been used to compare bioaccumulation factors among sites.
Determining bioaccumulation factors for Hg include sampling a suite of waterbodies that encompass landscape or geochemical variety, spatial or landscape gradients, or food-chain length and content variability (Chételat et al., 2011; Gorski et al., 2003; Mason et al., 2000; Parkman & Meili, 1993; Riva-
Murray et al., 2011; Tremblay & Lucotte, 1997; Tremblay et al., 1996; Wyn et al., 2009). These studies usually have many physical, chemical, and biological environmental factors that were measured and statistically tested to work toward predicting and understanding bioaccumulation in biota. For example,
Chételat et al. (2011) measured a suite of primary consumer insects, pelagic zooplankon, and secondary consumer macroinvertebrates to determine which environmental factors (e.g. pH, DOC, total phosphorus, lake depth, lake area, etc.) influenced MeHg concentrations in these lake invertebrates.
Other studies indicated sources of organic matter in diet that in turn influenced Hg concentrations in invertebrates (Jardine et al., 2012) or the location within the waterbody in which predators consume prey, either benthic or pelagic (Karimi et al., 2016) (Table 1.3). These studies piece together many environmental characteristics and multiple trophic levels to begin to understand bioaccumulation in waterbodies of varying biogeochemical characteristics.
Field collection methods differ by dragonfly life stage and study goals
20
All of the studies presented here using dragonfly nymphs as Hg biosentinels captured nymphs via a dip net. This is perhaps not surprising given that minimal equipment and training in field protocols are needed and the method is effective at sampling ample and diverse habitat in a short time period.
Capturing dragonfly adults requires more specialized equipment and can be more time consuming than capturing nymphs. A popular method of capturing emergent aquatic insects is through the use of emergence traps (Chumchal et al., 2017; Davies, 1984; Jones et al., 2013; Speir et al., 2014; Tweedy et al., 2013; Williams et al., 2017). Emergence traps are pyramid or cone shaped nets that funnel emergent insects (here after, adult insects) into a sampling cup containing a preservative (e.g., ethanol). Traps may either float on the water’s surface in natural habitats or completely enclose the openings of artificial aquaria or mesocosms. Advantages of using emergence traps is that researchers can obtain data without being physically present at the sampling site, data are collected within a given area which enables the calculation of emergence density, and collection cups can be emptied relatively easily without disturbing sampling habitat. However, a disadvantage of emergence traps in the field is that some aquatic insects do not swim within the water column to the water’s surface to emerge. For example, dragonfly nymphs will emerge on a variety of natural habitat including reeds, sticks, banks and vegetation, therefore emergence traps, especially smaller ones, are relatively ineffective at capturing adult dragonflies and a substantial portion of emergents can be missed. Therefore, it is important for maximizing capture in the field to have emergence traps placed in locations where there is structure for the nymph to crawl out, for example, encapsulating reeds and vegetation within emergence traps.
Emergence traps are more effective in capturing dragonfly adults when used to enclose an entire mesocosm, as implemented in most adult Hg studies (Chumchal et al., 2017; Jones et al., 2013; Speir et al., 2014; Tweedy et al., 2013). Additionally, it is important to consider the type of preservative used when using emergence traps to capture adult dragonflies for Hg analysis to avoid contaminating the sample. Ethanol can be used if analysis will be done shortly after collection, whereas Hot Shot No Pest
21
Strips can be used if humidity or time of collection is not an issue, to preserve samples for later Hg analysis (Chumchal et al., 2017: Supplemental Data). However, it should be noted that larger dragonfly adults may not move into collection cups, so thorough examination of the mesh inside the trap should be undertaken and dragonfly adults should be hand-picked from the sides if necessary (Speir et al.,
2014). Methylmercury flux is the cumulative mass of MeHg that is transferred from the aquatic to terrestrial environment, and is the product of biomass and MeHg concentrations. Standardizing the known areas of emergence traps can enable estimates of Hg flux per unit area of a waterbody, ultimately facilitating extrapolations to a stream reach or lake.
Dragonfly emergence can also be estimated by quantifying exuviae in a location (Wissinger,
1988). Most simply, exuviae can be collected on the edges of aquatic habitats at set time intervals to measure emergence rate. Alternatively, emergence platforms can be installed which are connected to the sediment but extend to the water’s surface, providing structure for nymphs to crawl up and emerge upon (Williams et al., 2017). Exuviae may be picked off the platforms during set time intervals to quantify emergence biomass over time (Williams et al., 2017). However, a subset of emerged adults is still needed to obtain a Hg sample for analysis to extrapolate exuvial density to Hg flux to the terrestrial environment.
Two studies indicate the use of hand-held entomological nets to capture perching adult dragonflies (Lesch & Bouwman, 2018; Tsui et al., 2012). While this is an effective way to specifically target adult dragonflies, especially when abundances are low, it is more time consuming in terms of person-hours and difficult to obtain samples because of the tremendous agility of adult dragonflies.
Furthermore, estimates of adulty density or biomass flux are compromised by difficulties in quantifying the sampling area. Each method for dragonfly collection (dip nets, emergence traps, exuvia counts, and
22 entomological nets) has advantages and disadvantages and the choice of method depends on the question and goals of the study.
Methodological considerations
Appropriate taxonomic identification is an important aspect for ensuring comparable results for interpretation. Life history, foraging ecology, and physiology/metabolism are important determinants of
Hg bioaccumulation (Becker et al., 2018; Riva-Murray et al., 2013; Wiener et al., 2013; Yung et al., 2019) and also are common distinguishing characteristics of different taxa. For dragonflies, family-level identification is the most common level of resolution (Beganyi & Batzer, 2011; Eagles-Smith, Nelson, et al., 2016; Haines et al., 2003; Haro, 2014; Hothem, 2008; Jardine et al., 2012; Karimi et al., 2016; Nelson
& Flanagan Pritz, 2014; Riva-Murray et al., 2011; Rolfhus et al., 2015; Sinclair et al., 2012; Speir et al.,
2014; Williams et al., 2017). This is because it is relatively easy to identify dragonfly nymphs to family level compared to other aquatic insects and it provides more life history information than order level.
Differences in taxonomic resolution (e.g. family, genus, species) can result in different tissue concentrations of Hg (Buckland-Nicks et al., 2014; Fletcher et al., 2017) and taxa with different life histories and foraging modes often differ in Hg burdens even within a sampling location (Eagles-Smith,
Nelson, et al., 2016).
As with any study, sample size for Hg monitoring with dragonflies must be adequate for hypothesis testing or detecting change across space or through time. A power analysis is an effective way of determining the minimum sample size needed to answer research goals (VanderMeulen et al.,
2018). For example, using power analyses, studies from National Parks in the Great Lakes region determined a range in minimum sample size of 4–27 dragonfly nymphs per waterbody (VanderMeulen et al., 2018). The power analyses determined an 80% probability of detecting a 20% change in THg within dragonfly nymphs over ten years (with Type I error of 0.05)(VanderMeulen et al., 2018). The data
23 and citations presented herein should be useful for practitioners seeking to estimate sample variance and effect sizes in Hg burdens.
Once nymphs are collected, samples can either be analyzed individually or with multiple individuals combined into one composite sample. Composite samples are useful for very small dragonfly nymphs to obtain a sufficient mass for Hg analysis. For example, Gorski et al. (2003) composited three or more dragonflies together when analyzing for Hg due to mass specifications needed for analysis (Gorski et al., 2003). However, with the continued increases in instrument sensitivity, this is less of a concern than it has been in the past. Composite sampling can reduce analytical costs because fewer samples are analyzed. Given that costs can constrain the number of Hg determinations, a greater number of nymphs in composites (opposed to individuals) can provide a better overall estimate of the mean Hg concentration as compared with a small number of individual samples. However, information on variability in Hg among individual nymphs is lost when they are composited, and combining multiple individuals may result on reduced taxonomic sensitivity to analyses depending on how individuals are selected for analyses. As Hg instrumentation has evolved, detection limits have generally declined and thus, less mass is needed to analyze for Hg; this has allowed more studies to analyze individuals instead of composites. Analyzing dragonfly individuals typically also produces greater statistical power through increased replication within a given site/sample time. Analyzing individuals may mean targeting larger individuals and can provide a better chance of identifying individuals to a lower taxonomic level (i.e. species). Lastly, analysis of individuals can allow for biological questions to be answered involving Hg bioaccumulation, including length and mass comparisons in relation to Hg concentration. For example, it is generally accepted that Hg concentrations in fish increase with body mass, however this relationship has not been tested in aquatic insects (Buckland-Nicks et al., 2014). Ultimately, there are advantages and disadvantages to analyzing individual dragonflies or composite samples and the cost and benefits should be weighed based on the research goal.
24
Research needs
Our review highlights areas that would benefit from further research. In particular, differences in Hg concentrations have been documented between dragonfly families which suggests ecological factors may influence differences in Hg accumulation (Buckland-Nicks et al., 2014). Diet is arguably the most important factor that determines Hg concentrations.
On average, dragonflies of the families Aeshnidae, Cordulegastridae and Macromiidae have the largest bodies and also happen to have the largest THg concentrations nation-wide (Eagles-Smith et al. in prep). However, we do not see increasing THg concentrations with increasing body mass (Buckland-
Nicks et al., 2014). Although, a larger bodied nymph typically has a larger labium and is able to catch and consume larger prey. Larger prey items typically occupy a higher trophic level than small prey
(Woodward et al., 2005) and thus, may contain higher MeHg concentrations. To test this assertion, further research could determine whether head capsule width (labium size) correlated with THg concentrations. Ultimately, using carbon and nitrogen stable isotope analysis may well illuminate the complex relationships between dragonfly size, diet, trophic position and Hg differences between dragonfly taxa or individuals.
Environmental Hg can vary temporally, thus it is important to quantify how dragonfly nymphs as biosentinels integrate this variability, and how they are influenced by seasonal abiotic changes. For example, an increase in water temperature (i.e. summer months) can increase metabolic rate, increase feeding, and thus increase MeHg burden in invertebrates and fish (Buckman et al., 2019; Dijkstra,
Buckman, Ward, Evans, & Dionne, 2013). However, growth dilution, in which Hg concentrations decrease during rapid growth, can counteract increased Hg from increased feeding and may play a significant role interpreting individual Hg burdens (Karimi, Fisher, & Folt, 2010). Unraveling these mechanisms highlights a need to collect data on dragonfly nymph THg over time to determine if and
25 how THg concentrations vary throughout dragonfly growth or seasonal habitat changes. This information would ultimately be beneficial for monitoring programs to indicate the best time to sample for robust Hg estimates for baseline data or determination of risk to wildlife.
Lastly, dragonfly adults can potentially transfer their Hg to terrestrial habitats upon emergence.
Some studies have used dragonfly MeHg concentrations and their emergence rates to estimate MeHg flux from dragonflies in aquatic environments (Chumchal et al., 2017; Jones et al., 2013; Williams et al.,
2017). While these methods are standard for research on emergence and cross ecosystem subsidy studies, using adult dragonflies Hg concentrations for terrestrial Hg monitoring is much less studied; we only identified five such studies in our review (Lesch & Bouwman, 2018; Ortiz et al., 2015; Tsui et al.,
2012; Z.-S. Zhang et al., 2009; Z. Zhang et al., 2012). Adult dragonflies have specifically been tested as indicators of environmental metallic elements, however, Hg analysis was only one of the 33 elements that were analyzed and not the focus of the study (Lesch & Bouwman, 2018). Adult dragonflies have been analyzed for Hg as baseline concentrations but concentrations were very high and therefore, it is important to investigate why concentrations are elevated (0.6 – 151,700 ng/g dw) (Lesch & Bouwman,
2018; Z.-S. Zhang et al., 2009; Z. Zhang et al., 2012). Because adult dragonflies are a relatively new terrestrial Hg biosentinel, more research is needed to interpret the applicability of dragonflies to this use.
Conclusions
This review documented the use of dragonflies in mercury research and monitoring. I identified a body of research (53 papers) supporting the use of dragonfly nymphs as an effective biosentinel to monitor Hg concentrations because they contain most of their Hg as MeHg, their MeHg concentrations positively correlate with aqueous MeHg concentrations and/or fish MeHg concentrations, and they are abundant and easy to collect across multiple ecosystem types (Haro et al., 2013; Jeremiason et al., 2016;
26
Nelson et al., 2015). Dragonfly nymphs have been used throughout the world to provide baseline Hg concentrations within waterbodies, analyze factors affecting bioaccumulation in invertebrates, and determine Hg concentrations within food webs. Dragonfly adults have been used for monitoring Hg concentrations into and among terrestrial habitats and food webs, but nymphs were the most commonly studied life-stage. There has been an increase in the use of dragonflies for Hg monitoring in the last 10 years, with more than half of the studies in this review conducted after 2009. Therefore, field collection protocols, lab protocols, and EPA methods have been recently solidified to effectively and accurately measure Hg concentrations in dragonfly nymphs (VanderMeulen et al., 2018) (Table A.1). In order to achieve an integrated understanding of dragonfly use in Hg monitoring, further research should focus on understanding how ecology and natural history influence Hg accumulation in dragonflies.
27
CHAPTER 2
TEMPORAL VARIATION IN MERCURY CONCENTRATIONSIN DRAGONFLY NYMPHS FROM TWO
STREAMS IN ACADIA NATIONAL PARK, MAINE, USA
Introduction
Mercury (Hg) is a global concern for aquatic ecosystems because it is a globally-distributed pollutant and biomagnifies through food webs. It is widely distributed onto the landscape via anthropogenic emissions (Krabbenhoft & Sunderland, 2013; United Nations Environment Programme,
2013). It then can biomagnify through food webs when converted to methylmercury (MeHg), posing neurotoxic risk to biota of higher trophic levels, including humans (Compeau & Bartha, 1985; Wiener,
Krabbenhoft, Heinz, & Scheuhammer, 2003; Wiener & Spry, 1996). Thus, it is important to monitor Hg in water bodies across the United States to assess Hg risk to wildlife and humans.
Mercury monitoring at the national scale has largely occurred in abiotic media such as atmospheric deposition (e.g., NADP-MDN) and lake or stream water, and in a few broad-scale surveys of fish (Brigham, Wentz, Aiken, & Krabbenhoft, 2009; Eagles-Smith, Wiener, et al., 2016; U.S.
Environmental Protection Agency, 2016). Bioindicators like fish provide an integrated assessment of Hg that reflects higher trophic level Hg exposure and are important for fish consumption advisories.
Bioindicators are valuable for better understanding ecological exposure and risk. However, uniform fish species presence across broad geographic extents are rare, making it difficult to consistently compare at broad scales. Dragonfly nymphs are widespread and have increasingly been used to monitor Hg in aquatic systems (Eagles-Smith, Nelson, et al., 2016; Haro et al., 2013; Jeremiason et al., 2016; Nelson et al., 2015; VanderMeulen et al., 2018; Wiener et al., 2013). Dragonfly nymphs are considered top aquatic
28 insect predators and thus can contain high concentrations of Hg, most of which is in the MeHg form
(>90%; Eagles-Smith et al. in prep). They also serve as a key link in the trophic transfer of Hg between primary consumers and larger predators such as fish and birds (Corbet, 1999; Haro et al., 2013; Williams et al., 2017), suggesting that elevated Hg burdens in dragonflies inhabiting some waterbodies could be of concern for the health of their predators.
Mercury cycling and MeHg bioaccumulation are temporally dynamic processes that can vary over relatively short time periods. Multiple environmental factors influence temporal variation in Hg including precipitation, hydrologic discharge, and temperature. For example, stream discharge has been positively correlated with aqueous THg concentrations in lotic systems (Brigham et al., 2009; Johnson et al., 2007; Shanley, Kamman, Clair, & Chalmers, 2005) and increased water temperature results in increased MeHg bioaccumulation in invertebrates and fish (Buckman et al., 2019; Dijkstra et al., 2013;
Sokolova & Lannig, 2008). This temporal Hg variation has been documented as seasonal differences in
Hg burdens of aquatic bioindicators. Phytoplankton sampled in the spring had the highest concentrations of Hg and MeHg while zooplankton sampled in the autumn exhibited the highest Hg concentrations in Long Island Sound (Gosnell, Balcom, Tobias, Gilhooly, & Mason, 2017). Additionally,
George and Batzer (2008) documented significant Hg concentrations changes in amphipods
(Crangonyctidae) over six sampling dates from December 1998–August 2000 in 32 sampling locations
(George & Batzer, 2008). Further up the food chain, established fish monitoring programs recommend or require that fish samples be collected within the same season each year because Hg concentrations in small fish can differ among seasons (Eagles-Smith & Ackerman, 2009; Mason et al., 2000). However, to our knowledge, there has been no research of within- or among-season variability in dragonfly nymph
Hg concentrations. High temporal variability of Hg concentrations in dragonfly nymphs would indicate that point-in-time sampling events may not reflect the overall risk to upper-level predators. Thus, it’s
29 important to understand the temporal variation in dragonflies in order to more effectively interpret their utility as bioindicators.
We aimed to examine the magnitude of temporal variation in dragonfly nymphs throughout the year by sampling similar sized cordulegastrid nymphs every month for one year in two streams of Acadia
National Park. Understanding this temporal variability provides important insights into the factors influencing Hg bioaccumulation in these biosentinels and will inform optimal timing of future sampling to best represent stream-specific Hg risk. This investigation helps to define the utility of this important biosentinel for broad scale monitoring programs.
Methods
Study Streams
Acadia National Park is located on Mount Desert Island off the coast of Maine, USA (Figure 2.1).
The island covers an area of 27,972 hectares containing 26 mountains, 24 ponds, at least two dozen streams, and 10 named wetland areas. We sampled cordulegastrid dragonfly nymphs from two streams located within ANP, Hunters Brook 44°18’ 20.7”N 68°13’18.6”W) and Marshall Brook 44°17’ 00.9”N
68°20’55.7”W) Figure 2.1). These two streams have similar pH, temperature, dissolved oxygen, but may differ in dissolved organic carbon, total phosphorus, and nitrogen (Gawley & Wiggin, 2016) .
Additionally, Marshall Brook has lower discharge and higher dissolved organic carbon than Hunters
Brook (Gawley & Wiggin, 2016). At each stream we selected similar 100–meter reaches with ample dragonfly nymphs for sampling.
30
Figure 2.1. Sampling locations for dragonfly nymphs. Mount Desert Island located on the Downeast coast of Maine showing the watersheds of Marshall Brook (left) and Hunters Brook (right). Hashed lines indicate Acadia National Park boundaries.
31
Sample species
We focused our sampling on dragonflies from the Cordulegastridae family because they can dominate lotic odonate assemblages, are easy to identify to species, and are long-lived. Larval development of cordulegastrids can range to six years (i.e. merovoltine). Cordulegastrids are only found in lotic systems and both streams contain an abundance of delta-spotted spiketail (Cordulegaster diastatops) dragonflies and a smaller population of twin-spotted spiketail (Cordulegaster maculata) dragonflies. Cordulegaster species tend to inhabit backwater depositional areas of streams or rivers and conceal their bodies under the sediment (i.e. burrowing guild) (Corbet, 1999). They are sit-and-wait predators with a large scooped labium to capture prey effectively and are usually the top invertebrate predator (Corbet, 1999).
Dragonfly collection and preparation
Once a month between March 2018 and February 2019 we collected dragonfly nymphs via dip nets swept through the benthic substrates of each stream. Five nymphs from Hunters Brook and eight nymphs from Marshall Brook were sampled each month to obtain an 80% probability of detecting 20% change in mean THg concentrations annually with a Type I error of 0.05 (VanderMeulen et al., 2018).
The same 100m stretch of stream was sampled each month to attempt to eliminate habitat bias. To reduce size or age bias, we used head capsule width as an estimator of instar in the field; our goal was to collect dragonfly nymphs of similar cohorts (F-3 or F-2 stadia) every month. Nymphs were handled via
“clean-hands” procedures, individually bagged, and kept on ice in the field until they could be transported to the lab and frozen at -20C within 8 hours (VanderMeulen et al., 2018).
In the laboratory, head width (from the edge of eye to eye in dorsal view) and hind wing sheath length were obtained to the nearest millimeter from images taken with Nikon SMZ800 microscope with a Zeiss Axiocam ERC 5s camera attachment. Instar was determined by dividing hind wing sheath by head
32 capsule width to determine a ratio. Our ratios were then compared with the Cordulegaster maculata ratio presented in Tennessen (2017) to determine instar (Tennessen, 2017). Thawed nymphs were blotted dry and weight wet determined to the nearest 0.00001g. We then dried samples for 48 hours at
50°C, and recorded a dry weight. Whole and aliquot samples of dragonfly nymphs were then dried and ground to a fine powder using a porcelain mortar and pestle, or glass rod.
Mercury analysis
Each individual dragonfly nymph (or an aliquot thereof, for larger samples) was analyzed for total mercury (THg) concentration following EPA Method 7473 (U.S. Environmental Protection Agency,
2007), at the USGS Contaminant Ecology Research Lab in Corvallis, Oregon on a Nippon MA-3000
Mercury Analyzer. Laboratory QA/QC procedures include analysis of certified reference materials
(National Research Council Canada dogfish muscle [DORM-4] and lobster hepatopancreas [TORT-3]), instrument duplicates, laboratory tissue standards, liquid calibration standards, matrix spikes, matrix spike duplicates, and reagent blanks. Recoveries averaged 100.8% ± 6.9 % (n=16) for certified reference materials.
Total Hg concentrations are typically strongly correlated with MeHg in dragonflies and most of the THg is comprised of MeHg (Eagles-Smith et al. 2019 in prep). We also measured MeHg for a subset of individuals to ensure this was also the case for our samples. Methylmercury was analyzed via methods described elsewhere (Eagles-Smith et al. 2019 in prep).
Statistical methods
All analyses were conducted using R v. 3.5.0 statistical software (R Core Team, 2019). Nymph total Hg data were log-transformed to normalize residuals and reduce heteroscedasticity. Total Hg results are presented as geometric means (ng/g dw) with geometric standard errors estimated using the
33 delta method (B. K. Williams, Nichols, & Conroy, 2002). We aimed to ensure that we minimized biological differences (i.e. sampled the same family and similar sized nymphs each month) from samples among and between each month. We used a one-way analysis of variance (ANOVA) to, first, determine if nymph instar (age) had an effect on nymph THg concentrations. Because there were no significant differences of THg concentrations among instars (see Results), we grouped all instars and proceeded to evaluate the temporal differences on nymph THg concentrations. Next, we used a two-way ANOVA
(Type III) with month and stream as main effects and a month x stream interaction to evaluate variation in THg concentrations. We exclude the last sampling event for each stream because they only contained a sample from one stream and not both, thus the model with an interaction failed to converge. Multiple comparisons were accessed using pair-wise contrasts among months within each stream and between streams in each month with a false discovery rate p-value adjustment.
We also evaluated the effects of season and stream on THg concentrations using a two-way
ANOVA with a type III sum of squares. The spring season compiled data from March, April and May; the summer season was June, July and August; autumn consisted of September, October and November, and winter compiled data from December, January and February.
Results
We sampled a total of 170 cordulegastrid dragonfly nymphs, 100 nymphs from Marshall Brook and 70 nymphs in Hunters Brook, in Acadia National Park between March 2018 and April 2019. Nymph size was similar between both streams; head widths, a proxy of age, ranged 2.46–5.85 mm (mean ± SD =
3.63 mm ± 0.673, n= 70) at Hunters Brook and 2.53–6.66mm (mean ± SD = 3.98 ± 0.913 mm, n= 100) at
Marshall Brook. Instar ranged from F-5 to F-1; however, the majority (92%) of nymphs that were sampled were F-4 (23%), F-3 (51%), and F-2 (18%) instars. Total Hg concentrations did not differ among
34 instars (F4,165 = 0.121, p = 0.975, Figure A.1), therefore data from all instars were combined for use in subsequent analyses.
Across both streams and all sampling events, dragonfly nymph THg concentrations ranged from
209–852 nanograms per gram dry weight (ng/g dw), with an arithmetic mean (± SE) of 366 ± 7.79 ng/g dw (Table 2.1). Geometric means of dragonfly THg concentrations across months (± geometric standard error) were 351 ± 12.19 ng/g dw in Hunters Brook and 357 ± 7.85 ng/g dw in Marshall Brook. A total of
13 nymphs from Marshall Brook were analyzed for MeHg from June (N = 4), July (N=4) and January (N=5)
(Table 2.1), and there was a strong relationship between THg and MeHg concentration, indicating that
THg concentrations adequately characterized MeHg concentrations (R2 = 0.931, p < 0.0001)(Figure
A.2.2). Additionally, the mean percentage (± SE) of MeHg as THg was 86.5 ± 1.58% (N=13) (Table 2.1).
Our final model with stream and month as fixed effects, and a stream x month interaction, revealed that dragonfly THg concentrations differed among months (F10,132 = 2.82, p = 0.003) but not between streams (F1,132 = 3.35, p = 0.070), and the interaction indicated that THg concentrations varied differently between streams (F10,132 = 3.23, p < 0.001)(Table 2.2). Multiple comparisons revealed that
THg differed between streams in both June and November (Figure 2.1), but not during other months. In
June, THg concentrations from Marshall Brook (358 ± 18.6 ng/g dw) were significantly greater than those at Hunters Brook (269 ± 13.1 n/g dw, p = 0.031) (Figure 2.1). By contrast, in November, THg concentrations from Marshall Brook (272 ± 17.3 ng/g dw) were significantly lower than THg concentrations at Hunters Brook (376 ± 46.0 ng/g dw, p = 0.005) (Figure 2.1).
35
Table 2.1. Geometric mean total mercury (THg) concentrations in cordulegastrid dragonfly nymphs (ng/g ± geometric standard error) for each month in two streams of Acadia National Park, Maine from March 2018 to April 2019.
Methylmercury (MeHg) concentrations (ng/g), and thus MeHg percentage from
THg, were also analyzed for a subset of dragonfly nymphs from Marshall Brook in
June, July, and January.
Hunters Brook Marshall Brook Marshall Brook Month THg (ng/g) THg (ng/g) MeHg (ng/g)
March 2018 316 ± 34.5 389 ± 26.9 April 266 ± 20.1 338 ± 23.7 May 330 ± 29.0 378 ± 13.6 June 269 ± 13.1 358 ± 18.6 319 ± 27.5 July 389 ± 46.1 321 ± 14.6 266 ± 30.0 August 332 ± 46.9 415 ± 21.3 September 437 ± 36.4 372 ± 19.2 October 450 ± 81.8 350 ± 32.4 November 376 ± 46.0 272 ± 17.3 December 359 ± 45.6 352 ± 33.6 January 2019 364 ± 35.5 435 ± 28.2 413 ± 20.9 February - 313 ± 18.5 April 339 ± 30.9 -
36
Figure 2.2. Total mercury (THg) concentrations among twelve months in cordulegastrid dragonfly nymphs from two streams in Acadia National Park, Maine, 2018-2019. THg concentrations are reported as the geometric means of each season and error bars represent ± standard error with a LOESS smoothed line. Total Hg concentrations were different between streams in June and November
(indicated with *).
37
Table 2.2. Statistical results (degrees of freedom (Df) and residuals, F statistics, and p-values) from ANOVAs testing for main effects of stream and month and a stream and month interaction effect on natural log total mercury concentrations in Cordulegastridae dragonfly nymphs.
Response Effects Df F p-value Log(THg) Stream 1,132 3.35 0.070 Month 10,132 2.82 0.003 Stream:Month 10,132 3.23 < 0.001 Log (THg) Stream 1,162 9.01 0.003 Season 3,162 4.53 0.004 Stream:Season 3,162 5.05 0.002
38
Table 2.3. Significant statistical results (degrees of freedom (Df), T ratio, and p-values) from multiple comparisons testing for THg differences between streams in the same month and THg differences among months within each stream. THg = total mercury concentrations in Cordulegastridae dragonfly nymphs. HB = Hunters Brook. MB = Marshall Brook.
Response Contrasts Df T-ratio p-value June THg HBvMB 132 -2.18 0.031 November THg HBvMB 132 2.83 0.005 Hunters Brook THg April - October 132 -3.62 0.015 April - September 132 -3.42 0.015 June - October 132 -3.53 0.015 June - September 132 -3.33 0.015 Marshall Brook THg August - November 132 3.69 0.009 January - November 132 4.32 0.002 March - November 132 3.12 0.041
39
However, the interaction between stream and month indicated that relative differences among months were not consistent between streams (F10,132 = 3.23, p < 0.001)(Table 2.2). In Marshall Brook,
THg concentrations in November were lower than THg concentrations in March (p = 0.041), August (p =
0.009), and January (p = 0.002), but all other months were the same (Table 2.3). In Hunters Brook, THg concentrations in April and June were 1.6x fold lower than in October (p = 0.015) and September (p =
0.015) (Table 2.3).
Our model with stream and season as fixed effects, and a stream x season interaction, revealed that dragonfly THg concentrations differed among streams (F1,162 = 9.01, p = 0.003) and between seasons
(F3,162 = 4.53, p = 0.004)(Table 2.2). However, the interaction term indicated that, THg concentration differences among seasons were not consistent between streams (F3,162 = 5.04, p < 0.002) (Table 2.2,
Figure 2.2). Multiple comparisons revealed that THg concentrations differed between streams within the same season. The THg concentrations in nymphs from the autumn were greater at Hunters Brook than
Marshall Brook (contrast, p = 0.003) (Table 2.4). In contrast, THg concentrations in the spring were greater at Marshall Brook than Hunters Brook (contrast, p = 0.031; Table 2.4). Multiple comparisons also revealed that THg concentrations in dragonflies differed among seasons within a stream. Total Hg concentrations in Hunters Brook were higher in autumn than spring (p = 0.003) and summer (p = 0.026), whereas THg concentrations in Marshall Brook were similar throughout seasons (Table 2.4, Figure 2.2).
40
Table 2.4. Statistical results (degrees of freedom (df), T ratio, and p-values) from multiple comparisons testing for THg differences between streams in the same season
and THg differences among seasons within each stream. THg = total mercury concentrations in Cordulegastridae dragonfly nymphs. HB = Hunters Brook. MB =
Marshall Brook.
Response Contrasts Df T-ratio p-value
Autumn THg HB - MB 162 3.00 0.003 Spring THg HB - MB 162 -2.18 0.031
Summer THg HB - MB 162 -1.20 0.232
Winter THg HB - MB 162 -0.180 0.857 Hunters Brook
THg Autumn – Spring 162 3.51 0.004 Autumn – Summer 162 2.65 0.027
Autumn – Winter 162 1.47 0.216
Spring – Summer 162 -0.600 0.549 Spring – Winter 162 -1.645 0.204 Summer – Winter 162 -1.006 0.379 Marshall Brook THg Autumn – Spring 162 -1.620 0.308 Autumn – Summer 162 -1.43 0.308 Autumn – Winter 162 -1.66 0.308 Spring – Summer 162 0.187 0.985 Spring – Winter 162 0.019 0.985 Summer – Winter 162 -0.175 0.985
41
Figure 2.3. Total mercury (THg) concentrations among seasons in cordulegastrid dragonfly nymphs from two streams in Acadia National Park, Maine, March 2018-April 2019. THg concentrations are reported as the geometric means of each season and error bars represent ± standard error. Letters indicate significant differences among seasons in Hunters Brook determined by multiple comparisons. There were no significant differences among seasons in Marshall Brook. Spring: March, April, May. Summer:
June, July, August. Autumn: September, October, November. Winter: December, January, February.
42
To better visualize temporal shifts in Hg, we calculated stream-specific means and present monthly concentrations as differences from these means. Over the 12-month sampling period, we observed variance from the stream-specific mean each month in both streams (Figure 2.3). The dragonfly nymph THg data from Marshall Brook are randomly distributed from the stream-specific mean throughout the year (Figure 2.3). In Hunters Brook, we observed lower than average mean monthly THg concentrations in nymphs in the spring and higher than average mean monthly THg concentrations in nymphs in early autumn (Figure 2.3).
Figure 2.4. Percent difference in monthly geometric mean THg cordulegastrid dragonfly nymph from the stream-specific annual THg geometric means across twelve months from two streams in Acadia National
Park, Maine, 2018-2019.
43
Discussion
We sampled cordulegastrid dragonfly nymphs of similar instars across two streams monthly during 2018–2019 to determine whether there were temporal patterns in THg throughout the year in dragonfly nymphs. The sampling streams were located in the northeastern US, a temperate location with strongly varying seasonal conditions. We found that dragonfly THg concentration did not differ with age (instar) and their THg concentration was an appropriate proxy for MeHg (MeHg:THg was ~88%)
(Table 2.1). Our two streams did not show overall differences in dragonfly nymph THg concentrations, as the annual mean was nearly the same in both streams (Table 2.2). However, there were significantly different THg concentrations during two months (June and November) between streams (Table 2.3).
This, along with the stream by month interaction, reinforces that the trajectories of THg concentrations throughout the year within streams were different. However, the temporal differences within each stream were limited to only a few sampling periods; November from March, August, January in Marshall
Brook and June and April from October and September in Hunters Brook (Table 2.3).
Consequently, the THg concentrations in autumn and spring were different between streams and it followed that the seasonal dragonfly THg concentration changes were not consistent between streams throughout the year (Figure 2.2, 2.3). Autumn had significantly higher THg concentrations than spring and summer in Hunters Brook. However, there were no differences in THg concentrations among seasons in Marshall Brook. The reasons for the fluctuation in dragonfly THg concentrations are unclear, but may be due to a combination of stream characteristics involving temporal changes in biogeochemistry and available prey items.
44
Dragonflies as bioindicators for temporal Hg in aquatic environments
We did not observe THg differences among instars; however, the study was designed to minimize the influence of dragonfly biological factors across the year-long study. This was done by focusing on one dragonfly family, Cordulegastridae, that were similar in age. Cordulegastrid nymphs molt approximately 13 times (13 instars) throughout larval development (Corbet, 1999) and 92% of our samples were within three of those molts (instars F-4, F-3, and F-2), therefore they were fairly close in age. There were no significant differences in THg concentrations among the F-1 through F-5 instars we collected (Figure A. 2.1). Thus, development stage was not a strong factor influencing changes in temporal THg concentrations. Because cordulegastrids are merovoltine and live up to six years in the aquatic environment, we could not determine the growth rates of the populations in the scope of this study. However, it is unlikely that a new cohort of individuals would enter our sampling size range and exhibit lower THg concentration from the previous month because similarly aged instars (F-4, F-3, F-2) did not exhibit differing THg concentrations in any given month. Therefore, there is flexibility in sampling instars F-5 through F-1 because the THg concentrations in these later-aged nymphs did not differ between instars at any time of the year.
Additionally, a subset of samples from three different months were analyzed for MeHg to confirm that THg was representative of bio-available Hg. Dragonflies in our study had similar MeHg concentrations in all three months (June, July and January) and the mean was 86 ± 2.93 % MeHg of THg
(Table 2.1). This result aligns with other studies that have shown that percent MeHg ranged 63-110%
(Bates & Hall, 2012; Mason et al., 2000; Rolfhus et al., 2015; Sinclair et al., 2012; Speir et al., 2014).
However, this was the first time dragonflies were measured for MeHg among months and we obtained a small sample size, therefore we cannot directly say that the MeHg concentrations were not different in these growing season and winter months.
45
Temporal differences in THg within each stream provide evidence that temporal THg variation depends on stream characteristics
When we accounted for stream, there was no overall temporal pattern in dragonfly THg concentrations throughout the year. However, our model indicated that monthly and seasonal dragonfly
THg concentration changes were not consistent between streams and graphical inspection revealed that dragonfly THg concentrations varied differently over time in each stream (Figure 2.1). Therefore, we investigated stream-specific factors that could result in the different THg trajectory throughout the year.
There was a more distinct temporal pattern of dragonfly THg concentrations in Hunters Brook than in Marshall Brook, upon visual inspection of results (Figure 2.1, Figure 2.3). This pattern in Hunters
Brook could be indicative of increased feeding during the summer months and seasonal prey shifts.
Graphically, we observed an increase of THg concentrations from spring throughout the summer months, which peaked during the autumn in Hunters Brook. This timeframe corresponds to warming water temperatures, which increases dragonfly nymph metabolism (CITE). The energetic requirements to maintain a higher metabolism are generally met via increasing consumption rates, which could enhance MeHg bioaccumulation if the not offset by similarly increasing growth rates. Other studies have shown that elevated water temperature can increase MeHg bioaccumulation in invertebrates and fish
(Buckman et al., 2019; Dijkstra et al., 2013; Sokolova & Lannig, 2008). However, dragonfly nymph THg concentrations were not correlated with water temperature from Hunters Brook (Figure A. 2.4). This suggests that other factors, such as seasonal variation in dragonfly nymph diet may be responsible for the temporal variation in dragonfly THg concentrations.
Aquatic insect diet can vary temporally based on changes in prey availability and shifts in prey diversity, which can influence their intake of Hg through prey items. For example, seasonally changing feeding habits in the caddisfly, Hydropsyche morosa, were associated with changing Hg concentrations within the larvae throughout the year (Snyder & Hendricks, 1995). As obligate predators, dragonfly
46 nymphs will consume a broad range of prey items (Corbet, 1999; Merrill & Johnson, 1984). Moreover, cordulegastrid nymphs generally have the broadest habitat flexibility compared to other dragonfly families and thus a higher likelihood of encountering a more diverse range of prey (Burcher & Smock,
2002). Prey availability can shift throughout the year. For example, a fish predator can feed on smaller insects in the spring when they are more readily available and shift toward larger insects in the summer as additional insect prey begin to increase in size and abundance. However, we cannot be sure if diet shifts occurred in dragonfly nymphs throughout the year in Hunters Brook without additional analyses such as isotopic analysis or gut content analysis of our specimens or their gut contents.
Temporal THg variation in Marshall Brook was much more random and graphically, there was no real overall temporal pattern in dragonfly THg concentrations. November had lower THg concentrations than March, August, and January but there were no overall differences in THg concentrations among seasons (Table 2.3, 2.4). Based on monthly observations, Marshall Brook had slower water velocity and less discharge than Hunters Brook. During spring snowmelt and rain events, Marshall Brook exhibited increased inundation of the marshy banks instead of increasing discharge downstream as occurred at
Hunters Brook. Therefore, Marshall Brook might always be saturating its waters with Hg from the terrestrial environment around it, which may be indicative of the roughly 7x higher DOC in Marshall
Brook than Hunters Brook (Gawley & Wiggin, 2016; Peckenham, Kahl, Nelson, Johnson, & Haines, 2007;
St Louis et al., 1994). Additionally, DOC in streams of New Hampshire have shown to be correlated with mean streamwater MeHg (Chaves-Ulloa et al., 2016). The high availability of DOC and potentially MeHg in streamwater at Marshall Brook could therefore dampening variability in THg concentrations in dragonflies seasonally.
47
Monthly THg differences between streams suggests benefits of multiple samples per year to compare streams
Our streams are of similar watershed size and characteristics, subject to the same climate, and are not influenced by any known point sources of Hg. There was a difference in THg concentrations between streams in June and November; however, there was no overall difference in dragonfly nymph
THg concentrations when we accounted for month. Therefore, multiple sampling events throughout the year could yield a more robust measurement of THg concentrations within the streams than a single sample per year. For example, if a single yearly sample would have been taken in June or November, we might have concluded an individual stream had higher or lower dragonfly THg concentration than the other. Additionally, THg concentrations in autumn and spring were different between streams and likewise, the seasonal dragonfly THg concentration changes were not consistent between streams throughout the year (Figure 2.2, 2.3). Two sampling events per year using dragonfly nymph biosentinels, perhaps summer and autumn sampling events, could be beneficial in determining THg concentrations in a waterbody and comparing THg concentrations among waterbodies.
Conclusion
This study evaluated temporal Hg variability in dragonfly nymph biosentinels, which can have important implications for monitoring programs and risk assessment. The variation of THg concentrations in nymphs throughout the year was not clearly directional and differed among months and among seasons depending on the stream sampled. Temporal studies provide important insight for monitoring programs because risk to wildlife may be under- or overestimated due to offsets between sample timing and representative life-history events or body burden variation. Therefore, depending on the stream being sampled, multiple collections per year could be warranted to obtain a robust estimate
48 of Hg concentrations within a lotic ecosystem. In our study of two streams at Acadia National Park, there were limited temporal THg differences within Marshall Brook and a seasonal temporal THg pattern in
Hunters Brook. Samples in different seasons would permit a more representative estimate of THg concentrations within a waterbody (Figure 2.2). Additionally, because both interactions of (1) stream and month and (2) stream and season were significant, the stream location, for temperate streams, could be the most important factor when determining if THg will change throughout the year on a monthly or seasonal scale. Temporal variation in dragonfly nymphs would need to be further studied in different lotic systems and lentic systems to determine what is driving temporal changes in dragonfly nymph Hg biosentinels.
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CHAPTER 3
AQUATIC INSECT EMERGENCE AND METHYLMERCURY FLUX
IN TWO COASTAL MAINE STREAMS
Introduction
Many insect species spend the majority of their lifecycle as aquatic juveniles, incorporating the nutrients and energy from aquatic ecosystems into their tissues. Upon emergence as terrestrial adults, they provide a cross-ecosystem subsidy of nutrients and energy to terrestrial systems. The timing and duration of these subsidies differ among aquatic insect species, which has implications for consumers in the recipient community (Baxter, Fausch, & Saunders, 2005; Corbet, 1964; Judd, 1962; Nakano &
Murakami, 2001). For example in temperate locations, some aquatic insects like chironomid midges
(Diptera: Chironomidae) emerge all year, whereas most other insects like mayflies (Ephemeroptera), stoneflies (Plecoptera), and caddisflies (Trichoptera) exhibit pulsed emergence during the warmer months (Corbet, 1964; Sweeney & Vannote, 1982). This translates to differences in subsidy biomass, and thus energy and nutrients, throughout the year, which influences the foraging and phenology of terrestrial consumers and the stability of their food webs (Nakano & Murakami, 2001; Takimoto, Iwata,
& Murakami, 2009; Takimoto, Omoya Wata, & Asashi Urakami, 2002). Thus, it is important to examine the emergence patterns of aquatic insects to be able to (1) estimate the timing and (2) predict the biomass of this important energy and nutrient subsidy from different aquatic ecosystems.
In addition to being a source of cross-ecosystem subsidies, emerging aquatic insects also are a conduit for toxicants that can bioaccumulate during freshwater larval development (Chumchal &
Drenner, 2015; Menzie, 1980; Paetzold, Smith, Warren, & Maltby, 2011; Walters, Fritz, Otter, Luther, &
50
Drive, 2008). For example, mercury (Hg) in the freshwater environment is methylated to its more bio- available form, methylmercury (MeHg), which is a potent neurotoxin that biomagnifies through food webs (Wiener and Spry, 1996; Wiener et al. 2002). Aquatic insect larvae serve as an important link in the
MeHg pathway from their diet (e.g. sulfate reducing bacteria, diatoms, green algae) to larger aquatic predators (i.e. fish) and also terrestrial consumers after emergence (Becker et al., 2018; Kidd, Clayden, &
Jardine, 2012; Pennuto & Smith, 2015; Raikow, Walters, Fritz, & Mills, 2011; Speir et al., 2014; Williams et al., 2017). Emergent aquatic insects make up a substantial proportion of the diet of many riparian terrestrial consumers (Sabo and Power 2002; Sanzone et al. 2003; Speir et al. 2014; Pennuto and Smith
2015) and these feeding relationships are likely the primary mechanism by which MeHg is incorporated into terrestrial systems (Gerrard and Louis 2001; Cristol et al. 2008). Understanding spatial and temporal variation in aquatic insect emergence is therefore an important aspect of understanding and monitoring
Hg dynamics.
The transfer of Hg and other toxicants to terrestrial vertebrate predators (e.g., birds and bats) can occur through either direct consumption of aquatic insects or indirectly via consumption of prey that consume aquatic insects (e.g., riparian spiders). For example, spiders (Araneae) that obtain a large portion of their Hg burdens from emergent aquatic midges (Chaves-Ulloa et al., 2016; Ortega‐Rodriguez et al., 2019; Pennuto & Smith, 2015; Speir et al., 2014; Tweedy et al., 2013) can be a significant Hg source for their songbird predators (Cristol et al., 2008; Gann, Powell, Chumchal, & Drenner, 2015).
Evidence for direct consumption has been documented in Rusty blackbirds (Euphagus carolinus) that accumulated most of their Hg in summer months when they were actively feeding on high abundances of emerging invertebrates (Edmonds et al. 2010). The potential for both direct and indirect transfer of
MeHg from aquatic emergent insects to vertebrate predators motivates the need to study Hg burdens of a wide range of emergent aquatic insects that are potential prey for terrestrial consumers.
51
Aquatic insect emergence and their role as a vector of aquatic-derived MeHg to terrestrial environments has not yet been examined in northeastern, temperate streams where seasonality plays a significant role in insect phenology. We aimed to determine (1) taxonomic composition, (2) biomass, and (3) temporal trends of emerging aquatic insects in two coastal streams in Maine using weekly emergence trap collections. We also collected Tetragnathidae, a riparian aquatic insect predatory spider, on our emergence traps to estimate the biomass of intermediate terrestrial consumers that could form an important indirect pathway of Hg to riparian consumers. We then (4) analyzed the insect orders for MeHg concentrations each month to estimate temporal MeHg flux to the terrestrial environment from these two streams.
Methods
Study sites
This study was conducted in two streams of Acadia National Park, Maine, USA: Hunters Brook
44°18’ 20.7”N 68°13’18.6”W) and Marshall Brook 44°17’ 00.9”N 68°20’55.7”W) Figure 2.1., Figure .1,
Figure C.2.). Stream sampling reaches (~100m) selected were representative of their stream. The
Hunters Brook stream reach has a series of pools, riffles and backwater areas of cobble substrate whereas the Marshall Brook sampling reach contained slow moving to still water with high deposition of fine sediment (Figure C.1., Figure C.2.). Daily mean water temperatures were collected via Hobo pendant temperature loggers (Onset Computer) every two hours and were averaged daily from June to
November 2018. A temperature logger was deployed in Hunters Brook on June 14 and in Marshall Brook on July 18.
Emergent aquatic insect collection and preparation
Emergent aquatic insects were collected weekly at both streams from April 6th, 2018 until
November 9th, 2018. Six 1mm mesh cone-shaped floating emergence traps were installed within 100m
52 reach in of each stream and placed in various aquatic microhabitats that ensured representative sampling of emergence at each site. Each trap sampled a 0.5m X 0.5m area (0.25m2) and was held in place for the duration of the study with one or two 2cm diameter rebar stakes pounded into the sediment (Figure C.1, Figure C.2.). The traps funneled emerging insects into a collecting bottle containing
Hot-Shot Pest Strip® fumigant. Aspirators were used to collect any insects that were not funneled into the collection bottle but were inside the traps, and any riparian spiders that were on or within the traps.
All insects from each trap were contained in a single zipper-seal bag and frozen each week. Because of the heterogeneity of the streams and trap placements, not every week had collections from all six traps.
Each emergent aquatic insect and spider collected was identified to order (Merritt, Cummins, &
Berg, 2008) in the laboratory and photographed via a Nikon SMZ800 microscope equipped with a Zeiss
Axiocam ERC 5s camera. The Diptera, Ephemeroptera, Plecoptera and Araneae were identified to family; some Trichoptera were able to be identified to genus. Body length was measured to the nearest millimeter with the distance tool in Adobe Acrobat Pro. Dry weights of each emergent aquatic insect were calculated using length-weight regressions and are reported as biomass in milligrams (Klemmer,
2015; Sabo, Bastow, & Power, 2002; Stagliano, Benke, & Anderson, 1998).
After each insect was photographed, individuals were composited into vials by site and order per month and each vial’s content was analyzed for MeHg concentrations. Trichoptera from Marshall
Brook in July were separated into three different vials, Limnephilidae, Phryganeidae, and other (smaller)
Trichoptera taxa, each of which were analyzed separately for MeHg. Each composited vial was measured for wet weight, dried for 48 hours in a 50°C oven, and dry weight recorded at 24 and 48 hours. Samples were then shipped dry to the U.S. Geological Survey Forest and Rangeland Ecosystem Science Center
(USGS-FRESC) in Corvallis, Oregon for MeHg analysis.
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Methylmercury analysis
Prior to MeHg analysis, each emergent aquatic insect and spider vial was dried to verify desiccation and ground to a fine powder to homogenize tissues. Following EPA Method 1630 (U.S.
Environmental Protection Agency, 1998), each composite vial was analyzed for MeHg at the USGS
Contaminant Ecology Research Lab (Corvallis, OR) via cold-vapor atomic fluorescence spectrometry with a MERX-M automated methylmercury analyzer (Brooks Rand Instruments, Seattle, Washington, USA).
The quality assurance entailed certified reference materials (scallop tissue [IAEA-452; International
Atomic Energy Agency, Vienna, Austria], and lobster hepatopancreas [TORT-3; National Research Council of Canada, Ottawa, Canada]), instrument triplicates, liquid calibration standards, matrix spikes, matrix spike duplicates, and reagent blanks following USGS standard operating protocol (Willacker & Eagles-
Smith, 2017). Recoveries averaged 103.9% ± 7.414% (n=35) for certified reference materials.
Statistical analyses
All analyses were conducted using R v. 3.5.0 statistical software (R Core Team, 2019). High spring and late fall flows dislodged some traps and lowered sampling efficacy so we restricted analysis to the 20 collections from May 5, 2018 (Julian Day: 123) to September 21, 2018 (Julian day: 264).
Total biomass
Insect dry weights (mg), based on length–mass regressions, were summed by order for each of the six replicate traps per stream. Each biomass estimate was then log-transformed to reduce heteroscedasticity and normalize residuals. We tested whether biomass varied among orders and across streams by fitting a linear model with the log-transformed total biomass per trap as the response variable and stream and insect order and their interaction as fixed effects. We examined the differences
54 in total insect order biomass in traps between streams using analysis of variance (ANOVA) and Tukey post hoc test, with trap as the statistical unit.
Temporal biomass
Because there were different percentages of insect orders emerging in each stream, we analyzed them separately. To do so, insect dry weights (g) in each stream were totaled by order in each trap per month. When an order was not observed in a particular stream/order/trap/month the emergence value was 0. Variables were log (x+1)-transformed prior to analysis. We examined if there was a temporal difference in each insect order biomass over time within each stream using multiple contrasts of insect orders’ biomass within months and between months with a Tukey p-value adjustment.
Methylmercury
Across all sampled months, a total of 1,020 insects were composited by site and order, and analyzed for MeHg concentrations. Insect order MeHg concentrations are reported as geometric means in nanograms per gram dry weight (ng/g dw) with backtransformed geometric standard errors estimated using the delta method (B. K. Williams et al., 2002). We used stream and insect order as fixed effects in the linear model of MeHg concentrations. Next, we did a least square means comparison using
Tukey HSD post-hoc test to evaluate differences among specific orders.
Methylmercury flux
We estimated monthly MeHg flux of each taxa by multiplying the total emergent biomass (g) of each taxa by the concentration of MeHg in their tissues (ng/g dw). All MeHg flux data from all emergence traps (1.5m2) per site each month were multiplied by 2/3 to report as monthly Hg flux per meter squared (i.e. ng/g dw/month/m2)
55
Results
Water temperature
Water temperature in Hunters Brook ranged from 14°C to 21°C over the sampling period, with monthly averages of 14.5°C in June and 17.2°C in July. Water temperature at Hunters Brook was on average 2.2°C colder than Marshall Brook over the period from July 19 (logger deployment) until
September 21 (end of insect collections; Figure 3.1). Water temperatures peaked on August 8 with a water temperature of 24°C at Marshall Brook and 21°C at Hunters Brook. On September 11 (Julian date:
254), water temperatures started to converge between sites and they were similar temperature until
November (Julian date: 305)(Figure 3.1).
Figure 3.1. Water temperature (C) from the end of July to the middle of November in two streams, Hunters Brook and Marshall Brook, from Acadia National Park, Maine.
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Stream discharge
We did not instrument streams with discharge loggers, so we relied on a nearby stream gauge operated by the US Geological Survey (Otter Creek on Mount Desert Island, USGS: 01022840) as an index of small watershed discharge for Acadia National Park (Figure 3.2). Except for an unusual winter rainstorm in late January 2018, streamflow patterns were typical for a small, temperate forested stream watershed. Specifically, snowmelt during March and April led to peakflow (13.3 cfs) for 2018 (Figure
3.2). Low summer flows occurred from May through a minimum in August (0.170 cfs), with discharge again increasing during autumn rain in September through November (Figure 3.2).
Figure 3.2. Stream discharge (cfs) from January to December 2018, from Otter Creek in Acadia National Park,
Maine. Data source: USGS NWIS, Otter Creek on Mount Desert Island: 01022840.
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Taxonomic composition of insects
Adults collected during the May – September trapping period included the orders
Ephemeroptera (mayflies), Plecoptera (stoneflies), Trichoptera (caddisflies), Diptera (true flies), and
Araneae (spiders) (Table 3.1). The stoneflies contained the families Leuctridae and Chloroperlidae and the mayflies were composed of Siphlonuridae, Heptageniidae, and Baetidae (Table 3.1). Caddisflies included Limnephilidae, Hydropsychidae, Phryganeidae, and Polycentropodidae with several additional individuals that could not be identified to family (Table 3.1). The dipterans belonged to the families
Ceratopogonidae, Simullidae, Tipulidae, but the vast majority (> 95% from either stream) were
Chironomidae midges (Table 3.1). All but one spider collected in our traps were Tetragnathidae
(longjawed orbweavers) (Table 3.1).
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Table 3.1. Taxonomic composition of emergent insects and Araneae collected from two streams of Acadia National Park, Maine from May–September 2018. A = abundance percentage of emergent aquatic insects in each stream.
Insect Order Insect Family Hunters A (%) Marshall A (%) Brook (N) Brook (N)
Diptera 236 73.9% 487 89.7% Chironomidae 225 467 Ceratopogonidae 4 7 Simullidae 4 13 Tipulidae 2 0
Plecoptera 42 13.2% 3 0.552% Chloroperlidae 25 1 Leuctridae 17 2
Ephemeroptera 22 7.89% 6 1.10% Baetidae 15 4 Heptageniidae 4 2 Siphlonuridae 3 0
Trichoptera 19 5.95% 47 8.65% Hydropsychidae 5 14 Polycentropodidae 5 5 Limnephilidae Platycentropus 0 5 Unknown 0 5 Phryganidae Oligostomis 0 2 Unknown 9 16
Araneae Tetragnathidae 115 45 Pisauridae 0 1
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Emergent aquatic insect biomass
Marshall Brook had nearly double (1.84 times) the total biomass per unit area of aquatic insect emergence as Hunters Brook over the duration of the study (Table 3.2). The aquatic insect orders that contributed to this biomass differed between sites (Figure 3.3), which was supported by a significant order x site interaction term (F3,34 = 4.22, p = 0.012, Table 3.3). Plecoptera dominated (47.7%) the emerging biomass in Hunters Brook, with both Chloroperlidae and Leuctridae families being important components of this emergence (Table 3.1). In contrast, Trichoptera (77.5%) dominated biomass in
Marshall Brook (Table 3.2) and this was split across four families: Polycentropodidae, Hydropsychidae,
Limnephilidae and Phryganeidae (Table 3.1). Tukey HSD was supportive (p = 0.065) of a difference in the
Trichoptera biomass between Hunters Brook and Marshall Brook (Tukey HSD, p = 0.065). Diptera emergence as a percentage of total aquatic insect biomass was similar in each stream but there was greater abundance of Ephemeroptera in Hunters Brook as compared to Marshall Brook (Table 3.2).
Table 3.2. Biomass (mg dry mass/m2) for aquatic insect orders from each stream with all trap data combined. N = number of individual insects collected and % = the percent biomass of each order from the total biomass emerged from the stream
Insect Order N Hunters % N Marshall Brook % Brook total biomass total biomass (mg/m2) (mg/m2) Diptera 239 19.6 18.6 488 32.4 16.7 Ephemeroptera 22 20.0 19.0 6 6.41 3.31 Plecoptera 42 50.1 47.7 3 4.75 2.46 Trichoptera 19 15.4 14.6 47 150 77.5 Total biomass 105.1 193.56 (mg/m2)
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Figure 3.3. The distributions of total biomass of each aquatic insect order collected in each of six traps from Hunters Brook and Marshall Brook, Acadia National Park, Maine from May-September 2018. There were no significant differences in biomass among orders in either stream.
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Table 3.3. Statistical results (degrees of freedom
(Df), F statistics, and p-values) from ANOVAs testing
for main effects of stream and order and a stream
and order interaction effect on log total biomass by
insect order.
Effects Df F p-value Stream 1, 34 1.45 0.237 Order 3, 34 0.268 0.848 Stream:Order 3, 34 4.22 0.012
Temporal emergence of insect orders between months
In Marshall Brook, we caught emerging insects in our traps earlier in May and later in September than at Hunters Brook (Figure 3.4), and the largest emergence biomass was in July within both streams
(Figure 3.5). Multiple comparisons indicated significant differences in biomasses of emerging aquatic insect orders over time within each stream (Table 3.4, 3.5). In Hunters Brook, there was steady emergence of Diptera and Plecoptera across the sampling period, but Plecoptera emergence transitioned from Chloroperlidae in May to July to Leuctridae from July to September (Table 3.5, Figure
3.4, 3.6). By contrast, Diptera emergence in Marshall Brook increased 3.7-fold from June to August but no temporal differences were detected among Plecoptera, largely because only three individuals were captured in traps at the end of May (Figure 3.4, Table 3.4). There was a significant peak in
Ephemeroptera emergence in Hunters Brook in July (Figure 3.4, Table 3.5) encompassing three families,
Heptageniidae, Siphlonuridae, and Baetidae. Some Baetidae mayflies also emerged later in September
(Figure 3.4). However, in Marshall Brook, there were no temporal differences in Ephemeroptera biomass
(Table 3.4), and only six individuals were captured consisting of Heptageniidae in May and Baetidae in
September. Trichoptera emerged from Hunters Brook in July and August and were not observed in other
62 months (Figure 3.4, 3.5, Table 3.5). Trichoptera biomass was significantly different among months in
Marshall Brook, with an increase from May to July and decrease from July to September but the emergence throughout the study varied across the four Trichoptera taxa (Table 3.4, Figure 3.4, 3.7).
Polycentropodidae emerged first in June followed by Hydropsychidae later in June and into July (Figure
3.7). The large Phryganeids and Limnephilids emerged in July and some unknown Trichoptera emerged in July through September (Figure 3.7).
Figure 3.4. The distributions of total biomass (mg/m2/week) of each aquatic insect order (Diptera,
Ephemeroptera, Plecoptera, Trichoptera) per week collected in emergence traps from Hunters Brook and Marshall Brook, Acadia National Park, Maine from May–September 2018.
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Figure 3.5. Total biomass (mg/m2/month) of aquatic insect orders per month in Hunters Brook and
Marshall Brook.
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Table 3.4. Significant statistical results (degrees of freedom (df), T-ratio, and p-values) from multiple comparisons testing for insect order biomasses differences within months and insect order biomass differences among months from Marshall Brook. P-value adjustment using Tukey method.
Marshall Brook Response Comparisons Df T-ratio p-value (Biomass) June Diptera - Trichoptera 100 -3.32 0.007 Ephemeroptera - Trichoptera 100 -4.12 <0.001 Plecoptera - Trichoptera 100 -4.12 <0.001 July Diptera - Trichoptera 100 -5.49 <0.001 Ephemeroptera - Trichoptera 100 -7.00 <0.001 Plecoptera - Trichoptera 100 -7.00 <0.001 August Diptera - Ephemeroptera 100 3.94 <0.001 Diptera - Plecoptera 100 3.94 <0.001 Diptera June - August 100 -3.14 0.018 Trichoptera May - June 100 -4.12 <0.001 May - July 100 -7.00 <0.001 June -July 100 -2.88 0.038 June - September 100 3.53 0.006 July - August 100 5.52 <0.001 July - September 100 6.41 <0.001
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Table 3. 5. Significant statistical results (degrees of freedom (df), T-ratio, and p-values) from multiple comparisons testing for insect order biomasses differences within months and insect order biomass differences among months from Hunters Brook. P-value adjustment using Tukey method.
Hunters Brook Response Comparisons Df T-ratio p-value (Biomass) June Ephemeroptera - Plecoptera 100 -2.73 0.036 Plecoptera - Tricoptera 100 2.73 0.036 September Plecoptera - Tricoptera 100 2.69 0.041 Ephemeroptera August - July 100 -4.26 <0.001 July - May 100 4.26 0.001 July - June 100 4.26 <0.001 Trichoptera July - June 100 3.47 0.007 July - May 100 3.47 0.007 July - September 100 3.47 0.007
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Figure 3.6. The distributions of total biomass (mg/m2/week) of Plecoptera collected in six traps weekly from Hunters Brook, Acadia National Park, Maine from May-September 2018.
67
Figure 3.7. The distributions of total biomass (mg/m2/week) of Trichoptera collected in six traps weekly from Marshall Brook, Acadia National Park, Maine from May-September 2018.
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Multiple comparisons also revealed significant differences in biomass among insect orders within specific months in each stream (Tables 3.4, 3.5). In Marshall Brook, peak Trichoptera biomass during June and July was significantly higher than all other orders (Figure 3.4, 3.5, Table 3.4).
Additionally, there was very little emergence of Plecoptera and Ephemeroptera in August and thus, there was significantly greater biomass of Diptera in that month (Figure 3.4, Table 3.4). In Hunters Brook in June, Plecoptera had greater biomass than Ephemeroptera and Tricoptera, but not Diptera (Table
3.5). Additionally, in Hunters Brook in September, there was no emergence of Trichoptera and thus
Plecoptera biomass was 4.6x fold higher than Tricoptera (Table 3.5).
Araneae frequently inhabited the traps at Hunters Brook during June, July and August, but did not appear in our traps until August through September at Marshall Brook (Figure 3.8). Araneae biomass was significantly greater in July at Hunters Brook compared to any other month and was significantly greater in September than May, June, and July at Marshall Brook (Table 3.6).
Figure 3.8. The distributions of total biomass (mg/m2/week) of Tetragnathids per week collected in six traps from Hunters Brook and Marshall Brook, Acadia National Park, Maine from May-September 2018.
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Table 3.6. Significant statistical results (degrees of freedom (df), T-ratio, and
p-values) from multiple comparisons testing for Araneae biomasses
differences among months from both streams. P-value adjustment using
Tukey method.
Stream Comparisons Df T-ratio p-value Marshall Brook May - September 25 -3.05 0.039 June – September 25 -3.05 0.039 July – September 25 -3.05 0.039
Hunters Brook July – May 100 5.60 <0.001 July – June 100 4.27 <0.001 July - August 100 3.49 0.006 July - September 100 3.56 0.005
Methylmercury concentrations in insects
Our model with stream and insect order as fixed effects explained 44% of the variance of MeHg in insects (p = 0.003). Methylmercury concentrations in insects were significantly different among orders
(F4,28 = 5.48, p = 0.002) but not between streams (F1,28 = 0.681, p = 0.416) (Figure 3.9). With both streams combined, MeHg in aquatic emergent insects ranged from 73.8 to 1007 ng/g dw (Table 3.7). The highest geometric mean and standard error was in Trichoptera (338 ± 80.5 ng/g) and the lowest geometric mean was in Plecoptera (142 ± 26.7 ng/g) (Table 3.7, Figure 3.9B). Methylmercury concentrations in
Plecoptera of Hunters Brook and Trichoptera of Marshall Brook were highly taxa driven as we observed differences in MeHg concentrations between families within each order (Appendix C). Diptera had a geometric mean of 184 ± 18.7 ng/g dw and Ephemeroptera had a geometric mean of 225 ± 47.3 ng/g dw
(Table 3.7). Methylmercury concentrations in long-jawed orb weaver spiders were greater than MeHg concentrations in any emergent insect order, ranging from 363–644 ng/g dw with a mean of 498 ± 48.9
70 ng/g dw (Table 3.7). When controlling for the effect of stream, Araneae MeHg concentrations were 2.7x and 3.5x fold higher than Diptera and Plecoptera, respectively (Table 3.7, Figure 3.9B). Additionally,
Trichoptera MeHg concentrations were 2.4x fold higher than Plecoptera (Tukey HSD, p = 0.038, Table
3.8) (Figure 3.9B).
Figure 3.9. A) Total insect MeHg concentrations between Hunters Brook and Marshall Brook and B)
MeHg concentrations among insect orders while accounting for site.
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Table 3.7. Summary statistics of MeHg concentration data for insect orders from Hunters Brook and Marshall Brook, Acadia National Park, Maine. N = the number of composted insect vials that were analyzed for MeHg. Mean = geometric mean. SE = calculated as geometric standard error.
Insect Order N Hunters SE N Marshall SE N Stream SE Brook Brook combined mean mean mean MeHg MeHg MeHg (ng/g dw) (ng/g dw) (ng/g dw) Araneae 4 571 ± 40.2 2 379 ± 16.0 6 498 ± 48.9 Diptera 4 225 ± 30.8 5 156 ± 16.4 9 184 ± 18.7 Ephemeroptera 2 320 ± 26.9 2 158 ± 16.6 4 225 ± 47.3 Plecoptera 6 159 ± 28.5 1 73.8 - 7 142 ± 26.7 Trichoptera 2 280 ± 35.9 6 358 ± 130 8 337 ± 90.8 MeHg site mean 263 ± 34.8 227 ± 42.0
Table 3.8. Tukey HSD output comparing MeHg concentrations means by insect order.
Araneae Diptera Ephemeroptera Plecoptera Diptera 0.013 Ephemeroptera 0.141 0.989 Plecoptera 0.009 0.995 0.942 Trichoptera 0.907 0.061 0.404 0.039
Methylmercury flux via emergent aquatic insects
The total MeHg flux (ng/m2/month) in emergent insects from Marshall Brook was 1.7 times that from Hunters Brook (Table 3.9). Trichoptera contributed the greatest (~83%) MeHg flux out of all aquatic emergent insect orders collected at Marshall Brook (Table 3.9, Figure 3.10). In contrast, MeHg flux was more evenly spread across insect orders at Hunters Brook, with the greatest contributions from
72
Plecoptera (~33%) throughout the study period (Table 3.9, Figure 3.9). Monthly patterns of aquatic insect-mediated MeHg flux corresponded to changes in emerging insect biomass over time (Figure 3.5).
Table 3.9. Summary statistics of total MeHg flux data for insect orders from
Hunters Brook and Marshall Brook, Acadia National Park, Maine.
Insect Order Hunters Brook Marshall Brook MeHg flux MeHg flux (ng/m2/month) (ng/m2/month) Diptera 4.23 4.97 Ephemeroptera 6.70 1.05 Plecoptera 7.54 0.351 Trichoptera 4.68 31.9 Total MeHg 23.2 38.3 (ng/m2/month)
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Figure 3.10. Emergent aquatic insect total MeHg flux (ng/m2/month) from two streams, Hunters Brook and Marshall Brook, in Acadia National Park.
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Discussion
Emergence in Hunters Brook and Marshall Brook was dominated by taxa typical of coastal temperate Maine streams: Diptera, Ephemeroptera, Plecoptera, Trichoptera, and Araneae. Emergence biomass was more evenly spread across taxa at Hunters Brook, while Trichoptera dominated emergence from Marshall Brook throughout the study period, which was a reflection of each stream’s habitat and hydrologic characteristics. We observed a change in aquatic insect taxa biomass emerging throughout the study period with peak emergence occurring in July. We measured insect MeHg concentrations per order per month per stream; results indicated that trophic level and diet were likely important determinants of MeHg concentrations within emerging insects. Lastly, we describe important aquatic insect taxa that contribute to MeHg flux to the terrestrial environment within these streams.
Taxonomic composition and biomass differences between stream reaches
Our two study streams are located on Mount Desert Island, Maine, experience the same temperate seasonal variation, and have similar watershed characteristics including size and forest type.
We collected taxa in both streams that are typical for northeastern U.S. streams: Diptera,
Ephemeroptera, Plectoptera and Trichoptera (Breen, 2004; Cheney, Roy, Smith, & Dewalt, 2019;
Giberson & Garnett, 1996; Maine Department of Environmental Protection, 2016, 2017). For example,
Trichoptera, Ephemeroptera, and Diptera were the most dominant benthic aquatic insect taxa in
Marshall Brook during previous biomonitoring surveys (Maine Department of Environmental Protection,
2016). While we observed Trichoptera and Diptera emerging from Marshall Brook, the lack of
Ephemeroptera was puzzling, given previous monitoring that concluded that Maccaffertium
(Heptageniidae) mayflies were the second most abundant aquatic benthic taxa within this stream
(Maine Department of Environmental Protection, 2016). Likewise, Diptera, Plecoptera, and Trichoptera were the most dominant benthic aquatic insect taxa in surveys of Hunters Brook (Maine Department of
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Environmental Protection, 2016) and while we observed these taxa in our emergence, we also had detectable emergence of Ephemeroptera. Thus benthic survey data may not be indicative of the taxonomic composition of emerging aquatic insects, and additional benthic survey methods coupled with other sampling techniques for emergence, including malaise traps, may produce a more complete picture.
We detected considerable differences in biomass of emerging aquatic insect orders between streams that could be explained by differences in our stream reach habitat. Based on our monthly observations during the course of the preceding year, there is slower discharge, water is nearly stagnant, and there is more detritus and woody debris in our Marshall Brook stream reach (Appendix C) which may explain why aquatic insect emergence biomass was dominated by Trichoptera as they are known to emerge in slower discharge habitats and graze on detritus (Castro-Rebolledo & Donato-
Rondon, 2015). Based on monthly observations during the preceding year, Hunters Brook has greater discharge and colder average water temperature than Marshall Brook (Figure 3.1), and the Hunters
Brook stream bottom is a series of riffles, runs, and pools that are mostly cobblestone substrate (Maine
Department of Environmental Protection, 2017). Course substrate is preferred by many larval stream insects (Roy, Rosemond, Leigh, Paul, & Wallace, 2003), which may be why we observed that emergence was distributed among insect orders at Hunters Brook.
Temporal variation of insect biomass within streams
Seasonal differences in hydrological regime of temperate streams dictate temporal patterns in insect emergence. More specifically, water temperature and flow variation change throughout the seasons and can influence aquatic insect development and emergence typically via accumulated degree days (Anderson, Albertson, & Walters, 2019; Castro-Rebolledo & Donato-Rondon, 2015; Chumchal et al.,
2018; Gillooly, harnov, West, Savage, & Brown, 2002; Ivković, Miliša, Previšić, Popijač, & Mihaljević,
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2013; Jannot, 2009). The largest biomass emergence of aquatic insects was in July (Figure 5) and there was roughly a 3°C increase in average water temperature from June to July (Figure 3.1), which is consistent with water temperature thresholds that trigger emergence. Lower flows that occurred synchronously with the accumulated degree day thresholds are also known to initiate emergence for some aquatic insect taxa, like Trichoptera (Castro-Rebolledo & Donato-Rondon, 2015).
Insect taxa often vary in their cues for emergence and therefore it is common to observe different insect orders emerging at different times of the year. Dipterans emerged throughout the study period in both streams (Figure 3.4, 3.5), which is not surprising, as they are a very diverse order of aquatic insects and can withstand nearly any water condition including varied temperature, flow, and dissolved oxygen content (Milner, Brittain, Castella, & Petts, 2001; Pritchard, 1991). In contrast,
Plecoptera emergence in Hunters Brook transitioned from Chloroperlidae emerging in spring to
Leuctridae later in the summer (Figure 3.6), which is similar to observations in other studies (Giberson &
Garnett, 1996; Harper & Pilon, 1970; López-Rodríguez, Tierno De Figueroa, Bo, Mogni, & Fenoglio,
2012). This patterns is consistent with voltinism differences among taxa; univoltine Leuctridae could require warmer temperatures in the summer to complete development (López-Rodríguez et al., 2012) compared to semivoltine Chloroperlidae that emerge after overwintering as late instar nymphs (Tierno
De Figueroa & López-Rodríguez, 2019). Trichoptera emergence peaked in July at both streams regardless of a 3°C water temperature difference between sites (Figure 3.4, 3.5, Table 3.4, 3.5). Similar patterns of
Trichoptera emergence were found in two streams of lower Michigan with date shown to be a greater predictor of 27 Trichoptera species emergence than water temperature (Houghton, 2015). This can be indicative of photoperiod and low summer flows driving Trichoptera emergence timing (Figure 3.4).
There were less clear patterns of emergence of Ephemeroptera in our streams and because of the low emergence of Ephemeroptera in Marshall Brook, not many conclusions can be made.
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Methylmercury concentrations in insect orders
To our knowledge, we are among the first to examine MeHg concentrations from emergent aquatic insects from natural stream habitats in the northeastern US. We observed differences in MeHg concentrations among insect orders (Figure 3.9B), which likely reflect trophic position, diet, and feeding modes. First, trophic position influences MeHg concentrations in insects by controlling the potential for biomagnification. For example, Araneae had significantly greater MeHg concentrations than Diptera and
Plecoptera, which we expected because these riparian spiders feed heavily on emergent aquatic insects and MeHg biomagnifies with trophic level (Chaves-Ulloa et al., 2016; Krell et al., 2015; Ortega‐Rodriguez et al., 2019; Pennuto & Smith, 2015; Tsui et al., 2012). MeHg concentrations in shoreline spiders were 2 to 5 times greater than MeHg concentrations in dipterans in studies from Texas and New York (Pennuto
& Smith, 2015; Speir et al., 2014; Tweedy et al., 2013), which is similar to what we found in our study.
Second, diet, and thus carbon source, has been shown to be an important factor that influences
MeHg concentrations within aquatic insect larvae (Jardine et al., 2012; Riva-Murray et al., 2013).
Trichoptera had the highest mean and variation in MeHg concentration of all emergent aquatic insects in our study (Figure 9B) and this taxon was dominated by omnivorous Hydropsychidae (net spinning caddisfly) and detritivorous Limnephilidae (Table 3.1, Figure C.3.). Hydropsychidae larvae had consistently higher MeHg concentrations compared to larval Limnephilidae, which was a reflection of greater nitrogen isotope ratios (15N) and lower carbon isotope ratio (13C) in hydropsychids compared to limnephilids in three streams of New York (Riva-Murray et al., 2013). Additionally within the Plecoptera,
Leuctridae had significantly lower MeHg concentrations than Chloroperlidae (Figure C.2.). Larval
Chloroperlidae (green stoneflies) are mostly predatory or omnivorous (Tierno De Figueroa & López-
Rodríguez, 2019; Zwick, 1980) whereas Leuctridae (rolled-winged stoneflies) are mostly detritivorous collector-gatherers also known to feed on algae (López-Rodríguez et al., 2012; Tierno De Figueroa &
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López-Rodríguez, 2019). These dietary differences are consistent with differences in MeHg concentrations among these families, and speaks to a broader role of carbon source in determining
MeHg concentrations in emerging aquatic insects.
These familial differences in MeHg concentrations were also reflected in ordinal MeHg.
Trichoptera had significantly greater MeHg than Plecoptera (Figure 3.9B). Hydropsychids had the greatest MeHg concentrations among Trichoptera and greatly increased the mean MeHg for this order.
Additionally, nearly half of our Plecoptera MeHg samples consisted of primary consumer Leuctridae which could contribute to significantly lower MeHg in Plecoptera. There were no other differences in
MeHg concentrations among orders, perhaps because Ephemeroptera, Diptera, and Plecoptera are all lower trophic levels than Trichoptera. However, we might have observed a difference with a larger
MeHg sample size of Ephemeroptera as we only had two MeHg samples of these orders within streams.
Besides one study in New Hampshire, the MeHg concentrations from Trichoptera, Diptera,
Ephemeroptera and Araneae collected from our natural streams were higher than MeHg reported for these adult taxa elsewhere (Tweedy et al. 2013; Pennuto and Smith 2015; Chumchal et al. 2017, 2018;
Ortega‐Rodriguez et al. 2019; Table 3.10). For example in our study, we observed MeHg concentrations in Trichoptera ranging from 112 to 1,007 ng/g dw in our two study streams with an average of 429 ng/g dw, whereas, the average MeHg concentrations in caddisflies from Texas experimental ponds were 58.2 ng/g dw in micro-caddisflies and 63.0 ng/g dw in long-horned caddisflies (Chumchal et al., 2017). Our
Diptera MeHg concentrations (range: 127–247 ng/g dw) were greater than Diptera from experimental ponds in Texas (range: 40–79 ng/g dw) (Chumchal et al., 2017, 2018) and THg concentrations in midges from the Buffalo River Area of Concern, classified by the US EPA, in New York (range: ~ 75–100 ng/g dw)
(Pennuto & Smith, 2015). Additionally, our long-jawed orb weavers MeHg concentrations are roughly 3 times greater than long-jawed orbweavers MeHg concentrations from Texas experimental ponds and
79
THg concentrations in spiders the Buffalo River Area of Concern in New York Ortega‐Rodriguez et al.,
2019; Pennuto & Smith, 2015) but much lower than long-jawed orbweavers’ from streams in New
Hampshire (Chaves-Ulloa et al., 2016). Our Ephemeroptera MeHg concentrations ranged from 152–348 ng/g dw, whereas the average MeHg concentration Ephemeropta from Texas was 68.1 ng/g dw. These elevated MeHg concentrations may be a product of long-range transport and atmospheric deposition patterns (Downs, Macleod, & Lester, 1998; Fitzgerald, William, Engstrom, Mason, & Nater, 1998; Morel,
Kraepiel, & Amyot, 1998). Mercury concentrations are commonly elevated in forested streams and biota of the eastern U.S. (Bank et al., 2007; Driscoll et al., 2007; Riva-Murray et al., 2011), which could explain high MeHg concentration in our Maine study and in New Hampshire streams (Chaves-Ulloa et al.,
2016)(Table 3.10). Additional research would be helpful in comparing MeHg flux from aquatic insect taxa from other natural stream settings in other parts of the United States, including the Northeast.
Table 3.10. Comparing MeHg concentrations with other adult insects from other studies.
Study Our study Chaves-Ulloa Pennuto Chumchal Gann et Ortega-Rodriguez et al. 2016, and et al. 2017 al. 2015 et al. 2019 Ave. MeHg Appendix 4 Smith (Ave. (estimated MeHg ng/g dw (Mean MeHg 2015 MeHg ng/g wet concentration in (range) ng/g*) (THg ng/g dw) weight whole body ng/g ng/g dw) dw) State Maine New New Texas Texas Texas Hampshire York Diptera 142 ± 90.8 67.4-393.3 ~75-100 40.1-79.4 39.1 - 67.3 (127-318) Ephemeroptera 184 ± 18.7 112.4-332.9 68.1 (142-348) Trichoptera 225 ± 47.3 106.5-473.5 58.2-63.0 85.7 ± 17.0 (112-1007) Plecoptera 142 ± 26.7 62.8-125.2 (73.8-286) Araneae 498 ± 48.9 ~200-2,300 ~160- 19.4 - 159 ± 14.8 (363-644) 260 256 *Dry weight or wet weight basis for concentrations was not specified in the paper
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Methylmercury flux from emergent aquatic insects
While there were taxonomic differences in biomass and concentration, Hg flux was consistent and relatively high. Our Diptera, comprised mostly of chironomid midges, had a steady emergence from both sites throughout the year with similar MeHg concentrations between sites. This could translate to a steady ‘baseline’ flux of MeHg throughout the year in both sites of ~4-5 ng/m2/month within each stream (Table 3.9). In Texas, dipteran-mediated MeHg flux increased with water temperature throughout the summer (Chumchal et al., 2018). We did see an increase of Diptera emergence from
June to August in Marshall Brook which may be because of increase in water temperature. However, we did not observe this trend in Hunters Brook as there was no temporal difference in emergence, possibly because we were observing Diptera in a greater discharge stream where emergence could also be influenced by water velocity (Castro-Rebolledo & Donato-Rondon, 2015). Regardless, Dipterans provide a low but constant flux of Hg to terrestrial predators throughout May–September in these streams.
Alternatively, within both streams, MeHg flux was greatest in July which was contributed via
Trichoptera, Plecoptera, and Ephemeroptera (Figure 3.10).
Our study indicates some high MeHg concentrations in Trichoptera, especially small-bodied taxa, suggesting that caddisflies may be overlooked as important components of MeHg flux to the riparian environment. For example, previous studies have focused on large-bodied aquatic insects, like
Odonates (dragonflies and damselflies), as acting as the greatest MeHg flux (Chumchal & Drenner,
2015). We did not catch any odonates in our traps; however, dragonfly nymph MeHg concentrations in this stream range from 266 to 413 ng/g dw (Hess et al. 2019 in prep), and MeHg concentrations are not significantly different from nymph to adult dragonflies (Buckland-Nicks et al., 2014). Therefore, dragonfly MeHg flux may be estimated between 266–413 ng/g dw in Marshall Brook whereas
Trichoptera MeHg concentrations range from 112–1007 ng/g dw in Marshall Brook with a mean of 478 ng/g dw. Likewise, Trichoptera was the least often captured insect order in Hunters Brook (Table 3.1)
81 but they contributed roughly the same amount of MeHg flux in two months as Dipterans did throughout the study period (Table 3.9). This indicates that Trichoptera, while not the most abundant, can nonetheless be a large component of aquatic insect-mediated MeHg flux in these streams.
MeHg flux over time correlated with patterns in emergence biomass (Figure 3.5); however,
MeHg concentrations were dependent on insect order. Therefore, emergence biomass alone does not yield an accurate prediction of MeHg flux. For example in Marshall Brook, large bodied Trichoptera
(Limnephilidae and Phryganeidae) generated high emergence biomass but contained lower MeHg concentrations, whereas smaller-bodied Trichoptera (Hydropsychidae and Polycentropodidae) contributed less biomass to Trichoptera emergence but contained nearly double the amount of MeHg
(Figure C.1.). Likewise, in Hunters Brook, there was about half the abundance of Ephemeroptera emerging than Plecoptera throughout the study period (Table 3.1), but Ephemeroptera contributed a similar amount to total MeHg flux as Plecoptera (Figure 3.10, Table 3.9). These differences also occur within orders. For example, Plecoptera were abundant in Hunters Brook and their MeHg concentrations ranged from 101–286 ng/g dw (Figure 3.9B, Figure C.4.). However, spring emerging Chloroperlidae had roughly 2x higher MeHg concentrations than Leutricidae (Figure C.4.) and generated a higher pulse of
Plecoptera MeHg flux during this time period compared to later in the summer (Figure 3.10). These examples highlight the limitations of using body mass and emergence biomass to estimate MeHg flux in aquatic insects, and the importance of acknowledging taxonomic differences in trophic level and diet that can mediate MeHg concentrations.
Lastly, the Tetragnathid riparian spiders inhabited our emergence traps at different times throughout the study. Spiders were more prominent in July at Hunters Brook and September at Marshall
Brook (Figure 3.8). Between streams, this could indicate different times when emergent aquatic insects were being consumed via riparian spiders, and alther the timing of subsequent transfer of MeHg from
82 the aquatic environment. There was a positive correlation between MeHg in riparian spiders and MeHg in emerging midges in Texas experimental ponds (Speir et al., 2014; Tweedy et al., 2013). The differences in timing of peak abundance of spiders between streams also indicates different times when riparian spiders were available to terrestrial predators (e.g., birds). In riverine habitats, MeHg concentrations in long-jawed orb weaver spiders (Tetragnatha sp.) collected in Caddo Lake on the border of Texas and Louisiana exceeded wildlife values for Carolina chickadee (Peocile carolinensis)
(Gann et al., 2015). Therefore, Tetragnathidae may decrease MeHg flux via emergent aquatic insects to other terrestrial predators but may also be a large food source themselves to terrestrial predators during July in Hunters Brook and September in Marshall Brook.
Conclusion
Our aim was to provide robust measurements of energy (i.e. insect biomass) and MeHg flux to the terrestrial environment and examine if there were any differences between the two streams. These two streams are similar in many ways but the different stream reaches harbored different aquatic insect communities. Thus, we found differences in cross-ecosystem subsidies measured as biomass, and in temporal flux of MeHg with respect to aquatic insect emergence. These differences inform which organisms provided the largest MeHg exposure to consumers. We found that Trichoptera and
Plecoptera played a larger role in MeHg flux relative to findings of other studies. We also investigated the timing of MeHg pulses to the terrestrial environment in stream systems, finding that the greatest
MeHg flux was in July via greater biomass of emerging aquatic insects. Additionally, we saw temporal differences in the presence of riparian spiders between the two streams, indicating times when MeHg uptake to spiders via insects may be high, and also spider abundance could be greater for other terrestrial prey that feed upon them. Lastly, this study indicated that remote protected natural streams, such as those in Acadia National Park, can yield high MeHg concentrations in biota and thus, large
83 amounts of MeHg flux to the terrestrial environment and predators. Therefore, additional studies from natural streams with high sampling frequency (i.e. at least weekly) should be completed to provide robust measurements of MeHg flux via emergent aquatic insects in other locations in the U.S., not just known contaminated sites. Moreover, emergence studies that are coupled with various terrestrial predator Hg concentrations and Hg concentrations within specific diets (i.e. what insect order and how many insects are being consumed) would be important in examining the impacts of insect mediated
MeHg flux in natural systems.
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BIBLIOGRAPHY
Allen, E. W., Prepas, E. E., Gabos, S., Strachan, W. M. J., & Zhang, W. (2005). Methyl mercury concentrations in macroinvertebrates and fish from burned and undisturbed lakes on the Boreal Plain. Canadian Journal of Fisheries and Aquatic Sciences, 62(9), 1963–1977. https://doi.org/10.1139/f05-103
Anderson, H. E., Albertson, L. K., & Walters, D. M. (2019). Thermal variability drives synchronicity of an aquatic insect resource pulse. Ecosphere, 10(8), 1–11. https://doi.org/10.1002/ecs2.2852
Arribére, M. A., Campbell, L. M., Rizzo, A. P., Arcagni, M., Revenga, J., & Guevara, S. R. (2010). Trace Elements in Plankton, Benthic Organisms, and Forage Fish of Lake Moreno, Northern Patagonia, Argentina. Water, Air, & Soil Pollution, 212, 167–182. https://doi.org/10.1007/s11270-010-0330-3
Bank, M. S., Burgess, J. R., Evers, D. C., & Loftin, C. S. (2007). Mercury Contamination of Biota from Acadia National Park, Maine: A Review. Environmental Monitoring and Assessment, 126(1–3), 105– 115. https://doi.org/10.1007/s10661-006-9324-4
Barros Miguel, T., Max, J., Oliveira-Junior, B., Ligeiro, R., & Juen, L. (2017). Odonata (Insecta) as a tool for the biomonitoring of environmental quality. Ecological Indicators, 81, 555–566. https://doi.org/10.1016/j.ecolind.2017.06.010
Bates, L. M., & Hall, B. D. (2012). Concentrations of methylmercury in invertebrates from wetlands of the Prairie Pothole Region of North America. Environmental Pollution, 160, 153–160. https://doi.org/10.1016/j.envpol.2011.08.040
Baxter, C. V., Fausch, K. D., & Saunders, W. C. (2005). Tangled webs: Reciprocal flows of invertebrate prey link streams and riparian zones. Freshwater Biology, 50(2), 201–220. https://doi.org/10.1111/j.1365-2427.2004.01328.x
Becker, D. J., Chumchal, M. M., Broders, H. G., Korstian, J. M., lare, E. L., Rainwater, T. R., … Fenton, M. B. (2018). Mercury bioaccumulation in bats reflects dietary connectivity to aquatic food webs *. Environmental Pollution, 233, 1076–1085. https://doi.org/10.1016/j.envpol.2017.10.010
Beeby, A. (2001). What do sentinels stand for? Environmental Pollution, 112, 285–298. https://doi.org/10.1016/S0269-7491(00)00038-5
Beganyi, S. R., & Batzer, D. P. (2011). Wildfire induced changes in aquatic invertebrate communities and mercury bioaccumulation in the Okefenokee Swamp. Hydrobiologia, 669(1), 237–247. https://doi.org/10.1007/s10750-011-0694-4
Blancher, P. J., & McNicol, D. K. (1991). Tree swallow diet in relation to wetland acidity. Canadian Journal of Zoology, 69(10), 2629–2637. https://doi.org/10.1139/z91-370
Breen, R. M. (2004). Benthic Stream Macroinvertebrate Monitoring Report: Acadia National Park 2000- 2002.
85
Brigham, M. E., Wentz, D. A., Aiken, G. R., & Krabbenhoft, D. P. (2009). Mercury Cycling in Stream Ecosystems. 1 . Water Column Chemistry and Transport. Environmental Science and Technology, 43(8), 2720–2725. https://doi.org/10.1021/es802694n
Buckland-Nicks, A., Hillier, K. N., Avery, T. S., & O’Driscoll, N. J. 2014). Mercury bioaccumulation in dragonflies (Odonata: Anisoptera): Examination of life stages and body regions. Environmental Toxicology and Chemistry, 33(9), 2047–2054. https://doi.org/10.1002/etc.2653
Buckman, K. L., Seelen, E. A., Mason, R. P., Balcom, P., Taylor, V. F., Ward, J. E., & Chen, C. Y. (2019). Sediment organic carbon and temperature effects on methylmercury concentration: A mesocosm experiment. Science of the Total Environment, 666, 1316–1326. https://doi.org/10.1016/j.scitotenv.2019.02.302
Bulankova, E. (1997). Dragonflies (Odonata) As bioindicators of environment quality. Biologia, 52(2), 177–180.
Burcher, C. L., & Smock, L. A. (2002). Habitat Distribution, Dietary Composition and Life History Characteristics of Odonate Nymphs in a Blackwater Coastal Plain Stream Characteristics of Odonate Nymphs in a Blackwater Coastal Plain Stream. Source: The American Midland Naturalist, 148(1), 75–89. Retrieved from http://www.jstor.org/stable/3082962
Burgess, J. R. (1997). Mercury contamination in fishes of Mount Desert Island: A comparative food chain mercury study. Unvierstiy of Maine, Orono, Maine, USA.
Camci-Unal, G., & Pohl, N. L. B. (2010). Quantitative Determination of Heavy Metal Contaminant Complexation by the Carbohydrate Polymer Chitin. Journal of Chemical & Engineering Data, 55(3), 1117–1121. https://doi.org/10.1021/je900552w
Castro-Rebolledo, M. I., & Donato-Rondon, J. C. (2015). Emergence patterns in tropical insects: the role of water discharge frequency in an Andean Stream. International Journal of Limnology, 51, 147– 155. https://doi.org/10.1051/limn/2015011
Chaves-Ulloa, R., Taylor, B. W., Broadley, H. J., ottingham, K. L., Baer, N. A., Weathers, K. ., … hen, . Y. (2016). Dissolved organic carbon modulates mercury concentrations in insect subsidies from streams to terrestrial consumers HHS Public Access. Ecological Applications, 26(6), 1771–1784. https://doi.org/10.1890/15-0025.1
Cheney, K. N., Roy, A. H., Smith, R. F., & Dewalt, R. E. (2019). Effects of Stream Temperature and Substrate Type on Emergence Patterns of Plecoptera and Trichoptera From Northeastern United States Headwater Streams. Environmental Entomology, XX(XX), 1–11. https://doi.org/doi: 10.1093/ee/nvz106
Chételat, J., Amyot, M., & Garcia, E. (2011). Habitat-specific bioaccumulation of methylmercury in invertebrates of small mid-latitude lakes in North America. Environmental Pollution, 159(1), 10–17. https://doi.org/10.1016/j.envpol.2010.09.034
Chumchal, M. M., & Drenner, R. W. (2015). An Environmental Problem Hidden in Plain Sight? Small Human-made Ponds, Emergent Insects, and Mercury Contamination of Biota in the Great Plains. Environmental Toxicology and Chemistry, 34, 1197–1205. https://doi.org/10.1002/etc.2954
86
Chumchal, M. M., Drenner, R. W., Greenhill, F. M., Kennedy, J. H., Courville, A. E., Gober, C. A. A., & Lossau, L. O. (2017). Recovery of Aquatic Insect-Mediated Methylmercury Flux from Ponds Following Drying Disturbance. Environmental Toxicology and Chemistry, 9999(9999), 1–5. https://doi.org/10.1002/etc.3734
Chumchal, M. M., Drenner, R. W., Hall, M. N., Polk, D. K., Williams, E. B., Ortega-Rodriguez, C. L., & Kennedy, J. H. (2018). Seasonality of dipteran-mediated methylmercury flux from ponds. Environmental Toxicology and Chemistry, 37(7), 1846–1851. https://doi.org/10.1002/etc.4134
Compeau, G. C., & Bartha, R. (1985). Sulfate-Reducing Bacteria: Principal Methylators of Mercury in Anoxic Estuarine Sedimentt. Applied and Environmental Microbiology, 50(2), 498–502. Retrieved from https://www.ncbi.nlm.nih.gov/pmc/articles/PMC238649/pdf/aem00143-0322.pdf
Corbet, P. S. (1964). Temporal Patterns of Emergence in Aquatic Insects. The Canadian Entomologist, 96, 264–279. https://doi.org/10.4039/Ent96264-1
Corbet, P. S. (1999). Dragonflies: behavior and ecology of Odonata. Ithaca, New York: Cornell University Press.
rdoba-Aguilar, A. (2008). Dragonflies and damselflies : model organisms for ecological and evolutionary research. Oxford University Press.
ristol, D. A., Brasso, R. L., ondon, A. M., Fovargue, R. E., Friedman, S. L., Hallinger, K. K., … White, A. E. (2008). The Movement of Aquatic Mercury through Terrestrial Food Webs. Source: Science, New Series, 320(5874), 335. https://doi.org/10.1126/science.ll54082
Davies, I. J. (1984). Sampling Aquatic Insect Emergence. In J. A. Downing & F. H. Rigler (Eds.), A manual on methods for the assessment of secondary productivity in fresh waters (pp. 161–227). Oxford, UK: Blackwell Scientific Publications. Retrieved from https://downing.public.iastate.edu/tier 2/jadpdfs/1984 IBP 17 Chapter 6 Emergence.pdf
Dijkstra, J. A., Buckman, K. L., Ward, D., Evans, D. W., & Dionne, M. (2013). Experimental and Natural Warming Elevates Mercury Concentrations in Estuarine Fish. PLoS ONE, 8(3), 58401. https://doi.org/10.1371/journal.pone.0058401
Downs, S. G., Macleod, C. L., & Lester, J. N. (1998). Mercury in precipitation and its relation to bioaccumulation in fish: a literature review. Water, Air, and Soil Pollution, 108, 149–187.
Driscoll, C. T., Han, Y.-J., hen, . Y., Evers, D. ., Lambert, K. F., Driscoll, . T., … Munson, R. K. 2007). Mercury contamination in forest and freshwater ecosystems in the Northeastern United States. Bioscience, 57(1), 17–28. https://doi.org/10.1641/b570106
Eagles-Smith, C. A., & Ackerman, J. T. (2009). Rapid Changes in Small Fish Mercury Concentrations in Estuarine Wetlands: Implications for Wildlife Risk and Monitoring Programs. Environmental Science & Technology, 43, 8658–8664. https://doi.org/10.1021/es901400c
87
Eagles-Smith, C. A., Nelson, S. J., Willacker Jr., J. J., Flanagan Pritz, C., & Krabbenhoft, D. P. (2016). Dragonfly Mercury Project—A Citizen Science Driven Approach to Linking Surface-Water Chemistry and Landscape Characteristics to Biosentinels on a National Scale. https://doi.org/http://dx.doi.org/10.3133/fs20163005
Eagles-Smith, C. A., Wiener, J. G., Eckley, C. S., Willacker, J. J., Evers, D. C., Marvin-Dipasquale, M., … Ackerman, J. T. (2016). Mercury in western North America: A synthesis of environmental contamination, fluxes, bioaccumulation, and risk to fish and wildlife. Science of the Total Environment, 568, 1213–1226. https://doi.org/10.1016/j.scitotenv.2016.05.094
Fitzgerald, William, F., Engstrom, D. R., Mason, R. P., & Nater, E. A. (1998). Critical Review The Case for Atmospheric Mercury Contamination in Remote Areas. Environmental Science & Technology Technology, 32(1), 1–7. Retrieved from https://pubs.acs.org/sharingguidelines
Flanagan Pritz, C., Eagles-Smith, C., & Krabbenhoft, D. (2014). Mercury in the National Parks. The George Wright Forum, 31(2), 168–180. https://doi.org/http://www.jstor.org/stable/43598338.
Flanagan Pritz, C., & Nelson, S. J. (2017). Collecting Data on Charismatic Mini-Fauna: Public Participation and the Dragonfly Mercury Project. Maine Policy Review, 26(2), 50–54. https://doi.org/http://digitalcommons.library.umaine.edu/mpr/vol26/iss2/9.
Flanagan Pritz, C., Nelson, S. J., & Eagles-Smith, C. A. (2014). The call to collect dragonflies: citizen scientists study mercury contamination in National Parks. Park Science, 31(1), 74–77.
Fletcher, D. E., Lindell, A. H., Stillings, G. K., Blas, S. A., & McArthur, J. V. (2017). Trace element accumulation in lotic dragonfly nymphs: Genus matters. PLOS One, 12(2), 1–27. https://doi.org/10.1371/journal.pone.0172016
Gann, G. L., Powell, C. H., Chumchal, M. M., & Drenner, R. W. (2015). Hg-Contaminated Terrestrial Spiders Pose a Potential Risk to Songbirds at Caddo Lake (Texas/Louisiana, USA). Environmental Toxicology and Chemistry, 34(2), 303–306. https://doi.org/10.1002/etc.2796
Gawley, W. G., & Wiggin, S. P. (2016). Water quality monitoring at Acadia National Park: Northeast Temperate Network 2014 summary report. Natural Resource Data Series NPS/NETN/NRDS— 2016/1021. Fort Collins, Colorado, USA. Retrieved from https://irma.nps.gov/DataStore/DownloadFile/554568
George, B. M., & Batzer, D. (2008). Spatial and temporal variations of mercury levels in Okefenokee invertebrates: Southeast Georgia. Environmental Pollution, 152(2), 484–490. https://doi.org/10.1016/j.envpol.2007.04.030
Giberson, D. J. M., & Garnett, H. L. (1996). Species composition, distribution, and summer emergence phenology of stonef lies (Insecta: Plecoptera) from Catamaran Brook, New Brunswick. Canadian Journal of Zoology, 74, 1260–1267. Retrieved from www.nrcresearchpress.com
Gillooly, J. F., Charnov, E. L., West, G. B., Savage, V. M., & Brown, J. H. (2002). Effects of size and temperature on developmental time. Nature, 417(2), 70–73. Retrieved from www.nature.com
88
Gorski, P. R., Cleckner, L. B., Hurley, J. P., Sierszen, M. E., & Armstrong, D. E. (2003). Factors affecting enhanced mercury bioaccumulation in inland lakes of Isle Royale National Park, USA. The Science of the Total Environment, 304, 327–348. Retrieved from https://s3.amazonaws.com/academia.edu.documents/42441831/Factors_affecting_enhanced_me rcury_bioac20160208-7077- diw052.pdf?AWSAccessKeyId=AKIAIWOWYYGZ2Y53UL3A&Expires=1548271624&Signature=Srnh6 RPVxhHl1pvmtU3q%2BRVw%2Bls%3D&response-content-disposition=inli
Gosnell, K. J., Balcom, P. H., Tobias, C. R., Gilhooly, W. P., & Mason, R. P. (2017). Spatial and temporal trophic transfer dynamics of mercury and methylmercury into zooplankton and phytoplankton of Long Island Sound. Limnology and Oceanography, 62(3), 1122–1138. https://doi.org/10.1002/lno.10490
Goutner, V., & Furness, R. W. (1997). Mercury in Feathers of Little Egret *Egretta garzetta* and Night Heron *Nycticorax nycticorax* Chicks and in Their Prey in the Axios Delta, Greece. Archives of Environmental Contamination and Toxicology, 32(2), 211–216. https://doi.org/10.1007/s002449900177
Grof-Tisza, P., LoPresti, E., Heath, S. K., & Karban, R. (2017). Plant structural complexity and mechanical defenses mediate predator–prey interactions in an odonate–bird system. Ecology and Evolution, 7(5), 1650–1659. https://doi.org/10.1002/ece3.2705
Haines, T. A., May, T. W., Finlayson, R. T., & Mierzykowski, S. E. (2003). Factors Affecting Food Chain Transfer of Mercury in the Vicinity of the Nyanza Site, Sudbury River, Massachusetts. Environmental Monitoring and Assessment, 86(3), 211–232. https://doi.org/https://doi.org/10.1023/A:1024017329382
Hall, B. D., Rosenberg, D. M., & Wiens, A. P. (1998). Methyl mercury in aquatic insects from an experimental reservoir. Canadian Journal of Fisheries and Aquatic Sciences, 55(9), 2036–2047. https://doi.org/10.1139/cjfas-55-9-2036
Haro, R. J. (2014). All along the watchtower: Larval dragonflies are promising biological sentinels for monitoring methylmercury contamination. Park Science, 31(1), 70–73. Retrieved from https://www.nature.nps.gov/ParkScience/Archive/PDF/Article_PDFs/ParkScience31(1)SpecialIssue 2014_70-73_Haro_3793.pdf
Haro, R. J., Bailey, S. W., Northwick, R. M., Rolfhus, K. R., Sandheinrich, M. B., & Wiener, J. G. (2013). Burrowing Dragonfly Larvae as Biosentinels of Methylmercury in Freshwater Food Webs. Environmental Science & Technology, 47, 8148–8156. https://doi.org/10.1021/es401027m
Harper, P. P., & Pilon, J.-G. (1970). Annual patterns of emergence of some Quebec stoneflies (Insecta: Plecoptera). Canadian Journal of Zoology, 48(4), 681–694. https://doi.org/10.1139/z70-126
Hothem, R. L. (2008). Mercury contamination in Foothill Yellow-legged Frogs (Rana boylii) and Invertebrates from Harley Gulch, California, 2007. Sacramento, CA.
Houghton, D. C. (2015). A 5-Year Study of the Adult Flight Periodicity of 27 Caddisfly (Trichoptera) Species in Forest and Meadow Habitats of a First-Order Lower Michigan (USA) Stream. Environmental Entomology, 44(6), 1472–1487. https://doi.org/10.1093/ee/nvv138
89
Ivković, M., Miliša, M., Previšić, A., Popijač, A., & Mihaljević, Z. 2013). Environmental control of emergence patterns: Case study of changes in hourly and daily emergence of aquatic insects at constant and variable water temperatures. International Review of Hydrobiology, 98(2), 104–115. https://doi.org/10.1002/iroh.201301483
Jannot, J. E. (2009). Life history plasticity and fitness in a caddisfly in response to proximate cues of pond-drying. Oecologia, 161(2), 267–277. https://doi.org/10.1007/s00442-009-1389-7
Jardine, T. D., Kidd, K. A., & Rasmussen, J. B. (2012). Aquatic and terrestrial organic matter in the diet of stream consumers: Implications for mercury bioaccumulation. Ecological Applications, 22(3), 843– 855. https://doi.org/10.2307/23213921
Jeremiason, J. D., Reiser, T. K., Weitz, R. A., Berndt, M. E., & Aiken, G. R. (2016). Aeshnid dragonfly larvae as bioindicators of methylmercury contamination in aquatic systems impacted by elevated sulfate loading. Ecotoxicology, 25(3), 456–468. https://doi.org/10.1007/s10646-015-1603-9
Johnson, K. B., Haines, T. A., Kahl, J. S., Norton, S. A., Amirbahman, A., Sheehan, K. D., & Amirbahman, A. (2007). Controls on Mercury and Methylmercury Deposition for Two Watersheds in Acadia National Park, Maine. Environ Monit Assess, 126, 55–67. https://doi.org/10.1007/s10661-006- 9331-5
Jones, T. A., Chumchal, M. M., Drenner, R. W., Timmins, G. N., & Nowlin, W. H. (2013). Bottom-up nutrient and top-down fish impacts on insect-mediated mercury flux from aquatic ecosystems. Environmental Toxicology and Chemistry, 32(3), 612–618. https://doi.org/10.1002/etc.2079
Judd, W. W. (1962). A Study of the Population of Insects Emerging as Adults from Medway Creek at Arva, Ontario. Source: The American Midland Naturalist, 68(2), 463–473. https://doi.org/130.111.46.54
Karimi, R., Chen, C. Y., & Folt, C. L. (2016). Comparing nearshore benthic and pelagic prey as mercury sources to lake fish: the importance of prey quality and mercury content. Science of the Total Environment, 565, 211–221. https://doi.org/10.1016/j.scitotenv.2016.04.162
Karimi, R., Fisher, N. S., & Folt, C. L. (2010). Multielement Stoichiometry in Aquatic Invertebrates: When Growth Dilution Matters. The American Naturalist, 176(6), 699–709. https://doi.org/10.5061/dryad.1858
Karimi, R., Fisher, N. S., & Meliker, J. R. (2014). Mercury-nutrient signatures in seafood and in the blood of avid seafood consumers. Science of the Total Environment, 496, 636–643. https://doi.org/10.1016/j.scitotenv.2014.04.049
Kidd, K., Clayden, M., & Jardine, T. (2012). Bioaccumulation and biomagnification of mercury through food webs. In G. Liu, Y. ai, & N. O’ Driscoll Eds.), Environmental Chemistry and Toxicology of Mercury (pp. 474–480). Hoboken, New Jersey: A John Wiley & Sons, Inc.
Klemmer, A. J. (2015). Unravelling the effects of multiple cross-ecosystem subsidies on food webs. University of Canterbury, Christchurch, New Zealand.
90
Knights, B. C., Wiener, J. G., Sandheinrich, M. B., Jeremiason, J. D., Kallemeyn, L. W., Rolfhus, K. R., & Brigham, M. E. (2005). Final Report for Project No. 02-01.
Krabbenhoft, D. P., & Sunderland, E. M. (2013). Global Change and Mercury. Science, 341, 1457–1458. https://doi.org/10.1126/science.1242838
Krell, B., Röder, N., Link, M., Gergs, R., Entling, M. H., & Schäfer, R. B. (2015). Aquatic prey subsidies to riparian spiders in a stream with different land use types. Limnologica, 51, 1–7. https://doi.org/10.1016/j.limno.2014.10.001
Lesch, V., & Bouwman, H. (2018). Adult dragonflies are indicators of environmental metallic elements. Chemosphere, 209, 654–665. https://doi.org/10.1016/j.chemosphere.2018.06.029
López-Rodríguez, M. J., Tierno De Figueroa, J. M., Bo, T., Mogni, A., & Fenoglio, S. (2012). Living apart together: On the biology of two sympatric Leuctra species (Plecoptera, Leuctridae) in an Apenninic stream, Italy. International Review of Hydrobiology, 97(2), 117–123. https://doi.org/10.1002/iroh.201111413
Lusilao-Makiese, J. G., Cukrowska, E. M., Tessier, E., Amouroux, D., & Weiersbye, I. (2013). The impact of post gold mining on mercury pollution in the West Rand region, Gauteng, South Africa. Journal of Geochemical Exploration, 134, 111–119. https://doi.org/10.1016/j.gexplo.2013.08.010
Maine Department of Environmental Protection. (2016). Biological Monitoring Program: Aquatic Life Classification Attainment Report.
Maine Department of Environmental Protection. (2017). Biological Monitoring Program: Aquatic Life Classification Attainment Report.
Mason, R. P., Laporte, J. M., & Andres, S. (2000). Factors Controlling the Bioaccumulation of Mercury, Methylmercury, Arsenic, Selenium, and Cadmium by Freshwater Invertebrates and Fish. Archives of Environmental Contamination and Toxicology, 38(3), 283–297. https://doi.org/10.1007/s002449910038
Menzie, C. A. (1980). Potential significance of insects in the removal of contaminants from aquatic systems. Water, Air, & Soil Pollution, 13(4), 473–479.
Merrill, R. J., & Johnson, D. M. (1984). Dietary niche overlap and mutual predation amond coexisting larval Anisoptera. Odonatologica, 13(3), 387–406.
Merritt, R. W., Cummins, K. W., & Berg, M. B. (2008). An Introduction to the Aquatic Insects of North America. Kendall Hunt Publishing.
Milner, A. M., Brittain, J. E., Castella, E., & Petts, G. E. (2001). Trends of macroinvertebrate community structure in glacier-fed rivers in relation to environmental conditions: a synthesis. Freshwater Biology, 46, 1833–1847. Retrieved from https://bb.courses.maine.edu/bbcswebdav/pid-2941659- dt-content-rid-6148797_2/courses/1810.UMS05-C.0169.1/Milner-Brittain-Petts 2001_unedited.pdf
91
Molina, C. I., Gibon, F.-M., Duprey, J.-L., Dominguez, E., Guimarães, J.-R. D., & Roulet, M. (2010). Transfer of mercury and methylmercury along macroinvertebrate food chains in a floodplain lake of the Beni River, Bolivian Amazonia. Science of the Total Environment, 408, 3382–3391. https://doi.org/10.1016/j.scitotenv.2010.04.019
Morel, F. M., Kraepiel, A. M. L., & Amyot, M. (1998). The Chemical Cycle and Bioaccumulation of Mercury. Annu. Rev. Ecol. Syst. , 29, 542–566. Retrieved from https://www.researchgate.net/profile/Marc_Amyot/publication/229139013_The_chemical_cycle_ and_bioaccumulation_of_mercury/links/00b49533362c463877000000/The-chemical-cycle-and- bioaccumulation-of-mercury.pdf
Nakano, S., & Murakami, M. (2001). Reciprocal subsidies: Dynamic interdependence between terrestrial and aquatic food webs. Proceedings of the National Academy of Sciences, 98(1), 166–170. https://doi.org/10.1073/pnas.98.1.166
Nasirian, H., & Irvine, K. N. (2017). Odonata larvae as a bioindicator of metal contamination in aquatic environments: application to ecologically important wetlands in Iran. Environmental Monitoring and Assessment, 189(9), 1–18. https://doi.org/10.1007/s10661-017-6145-6
Nelson, S. J., Chen, C., Kahl, J. S., & Krabbenhoft, D. P. (2014). Validating Landscape Models for Mercury in Northeast Lakes using Dragonfly Nymphs as Mercury Bio-sentinels. Orono, Maine, USA. https://doi.org/https://digitalcommons.library.umaine.edu/orsp_reports/52
Nelson, S. J., & Flanagan Pritz, C. (2014). Six-legged Scouts: Dragonfly larvae provide baseline data to evaluate mercury in parks nationwide.
Nelson, S. J., Webber, H. M., & Flanagan Pritz, C. (2015). Citizen Scientists Study Mercury in Dragonfly Larvae: Dragonfly larvae provide baseline data to evaluate mercury in parks nationwide. National Resources Data Series NPS/GLKN/NRDS. Fort Collins, Colorado, USA. Retrieved from https://irma.nps.gov/DataStore/DownloadFile/521133
Oosthuizen, M. A., John, J., & Somerset, V. (2010). Mercury exposure in a low-income community in South Africa. South African Medical Journal, 100(6), 366–371. Retrieved from http://www.scielo.org.za/pdf/samj/v100n6/v100n6a17.pdf
Ortega‐Rodriguez, . L., humchal, M. M., Drenner, R. W., Kennedy, J. H., Nowlin, W. H., Barst, B. D., … Basu, N. (2019). Relationship Between Methyl Mercury Contamination and Proportion of Aquatic and Terrestrial Prey in Diets of Shoreline Spiders. Environmental Toxicology and Chemistry, 00(00), 1–6. https://doi.org/10.1002/etc.4579
Ortiz, C., Weiss-Penzias, P. S., Fork, S., & Flegal, A. R. (2015). Total and monomethyl mercury in terrestrial arthropods from the central california coast. Bulletin of Environmental Contamination and Toxicology, 94(4), 425–430. https://doi.org/10.1007/s00128-014-1448-6
Paetzold, A., Smith, M., Warren, P. H., & Maltby, L. (2011). Environmental impact propagated by cross- system subsidy: Chronic stream pollution controls riparian spider populations. Ecology, 92(9), 1711–1716.
92
Parkman, H., & Meili, M. (1993). Mercury in Macroinvertebrates from Swedish Forest Lakes: Influence of Lake Type, Habitat, Life Cycle, and Food Quality. Canadian Journal of Fisheries and Aquatic Sciences, 50(3), 521–534. https://doi.org/10.1139/f93-061
Peckenham, J. M., Kahl, J. S., Nelson, S. J., Johnson, K. B., & Haines, T. A. (2007). Landscape Controls on Mercury in Streamwater at Acadia National Park, USA. Environmental Monitoring and Assessment, 126, 97–104. https://doi.org/10.1007/s10661-006-9334-2
Pennuto, C. M., & Smith, M. (2015). From Midges to Spiders: Mercury Biotransport in Riparian Zones Near the Buffalo River Area of Concern (AOC), USA. Bulletin of Environmental Contamination and Toxicology, 95(6), 701–706. https://doi.org/10.1007/s00128-015-1658-6
Pritchard, G. (1991). Insects in thermal springs. Memoirs of the Entomologicial Society of Canada, 155, 89–106. https://doi.org/10.4039/entm123155089-1
R Core Team. (2019). R: A language and environment for statistical computing. Vienna, Austria: R Foundation for Statistical Computing. Retrieved from https://www.r-project.org
Raikow, D. F., Walters, D. M., Fritz, K. M., & Mills, M. A. (2011). The distance that contaminated aquatic subsidies extend into lake riparian zones. Ecological Applications, 21(3), 983–990. Retrieved from https://esajournals-onlinelibrary-wiley-com.prxy4.ursus.maine.edu/doi/pdf/10.1890/09-1504.1
Riva-Murray, K., Bradley, P. M., hasar, L. ., Button, D. T., Brigham, M. E., Scudder Eikenberry, B. ., … Lutz, M. A. (2013). Influence of dietary carbon on mercury bioaccumulation in streams of the Adirondack Mountains of New York and the Coastal Plain of South Carolina, USA. Ecotoxicology, 22, 60–71. https://doi.org/10.1007/s10646-012-1003-3
Riva-Murray, K., Chasar, L. C., Bradley, P. M., Burns, D. A., Brigham, M. E., Smith, M. J., & Abrahamsen, T. A. (2011). Spatial patterns of mercury in macroinvertebrates and fishes from streams of two contrasting forested landscapes in the eastern United States. Ecotoxicology, 20(7), 1530–1542. https://doi.org/10.1007/s10646-011-0719-9
Rizzo, A., Arcagni, M., Arribére, M. A., Bubach, D., & Ribeiro Guevara, S. (2011). Mercury in the biotic compartments of Northwest Patagonia lakes, Argentina. Chemosphere, 84, 70–79. https://doi.org/10.1016/j.chemosphere.2011.02.052
Rolfhus, K. R., Wiener, J. G., Haro, R. J., Sandheinrich, M. B., Bailey, S. W., & Seitz, B. R. (2015). Mercury in streams at Grand Portage National Monument (Minnesota, USA): Assessment of ecosystem sensitivity and ecological risk. Science of the Total Environment, 514, 192–201. https://doi.org/10.1016/j.scitotenv.2015.01.079
Roy, A. H., Rosemond, A. D., Leigh, D. S., Paul, M. J., & Wallace, J. B. (2003). Habitat-specific responses of stream insects to land cover disturbance: biological consequences and monitoring implications. Journal of the North American Benthological Society, 22(2), 292–307. https://doi.org/10.2307/1467999
Sabo, J. L., Bastow, J. L., & Power, M. E. (2002). Length–mass relationships for adult aquatic and terrestrial invertebrates in a California watershed. Journal of the North American Benthological Society, 21(2), 336–343. https://doi.org/10.2307/1468420
93
Shanley, J. B., Kamman, N. C., Clair, T. A., & Chalmers, A. (2005). Physical Controls on Total and Methylmercury Concentrations in Streams and Lakes of the Northeastern USA. Ecotoxicology, 14, 125–134. Retrieved from https://link-springer- com.prxy4.ursus.maine.edu/content/pdf/10.1007%2Fs10646-004-6264-z.pdf
Silva, D. de paiva, De Marco, P., & Chaves Resende, D. (2010). Adult odonate abundance and community assemblage measures as indicators of stream ecological integrity: A case study. Ecological Indicators, 10, 744–752. https://doi.org/10.1016/j.ecolind.2009.12.004
Sinclair, K. A., Xie, Q., & Mitchell, C. P. J. (2012). Methylmercury in water, sediment, and invertebrates in created wetlands of Rouge Park, Toronto, Canada. Environmental Pollution, 171, 207–215. https://doi.org/10.1016/j.envpol.2012.07.043
Slotton, D. G., Ayers, S. M., Reuter, J. E., & Goldman, C. R. (1995). Gold Mining Impacts on Food Chain Mercury in Northwestern Sierra Nevada Streams. Davis, CA. Retrieved from https://escholarship.org/uc/item/58h617dr
Snyder, C. D., & Hendricks, A. C. (1995). Effect of seasonally changing feeding habits on whole-animal mercury concentrations in Hydropsyche morosa (Trichoptera: Hydropsychidae). Hydrobiologia, 299(2), 115–123.
Sokolova, I. M., & Lannig, G. (2008). Interactive effects of metal pollution and temperature on metabolism in aquatic ectotherms: implications of global climate change. Climate Research, 37, 181–201. https://doi.org/10.3354/cr00764
Speir, S. L., Chumchal, M. M., Drenner, R. W., Cocke, W. G., Lewis, M. E., & Whitt, H. J. (2014). Methyl mercury and stable isotopes of nitrogen reveal that a terrestrial spider has a diet of emergent aquatic insects. Environmental Toxicology and Chemistry, 33(11), 2506–2509. https://doi.org/10.1002/etc.2700
St Louis, V. L., Rudd, J. W. M., Kelly, C. A., Beaty, K. G., Bloom, N. S., & Flett, R. J. (1994). Importance of wetlands as sources of methyl mercury to boreal forest ecosystems. Canadian Journal of Fisheries and Aquatic Sciences, 51(5), 1065–1076. https://doi.org/10.1139/f94-106
Stagliano, D. M., Benke, A. C., & Anderson, D. H. (1998). Emergence of Aquatic Insects from 2 Habitats in a Small Wetland of the Southeastern USA: Temporal Patterns of Numbers and. Journal of the North American Benthological Society, 17(1), 37–53. Retrieved from https://www.jstor.org/stable/1468050
Sweeney, B. W., & Vannote, R. L. 1982). POPULATION SYN HRONY IN MAYFLIES : A PREDATOR SATIATION HYPOTHESIS The larvae of mayflies Order : Ephem- eroptera ) inhabit most permanent fresh- water environments ( rivers , lakes , ponds ). The duration of the mayfly life cycle varies from a few week, 36(4), 810–821.
Takimoto, G., Iwata, T., & Murakami, M. (2009). Timescale Hierarchy Determines the Indirect Effects of Fluctuating Subsidy Inputs on In Situ Resources. Source: The American Naturalist, 173(2), 200–211. https://doi.org/10.1086/595759
94
Takimoto, G., Omoya Wata, T. I., & Asashi Urakami, M. M. (2002). Seasonal subsidy stabilizes food web dynamics: Balance in a heterogeneous landscape. Ecological Research, 17, 433–439. https://doi.org/10.1046/j.0912-3814.2002.00502.x
Tennessen, K. (2017). A Method for Determining Stadium Number of Late Stage Dragonfly Nymphs (Odonata: Anisoptera). Entomological News, 126(4), 299–306. https://doi.org/10.3157/021.126.0407
Tierno De Figueroa, J. M., & López-Rodríguez, & M. J. (2019). Trophic ecology of Plecoptera (Insecta): a review. The European Zoological Journal, 86(1), 79–102. https://doi.org/10.1080/24750263.2019.1592251
Tremblay, A., & Lucotte, M. (1997). Accumulation of total mercury and methyl mercury in insect larvae of hydroelectric reservoirs. Canadian Journal of Fisheries and Aquatic Sciences, 54, 832–841. https://doi.org/10.1139/f96-339
Tremblay, A., Lucotte, M., Meili, M., Cloutier, L., & Pichet, P. (1996). Total Mercury and Methylmercury Contents of Insects from Boreal Lakes: Ecological, Spatial and Temporal Patterns. Water Quality Research Journal of Canada, 31(4), 851–873. Retrieved from https://www.researchgate.net/publication/265524690
Tsui, M. T. K., Blum, J. D., Kwon, S. Y., Finlay, J. C., Balogh, S. J., & Nollet, Y. H. (2012). Sources and Transfers of Methylmercury in Adjacent River and Forest Food Webs. Environmental Science & Technology, 46, 10957–10964. https://doi.org/10.1021/es3019836
Tweedy, B. N., Drenner, R. W., Chumchal, M. M., & Kennedy, J. H. (2013). Effects of fish on emergent insect-mediated flux of methyl mercury across a gradient of contamination. Environmental Science & Technology Technology, 47(3), 1614–1619. https://doi.org/10.1021/es303330m
U.S. Environmental Protection Agency. (1998). Method 1630, Methyl Mercury in Water by Distillation, Aqueous Ethylation, Purge and Trap, and Cold-Vapor Atomic Fluorescence Spectrometry, (4303), 1–55.
U.S. Environmental Protection Agency. (2007). EPA Method 7473 (SW-846): Mercury in Solids and Solutions by Thermal Decomposition, Amalgamation, and Atomic Absorption Spectrophotometry. Methods. Retrieved from https://www.epa.gov/sites/production/files/2015-07/documents/epa- 7473.pdf
U.S. Environmental Protection Agency. (2016). National Lakes Assessment 2012: A Collaborative Survey of Lakes in the United States, (December), 40. Retrieved from https://nationallakesassessment.epa.gov/
United Nations Environment Programme. (2013). Global Mercury Assessment 2013: Sources, Emissions, Releases and Environmental Transport. Geneva, Switzerland. Retrieved from http://www.unep.org/hazardoussubstances/Mercury/Informationmaterials/ReportsandPublicatio ns/
95
USGS. (2019). National Water Information System data available on the World Wide Web (USGS Water Data for the Nation). Retrieved June 23, 2019, from https://waterdata.usgs.gov/nwis/uv?site_no=01022840
VanderMeulen, D. D., Route, B., Wiener, J., Haro, R., Rolfhus, K., Sandheinrich, M., … Willacker, J. 2018). Protocol for Monitoring Mercury in Dragonfly Larvae and Fish (Version 1.0). Fort Collins, Colorado, USA. https://doi.org/10.13140/RG.2.2.24166.47682
Vermeer, K., Armstrong, F. A. J., & Hatch, D. R. M. (1973). Mercury in Aquatic Birds at Clay Lake, Western Ontario. The Journal of Wildlife Management, 37(1), 58–61. Retrieved from https://www-jstor- org.prxy4.ursus.maine.edu/stable/pdf/3799738.pdf?seq=1
Walters, D. M., Fritz, K. M., Otter, R. R., Luther, W. M., & Drive, K. 2008). The Dark Side of Subsidies : Adult Stream Insects Export Organic Contaminants to Riparian Predators. Ecological Applications, 18(8), 1835–1841.
Wiener, J. G., Haro, R. J., Rolfhus, K. R., Sandheinrich, M. B., Bailey, S. W., Northwick, R. M., & Gostomski, T. J. (2013). Bioaccumulation of Contaminants in Fish and Larval Dragonflies in Six National Park Units of the Western Great Lakes Region , 2008-2009. National Resources Data Series NPS/GLKN/NRDS.
Wiener, J. G., Krabbenhoft, D. P., Heinz, G. H., & Scheuhammer, A. M. (2003). Ecotoxicology of mercury. In D. J. Hoffman, B. A. Rattner, G. A. J. Burton, & J. J. Cairns (Eds.), Handbook of ecotoxicology (2nd ed., pp. 409–463). Boca Raton, FL: Lewis Publishers.
Wiener, J. G., & Shields, P. J. (2000). Mercury in the Sudbury River (Massachusetts, U.S.A.): pollution history and a synthesis of recent research. Canadian Journal of Fisheries and Aquatic Sciences, 57, 1053–1061. https://doi.org/10.1139/f00-039
Wiener, J. G., & Spry, D. J. (1996). Toxicological significance of mercury in freshwater fish. In W. N. Beyer, G. H. Heinz, & A. W. Redmond-Norwood (Eds.), Environmental contaminants in wildlife: interpreting tissue concentrations (pp. 297–339). Boca Raton, FL: CRC Press.
Wikelski, M., Moskowitz, D., Adelman, J. S., Cochran, J., Wilcove, D. S., & May, M. L. (2006). Simple rules guide dragonfly migration. Biology Letters, 2, 325–329. https://doi.org/10.1098/rsbl.2006.0487
Willacker, J. J., & Eagles-Smith, C. A. (2017). STANDARD OPERATING PROCEDURES FOR THE ANALYSIS OF METHYLMERCURY IN BIOLOGICAL TISSUE VIA COLD VAPOR ATOMIC FLUORESCENCE DETECTION WITH THE BROOKS-RAND “MERX” SYSTEM. U.S. Geological Survey: Forest and Rangeland Ecosystem Science Center Contaminant Ecology Research Lab.
Williams, B. K., Nichols, J. D., & Conroy, M. J. (2002). Analysis and Management of Animal Populations. Academic Press.
Williams, E. B., Chumchal, M. M., Drenner, R. W., & Kennedy, J. H. (2017). Seasonality of odonate- mediated methylmercury flux from permanent and semipermanent ponds and potential risk to red-winged blackbirds (Agelaius phoeniceus). Environmental Toxicology and Chemistry, 9999(9999), 1–5. https://doi.org/10.1002/etc.3844
96
Wissinger, S. A. (1988). Life history and size structure of larval dragonfly populations. Journal of the North American Benthological Society, 7(1), 13–28.
Wolfe, J. D., Lane, O. P., Brigham, R. M., & Hall, B. D. (2018). Mercury exposure to red-winged blackbirds (Agelaius phoeniceus) and dragonfly (Odonata: Aeshnidae) nymphs in Prairie Pothole wetlands. Facets, 3, 174–191. https://doi.org/10.1139/facets-2017-0086
Woodward, G., Ebenman, B., Emmerson, M., Montoya, J. M., Olesen, J. M., Valido, A., & Warren, P. H. (2005). Body size in ecological networks. Trends in Ecology and Evolution, 20(7), 402–409. https://doi.org/10.1016/j.tree.2005.04.005
Wyn, B., Kidd, K. A., Burgess, N. M., & Curry, R. A. (2009). Mercury biomagnification in the food webs of acidic lakes in Kejimkujik National Park and National Historic Site, Nova Scotia. Canadian Journal of Fisheries and Aquatic Science, 66, 1532–1545. https://doi.org/10.1139/F09-097
Yung, L., Bertheau, C., Cazaux, D., Regier, N., Slaveykova, V. I., & Chalot, M. (2019). Insect Life Traits Are Key Factors in Mercury Accumulation and Transfer within the Terrestrial Food Web. Cite This: Environ. Sci. Technol, 53, 11122–11132. https://doi.org/10.1021/acs.est.9b04102
Zhang, Z.-S., Lu, X.-G., Wang, Q.-C., & Zheng, D.-M. (2009). Mercury, Cadmium and Lead Biogeochemistry in the Soil-Plant-Insect System in Huludao City. Bulletin of Environmental Contamination and Toxicology, 83, 255–259. https://doi.org/10.1007/s00128-009-9688-6
Zhang, Z., Song, X., Wang, Q., & Lu, X. (2012). Mercury bioaccumulation and prediction in terrestrial insects from soil in Huludao City, Northeast China. Bulletin of Environmental Contamination and Toxicology, 89(1), 107–112. https://doi.org/10.1007/s00128-012-0649-0
Zwick, P. (1980). Plecoptera (Steingliegan). In M. Beier (Ed.), Hanbuch der Zoologic (4th ed., pp. 1–115). Berlin: Walter de Gruyter.
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APPENDIX A: ADDITIONAL INFORMATION FROM CHAPTER 1
Text A. 1 Reported field and laboratory methods for dragonfly collection and analysis
Nearly all of the studies presented here using dragonfly nymphs as Hg biosentinels captured nymphs with a dip net or hand picking individuals from the substrate with a forceps. Most studies collected any size dragonfly nymphs that were available, but a few studies stratified collections to larger nymphs that were closer to the end of their aquatic lifecycle and allowed to depurate overnight to clear any gut contents that may influence Hg concentrations measured (Haro et al., 2013; Rolfhus et al.,
2015). However, using a one-way ANOVA, there was not a significant difference between depurated and non-depurated nymphs in a pilot study containing one species of dragonfly (Nelson et al., 2015). Studies held dragonfly nymphs on wet ice in the field and then froze them when returning to the lab if immediate processing of samples was not an option. Later, a wet weight was obtained and then dragonflies were freeze-dried or oven dried (~50°C) for 48-72 hours. Dry weights were obtained before tissues were homogenized with mortar and pestle before analysis.
There are several methods for Hg analysis and the species measured could be THg, MeHg, or inorganic Hg (IHg). Many of the literature sources in this review (~49%) used a form of atomic fluorescence spectroscopy (AFS) detection when analyzing MeHg or THg concentrations (Table A. 1). The second most common method (~21%) was inductively coupled plasma – mass spectroscopy (ICP-MS) and other methods were atomic absorption spectrophotometry (AAS), instrumental neutron activation analysis (INAA), inductively coupled plasma-optical emission spectrometry (ICP-OES) (Table A. 1). These different analytical methods can be used to measure different Hg species (Table A. 1). For example, for
ICP-MS, MeHg and IHg can be measured and then added together to compute THg.
A variety of instruments can be used within each analytical method. US EPA provides several approved methods for Hg analysis based on whether water, bed sediment or soil, or plant and animal
98 tissue is being examined. The EPA hazardous waste test methods documents the protocol and standards that should be followed while running samples for different Hg species in biological tissues (Table A. 1).
Table A.1. The environmental protection agency (EPA) hazardous waste test methods (SW-
846) that documents the protocol and standards for measuring inorganic mercury Hg(II), total mercury (THg: inorganic and organic) and organic methylmercury (MeHg) used in biological tissues.
Hg EPA Process examples species method Hg(II) 6010D Inductively Coupled Plasma-Atomic Emission Spectrometry (ICP-AES) 6020A Inductively Coupled Plasma-Mass Spectrometry (ICP-MS) THg 7473 Thermal Decomposition, Amalgamation, and Atomic Absorption Spectrophotometry (AAS) 7474 Atomic Fluorescence Spectrometry (AFS) MeHg 1630 Aqueous Phase Ethylation, Followed by Gas Chromatographic Separation with Cold Vapor Atomic Fluorescence Detection (GC- CVAFS) 1630 Inductively Coupled Plasma-Mass Spectrometry (ICP-MS)
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APPENDIX B: ADDITIONAL INFORMATION FROM CHAPTER 2
Text B.1. Dragonfly collection and preparation
An ANOVA power analysis, accounting for nymph family (Cordulegastridae) and location
(Hunters Brook and Marshall Brook), was completed using previous Dragonfly Mercury Project (DMP) cordulegastrid nymph Hg data. The power analysis was used to determine the minimum number of samples needed from each stream to obtain a representative Hg sample per month for one year
(Vandermeulen et al. 2018). Five nymphs from Hunters Brook and eight nymphs from Marshall Brook were needed each month to obtain an 80% probability of detecting 20% change in mean THg concentrations annually with a Type I error of 0.05.
Text B.2. Environmental data collection
The HOBO water temperature loggers were deployed in both stream reaches in July 2018 and retrieved in July 2019. Temperature was recorded every two hours when deployed. Climatic data were downloaded from the National Climatic Data Center. We used mean monthly precipitation from
Southwest Harbor (Station ID: US1MEHN0003) and mean monthly air temperature from McFarland Hill
(Station ID: USR0000MMCF). These two monitoring stations were chosen because they provided the most complete data that matched our sampling months and were local to Mount Desert Island, where weather is strongly marine-influenced. We used the concurrent months precipitation and the previous months precipitation to analyze precipitation influence on THg concentrations in dragonfly nymphs.
Additionally, Acadia National Park and the US Geological Survey provided stream discharge data from
Otter Creek located on Mount Desert Island, Maine (Station ID: 01022840) (USGS, 2019).
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Figure B.1. Total mercury among dragonfly nymph instars. There was no significant difference between the natural log of total mercury (THg) (ng/g) dry weight (dw) in cordulegastrid nymphs in relation to instar.
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Figure B.2. Relationship between total mercury (THg) and methylmercury (MeHg) concentrations in thirteen cordulegastrid dragonfly nymphs from Marshall Brook in Acadia National Park, Maine.
Cordulegastrid nymphs that were analyzed for both THg and MeHg, and represented here, were collected in June 2018, July 2018, and January 2019.
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Stream Hunters Brook Marshall Brook
A2.65 B2.65
2.60 2.60
2.55 2.55
) 2.50 2.50
w d
2.45 2.45
g
/
g n
( 50 100 150 200 50 100 150 200
g
H Precipitation (mm) Precipitation−1 Month (mm) T
(Southwest Harbor, ME) (Southwest Harbor, ME)
n a
e C2.65 D2.65
m
y
l 2.60 2.60
h
t n
o 2.55 2.55 M
2.50 2.50
2.45 2.45
0 5 10 20 30 40 50 60 70
Discharge (cfs) Air temperature (F) (Otter Creek, ME) (McFarland Hill, ME) Environmental Factor
Figure B.3. Relationship between environmental factors and monthly mean THg in dragonflies. There were no significant relationships between monthly mean dragonfly dragonfly nymph THg and the concurrent month’s precipitation, the one month lag precipitation, discharge, or air temperature from either stream independently or with both streams combined.
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Figure B.4. Relationship between monthly mean THg concentrations in dragonfly nymphs and water temperature in Hunters Brook. There were no significant relationships between monthly mean cordulegastrid dragonfly nymph THg and the concurrent stream water temperature in Hunters Brook (R2
= 0.036, p = 0.557).
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Table B.1. Cordulegastrid dragonfly nymph geometric mean total mercury (THg) concentrations in parts per billion (ng/g) ± geometric standard error for each season in two streams of Acadia National Park,
Maine from March 2018 to April 2019.
Stream Season THg ± SE (ppb) N
Hunters Brook Spring 315 ± 15.2 23 Summer 331 ± 23.7 16 Autumn 413 ± 31.3 18 Winter 362 ± 27.2 13
Marshall Brook Spring 368 ± 13.1 24 Summer 363 ± 12.7 24 Autumn 329 ± 15.9 24 Winter 368 ± 18.4 28
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APPENDIX C: ADDITIONAL INFORMATION FROM CHAPTER 3
Figure C. 1. Marshall Brook sampling reach with emergence traps and substrate picture.
Figure C. 2. Hunters Brook sampling reach with emergence traps and substrate picture.
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Text C.1. Methylmercury in Trichoptera from Marshall Brook
The MeHg concentrations in Trichoptera of Marshall Brook were driven by differences in family.
Because of the large amount of Trichoptera in Marshall Brook that emerged, we were able to analyze three different vials of taxa in July alone. Two vials were large Trichoptera individuals containing a mix of the family Phryganiedae, specifically Oligostomis sp. and Limnephilidae, specifically Platycentropus sp.
There were six Trichoptera in each of these vials and had an average biomass of 16.7 and 14.0 mg dry weight. The third vial contained twelve smaller bodied Trichoptera from July, and contained four
Hydropsychid while the other eight individuals were not identified to family. The twelve Trichoptera in this vial had an average biomass of 1.50 mg dry weight. The two vials containing the much larger bodied
Trichoptera contained MeHg concentrations of 113 ng/g dw and 142 ng/g dw while the vial containing the three smaller bodied Trichoptera had the MeHg concentration of 479 ng/g dw. All three vials contained Trichoptera that emerged in July from Marshall Brook, however, the specific taxa that emerged yielded very different MeHg concentrations. Hydropsychids are known to be omnivorous while
Limnephilids are mostly detrivores (see discussion). The unknown Trichoptera with high MeHg concentrations are most likely omnivorous.
A linear regression revealed that the average body mass of Trichoptera per vial analyzed for
MeHg explained 63.9% of MeHg concentration within the vial (R2 = 0.6385, p = 0.0565) (Figure A. 3. 1).
As average biomass per vial increased, MeHg concentration decreased (Figure A. 3. 1). This could be a factor of growth dilution or the life history characteristics, including differences in diet, of Trichoptera within the vials.
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Table C.1. Caddisflies that were analyzed for MeHg from Marshall Brook. Vial Month Number Family Genus Average MeHg length concentration (mm) (ng/g dw) 381 June 10 Hydropsychidae 352 5 Polycentropodidae 378 July 3 Limnephilidae Unknown 11.58 142 2 Limnephilidae Platycentropus 12.25 sp. 1 Phryganeidae Oligostomis sp. 13.81 380 July 5 Limnephilidae Platycentropus 12.03 113 sp. 1 Phryganeidae Oligostomis sp. 12.66 383 July 8 Unknown 3.89 479 4 Hydropsychidae 6.42 396 August 5 Unknown 5.55 776 403 September 3 Unknown 4.11 1007
Figure C.3. (A) Methylmercury concentrations within each Trichoptera vial, indicating which Trichoptera families are within each vial and (B) The average caddisfly biomass within each vial against the vials MeHg concentration. Colors in graph (A) are the same as the vials in graph (B).
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Text C.2. Methylmercury in Plecoptera from Hunters Brook
The MeHg concentrations in Plecoptera of Hunters Brook were taxa driven. Because of the large amount of Plecoptera of Hunters Brook that emerged, we were able to analyze three different vials of
Chloroperlidae and two vials of Leuctridae to compare MeHg concentrations between these two
Plecoptera families. Chloroperlidae had roughly 2x times higher MeHg concentrations than Leuctridae
(t-test, t = 3.55, df = 2.69, p = 0.045) (Figure A. 3. 2). Chloroperlidae are known to be predators of small chironomids while Leuctridae are primary consumers consuming algae (Tierno De Figueroa & López-
Rodríguez, 2019).
The average body mass of Plecoptera per vial explained 58.2% of MeHg concentration within the vial (linear regression, R2 = 0.5824, p = 0.07752) (Figure A. 3. 2). As average biomass per vial increased, MeHg concentration decreased (Figure A. 3. 2). This was most likely a factor of the different diet between Chloroperlidae and Leuctridae.
Figure C.4. (A) Methylmercury concentrations within Plecoptera (Chloroperlidae and Leuctridae) in
Hunters Brook and (B) The average Plecoptera biomass within each vial against the vials MeHg concentration.
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Table C.2. Plecoptera that were analyzed for MeHg from Hunters Brook.
Vial Month Number Family Average MeHg length concentration (mm) (ng/g dw) 391 May 3 Chloroperlidae 8.01 238
385 June 10 Chloroperlidae 7.15 180
389 June 9 Chloroperlidae 6.45 286
374 July 3 Chloroperlidae 4.78 103 2 Leuctridae 4.76
390 August 7 Leuctridae 4.14 101
404 September 8 Leuctridae 3.98 129
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BIOGRAPHY OF THE AUTHOR
Megan Hess was born in Berlin, Wisconsin on September 10, 1993. She grew up in a cozy farm house in the country in Wisconsin. She attended the University of Wisconsin – La Crosse and graduated with a Bachelor’s degree in Biology with environmental studies and chemistry minors. Besides taking classes during her undergraduate degree, she was involved in contaminant research, including mercury and triclosan, in the UW- La Crosse’s River Studies Center. After graduation, she worked as a research assistant at the Texas A&M extension center in Dallas, Texas researching endangered mussels for a year.
She moved to Maine and entered the Ecology and Environmental Sciences graduate program at the
University of Maine in the fall of 2017. Megan is a candidate for the Masters of Science degree in
Ecology and Environmental Sciences from the University of Maine in December 2019.
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