The Pennsylvania State University
The Graduate School
College of Agricultural Sciences
THE GREEN DRAKES OF SPRING CREEK: TOWARD REESTABLISHING A
POPULATION OF THE MAYFLY EPHEMERA GUTTULATA PICTET 1843
(EPHEMEROPTERA: EPHEMERIDAE)
A Dissertation in
Entomology
by
Hannah L. Stout
2012 Hannah L. Stout
Submitted in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
May 2012
ii
The dissertation of Hannah L. Stout was reviewed and approved* by the following:
Michael C. Saunders Professor of Entomology Dissertation Advisor Chair of Committee
James L. Frazier Professor of Entomology
Gregory A. Hoover Senior Extension Associate Ornamental Entomologist Special Member
Jay R. Stauffer, Jr. Distinguished Professor of Ichthyology
Ke Chung Kim Professor Emeritus of Entomology Curator of the Frost Entomological Museum Director of the Center for BioDiversity Research
Gary W. Felton Professor of Entomology Head of the Department of Entomology
*Signatures are on file in the Graduate School
iii ABSTRACT
The “eastern green drake” mayfly ( Ephemera guttulata ) can be found in eastern North
America, and is a large mayfly that is especially revered by anglers. Spring Creek (Centre
County, Pennsylvania) is a world-famous limestone trout-fishing stream that once had a viable population of E. guttulata , but numerous pollutants wreaked havoc on the stream’s biota. Some populations of macroinvertebrate species recovered, but E. guttulata has not been observed on
Spring Creek since the late 1950s. In the time since the extirpation of the “eastern green drake” from Spring Creek, much work has been done to improve the stream’s water quality; the question is whether it has improved enough to once again support an E. guttulata population.
The purpose of my research is to conduct a series of three feasibility studies that will help to answer the question: Can we reintroduce the “eastern green drake” to its historic location on
Spring Creek? Such preliminary studies are essential components of a scientifically-based reintroduction program. The three studies conducted were 1) chemical testing, 2) bioassessment and 3) community sampling.
In the first study, water and substrate of targeted locations on Spring and Penns Creeks were sampled, then tested for the presence of known pollutants. Water and substrate of two locations on Spring Creek were found to contain low levels of pollutants that were comparable to locations on Penns Creek that support a dense population of E. guttulata .
In the second study, I tested the >25-day toxicity of Spring Creek water and substrate using E. guttulata larvae collected from Penns Creek, and using Penns Creek substrate as the
Control. The survival and growth of E. guttulata larvae in Spring Creek water and substrate demonstrated that Spring Creek may be suitable for the mayfly species.
iv In the third study, I sampled the macroinvertebrate communities of targeted locations on
Spring and Penns Creeks, to look for taxa of equal or greater pollution intolerance (according to
the EPA ranking of mid-Atlantic macroinvertebrate populations). At all Spring Creek sites, I
identified several aquatic insect taxa that are rated as equally or more intolerant of organic pollution than E. guttulata .
Overall, the results of the three studies suggest that reintroduction of E. guttulata to
Spring Creek may be possible. Using two freshwater macroinvertebrate reintroduction programs
as a template, I designed a strategy for reintroduction, and began its implementation; results so far
have been encouraging.
Successfully restoring the “eastern green drake” mayfly to Spring Creek could not only
have a positive impact on the economy of Central Pennsylvania, but could benefit the advancing
science of aquatic insect reintroductions.
v
TABLE OF CONTENTS
LIST OF FIGURES ...... viiiii
LIST OF TABLES...... ix
ACKNOWLEDGEMENTS...... x
Chapter 1 Introduction...... 1
Spring Creek ...... 1 Physicochemical characteristics of Spring Creek...... 1 Past and present challenges for the Spring Creek watershed ...... 4 Research goals ...... 5 Goal 1: Describe the characteristics and ecological needs of E. guttulata ...... 6 Goal 2: Identify pollutants in Spring Creek that may be potentially lethal/sublethal to E. guttulata ...... 6 Goal 3: Test the effects of Spring Creek substrate on E. guttulata larvae ...... 6 Goal 4: Examine macroinvertebrate communities of Penns Creek – are "pollution intolerant" taxa present? ...... 7 Goal 5: If feasible, design a strategy for reintroduction of E. guttulata to Spring Creek ...... 7
Chapter 2 The “eastern green drake” mayfly ( Ephemera guttulata Pictet, 1843) ...... 8
Introduction...... 8 The order Ephemeroptera ...... 8 Larval stage ...... 8 Winged stages – Subimagos and imagos...... 10 Ecological importance ...... 11 Ephemera guttulata Pictet, 1843 ...... 12 Description ...... 12 Life history...... 15
Chapter 3 Chemical analysis of stream substrates: Spring Creek vs. Penns Creek ...... 18
Introduction...... 18 Organochlorine pesticides...... 18 Objectives...... 20 Methods...... 20 Selection of sampling sites...... 20 Collection of samples...... 23 Preparation and analysis of samples...... 24 Results...... 25
vi Spring Creek...... 25 Penns Creek...... 26 Discussion...... 28 Pollutant levels in Spring and Penns Creeks...... 28 Implications...... 29
Chapter 4 Bioassessment: The effect of Spring Creek substrate on survivorship and growth of E. guttulata larvae ...... 32
Introduction...... 32 Bioassays, bioassessments and viability experiments...... 32 Field vs. laboratory bioassessments ...... 33 Objectives...... 34 Methods...... 34 Design of laboratory equipment...... 35 Design of rearing cages...... 36 Collection of larvae ...... 37 Collection of substrate...... 38 Pre-assay measurement of larvae ...... 38 Viability and growth assay...... 39 Post-assay preservation and measurements...... 42 Results...... 42 Discussion...... 43 Survivorship and growth...... 43 Implications...... 44
Chapter 5 Sampling the macroinvertebrate communities of Spring Creek and Penns Creek...... 46
Introduction...... 46 Objective...... 49 Methods...... 50 Selection of sampling stations...... 50 Sampling design and equipment ...... 53 Collection and preservation methods ...... 55 Identification of specimens ...... 57 Results...... 57 Spring Creek...... 58 Penns Creek...... 59 Discussion...... 61 Implications for reintroduction...... 61
Chapter 6 Strategy for reintroduction of Ephemera guttulata to Spring Creek...... 63
Introduction...... 63 Reintroductions ...... 63 International Union for the Conservation of Nature and Natural Resources ...... 67 Strategy and process...... 68 Preliminary stage of reintroduction...... 68
vii Implementation...... 71 Post-implementation stage of reintroduction ...... 73 Discussion...... 75 Estimates of recolonization ...... 76 Implications...... 81 Looking forward ...... 81
References………...... 83
Appendix A Pollutants detected in substrate and water collected from Spring and Penns Creeks ...... 97
Appendix B Macroinvertebrate taxa collected from Spring Creek and Penns Creek...... 99
viii
LIST OF FIGURES
Figure 1-1: Map of towns through which Spring Creek flows...... 1
Figure 1-2: Map of limestone and dolomite distribution in Pennsylvania...... 3
Figure 2-1: Genital structures of male (left) and female (right) E. guttulata . Arrows point to genital forceps (“claspers”) of the male...... 10
Figure 2-2: Ephemera guttulata larva...... 13
Figure 2-3: Ephemera guttulata subimago (female)…………………………………………14
Figure 2-4: Ephemera guttulata imago (male)...... 15
Figure 4-1: Recirculation system used in vEXP Reps 1 – 11. Ichthyology Lab, School of Forest Resources. Rock Springs Agricultural Research Facility. Pennsylvania State University...... 34
Figure 4-2: Static system used in vEXP Reps 12 – 14. Private residence, State College PA...... 35
Figure 4-3: a. Rearing cage used in Reps 1 – 11 b. Illustration of floating foam support for rearing cages used in Reps 1 – 11...... 37
Figure 4-4: “Head capsule width”, as defined for measurement (pre- and post-assay)...... 39
Figure 4-5: Sorting tray with contents of a rearing cage and live E. guttulata larva (insert)...... 40
Figure 5-1: Location of Spring Creek community sampling sites...... 51
Figure 5-2: Location of Penns Creek community sampling sites...... 53
Figure 6-1: Number of E. guttulata larvae over time, with indefinite egg transplants (r = 0.3)...... 79
Figure 6-2: Number of E. guttulata larvae over time, if I = 0 as of year 4 (r = 0.3)...... 80
Figure 6-3: Number of E. guttulata larvae over time, if I = 0 as of year 11 (r = 0.6)...... 80
ix
LIST OF TABLES
Table 4-1: Design of Reps 1-11 of the viability experiment...... 40
Table 4-2: Design of Reps 12-14 of the viability experiment...... 41
Table 4-3: Number of survivors, percent survivorship and growth of surviving larvae for Reps 1 – 14 of the viability experiment...... 42
x
ACKNOWLEDGEMENTS
Over the course of this seven-year long journey, I have amassed quite a large number of people who deserve recognition and many, many, most sincere thank yous. First, I must thank Dr. Gary Felton, the Department Head of Entomology at PSU, for inviting me to the Department, and for his continued support of my research. Thanks to my Advisor, Mike Saunders. Mentor. “Assistant”. Master storyteller. Legend. Thanks to Andrea Hollnagel, Dr. Laura C. Klein and Dr. Dolores Maney for writing three lovely letters of recommendation for my graduate school application. Thanks to Ke Chung Kim, whose work in aquatics and in conserving biodiversity in Korea’s Demilitarized Zone inspired my graduate school application essay. Thanks to my each member of my Committee—Jim Frazier, Greg Hoover, Ke Chung Kim, and Jay Stauffer—for their invaluable wisdom and individual generosity. Thanks to my “unofficial” mentor, Bob Carline, a leading expert on Spring Creek and an all-around nice guy. Thanks to Tim Stecko and Rich Taylor, for assistance and advice at the Ichthyology Lab. Thanks to Frank Dorman, Seth Michalski, Beth Slaybaugh, and Jessica Westland of the Department of Forensic Science at PSU, for not only analyzing water and substrate samples, but for interpreting the results for this non-chemist. Thanks to Rick Lake and Jack Cochran of Restek, for so generously donating materials needed for the chemical analyses. Thanks to Carolyn Turgeon, the most glamorous photographer of mayflies EVER. Thanks to Chris Urban of the PA Fish & Boat Commission, for his approval (!) Thanks to Katie Ombalski of Clearwater Conservancy, for her enthusiasm throughout this project. Thanks to Rebecca Dunlap (Trout Unlimited) and John Nantz (USGS) for Spring Creek substrate and water quality data. Thanks to Rhiannon McClintock of the Centre County Historical Society, for background information on the Spring Creek watershed. Thanks to Rick Jacobsen of the USGS for fielding questions about our favorite mayfly. Thanks to Mark Vinson of the USGS, for discussing his own reintroduction project, and for the virtual high-fives. Thanks to The Feathered Hook (Coburn, PA) and the Kettle Creek Tackle Shop (Renovo, PA) for their personalized hatch reports. Thanks to Judi Sittler and Bill Brusse, the present and the past President of Spring Creek TU, for inviting me to discuss my research with them (while also providing food!). Thanks to Craig from Allied Mechanical & Electrical, for equipment repairs and mechanical advice. Thanks to my GDRP volunteers - Janet Saunders, Shelby and Barb Fleischer, Denice and Rick Wardrop, Dan Kerstetter, Barbara Kinne, Carlos Davis, Scott Harrison, Tom Bathgate, Alexis Barbarin, Sheena Sidhu, Jason Smith, Tom Bentley, Christina Harris, Maggie Douglas, Nina Stanczyk, Becky Heinig, Fernanda Penaflor, Amy Alesch and Sarah VanLandingham.
Special thanks to my parents, Michael and Laura Cave, for doing everything humanly (and superhumanly) possible to make sure I graduate.
xi Special thanks to my most gorgeous and most hilarious “seesters”, Ajia and Shalah Cave. Special thanks to my family - David and Sis Cave, Chris and Evynn Burnett, Taj Cave- Burnett; Bridget, Doug, Brandon and Shana Kennedy; Cheri Pearce; Kelin Linnan, Macie and Brenna Campbell; Catalina Hinchey-Trujillo; Jay Hinchey. Angeline and David Leeper; Amber and Seth Gamber; Frank, Dan and Nancy Stout; James, Nancy and Jim Morrissey. Special thanks to my friends - Maya Nehme, Jon Lelito and Kerry Mauck; Catherine Turgeon; Jeff Morelli; Steve Power; Chris and Michelle Craighead. Special thanks to my neighbors - Alan and Nancy Cameron; Lowie Bathgate Kerns; Jim and Lynda Stephenson.
Special thanks to my grandmother, Charlotte Pearce, who sadly is not here to read this.
MOST IMPORTANT AND MOST SPECIAL THANKS to Jason, Gryfinn and Thalia, for their patience, words of encouragement, and hugs when I needed them the most.
1
Chapter 1
Introduction
Spring Creek
Physicochemical characteristics of Spring Creek
Spring Creek (Centre County, Pennsylvania) is a 36-km long, 4 th order stream in the
Chesapeake Bay watershed via the Susquehanna River. Spring Creek originates in the outskirts of
Boalsburg, PA, flows through the towns of Lemont, Houserville and Bellefonte, and empties into
Bald Eagle Creek at Milesburg (Figure 1-1).
Figure 1-1 Map of towns through which Spring Creek flows. (Scale: 2.2cm = 5km)
2
Rich deposits of limestone and dolomite comprise the bedrock supporting the Spring
Creek watershed (Figure 1-2). This “karst” geology allows for a substantial amount of surface
water infiltration and groundwater storage, and the stream is unique in the large number of
limestone springs that feed it—the Pennsylvania Groundwater Association states that 86% of
Spring Creek’s total annual flow comes from groundwater. This constant input from springs
maintains the stream’s consistent flow and an average yearly water temperature of 10 ◦C.
In addition to its unique geological characteristics, Spring Creek is a Class A trout stream renowned for its abundant population of wild brown trout ( Salmo trutta ). Spring Creek has also been designated as a High Quality-Cold Water Fishery (HQ-CWF), which provides special protections via wastewater treatment requirements, water quality standards, and land use and development regulations. Spring Creek is home to the “world famous” flyfishing-only waters of
“Fisherman’s Paradise” (websites: The Alleghenies, PA Fish & Boat Commission). Spring Creek is also famous for the “green drake” mayflies that formerly inhabited the stream, and for the pollution that likely caused their disappearance decades ago.
3
Figure 1-2 Map of limestone and dolomite distribution in Pennsylvania. Spring Creek is located in the southcentral region of Centre County (source: DCNR)
4
Past and present challenges for the Spring Creek watershed
Once legendary, the population of the “eastern green drake” mayfly ( Ephemera guttulata ) in Spring Creek (Centre County, Pennsylvania) was first noticeably absent from Spring Creek in
1958, two years after sodium cyanide was flushed in a Pennsylvania State University laboratory drain, resulting in a fish kill that was dubbed “the Spring Creek massacre” (Daily News, 17 Dec
1956). The “eastern green drakes” of Spring Creek have not rebounded in the decades since.
Historically, the list of Spring Creek’s other major pollutants includes: raw sewage, Kepone
(chlordecone), mirex, chlorine and gasoline (Carline et al 2011). Siltation and effluent-induced eutrophication have declined within the last ten years, but continue to be of concern (Carline and
Walsh 2007).
Of these pollutants, the organochlorine (OC) pesticides Kepone and mirex are commonly blamed for the disappearance of E. guttulata . Organochlorine contamination of the groundwater that feeds Spring Creek most likely began in the early 1960s, when the Rütgers-Nease Organics
Corporation first documented the disposal of organic waste into onsite earthen lagoons at its State
College, Pennsylvania facility. In 1963, the company replaced the earthen lagoons with asphalt and concrete lagoons, but wastewater was sprayed onto onsite grassy areas. Offsite waste removal was initiated in 1972—the same year that the then-Pennsylvania Department of Environmental
Resources (PADER) ordered the clean-up of the lagoons. Following years of investigation, the
EPA placed the site on its National Priorities List in 1983, and assumed the lead role in the site’s remediation in 1986. A March 1989 Draft Work Plan for the site’s Remedial
Investigation/Feasibility Study (RI/FS) stated that soil samples taken onsite contained Kepone concentrations as high as 5300 µg/kg (ppb) and mirex as high as 1500 µg/kg (ppb) (USEPA
4/18/1994). These levels far exceeded the FDA’s action threshold of 0.1 mg/kg (ppm) by 5300%
5 and 1500%, respectively. In tests performed in 1990 and 1992, PADEP detected Kepone, mirex, and 27 other toxic compounds at the Superfund Site—namely volatile organic compounds (VOC) such as benzene and toluene. Remedial action began in 1999 (website: U.S. EPA Superfund Site
#PAD000436261).
The Pennsylvania Department of Environmental Protection (PADEP) has been monitoring organochlorine levels in Spring Creek’s brown trout population since 1976; high OC levels found in the trout were the impetus for the agency to enforce a strict “no-kill” policy from
1982 to 2001. Since 1998, the PADEP has found mirex levels to be below the FDA’s action threshold (0.1 mg/kg). Although the “no-kill” was lifted in 2001, the PA Fish and Boat
Commission continue to enforce a “no harvest” policy, in the interest of wildlife management rather than public health (website: PA Bull. 2001). With improvements in stream quality come speculation that perhaps a population of “green drakes” could be restored to Spring Creek. The intent of my research is to put an end to that speculation.
Research goals
To design a scientifically based reintroduction protocol for E. guttulata , it is essential to first conduct a number of feasibility studies. Eastern green drake mayflies spend their entire larval lives in stream substrate, thus the “quality” of that substrate is essential. Considering Spring
Creek’s history of pollution, toxicity of the streambed to E. guttulata larvae is possible, and must be ruled out before proceding with any attempts at reintroduction. The USEPA employs a
“Sediment Quality Triad Approach” (1992) that examines “Chemistry” (chemical analysis of sediment), “Toxicity” (experimental exposure of test organisms to sediment), and “Community”
(assessment of macroinvertebrate communities from area of sediment sampling). The USEPA uses this Approach to conclude whether chemical contamination is impacting benthic
6 communities; I found the components of this Approach well suited for predicting the likelihood of E. guttulata survival in Spring Creek. Each of these components is represented in later chapters, as each was used to achieve the goals of my research.
Goal 1: Describe the characteristics and ecological needs of E. guttulata
Chapter 2 - Conduct literature searches and field investigations, in order to become familiar with the characteristics and ecological needs of the “eastern green drake” mayfly. This knowledge is crucial for all aspects of this research, which include designing and conducting experiments, analyzing results, deciding whether reintroduction is feasible, and estimating the success or failure of a reintroduction program.
Goal 2: Identify pollutants in Spring Creek that may be potentially lethal/sublethal to E. guttulata
Chapter 3 - Analyze substrate and water collected from Spring Creek and Penns Creek
for the presence of two organochlorine pesticides (Kepone, mirex), and for a wide array of
environmentally toxic pollutants. The results of these chemical analyses will be used to make
inferences regarding substrate quality for both streams, and thus inferences regarding the
feasibility of reintroducing E. guttulata to Spring Creek.
Goal 3: Test the effects of Spring Creek substrate on E. guttulata larvae
Chapter 4 - Test the survivorship and growth effects of >25-day exposures to Spring
Creek and Penns Creek substrates for field-collected E. guttulata larvae. The results of this
7 viability experiment will be used to make inferences regarding substrate quality for both streams, and thus inferences regarding the feasibility of reintroducing E. guttulata to Spring Creek.
Goal 4: Examine macroinvertebrate communities of Spring Creek – are “pollution intolerant” taxa present?
Chapter 5 – Sample and identify the macroinvertebrate communities collected from
defined sections of Spring Creek and Penns Creek. The presence or absence of taxa equally or
more pollution-intolerant than E. guttulata (according to the EPA ranking of mid-Atlantic
macroinvertebrate populations) will be used to make inferences regarding the water and substrate
quality of Spring Creek, and thus inferences regarding the feasibility of reintroducing E. guttulata
to Spring Creek.
Goal 5: If feasible, design a strategy for reintroduction of E. guttulata to Spring Creek
Chapter 6 – When the first four goals have been met, I will examine the results to
evaluate the potential for a successful reintroduction of E. guttulata to Spring Creek. If I conclude that Spring Creek substrate is not likely suitable for this mayfly species, my findings will be used to make recommendations for further study to determine the reason(s), and to identify potential remedies. Should I determine that Spring Creek substrate does not appear to be unsuitable for E. guttulata , I will then design, implement and evaluate a long-term strategy for reintroduction.
8
Chapter 2
The “eastern green drake” mayfly ( Ephemera guttulata Pictet, 1843)
Introduction
The famously short lives and delicate reputation of mayflies belie the success of this ancient insect order. Originating in the early Devonian, the Ephemeroptera have been described as “the most basal, extant lineage of winged insects” (Grimaldi and Engel 2005). Mayflies pass through four stages: egg, larva, subimago (subadult, or “dun”) and imago (adult, or “spinner”).
The latter two are winged. Mayflies are the only insect s that molt after reaching a winged stage—likely a remnant ancestral trait, but functional as it facilitates the transition from aquatic to terrestrial environments, and may allow for lengthening of necessary structures (Domínguez et al
2006).
The order Ephemeroptera
Larval stage
The physical appearance of Ephemeroptera larvae is incredibly diverse, because the order as a whole has adapted to a wide range of lentic and lotic freshwater habitats. This is the longest of the four life stages (lasting several months to several years), and arguably the most important.
There are four categories of “habits” that represent not just mayfly larval locomotive behaviors,
9 but the habitats in which they dwell: “swimmers”, “crawlers”, “clingers” and “burrowers”
(Merritt et al 2008).
“Swimmers” are morphologically adapted for rapid and efficient darting movements through water. These characteristics include streamlined, piscatorial bodies, thin legs and dome- shaped heads, and cerci fringed with hairs that enable flipper-like propulsion (Edmunds et al
1976). Mayflies of the Baetidae family are classic examples of the “swimmer” type.
“Crawlers” (or “sprawlers”) are often described as looking like “bodybuilders”. Their stocky, somewhat squat build is ideal for creeping over substrate. Other features that exemplify this habit are operculate or plate-like gills, which shield the more delicate lobed ventral gills from particulate accumulation (Peckarsky et al 1990). “Crawlers” of the Ephemerellidae family also possess spines along the dorsal and lateral surfaces of the body—these uneven surfaces, and the
light coating of particles that cling to them, render ephemerellid larvae less conspicuous while
atop the streambed.
“Clingers” are dorsoventrally flattened and sprawled in appearance, with strong legs and
eyes positioned dorsally, rather than laterally. Some species possess ventral gills interlocked such
that they create an “attachment disk”, which enables larvae to adhere to surfaces within lotic
waters (Merritt et al 2008). The Heptageniidae family of mayflies best represents this habit.
“Burrowers” have long, often tube-shaped bodies and strong forelegs used for tunneling
through silt, sand, and gravel substrates. Gills are feathered, which maximizes surface area to
support a relatively high oxygen demand (Eriksen 1963). For most “burrowers”, the mandibles
are modified into tusks, and the legs (especially the forelegs) are wide; for some, a “frontal process” is present on the head (McCafferty 1975). Because of their large size, mayflies of the
Ephemeridae family are the most prominent examples of this type. The “eastern green drake”
mayfly, E. guttulata , belongs in this category.
10 Winged stages – Subimagos and imagos
Despite the great diversity displayed in larval morphology, the winged instars of
different Ephemeroptera species do not stray far from the standard form: the upright triangular
wings and curve of the body at rest, the vestigial mouthparts, the two or three cercal “tails”.
Because of their similarities, identification of adult specimens for many mayfly taxa is usually
made via differences in wing venation and male genital structures, and coloration. Adult mayflies
are sexed (i.e. categorized as either male or female) by genital structures—unlike females, males possess genital forceps, or “claspers” (indicated by arrows, Figure 2-1). (These structures are readily visible in E. guttulata larvae; therefore, they are also sexed according to these criteria).
Figure 2-1 Genital structures of male (left) and female (right) E. guttulata.Arrows point to genital forceps (“claspers”) of the male.
Subimagos (subadults, or “duns”) resemble imagos (adults, or “spinners”), but have clouded wings and are dulled in luster by a dense covering of microtrichia. These hairs repel water, which facilitates the mayfly’s transition from aquatic to terrestrial habitats. Subimaginal females of some species are capable of mating and producing viable offspring, but in general, mayflies in this stage are not reproductively mature.
The timing of subimago emergence is heavily influenced by environmental conditions.
Relatively warm/dry winters and springs tend to elicit earlier emergences (“hatches”). The duration of the subadult stage in mayflies varies between minutes and days, depending on the
11 species. Compared to the timing of emergence, the duration of the subimaginal instar appears to be influenced less by average temperatures and amount of winter/spring precipitation (personal
observation).
After their final molt, adult mayflies are clear-winged and free of microtrichia. As in the
subadult stage, adult mayflies lack functional mouthparts, and subsist solely on energy stores
accrued in the larval stage. These energy stores must sufficiently fuel male mating swarms,
male/female mid-flight copulation, and female oviposition (“spinner fall”). Males possess large
eyes to seek mates, and long forelegs, used in tandem with genital forceps to grasp and mate with
females. Once fertilized, different species of mayfly females oviposit using a variety of methods,
from dropping egg masses into the water, to attaching their eggs to the underside of submerged
rocks, to forgoing oviposition in favor of a two-week long period of ovoviviparity
(Sivaramakrishnan and Venkataraman 1985).
Ecological importance
One of most important characteristics of mayflies is their nutritional value. Mayflies are
consumed by at least 224 species of predators—exclusively so by an immature freshwater
stingray (Grant 2001)—and are food to economically important species such as fish and
waterfowl. Mayflies are also an important staple in some human diets, as they are not only easy to
catch, but also high in protein, minerals, B vitamins, essential amino acids, and “six times as
much iron as ox liver” (Shaxson et al 1979). For example, the peoples of the Lake Victoria and
Lake Malawi regions of Africa grind mayflies and other “lake flies” (chaoborid and chironomid
dipterans) into kungu paste (if boiled) or flour (if sun-dried), which they use to prepare dried
cakes. As these cakes are low in fat and moisture, they have a relatively long shelf life and are potentially a significant source of nutrition (Bergeron 1988).
12 Ephemera guttulata Pictet, 1843
Description
In his description of the species, François Jules Pictet distinguished imagoes of E. guttulata from other Ephemera species by their anterior wings—those of E. guttulata are covered with numerous spots (“ailes antérieurs couvertes de taches nombreuses”), while those of other
Ephemera sp. have no more than three or four spots (Pictet 1843. “g uttulatus ” is Latin for “little
spots”).
Larval stage – In its larval stage, E. guttulata presents as a large (12-32 mm), slender,
three-tailed burrowing mayfly with dorsally oriented gills (Figure 2-2). On the first abdominal
segment the gills are bifurcate and rudimentary; on the second through seventh abdominal
segments the gills are plumose and significantly larger. Like other Ephemera species, the
mandibles of E. guttulata are modified into generally spineless, upward-curved tusks; the prothoracic tibiae are distally rounded and without a process; and a bifurcate frontal process is present on the head. These characters are adaptations conducive to tunneling through substrate
(Needham 1935, Edmunds 1976, Merritt et al 2008).
13
Figure 2-2 Ephemera guttulata larva.
E. guttulata larvae are distinguished from other Ephemera species by the following characters: when viewed from above, the base of each tusk lies between an antenna and its adjacent frontal process, and the outer curve of the tusk is not visible. The spined basal half of each tusk and the basal two-thirds of each antennae possess long setae (Kennedy 1925,1926;
Needham 1935; McCafferty 1975). Abdominal stergites are uniformly pale and lack dark pigmentation (Brigham et al 1982; G. Hoover, personal communication). Also, the frontal process of this species is more deeply bifurcate than those of its congenerics (eg. E. simulans, E. varia) —
the deep bifurcation and the lack of abdominal pigmentation appear to be stable characters for
identification to species (G. Hoover, personal communication).
Subimaginal stage – The subadult E. guttulata presents as a large and visually striking mayfly, having a yellow head, a black and yellow thorax, a yellow abdomen with longitudinal black stripes and tipped with three long caudal filaments, and green opaque wings with black spots (Figure 2-3). Average wing length is 10-15mm (Edmunds 1976).
14
Figure 2-3 Ephemera guttulata subimago (female).
Imaginal stage – Adult “green drake” mayflies (Figure 2-4) are truly distinctive and easily seen, as they are large mayflies with a chalk-white abdomen and black thorax (Kennedy
1925, 1926; Needham 1935; McCaffterty 1998; Hoover 1978). At Penns Creek, the abdomens of adult males are much whiter than those of adult females (personal observation).
15
Figure 2-4 Ephemera guttulata imago (male).
Life history
The larva of E. guttulata dwells within erosional and depositional sand and gravel substrates of lotic systems, and coarse sand of pools and lakes (Kennedy 1925, 1926; Hynes
1970; McCafferty 1975; Edmunds 1976; Hoover 1978). The E. guttulata of Penns Creek are more readily found in riffles than in pools. Mature female larvae are larger than males, and there may be sex differences in substrate size preference—with females tending to inhabit substrate comprised of larger particles than by the males (Hoover 1978).
E. guttulata larvae of Pennsylvania are classified as collector-gatherers (Merritt et al
2008). Hoover (1978) performed gut analyses, and found presence of diatoms, plant detritus, and mineral material (for possible gizzard function, Cummins 1973). E. guttulata larvae are mainly nocturnal, and display pronounced photophobic behaviors, although they will occasionally dart
16 out of the substrate during the day if foraging or if disturbed (Kennedy 1925, 1926; personal
observation).
The voltinism of E. guttulata varies latitudinally—from semi- to partivoltine in its
northernmost range (Canada), to univoltine in its southernmost (Georgia). In central Pennsylvania
streams, E. guttulata is believed to be semivoltine, with yearly cohorts (i.e. overlapping
generations) (Hoover 1978). If true, then collectors should typically find two generations of
larvae at most times of the year. Sexing is critical, however, as these larvae exhibit strong sexual
dimorphism—two size classes do not necessarily indicate two cohorts. Further examinations of
the voltinism of this species may yield fascinating results.
In Pennsylvania, subimagoes emerge from late May through mid-June, depending on the
stream (Meck 1997), and depending on winter/spring total precipitation and average
temperatures. Males appear to emerge several hours to one day before females (personal
observation). Kennedy (1925) observed pronounced positive phototropism in E. guttulata
subimagoes, even after emergence, and noted that this behavior differs from other Ephemera
species. The total duration of the subimaginal stage is 1 to 2 days.
Imagoes of this species mate at twilight, from roughly 8pm until well after dark (the
“latest on the wing in the evening”, Kennedy 1925). The mating flight of the males consists of a
sequence of several wing strokes to ascend vertically, and then passive, floating descent to less
than 2 meters above the water’s surface (personal observation). Typically, adult females fly into
these swarms, and mating will occur in flight; however, mating pairs have been observed on the
underside of leaves (Edmunds 1976). After mating, Kennedy observed negative, photophobic behaviors in the females (Kennedy1925), as the females appeared to prefer the more shaded
regions of the stream for oviposition.
E. guttulata females produce roughly 3000-4000 eggs (Hoover 1978), and they skim just
above the water while dipping their abdomens several times to oviposit. Females die soon after
17 oviposition, and the water’s surface is littered with spent adults by nightfall. Egg development
time is temperature dependent—Hoover (1978) found that first instars emerge from the eggs after
27 to 32 days when water temperatures were held at 15.5 ◦ C. In the Ichthyology Lab at the Rock
Spring Agricultural Research Facility of the Pennsylvania State University, first instars were first
observed after 23 days, when water temperatures were kept at 17 + 3 ◦ C.
Mayflies are sensitive to high heat and low dissolved oxygen levels, but there is a significant range of organic pollution tolerance values, within the Order and between taxonomic levels, from fairly tolerant to near-absolute intolerance. According to the North Carolina Biotic
Index (NCBI), which rates sensitivity to “general pollution” (e.g. sediment, litter, fertilizers and pesticides) on a scale of 0 (very intolerant) to 10 (very tolerant), Ephemera spp. have an average
tolerance value of 2.2, but E. guttulata has a value of 0 (Lenat 1993). The Wisconsin Biotic Index
(WIBI), which rates sensitivity to organic pollution on a scale similar to the NCBI, lists
Ephemera spp. with an average tolerance of 1 (Hilsenhoff 1987). Because they burrow and feed
in substrate, burrowing mayflies in general are especially sensitive to siltation, and to lipophilic pollutants, as they will sink through the water column, then accumulate and persist in sediment.
Burrowing mayflies have been proposed as being especially useful indicators of water and sediment quality due to their historical abundance, intolerance of organic/”general” pollution and low dissolved oxygen concentrations, ability to recover following pollution abatement (e.g. the Hexagenia species of the Great Lakes), their ecological importance as “bioturbators” of sediment and as food for fish, and their highly visible mating flights (Edsall 2001). The “eastern green drake” mayfly is important for the above reasons, and with its sportfishing status has great potential as a representative of freshwater conservation in Pennsylvania.
18
Chapter 3
Chemical analysis of stream substrates: Spring Creek vs. Penns Creek
Introduction
Organochlorine pesticides
Organochlorine (OC) pesticides are a class of generally stable, environmentally persistent compounds which include the powerful insecticides DDT, chlordecone (Kepone), mirex (Declorane), endosulfan (Thiodan, Thionex), chlordane (Chlordan) and lindane (Kwell).
Most OC pesticides are highly lipophilic and will readily accumulate in fatty tissues—in the
1980s, the U.S. EPA estimated that 10.2% of the human population of the southern U.S. had measurable levels of mirex in their adipose tissue (Kutz 1985). Organochlorine pesticides are potent excitatory neurotoxins and are highly effective insecticides; however, in mammals, OCs
have been shown to act as teratogens and as endocrine disruptors, by binding to estrogen
receptors. In birds, OCs are thought to block calcium absorption, thereby weakening eggshells.
Due to their persistence and toxicity, many OC pesticides have been discontinued in developed
countries for over 20 years, but continue to be detected in living organisms, soils, and sediments.
Stream sediments act as sinks for persistent environmental contaminants, which heavily
impact sensitive benthic macroinvertebrates such as mayflies. Hose and Wilson (2005) proposed
that while average daily concentrations of endosulfan in Australia’s Namoi River were not toxic
to the leptophlebiid mayfly Jappa kutera , storm runoff, overspray from fields, and low-level
19 chronic exposures could raise endosulfan concentrations in the mayfly to 70 µg/L –an order of magnitude above its LC 50 value.
Studies of sublethal effects of OC contamination have also yielded significant results. In the the well-known model of food web biomagnification, pesticide levels are expected to increase with higher trophic levels. However, Campbell et al (2000) found that trophic level did not predict levels of contaminants as well as the relative lipid content of the organisms used in the study. Just as mammalian mothers accumulate OC pesticide residues in breast milk, vertical transmission of chlordane from females of the baetid mayfly, Centroptilum triangulifer to their eggs has been shown by Standley et al (1994). Interestingly, the mayfly’s chlordane “metabolite fingerprint” –the levels of individual metabolites present following a metabolic reaction—more closely resembled humans, seals, and porpoises than that of krill and plankton, which are organisms at the same trophic level as C. triangulifer (Standley et al 1994).
There is a substantial foundation of knowledge regarding OC pesticide impacts on higher trophic level organisms; however, the lethal/sublethal effects and dynamics of OCs within mayflies and other sensitive benthos are less well-known. Despite short-term tests of mirex toxicity that have revealed the resilience of several invertebrate taxa to 4-day exposures (Sanders et al 1981), latent mortality, bioaccumulation, and sub-lethal effects have been documented in aquatic invertebrates (Eisler 1985). Because of uncertain lethal levels, their potential sub-lethal effects, and their environmental persistence, the presence of OC pesticides in stream substrate is of great concern for burrowing mayflies.
The start of OC production by Rutgers-Nease Chemical (1959) and the disappearance of
Spring Creek’s “green drakes” in the late 1950s have led many in the local fishing community to assume that OC pesticides are responsible for the decades-long absence of E. guttulata from
Spring Creek. In order to determine whether present-day Spring Creek substrate and water are
20 potentially suitable for E. guttulata survival, it was first important to identify any environmentally hazardous compounds present in these media.
Objectives
The purpose of the chemical analyses was to test for measurable levels of Kepone and mirex, as well as for the presence of other environmental contaminants, in surface water and stream substrate samples collected from three sites on both Spring Creek and Penns Creek. Based on the persistence of OC pesticides, I hypothesized that the Spring Creek samples would contain measurable quantities of Kepone and mirex, and that Penns Creek will not. Based on Spring
Creek’s history, I also predicted that other environmental contaminants will be detected in Spring
Creek’s samples in greater concentrations than in Penns Creek’s samples.
Methods
Selection of sampling sites
Three substrate and water sampling stations were chosen from Spring Creek, and three from Penns Creek:
SC1 (“Fisherman’s Paradise”)
This site (40 ◦52’45.72” N, 77 ◦47’30.38” W) is approximately 186 meters upstream of the
metal footbridge at the Pennsylvania Fish and Boat Commission’s Bellefonte office, on Spring
Creek Road above “Fisherman’s Paradise” (Benner Township.). The location of “SC1” was
21 chosen because it is situated downstream of “Spring Creek Canyon”—a pristine 730 hectare (ha) landscape that has been under the protection of both the Rockville State Correctional Institution and the Pennsylvania Fish and Boat Commission since 1925.
SC2 (“the Rock”)
This site (40 ◦51’02.54” N, 77 ◦49’19.51” W) is approximately 12.8 meters upstream of the large parking area/metal bridge along Rock Road in Benner Township, and approximately 7.27 km upstream of SC1 . “SC2” is situated within the historic range of E. guttulata (Aurelio 1953). It was also the site of substrate collection for the bioassessment (see: Chapter 4), and 75.7 meters upstream of a community sampling station (see: Chapter 5) and the site of annual E. guttulata egg transplants, beginning in 2009 (see: Chapter 6).
SC3 (“Houserville”)
This site (40 ◦49’05.51” N, 77 ◦49’09.96” W) is immediately downstream of the
confluence of Thornton Spring and Spring Creek, approximately 32.8 meters downstream of the
intersection of PA State Route 26 and Houserville Road (College Township), and approximately
7.36 km upstream of SC2 . The location of “SC3” was chosen for its history of point- and non- point source pollutants (including Kepone, mirex, untreated sewage, gasoline, and state highway runoff).
22 PC1 (“end of Tunnel Rd.”)
This site (40 ◦50’56.85” N, 77 ◦27’3.86” W) is at the end of Tunnel Road in Penn
Township (Centre County), and approximately 58.8 meters upstream of a wooden foot bridge that crosses the stream. “PC1” was also the location of a community sampling station (see: Chapter
5).
PC2 (“cement bridge along Tunnel Rd.”)
This site (40 ◦51’0.35” N, 77 ◦27’20.19” W) is approximately 1.57 km upstream of PC1 ,
and approximately 79.9 meters upstream of a small cement bridge located on Tunnel Road in
Penn Township (Centre County), southeast of Coburn proper. Like PC1 , “PC2” was also the
location of a community sampling station (see: Chapter 5).
PC3 (“Coburn”)
This site (40 ◦51’39.23” N, 77 ◦27’42.33” W) is approximately 42.6 meters upstream of the Pine Creek/Penns Creek confluence in the town of Coburn (Penn Township, Centre County), and approximately 1.45 river mile upstream of PC2 . This location was selected because it is the area of Penns Creek from which a sufficient number of E. guttulata larvae have been collected for
14 replicates (reps) of a bioassessment (see: Chapter 4). “PC3” is approximately 100 meters
upstream of the location of a community sampling station (see: Chapter 5).
23 Collection of samples
Substrate was collected via a hand-held grab sampler, according to the procedures listed in section 7.2.1 of the USEPA “Sediment Sampling” guidance document (SOP #1215), and following the targeted sampling design as outlined in the “Methods for Collection, Storage and
Manipulation of Sediments for Chemical and Toxicological Analyses: Technical Manual” (EPA-
823-B-01-002). Surface water was collected using the dipping method described in Section 4 of the USEPA “Surface Water Sampling” operating procedure (SESDPROC-201-R1). Between each station, sampling equipment was decontaminated using the “Field Sampling Equipment Cleaning
Procedures” as described in the USEPA “Sampling Equipment Decontamination” guidance document (SOP #109). The number of samples collected from each site was determined according to the capabilities of the lab performing the sample preparation and analyses.
Substrate
All substrate grab samples were collected with a 500 mL stainless steel cup (Tomac,
#13850-005), at a minimum depth of approximately 30 cm. Three samples were collected at each station; these three samples were poured into one 600 mL Low Form glass Griffin beaker (Pyrex,
#70000-600) and mixed streamside using a PTFE-coated lab spoon (Lamson & Goodnow,
#21800). The homogenized substrate sample was then emptied into a 500 mL wide-mouth, clear- glass jar (VWR International, #89043-382) and sealed with its PTFE-coated lid.
Water
At each station, one surface water sample was collected directly into a 500 mL amber glass bottle (VWR International, #15900-140), then sealed with its PTFE-coated cap.
24 Preparation and analysis of samples
The complete procedure used for the preparation and analyses of substrate and water samples followed EPA Method 8081A – “Organochlorine Pesticides by Gas Chromatography”
(website: http://www.caslab.com/EPA-Methods), and was performed as a laboratory exercise and practice “case study” for a Separations course taught by Dr. Frank Dorman, Associate Professor of Biochemistry and Molecular Biology in the Department of Forensic Science at the
Pennsylvania State University. The three Forensic Science graduate students/ Environmental
Laboratory Analysts acting under the tutelage of Dr. Dorman were Seth E. Michalski, Beth
Slaybaugh, and Jessica Westland.
Surface water and stream substrate samples were retained in the Pennsylvania State
University’s Environmental Laboratory at 347 Davey Lab (University Park, PA), to allow for future analyses.
Substrate
Substrate samples and blanks were prepped using a Soxhelt extractor and concentrated down using a Kuderna-Danish apparatus. The resulting extracts were then analyzed using Gas
Chromatography (GC) with Mass Spectrometry (MS) detection.
Water
Water samples and blanks were first prepared via liquid-liquid extraction, then concentrated using a Kuderna-Danish concentrator. The resulting extracts were then analyzed using GC-MS detection.
25 Results
A tabular list of the compounds and their concentrations (when available) that were detected in the substrate and water samples from both streams is located in Appendix A:
“Pollutants detected in substrate and water collected from Spring and Penns Creeks ”.
Known environmental contaminants were detected in all three of the substrate samples collected
from Spring Creek, and two of the three substrate samples collected from Penns Creek. Known
contaminants were also detected in two of the three water samples for both streams. Due to poor
sample quality, water data are not available for The Rock (Spring Creek) and for the Cement
Bridge (Penns Creek).
Spring Creek
SC1 (“Fisherman’s Paradise”)
Substrate – There were no measurable quantities of Kepone or mirex found in the
substrate collected from the Fisherman’s Paradise site on Spring Creek, but there were
measurable quantities of five polycyclic aromatic hydrocarbon (PAH) compounds and one phenol
detected in the substrate sample.
Water – There were no measurable quantities of Kepone or mirex found in the water
samples collected from the Fisherman’s Paradise site. A tetrahydrobenzo[a]pyrene was detected
at an estimated concentration of 0.0003 ng/µL, or 0.3 ppb.
26 SC2 (“The Rock”)
Substrate – A measurable quantity of mirex was found in the substrate collected from The
Rock site on Spring Creek. There were also measurable quantities of twelve PAHs, one phenol, two halogenated organic compounds (HOC), one phenol/HOC, one nitroaromatic compound
(NAC), and one aldehyde detected in the substrate sample.
SC3 (“Houserville”)
Substrate – No measurable quantities of Kepone or mirex were found in the substrate collected from the Houserville site on Spring Creek. There were measurable quantities of fourteen PAHs, twelve phenols, one HOC, one phenol/HOC, one NAC, and one alcohol/polyol detected in the substrate sample.
Water – No Kepone or mirex was detected in the water samples collected from the
Houserville site. However, there were measurable quantities of one PAH (p-terphenyl-d14), one phenol (phenol-d6), one HOC (2-fluorobiphenyl), one phenol/HOC (2,4,6-tribromophenol) and
one NAC (nitrobenzene-d5) present in the Houserville water sample.
Penns Creek
PC1 (“End of Tunnel Rd.”)
Substrate – No mirex, Kepone or other environmental contaminants were detected in the
substrate sample collected from the Penns Creek site at the end of Tunnel Road.
27
Water – No mirex or Kepone was detected in the water sample collected at this site, but there was a measurable quantity of one phenol detected.
PC2 (“Cement bridge along Tunnel Rd.”)
Substrate – No mirex or Kepone was found in the substrate collected from the Cement
Bridge site on Penns Creek. However, there were measurable quantities of thirteen PAHs, one phenol, one HOC, one phenol/HOC, one aldehyde, and one ester detected in the substrate sample.
PC3 (“Coburn”)
Substrate – No mirex or Kepone was found in the substrate collected from the Coburn
site on Penns Creek. There were measurable quantities of five PAHs, three phenols, one HOC,
one phenol/HOC, one NAC, and one alcohol/polyol detected in the substrate sample.
Water – No mirex or Kepone was detected in the water sample collected at Coburn, but
there were measurable quantities of one PAH (p-terphenyl-d14), one phenol (phenol-d6), one
HOC (2-fluorobiphenyl), one phenol/HOC (2,4,6-tribromophenol) and one NAC (nitrobenzene-
d5) present in the Coburn water sample.
28 Discussion
Pollutant levels in Spring and Penns Creeks
Spring Creek sites
Organochlorines – Of the three Spring Creek sites, mirex was detected in the substrate collected from The Rock site, but at 0.018 ppb was 1.8E-04 of the FDA action threshold of 0.1 mg/kg. To protect aquatic life from any possible harmful effects, the EPA has set an upper limit of 0.001 ppb for mirex in surface water, but no mirex was detected in the surface water samples from either stream.
Other pollutants – The presence of numerous pollutants in the Houserville samples was not surprising, but the number and concentrations of PAHs, phenols, and other hazardous compounds detected at this site was alarming. The presence and prevalence of these compounds decreased further downstream from the sampling site.
The Houserville site is located immediately downstream from a busy intersection, so diesel exhaust may be at least partially responsible for the high levels of PAHs and phenols found at this site. The site is also immediately downstream from the location of a convenience store’s underground gasoline tanks—tanks that in 2006 leaked gasoline into Spring Creek (College
Township Ordinance O-06-14). Dr. Frank Dorman, who supervised the preparation and analyses of the samples, is interested in further study of this site, to triangulate and identify the source of these pollutants.
There were a total of 30 compounds detected at the Houserville site, with a mean concentration of 10472.0977 ppb. There were 19 compounds detected at The Rock site (mean =
1.3333 ppb), and six detected at Fisherman’s Paradise site (mean = 0.0689).
29 Penns Creek sites
Organochlorines – Neither Kepone nor mirex was detected in any of the Penns Creek water and substrate samples.
Other pollutants – Considering Penns Creek’s reputation as a pristine waterway, the presence of numerous pollutants in the Coburn samples, and the number and concentrations of
PAHs, phenols, and other hazardous compounds detected at this site were unexpected. The number of compounds was greatest at the Cement Bridge site (the middle site), but the concentrations were highest at the Coburn site (the most upstream site). According to the analyses, the furthest downstream site was the “cleanest” of all of the sites.
There were a total of 12 compounds detected at the Coburn site, with a mean concentration of 15878.6690 ppb. There were 18 detected at the Cement Bridge site (mean =
0.5725 ppb), and 0.0982 ppb of one compound was detected at the End of Tunnel Road site.
Implications
Stream biota and human health
Considering the high concentrations of toxic compounds recovered at Spring Creek’s
Houserville site and Penns Creek’s Coburn site, further research is essential to examine the effects of these pollutants, and at concentrations equal to those found by Dr. Dorman and his students. Also, additive effects are unknown for numerous combinations of compounds.
“Traditional” bioassays of compounds detected in Spring Creek and Penns Creek samples and using a number of native macroinvertebrate taxa would be ideal, though costly, next steps.
For this study, water and substrate were sampled on one day, and at three sites per stream. Sampling more frequently and at a greater number of sites would allow for a longitudinal
30 view of trends, and would compensate for possible “blips” due to temperature, precipation, stream flow, specific incidents, etc. Just as high levels are not indicative of a chronically polluted stream, absence or low levels of contaminants found in stream water and substrate do not necessarily mean that the waterway is “clean”. Insoluble or dense pollutants can settle out of the water column and accumulate in sediment. Uptake by stream biota can also result in false negatives, as well as magnification through ascending trophic levels. In the interest of gaining truer, more comprehensive knowledge of the contamination status of a waterway, examinations of fish and invertebrate tissues should be performed in addition to analyses of water and stream sediment. Unfortunately, such examinations could not be included in this study. Future studies should not make this omission.
The results of their findings has piqued the interest of another graduate student working with Dr. Dorman, and currently in progress is an “Environmental Forensics” research project examining potential sources of the high PAH levels in the vicinity the Houserville site. It will be interesting to compare my findings with any new data gleaned from that project.
Reintroduction of Ephemera guttulata
While the initial demise of the E. guttulata population in Spring Creek is attributed to a
variety of pollutants—most especially to cyanide and chlorine that entered the stream, and to the
leaching of OC pesticides into groundwater—there are undoubtedly a number of other factors that
could have prevented the species from repopulating the stream.
One such factor is the dispersal ability of a mayfly. “Eastern green drake” recolonization
of the Little Juniata River has been attributed to a rescue effect courtesy of the river’s tributaries.
Spring Creek’s few tributaries do not support E. guttulata populations, and the inter-catchment dispersal ability of mayflies is most likely only passive, which would be limited by physical
31 barriers such as mountains (Smith and Collier 2006) or culverts (Blakely et al 2006). The role of
endemism, as well as any other possible factors, should be explored further.
Though the upstream area of Spring Creek has been degraded by the effects of
urbanization and agricultural land-use, the area of Spring Creek located within the historic range
of the extirpated E. guttulata population appears to be free of OC pesticide residues. Also, the number and concentrations of non-OC contaminants detected at The Rock and at Fisherman’s
Paradise sites were much lower than at the Coburn site—an area which, despite PAH and phenol concentrations comparable to the Houserville site, supports a famously dense population of
“eastern green drake” mayflies. The results of the chemical analyses of substrate and water collected from Spring Creek and Penns Creek cannot explain the presence of E. guttulata at the
Coburn site, yet their absence at The Rock or at Fisherman’s Paradise sites. There is no indication that the amounts of OC pesticides or other pollutants detected in the species’ historic range in
Spring Creek would hinder the restoration of an “eastern green drake” mayfly population.
32
Chapter 4
Bioassessment: The effect of Spring Creek substrate on survivorship and growth of E. guttulata larvae
Introduction
Testing the biotic responses of organisms to the environment into which they will be transplanted is an essential step in the reintroduction process. Burrowing mayflies spend the majority of their larval stages within the substrate, feeding on benthic detrital, mineral and protist material (Cummins 1973; Hoover 1978); therefore, burrowing mayflies are especially sensitive to substrate quality.
Bioassays, bioassessments and viability experiments
A bioassay in the traditional sense is a series of toxicity tests using live organisms, in which they are exposed to a range of concentrations of a (suspected) toxicant. Survivorship data from these tests are then used to calculate the EC 50 (median effect concentration: the
concentration at which toxic effects such as reduced motility are present in exactly 50% of the
organisms) and the LC 50 (median lethal concentration: the concentration that is lethal to exactly
50% of the organisms) of that substance.
Not all bioassays follow this design. “Bioassay” has been more broadly defined as “the use of living organisms to detect the presence, or concentration of a toxic substance.” (Muirhead-
Thomson 1987). In this sense, “bioassays” can be used when the presence and identity of toxicants are unknown. Because this study is not a traditional “bioassay”, I avoid the semantic
33 and methodological arguments by instead deeming this as a “bioassassement”, or “viability experiment”.
For this study, the purpose of the bioassessment is to determine whether Spring Creek substrate is habitable, and is not toxic, to E. guttulata larvae. Because mirex was found in minute concentrations at only one of the three Spring Creek sites (Chapter 3), performing a “traditional” bioassay using graduated levels of these organochlorine compounds was not deemed necessary.
Field vs. laboratory bioassessments
There are two main bioassessment methods for aquatic taxa of lotic systems: laboratory bioassessment and in situ bioassessment. Laboratory methods offer the investigator greater
control over “stream” conditions, but can be cost-prohibitive and more prone to human error and
mechanical failure. Field methods are relatively inexpensive and expose organisms to more
natural conditions, but drawbacks include reduced flow within and deposition of fine sediments
caused by clogged mesh, and periphyton community changes resulting from the shading of
substrate (Peckarsky and Penton 1990). Field sites are also more vulnerable to vandalism and to
catastrophic events (eg. floods). Food availability may also be a confounding factor in field bioassessments using survival and growth as endpoint measurements (DeLange et al 2005).
Though initially the experiments were planned for the field, the opportunity to use Dr. Jay R.
Stauffer Jr’s Ichthyology Lab arose—all of the necessary equipment was available at the
Ichthyology Lab. Only minor modifications were needed for my purposes, so Dr. Stauffer’s very
generous offer was readily accepted.
34
Objectives
The goal of this bioassessment, or “viability experiment” (vEXP), is to predict the survival of E. guttulata larvae in Spring Creek. More specifically, the objective of the experiment
detailed in this chapter is to observe the effects of Spring Creek substrate on > 25-day survivorship and growth of E. guttulata larvae collected monthly from Penns Creek.
Methods
Replicates (“reps”) 1 through 11 of the vEXP were conducted in a recirculation system located in the Ichthyology Laboratory at Pennsylvania State University’s Rock Springs
Agricultural Research Facility (Figure 4-1). Reps 12 through 14 of the vEXP were conducted in a
static, temperature-controlled, and oxygenated system located at a private residence (Figure 4-2).
Figure 4-1 Recirculation system used in vEXP Reps 1 – 11. Ichthyology Lab, School of Forest Resources. Rock Springs Agricultural Research Facility. Pennsylvania State University.
35
Figure 4-2 Static system used in vEXP Reps 12 – 14. Private residence, State College PA.
Design of laboratory equipment
Reps 1 through 11
Each stream was represented by an interconnected series of eight ten-gallon (37.85 liter) aquaria that flowed through a 100-gallon (378.54 liter) cistern, and chillers placed in each cistern maintained water temperatures of 20 + 2 ◦C. Water from each cistern fed an outflow (and an overflow) pipe, and water from each tank passed through an inflow filter and fed into a common inflow pipe leading back to the cistern. Onsite water was used to fill the tanks and cisterns, and added to each system as needed.
36
Reps 12 through 14
Reps 12 through 14 of the vEXP were conducted in six individual 8-quart foam coolers
(Marine Metal Products, CB-11) located at a private residence. The foam’s insulative properties allowed the water in each cooler to remain at 20 + 2 ◦C for the duration of each rep. For each of
the six coolers, water was oxygenated via a 3-watt air pump (Clear-Seal, AC-9901) connected to
the 7/8” (22.23 mm) airstone that was included with the cooler.
The change from lab to private residence was due to several factors. In pilot studies, the percent survivorship of “residence” controls was more favorable than that of the “laboratory” controls of Reps 1 through 11. Also, the Ichthyology Laboratory and its equipment was needed for other experiments. Finally, conducting the final three reps in the private residence was more convenient for daily monitoring of the equipment and larvae.
Design of rearing cages
For reps 1 through 11, floating rearing cages (Figure 4-3a ) similar in design to Edmunds et al 1976 and Tseng 2003 were constructed from 24 oz (709.76 mL) transparent plastic cups
(Dart, #24CDART), with two 11 cm (top) X 9.4 cm (sides) X 9.1 cm (bottom) opposing sections cut 4.6 cm below the top lip of the cup. These sections were replaced with 0.011” diameter
(appx. 280 micron) fiberglass screen (New York Wire, #30207), affixed to the inside of the cup with 100% silicone sealant (GS Silicone II, #GE284). To support and float the cups inside each aquaria, six 9 cm diameter holes were drilled in a 2 x 3 configuration into 49.5 cm X 24.5 cm sheets of 1”(2.54 cm) thick extruded polystyrene (CodeBord, #270457) (Figure 4-3b ). Each sheet held four cups—one at the front, and three at the back. To facilitate circulation and aeration, the front right hole was cage-less to accommodate the outflow pipe, and a section of foam in the front left position was cut to fit around the inflow filter.
37
a. b.
Figure 4-3 a. Rearing cage used in Reps 1 – 11 b. Illustration of floating foam support for rearing cages used in Reps 1 – 11.
Collection of larvae
Ephemera guttulata larvae were collected from Penns Creek at the town of Coburn
(Centre County), as the population of this species is famously abundant at this location, and because previous collections at Coburn were consistently and reliably successful.
During the preliminary experiment, it was found that the most efficient and effective
method of collecting E. guttulata larvae requires two people: one person to use a garden shovel to
disturb gravel-sand deposits below rocks, and the other to hold a 4’ X 4’ (1.22 m X 1.22 m) kick
net immediately downstream of the disturbance. After several digs with the shovel, both
collectors carefully raise the kick net. One person holds the net flat and taut, and the other
transfers E. guttulata larvae from the net to a water-filled jar. When 10 or more larvae are
collected, the jar’s contents are carefully emptied into a thick, cylindrical 8-quart (7.57 L) polystyrene cooler (Marine Metal Products, #MMP-CB-11). Aeration of coolers during transport
38 was not necessary, as the turbulence created is sufficient to maintain suitable oxygenation. Upon return from the field, coolers were aerated by portable pumps and air stones. For the 24-hour holding period before the start of the bioassessment, coolers sat overnight in a secure and cool location. The following morning, larvae were measured, sexed (in Reps 5 and 7 – 14), then added to the substrate-filled rearing cages or coolers. Larvae that did not survive or that were apparently injured or unhealthy were retained in vials or jars of 90% ethyl alcohol as voucher specimens.
Collection of substrate
A mixture of gravel and sand substrate was collected from an area of Penns Creek known to support dense E. guttulata populations (Coburn), and from an area of Spring Creek within the mayfly’s historic range (“The Rock”). Preference for the size and composition of substrate to collect was confirmed when mature larvae were found in the Penns Creek substrate. In a preliminary experiment, substrate for the bioassessment was frozen for 10 days, thawed, then homogenized by removing large pebbles and other objects (eg. glass, shells) and mixing thoroughly by hand within a hard plastic “kiddie” pool. The purpose of this was to minimize the influence of other taxa (benthic or planktonic) on E. guttulata survivorship and/or growth. In these 14 vEXP reps, no such pre-experimental treatment to the substrate was applied, in order to gain a more realistic understanding of its influence on survivorship and growth.
Pre-assay measurement of larvae
Immediately before placement into a rearing cage or cooler, the head capsule width of each larva was measured at the 0.1 mm scale. “Head capsule width” was defined as the distance between the outer edges of the left and right eye (Figure 4-4). Larvae in Reps 1 – 4 and 6 were
not sexed pre-assay, but survivors were sexed post-assay. For Reps 5 and 7 – 14, larvae were
39 sexed pre-assay, and survivors were sexed post-assay. Dead larvae were sexed whenever possible
(i.e. when morphological characters were sufficiently intact for identification).
Figure 4-4 “Head capsule width”, as defined for measurement (pre- and post-assay).
Viability and growth assay
Reps 1 through 11 (Table 4-1) were conducted in the Ichthyology Laboratory at Penn
State’s Rock Springs Agricultural Research Facility. For these Reps, substrate collected from
Penns Creek was pooled, mixed by hand, and 11 oz (325.31 mL) of Penns Creek substrate was added to the rearing cages representing the Control treatment. The same was done with the Spring
Creek substrate, and an equal amount of Spring Creek substrate was added to the cages representing the Experimental treatment. Equipment, tanks, rearing cages and larvae were monitored periodically, at least weekly. Equipment (eg. chiller, water pump) and tank failures occurred occasionally, and repairs were made as quickly as possible. Larval monitoring consisted of removing and preserving (in 90% ethyl alcohol) any dead larvae, usually on the surface of the substrate (dying larvae tend to emerge from their burrows). For each rearing cage in which a dead larva was not found, the contents of the cage were emptied into a sorting tray, then examined for
E. guttulata larvae (Figure 4-5). Missing larvae were noted, but not yet presumed dead. In the case of live larvae, substrate was returned to the rearing cage, the rearing cage was returned to its position within the floating foam support, and the larva returned to the rearing cage. Larval
40 behavior when returned to the water was noted (eg. activity level, burrowing ability), but interaction was kept to a minimum, to limit stress.
Rep Larval Duration of Rep Larvae Number of Rearing Number of Sex Configuration Collection Date (Dates) sexed? Cages per Tank Larvae in Rep per Tank 1 4 May 2009 65 days no 4 (2 Control, 2 EXP) 32 (16 Control, 16 EXP) n/a (5 May - 8 Jul 2009) 2 20 June 2009 65 days no 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) n/a (21 Jun - 24 Aug 2009) 3 15 July 2009 50 days no 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) n/a (16 Jul - 3 Sep 2009) 4 16 August 2009 37 days no 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) n/a (17 Aug - 22 Sep 2009) 5 10 September 2009 30 days yes 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) 5 tanks: 1 M and 1 F (11 Sep - 10 Oct 2009) 3 tanks: 2 F 1 6 6 October 2009 31 days no 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) n/a (7 Oct - 6 Nov 2009) 7 12 November 2009 25 days yes 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) 4 tanks: 2 M (13 Nov - 7 Dec 2009) 4 tanks: 2 F 8 20 December 2009 26 days yes 2 (1 Control, 1 EXP) 12 (6 Control, 6 EXP) 2 3 tanks: 2 M (21 Dec 2009 - 15 Jan 2010) 3 tanks: 2 F 9 14 January 2010 32 days yes 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) 4 tanks: 1 M and 1 F (15 Jan - 15 Feb 2010) 2 tanks: 2 M 2 tanks: 2 F 10 21 February 2010 33 days yes 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) 1 M and 1 F (22 Feb - 26 Mar 2010) 11 25 March 2010 21 days yes 2 (1 Control, 1 EXP) 16 (8 Control, 8 EXP) 1 M and 1 F (26 Mar - 15 Apr 2010) 1 for Rep 5, an insufficient number of apparently healthy males was collected on the previous day 2 for Rep 8, the number of apparently healthy larvae collected was too low to utilize all 16 rearing cages
Table 4-1 Design of Reps 1-11 of the viability experiment.
Figure 4-5 Sorting tray with contents of a rearing cage and live E. guttulata larva (insert)
41
Reps 12 through 14 (Table 4-2) were conducted at a private residence. For these Reps, substrate collected from Penns Creek was pooled and mixed by hand, and 48 oz (1.42 L) of Penns
Creek substrate was added to the “Control” coolers, and the same amount of hand-mixed Spring
Creek substate was added to the “Experimental” coolers. Equipment, coolers and larvae were monitored daily; equipment failures did not occur during any of these three reps. As in the
Ichthyology Lab reps, any dead larva found on the surface of the substrate was immediately removed and preserved in 90% ethyl alcohol. For each cooler in which a dead larva was not found, substrate was gently disturbed with a finger—this method was usually sufficient to elicit swimming by a live larva. Missing larvae were noted, but not yet presumed dead.
Rep Larval Duration of Rep Larvae Number of Number of Sex Configuration Collection Date (Dates) sexed? Coolers Used Larvae in Rep per Cooler 12 28 April 2010 35 days yes 6 (3 Control, 3 EXP) 12 (6 Control, 6 EXP ) 1 M and 1 F (29 Apr - 2 Jun 2010) 13 14 July 2010 30 days yes 6 (3 Control, 3 EXP) 10 (5 Control, 5 EXP ) 4 coolers: 1 M and 1 F (15 Jul - 13 Aug 2010) 2 coolers: 1 F 1 14 9 December 2010 31 days yes 6 (3 Control, 3 EXP) 10 (5 Control, 5 EXP ) 4 coolers: 1 M and 1 F 1 (10 Dec 2010 - 9 Jan 2011) 2 coolers: 1 F 1 for Reps 13 and 14, an insufficient number of apparently healthy males was collected on the previous day
Table 4-2 Design of Reps 12-14 of the viability experiment.
42
Post-assay preservation and measurements
At the end of each rep of the viability and growth experiment, substrate in the rearing cages or coolers was emptied into a sorting tray, and examined for larvae. Post-assay, the head capsule width of the surviving larvae was measured at the 0.1 mm scale, then immediately preserved in 90% ethyl alcohol.
If no larva was found in either a rearing cage or a cooler, the substrate was examined for
larval remnants (eg. tusks, head capsules, exuvia). Substrate and containers were thoroughly
examined before deeming a larva as “missing –presumed dead”.
Results
The results of all 14 Reps of the viability experiment are shown in Table 4-3.
Table 4-3 Number of survivors, percent survivorship and growth of surviving larvae for Reps 1 – 14 of the viability experiment.
43
Discussion
Survivorship and growth
With larger sample sizes, the effect of either Spring Creek or Penns Creek substrate on
larval survival could be explored by comparing the survivorship of each rep—expressed as the percent of larvae per substrate treatment recovered alive at the end of the trial. Also, intra-
treatment differences in growth according to sex, and the effect of either Spring Creek or Penns
Creek substrate on larval growth could be explored by comparing the percent increase in pre- and post-experimental head capsule widths for each surviving larva in the rep. However, small sample
sizes could cause the appearance of similarities or differences between substrate treatments.
Without the benefit of large sample sizes, mean head capsule width increase could not be
calculated, within-treatment sexual differences could not be examined, and between-treatment
comparisons could not be made.
What is gained from this series of experiments is the knowledge that E. guttulata larvae
collected from Penns Creek can survive (and thrive, using growth as its proxy) in Spring Creek
water and substrate. In five of the Reps, larvae survived in Spring Creek substrate—in three of
these Reps, larval head capsule widths increased over the course of the experiment, and in one of
these Reps, survivorship in Spring Creek substrate was greater than in Penns Creek substrate.
Though not statistically sound, the results of this viability experiment suggest that E. guttulata
can survive in Spring Creek, and that reintroduction is feasible.
44 Implications
Future studies
The fact that survivorship in Penns Creek substrate was 0% in 6 of the 14 Reps casts serious doubt on the validity of the controls. Sample sizes were far too small to be considered truly representive. Future studies of the effects of substrate source on E. guttulata survivorship and growth should utilize a much larger number of larvae, to rule out the effects of small sample sizes. Future studies should follow protocols that include pre-assay physicochemical testing of the test substrate, to help reduce the influence of confounding variables.
Physiological changes associated with the growth and development of aquatic organisms can influence their sensitivity to pollutants (Anderson et al 1980; Stuijfzand et al 2000). For
mayflies, their response to toxicant exposure has been shown by Palmquist el al (2008) to differ
according to instar—an exceptional feat considering the notorious difficulty of determining
instars in Ephemeroptera. Simple frequency and allometric methods are unreliable, due to varied
growth rates and overlapping larval sizes. The most reliable means of identifying mayfly instars
is the “Palmen body method, which involves counting the number of chitin layers comprising the
Palmen body—a microscopic structure within the head capsule that is unique to Ephemeroptera
(Ruffieux et al 1997; Fink 2008). This method requires a considerable amount of time, and
specialized skills and equipment. If accurately determining mayfly instars could be made easier,
questions of voltinism and age effects could be better answered.
Instar data are essential for the development of life tables, which model the survivorship
of a wide variety of taxa, and are used to predict the most vulnerable life stage, or “limiting” age.
These data are essential in the design of viability experiments, as the life stage or age of larvae
used could impact results, and cause an over- or under-estimation of survivorship. Life table data
45 do not exist for E. guttulata , and its limiting age is unknown; however, we can make general predictions based on ecological studies of cofamilial (eg. Hexagenia spp.) and other mayfly taxa.
For example, it was estimated that only 9% of a population of baetid mayfly species would
survive from egg to final instar (Werneke & Zwick 1992). Age-specific growth and mortality data
for E. guttulata larvae are crucial for estimating population growth rates via life tables, and these estimations are themselves crucial in the design and implementation of a reintroduction project.
Reintroduction of Ephemera guttulata
For E. guttulata reared for > 25 days in Spring Creek substrate, I hypothesized that the
survivorship of E. guttulata larvae would be lower, and their growth would differ significantly
from E. guttulata reared for the same duration in their native, Penns Creek substrate. Though
survivorship was lower in Spring Creek substrate than in Penns Creek substrate for all but one of
the 14 Reps, larvae survived in Spring Creek substrate in five of the Reps. Judging by these
results, I can comfortably state that Spring Creek substrate collected from “the Rock” region of
Spring Creek was not found to be especially toxic (both lethally and sub-lethally) to the population of E. guttulata from the Coburn region of Penns Creek. The results of the viability experiment suggest that the restoration of Spring Creek’s “eastern green drake” mayfly population is achievable.
46
Chapter 5
Sampling the macroinvertebrate communities of Spring Creek and Penns Creek
Introduction
A third important step in estimating the feasibility of an aquatic insect reintroduction is to look at the macroinvertebrate communities of both the donor stream and the target stream.
According to Klemm et al (2002), Mid-Atlantic populations of Ephemera guttulata have a tolerance value (TV) of 2.3. The presence or absence of taxa of equal or higher “sensitivity” rankings (0 to 10 on the tolerance of organic pollution scale commonly used by the EPA) could hint at the chance of successful species restoration.
Characteristics of benthic macroinvertebrate communities can tell us much about their respective systems—the perceived ecological preferences and requirements of certain species are used to help define the types of habitats present, and the overall prevalence of certain species can be used to estimate the stream’s degree of impairment. For example, an apparent diversity of stoneflies can indicate nearly pristine waters, while a community dominated by Diptera species and lacking Plecoptera tends to raise red flags. Species “richness” is also a commonly used gauge of stream health: if there is significantly greater diversity in samples collected from the drift than from the benthos, this may indicate poorer substrate quality.
Besides water and substrate quality, other factors are known to affect the presence and prevalence of taxa collected in community sampling. Seasonal effects are well known, with
Slobodchikoff and Parrott (1977) stating that abundance is low in late spring to early summer, then higher in late summer to early autumn, then declines in winter. Mackay and Kalff (1969)
47 posit that overall diversity is highest in summer and winter . However, the consensus is that apparent macroinvertebrate species diversity is generally found to be the highest in the spring and the fall. In spring, most benthic invertebrates are nearing maturity and have not yet emerged as adults; in fall, the young of the year should have reached an adequate size for collection and identification.
Current speed and stream order are also factors in dictating aquatic insect communities, as oxygenation and the availability and abundance of food can vary greatly, and macroinvertebrates employ a wide range of morphological and behavioral adaptations to thrive in a myriad of conditions. To attach themselves to rocks and other rough substrate situated in fast currents, black flies (Simuliidae) rely on silken secretions and a proleg outfitted with crochets, while riffle beetles (Elmidae) and free-living caddisflies (Rhyacophilidae) possess “grappling hooks” at the tip of their abdomens. Body shapes are either streamlined (eg. Simuliidae, some
Elmidae) or flattened (eg. some Elmidae, Rhyacophilidae, Heptageniidae). In slower currents, where siltation poses a greater threat, adaptations to combat its suffocative properties include the hairy bodies of ephemerid mayflies, and the operculate gills of caenid mayflies. (Zimmerman
1961, Hynes 1970, Vannote et al 1980, Resh and Rosenberg 1984, Merritt et al 2008).
Apparent macroinvertebrate diversity has been found to vary according to the origin of the stream (“limestone” versus “freestone”). Less than 1% of Pennsylvania streams are
“limestone” (Botts 2009), yet with their often-prolific trout populations they contribute significantly to the state’s recreational revenue. Most input to limestone streams comes from limestone/dolomite springs, and these lowland streams are typically alkaline, perennial, productive, and hold at nearly-constant low temperatures year-round. The two most important elements in the definition of a “limestone” stream are alkalinity (> 140 mg/L year-round) and temperature (similar to the temperature of groundwater, and not veering outside the range of 4 ◦ to
18 ◦ C)—according to these and other criteria, Spring Creek is a “true” limestone stream. The
48 general lack of thermal and physicochemical diversity is thought to contribute to typically low macroinvertebrate diversity. The absence of wide temperature fluctuations limits macroinvertebrate diversity, as some taxa require higher or more varied water temperatures. Low diversity is compensated for by high density, and is attributed to limestone streams’ rich productivity. The taxa that most often dominate macroinvertebrate communities of limestone streams are the aquatic sowbug, Lirceus sp. , the amphipod, Gammarus sp. , the spiny crawler
mayfly, Ephemerella sp. , the riffle beetle, Optioservus sp. , and the non-biting midges of the
family Chironomidae in the order Diptera (Botts 2009).
Conversely, “freestone” streams tend to originate from runoff and smaller “feeder”
streams flowing down from sandstone/shale ridgetops. The hilly topography associated with
freestone streams generally lead to more intact riparian zones (Lindsey et al 1998). Penns Creek
is a stream with more freestone than limestone qualities. Freestone streams, and limestone
streams that receive inputs from freestone tributaries, tend to have more diverse habitats with
different temperature regimes, water chemistries, and productivity. As a result, these streams tend
to have much greater diversity of macroinvertebrates than limestone streams. Because of these
differences, an otherwise healthy limestone stream with low apparent macroinvertebrate diversity
may appear to be “impaired” when compared to a slightly-impaired freestone stream with higher
apparent diversity. To avoid this pitfall, in 2009 the Pennsylvania DEP developed an IBI
specifically for limestone streams, which included an adjusted threshold of impairment (Botts
2009).
In biotic investigations of “stream health”, chemical water quality and physical habitat
evaluations are often variables that are combined with macroinvertebrate community data to
generate impairment estimates of each sampling site. While informative, water chemistry data
and physical habitat assessments have their limitations. The relevance of single or synergistic
chemical parameters to the needs of most aquatic organisms is not known, and because of their
49 ever-changing flow regimes, “static” chemical profiles do not logically exist for lotic systems. To produce a sufficiently accurate chemical profile of each sampling site requires continuous or repeated chemical monitoring. Physical habitat evaluations are important tools for describing sampling sites and for determining multi-habitat sampling strategies, but tend to be untranslatable in terms of “ecosystem integrity” in the face of countless potential stressors and the temporal variations of macroinvertebrate community structure.
Because Spring Creek water and sediment did not prove to be toxic to E. guttulata larvae
in the viability experiment, chemical water quality and physical habitat assessments of each
sampling site in this macroinvertebrate community study were measured and recorded, but their potential effects on population density or apparent diversity were not explored further.
Objective
To estimate the feasibility of E. guttulata reintroduction to Spring Creek, I sampled the
macroinvertebrate communities of specific sections of Spring Creek and Penns Creek, to
determine the presence or absence of taxa of equal or greater “pollution intolerance”. For
Pennsylvania’s E. guttulata populations, the EPA has assigned a tolerance value of 2.3. The presence of taxa with tolerance values > 2.3 in Spring Creek macroinvertebrate community samples would suggest that the stream’s estimated degree of impairment would not sufficiently hinder the restoration of the “eastern green drake” mayfly.
50 Methods
Selection of sampling stations
Macroinvertebrate community sampling stations were selected from three locations on
Spring Creek (Figure 5-1), and three locations on Penns Creek (Figure 5-2).
Spring Creek
The three Spring Creek sampling stations are located within the historic range of the E. guttulata . From downstream to upstream:
“Spring 1 - 200 meters from Rock – area of power transformer” ( 40 ◦51'14.01" N,
77 ◦49'19.55" W. PA F&BC site ID# 9099). This site is approximately 400 meters downstream of the gated metal bridge at “the Rock”, which is accessible from Rock Road in Benner Township.
“Spring 2 - approximately 200 meters from Rock – area of fenced spring”
(40 ◦51'08.84" N, 77 ◦49'16.17" W. PA F&BC site ID# 9100) This site is approximately 207
meters downstream of the gated metal bridge at “the Rock”, in Benner Township, and approximately 193 meters upstream of “Spring 1”.
“Spring 3 - immediately downstream from falls at Rock” (40 ◦51'04.60” N,
77 ◦49'18.24" W. PA F&BC site ID# 9101) This site is approximately 58.3 meters downstream of
the gated metal bridge at “the Rock”, in Benner Township, and approximately 154 meters
upstream of “Spring 2”. This site is approximately 75.7 meters downstream of “SC2 ” – where substrate and water were collected for chemical analyses.
51
Figure 5-1 Location of Spring Creek community sampling sites.
Penns Creek
E. guttulata populations are abundant at each of the three Penns Creek sites. From downstream to upstream:
“Penns 1 - Tunnel Road at wooden footbridge” (40 ◦50’56.25” N, 77 ◦27’3.99” W. PA
F&BC site ID# 9096) This site is located at the end of Tunnel Road in Penn Township (Centre
County), and is approximately 34.3 meters upstream of a wooden foot bridge that crosses the stream. This community sampling station was approximately 20.7 meters downstream of substrate and water sampling station “ PC1 ”.
52
“Penns 2 - Tunnel Road at cement bridge” (40 ◦50’59.62” N, 77 ◦27’18.49” W. PA
F&BC site ID# 9097) This site is located approximately 1.54 kilometers upstream of “Penns 1”, and is approximately 29 meters upstream of a small cement bridge located on Tunnel Road in
Penn Township (Centre County), southeast of Coburn proper. “Penns 2” was approximately 49.5
meters downstream of substrate and water sampling station “ PC2 ”.
“Penns 3 - below Pine Creek confluence at Coburn” (40 ◦51’36.89” N, 77 ◦27’41.68”
W. PA F&BC site ID# 9098) This site is located approximately 51.4 meters downstream of the
Pine Creek/Penns Creek confluence in the town of Coburn (Penn Township, Centre County), and is approximately 1.40 kilometers upstream of “Penns 2”. “Penns 3” was approximately 99.5 meters downstream of the location of substrate and water sampling station “PC3”.
53
Figure 5-2 Location of Penns Creek community sampling sites.
Sampling design and equipment
As the abundance and diversity of stream communities follows a seasonal periodicity, sampling occurred within two seasons—fall (2009) and spring (2010). To increase the likelihood of collecting a greater number of taxa (and therefore the likelihood of finding pollution-intolerant taxa), the following modifications were applied to the EPA’s Rapid Bioassessment Protocol
(RBP, Plafkin et al. 1989):
54 Multi-habitat sampling
The RBP was originally designed to target a single habitat (riffle) only, as riffles have been found to be the regions of the stream with the greatest diversity of organisms. Because single-habitat sampling excludes many taxa and therefore cannot produce a complete picture of community diversity, more recent revisions of the RBP outline procedures for multi-habitat sampling—not just of riffles, but also pools, snags, submerged vegetation, and vegetated banks.
Semi-quantitative sampling procedures
Quantitative sampling methods via fixed surface area samplers (eg. Surber, modified
Hess) are the ideal for collecting population density data; however, Penns Creek’s depth and
heterogenous streambed are problematic for employing quantitative-type samplers. The use of
Surber samplers is restricted to depths less than 0.3 m, and modified Hess samplers to depths less
than 0.5 m (Barbour et al. 1999). To be used properly, the frames of Surber and modified Hess
samplers must penetrate the streambed uniformly—which is not possible when substrate
composition is coarse, rocky and irregular.
To combat this problem, “semi-quantitative” sampling methods have been developed, in
which qualitative-type samplers are used, but under stringent guidelines. These guidelines often
dictate a standard number of kicks into a net, within a standard sized area, and specific amounts
of time allotted to each action (eg. sampling, sorting). Tests of these methods performed by the
Mid-Atlantic Coastal Streams (MACS) Workgroup and by the Florida DEP found them to be
“scientifically valid for low-gradient streams” (Barbour et al. 1999). Therefore, my sampling protocol employed the following semi-quantitative strategies using qualitative-type sampling
equipment:
55
Standard number of sampling replications per site = 10
Standard number of kicks into D-frame kick net sampler = 20
Standard amount of time picking macroinvertebrates from samplers = 6 minutes.
For riffle/run sampling, organisms dislodged by kicks may evade capture due to the relatively small size of the D-frame kick net opening, or due to hydraulic properties that can cause backwash or excessive sediment disturbance (Merritt et al 2008). To intercept these organisms, a 0.91 m 2 hand net was positioned directly downstream of the D-frame kick net.
No sub-sampling
The RBP sets a limit on the number of organisms to include in analysis, via a fixed count or fixed ratio (Barbour et al. 1999). These limits are intended to increase efficiency, as sorting and identification of specimens are the most labor-intensive and costly components of conducting bioassessments (Plafkin et al 1989); however, this can often result in sorter bias against smaller and rarer organisms. For this study, all benthic macroinvertebrates collected were sorted and identified.
Collection and field preservation methods
Habitat types represented at each site were sampled proportionally. For example, if one site’s composition was approximately 50% riffle, 40% run, 10% pool and 10% snag, then my assistant and I collected four reps from riffles, three reps from runs, one rep from a pool, and one rep from a snag.
56 Riffles and runs
For riffle and run collections, one operator gave 20 vigorous kicks in an approximate area of 1 m2 (1 x 1 m) immediately upstream of a D-frame kick frame net (500 micron mesh), while a second operator held a 0.91 m 2 hand net (500 micron mesh) directly downstream of the first
operator, to intercept macroinvertebrates dislodged by kicks but missed by the D-frame kick net.
The D-frame kick net operator emptied the contents of his net into an airtight container half-filled
with 90% ethanol. The D-frame kick net and the hand net operators then spent exactly 6 minutes
transferring macroinvertebrates from their respective nets to similar airtight containers; after the
allotted time, any remaining organisms were returned to the stream.
Pools, snags, etc.
For non-riffle collections, the D-frame kick net operator performed a set of 15 sweeps of each habitat. After each set of sweeps, the D-frame kick net operator emptied the contents of his net into one of the airtight containers. While the D-frame kick net operator performed the non- riffle collections, the hand net operator evaluated the physical characteristics of the site, measured air and water temperatures, and made note of past/present weather conditions and any other known events relevant to the study. These data were recorded on a Habitat Assessment form
(Barbour et al 1999), for each site. A total of 10 kick and sweep samples were collected at each site, and composited into one or two of the airtight containers. For every site at both streams, all macroinvertebrate collections, physical measurements, and habitat assessments were performed similarly, and in the same sequence.
Samples were preserved with 90% ethyl alcohol (denatured), transported to the lab, and stored in a cool, dark room until analysis.
57 Identification of specimens
Each sample container was emptied into a sorting tray, and non-macroinvertebrate objects were removed. The remaining organisms were sequestered by order (or class, for non- insect taxa) into different regions of the same tray. Using Merritt et al (2008) and Peckarsky et al
(1990), macroinvertebrates were identified to:
Genus: non-chironomid insects, Amphipoda, Isopoda, and Bivalvia
Subfamily: Collembola and Chironomidae
Family: Planariidae
Suborder: Hydracarina
Subclass: Oligochaeta and Hirudinea
Class: Gastropoda
Results
A tabular list of the taxa, their EPA-assigned Tolerance Values (TV) for general organic pollution in Pennsylvania, and counts of individuals for each sampling station and each season is located in Appendix B : “Macroinvertebrate Taxa of Spring Creek and Penns Creek” .
58 Spring Creek
“Spring 1”
In both the fall and spring samples, two taxa with a tolerance value lower than 2.3 were collected at this station: The riffle beetle, Promoresia sp. (TV = 2, Elmidae: Coleoptera), and the spiny crawler mayfly, Ephemerella sp. (TV = 1, Ephemerellidae: Ephemeroptera).
“Spring 2”
The fall collection from this site yielded three taxa with a tolerance value lower than 2.3:
The black fly, Prosimulium sp. (TV = 2, Simuliidae: Diptera), as well as Promoresia sp. and
Ephemerella sp. The spring collection yielded two: The caddisfly, Rhyacophila sp. (TV = 1,
Rhyacophilidae: Trichoptera), and Ephemerella sp.
“Spring 3”
At this station, two taxa from the fall sample have a tolerance value lower than 2.3:
Promoresia sp. and Ephemerella sp. The spring sample also yielded two such taxa: Ephemerella sp. and Rhyacophila sp .
Summary – Spring Creek
When all three stations and both seasons are combined, there are four genera present
with a tolerance value less than that of E. guttulata : Promoresia , Prosimulium , Ephemerella, and
Rhyacophila .
59 Penns Creek
“Penns 1”
In the fall sample, 13 taxa with a tolerance value of less than 2.3 were collected: The watersnipe fly, Atherix sp. (Tolerance value = 2, Athericidae: Diptera), the mayflies, Ephemerella sp. (TV = 1, Ephemerellidae: Ephemeroptera), Teloganopsis sp. (TV = 2, Ephemerellidae:
Ephemeroptera) and Ephemera sp. (TV = 2, Ephemeridae: Ephemeroptera), the fishfly, Nigronia sp. (TV = 2, Corydalidae: Megaloptera), the stoneflies, Acroneuria sp. (TV = 0, Perlidae:
Plecoptera), Agnetina sp. (TV = 2, Perlidae: Plecoptera), Isoperla sp. (TV = 2, Perlodidae:
Plecoptera), and Pteronarcys sp. (TV = 0, Pteronarcyidae: Plecoptera), and the caddisflies,
Brachycentrus sp. (TV = 1, Brachycentridae: Trichoptera), Hydatophylax sp. (TV = 2,
Limnephilidae: Trichoptera), Psychomyia sp. (TV = 2, Psychomyiidae: Trichoptera), and
Rhyacophila sp. (TV = 1, Rhyacophilidae: Trichoptera). In the spring sample, there were also 13
genera: The spotted darner dragonfly, Boyeria sp. (TV = 2, Aeshnidae: Odonata), as well as
Atherix sp. , Ephemerella sp. , Teloganopsis sp., Ephemera sp., Nigronia sp., Acroneuria sp.,
Agnetina sp., Isoperla sp., Pteronarcys sp., Brachycentrus sp., Hydatophylax sp., and
Rhyacophila sp.
“Penns 2”
At this station, collections in the fall yielded 16 different taxa with a tolerance value less
than 2.3: the mayfly, Attenella sp. (TV = 2, Ephemerellidae: Ephemeroptera), the stonefly,
Paragnetina sp. (TV = 1, Perlidae: Plecoptera), the caddisfly, Glossosoma sp. (TV = 0,
Glossosomatidae: Trichoptera), Atherix sp. , Ephemerella sp. , Teloganopsis sp. , Ephemera sp. ,
Nigronia sp. , Boyeria sp. , Acroneuria sp. , Agnetina sp. , Isoperla sp. , Pteronarcys sp. ,
60 Brachycentrus sp. , Hydatophylax sp. and Rhyacophila sp. Collections in the spring yielded 12 different genera: Atherix sp. , Ephemerella sp. , Teloganopsis sp. , Ephemera sp., Nigronia sp. ,
Acroneuria sp. , Agnetina sp. , Paragnetina sp. , Isoperla sp. , Pteronarcys sp. , Brachycentrus sp.
and Rhyacophila sp.
“Penns 3”
In the fall sample, 10 different taxa with a tolerance value less than 2.3 were collected:
The mayfly, Paraleptophlebia sp. (TV = 1, Leptophlebiidae: Ephemeroptera), Attenella sp. ,
Ephemerella sp. , Teloganopsis sp. , Ephemera sp. , Acroneuria sp. , Agnetina sp. , Isoperla sp. ,
Brachycentrus sp. and Rhyacophila sp. . In the spring, 12 different genera were collected:
Attenella sp. , Ephemerella sp. , Teloganopsis sp. , Ephemera sp. , Paraleptophlebia sp. , Nigronia sp. , Acroneuria sp. , Agnetina sp. , Isoperla sp. , Pteronarcys sp. , Brachycentrus sp. and
Rhyacophila sp.
Summary – Penns Creek
When samples collected from all three stations and both seasons are combined, there are
18 different genera present with a tolerance value less than that of E. guttulata : Atherix , Attenella ,
Ephemerella , Teloganopsis , Ephemera , Paraleptophlebia , Nigronia , Boyeria , Acroneuria ,
Agnetina , Paragnetina , Isoperla , Pteronarcys , Brachycentrus , Glossosoma , Hydatophylax ,
Psychomyia and Rhyacophila.
61 Discussion
Implications for reintroduction
If there were no taxa present in the Spring Creek community samples with a Tolerance
Value equal to or less than 2.3, it could bode poorly for the reintroduction of E. guttulata to
Spring Creek. However, this was not the case. Four “less tolerant” taxa were collected from the
Spring Creek sampling stations — Promoresia sp. (Elmidae, TV = 2), Prosimulium sp.
(Simuliidae, TV = 2), Ephemerella sp. (Ephemerellidae, TV = 1), and Rhyacophila sp.
(Rhyacophilidae, TV = 1). Though these taxa are deemed as more sensitive to organic pollution
than E. guttulata , there are differences that may be significant. None of the four taxa are
“burrowers”—three are “clingers” and one is a “clinger/sprawler”—therefore substrate quality is
not likely to be relevant to their survival. The genera Prosimulium , Ephemerella and Rhyacophila
are univoltine. Promoresia is classified as semivoltine, but this may be due to their “protracted
oviposition period”—in which adults live underwater for one year. Compared to E. guttulata ,
Promoresia sp. eggs hatch quickly (6 to 10 days versus 23 to 30 days). In general, these four taxa
have more rapid generational turnovers and development times, which are reproductive strategies
for recolonizing streams after pollution events or other catastrophic occurrences. Despite this, the presence of such “sensitive” taxa in Spring Creek is suggestive of the stream’s recovery.
Substrate used in the viability experiment was collected approximately 75 meters
upstream of “Spring 3” . Though it is impossible to replicate the entire Spring Creek ecosystem in
a laboratory, the fact that Spring Creek substrate was not apparently toxic to E. guttulata larvae,
and that taxa with lower Tolerance Values are present in Spring Creek, suggests factors other than persistent pollution are responsible for their prolonged absence.
62 If the decades-long extirpation of the “eastern green drake” on Spring Creek is not due to persistent and/or significant levels of pollutants, the influence of biotic factors (eg. predation, competition) should be explored. Spring Creek is well known for its wild Brown Trout populations, which would be a likely source of predation. Of competition, Kolar et al (1997) noted that dense populations of chironomid larvae and mayflies in the genus Hexagenia tend not to coexist—the negative correlation of their densities within the same area suggests that one outcompetes the other for space and resources.
Abiotic factors are also possible. The lack of hydrological connectivity between Spring
Creek and a stream with an E. guttulata population could be hindering the mayfly’s recolonization. The Little Juniata River in central Pennsylvania has experienced a number of fish and macroinvertebrate kills in the last few decades; however, eventual recovery of those organisms are attributed to donor populations inhabiting the river’s tributaries. Spring Creek has no E. guttulata populations present in its tributaries, and the weak flying ability and short adult life span of this mayfly means Spring Creek is likely to be outside the potential dispersal range of populations from even the nearest stream (Brittain 1982; Alexander et al 2011).
Should predation or competition be strong factors in the persistent absence of E. guttulata from Spring Creek, then repopulation via reintroduction is less likely to be successful
than if geographic isolation is the stronger factor. Further study of the influence of these and
other potential biotic factors is needed, especially if restoration attempts fail. However, by finding
several taxa in Spring Creek that are classified as more pollution-intolerant than E. guttulata ,
there is no reason to assume that reintroduction of the “eastern green drake” mayfly to Spring
Creek would be ultimately unsuccessful.
63
Chapter 6
Strategy for reintroduction of Ephemera guttulata to Spring Creek
Introduction
As we become more aware of the looming threat and predicted effects of global biodiversity loss, a number of pre-emptive and retaliative strategies to combat species losses have been developed, explored, and employed. One such strategy that is becoming increasingly more
common is reintroduction.
Reintroductions
“Re-introduction” is defined as “an attempt to establish a species in an area which was once part of its historical range, but from which it has been extirpated or become extinct” (IUCN
1998). This does not include short-term translocations for sporting or commercial gain. To be successful, such attempts require sufficiently substantial planning, collaboration, funding and time (in fact, Kleiman et al 1994 suggested that failures were more strongly correlated with politics and financing than with scientific rigor); however, even when properly implemented, the majority of reintroduction attempts are unsuccessful.
A number of species re-introduction programs have been attempted across a range of animal and plant Classes. Some are deemed “successful”; most, however, are not. Among the
“successes” are:
- Grey wolf, Canis lupis (Idaho/Montana/Nevada. Carroll et al 2003)
64 - European beaver, Castor fiber (United Kingdom. South et al 2001) - Yellow-shouldered Amazon parrot, Amazona barbadensis (Venezuela. Sanz and Grajal 1998) - Eastern brown pelican, Pelecanus occidentalis carolinensis (Louisiana. Holm Jr. et al 2003) - Gharial, Gavialis gangeticus (India. Choudhury and Choudhury 1986) - Tuatara, Sphenodon punctatus (New Zealand. Soorae 2008) - Lake sturgeon, Acipenser fulvescens (Tennessee. Soorae 2008) - Yarqon bleak, Acanthobrama telavivensis (Israel. Soorae 2010) - Southern Damselfly, Coenagrion mercuriale (United Kingdom. Soorae 2010) - Banks Peninsula tree weta, Hemideina ricta (New Zealand. Soorae 2010) - Whibley Wattle shrub, Acacia whibleyana (Australia. Soorae 2010) - Sargent’s cherry palm, Pseudophoenix sargentii (Florida. Maschinski and Dusquesnel 2007)
Despite the diversity of taxa in the above examples, invertebrates are more disproportionately underrepresented than either plants or vertebrates. Philip Seddon and his colleagues calculated that of a list of 699 species targeted for reintroduction, only 9% of these projects focused on invertebrates; of this same list, mammalian projects comprised 61%.
Interestingly, this is the inverse of their estimated natural species richness (77% and 8%,
respectively) (Seddon et al 2005).
Polhemus (1993) projected aquatic insect species to be at less risk of extinction than
terrestrial insect species, reasoning that such species have greater tolerance for changes to the
terrestrial environs, and have the ability to disperse as adults. Conversely, Ricciardi and
Rasmussen (1999) estimated the peril facing aquatic fauna to be four to five times that of
terrestrials. Master et al (2000) and DeWalt et al (2005) each concluded that extinction risk was
far greater for aquatic insects—the latter authors basing their inference on Illinois’ extensive
records charting the decline in its Plecoptera species.
65 Unfortunately, this position is visible in neither global nor national policy, and conservation efforts for insects tend to favor terrestrial taxa disproportionately. Out of the 3338 insect species on the IUCN Red List, 2477 of these are aquatic insects—but 2446 of these are odonates, the “butterflies of the aquatic world” (DeWalt et al 2005). In 1991, the Federal Register of the Endangered Species Act (ESA) listed an estimated total fauna of 10,000; of these, 204 were aquatic insects (Polhemus 1993). None were considered to be endangered, and only one species out of these 204 was listed as “Threatened”, making it the only aquatic insect to be protected under the ESA at the time ( Ambrysus amergosus , the Ash Meadows naucorid). Currently, there are seven aquatic (out of 64 total) insect species on the Federal Register: The Ash Meadows
Naucorid, the Comal Springs dryopid beetle ( Stygoparnus comalensis , added in 1997), the Comal
Springs riffle beetle ( Heterelmis comalensis , also 1997), the “flying earwig Hawaiian damselfly”
(Megalagrion nesiotes , 2010), the Hine’s Emerald dragonfly ( Somatochlora hineana , 1995),
Hungerford’s crawling water beetle ( Brychius hungerfordi , 1994), and the Pacific Hawaiian damselfly ( Megalagrion pacificum , 2010) (website: USFWS-ESP). In 2006, an action plan was approved for the Hungerford beetle (USFWS 2006). Actions defined in this plan include: conservation of known sites, research to facilitate recovery efforts, additional surveys and monitoring of existing sites, development and implementation of public education and outreach, revision of criteria and actions as needed, and the development of a monitoring plan for after the
Hungerford beetle’s “delisting”. All of these actions are essential components of conservation programs—however, it is important to note that in this plan the term “restoration” is used in terms of habitat, and not of the actual populations.
The scientific development and implementation of aquatic insect restoration programs are scarce; more commonly, it is avid sportfishers and non-profit organizations who attempt these reintroductions. In 1945 and 1946, Charlie Fox and a group of fishermen attempted to transplant
E.guttulata larvae (1945) and eggs (1946) into LeTort Spring Run, Cumberland County,
66 Pennsylvania (Fox 1947). These attempts proved unsuccessful. A similar attempt to reintroduce
E. guttulata larvae to Spring Creek took place more than a decade ago. Dan Shields was part of a
group of local anglers who transplanted larvae and subimagos, and “thousands” of imagos from
another stream. This project lasted no longer than four years before it was deemed as
unsuccessful (Pittsburgh Post-Gazette, 6 Aug 2006).
More recently, the University of Utah’s National Aquatic Monitoring Center (or,
“BugLab”) has transplanted Blacksmith Fork River Pteronarcys californica and Pteronarcella badia (Plecoptera: Pteronarcyidae) larvae and adult females into the Logan River (website: Utah
State Today). As preliminary transplant experiments conducted in June 2001 had been successful, reintroductions began in 2004, and BugLab anticipates the eventual restoration of these
“salmonflies” to the waterway.
In 2009, the DeWalt lab of the University of Illinois-Urbana began experimental transplants of different populations of the stonefly Acroneuria frisoni (Plecoptera: Perlidae) into several “suitable” Illinois streams, in order to identify the best donor population(s), and the most effective methods of reintroduction (website: USDA)
Most promising to ephemerid mayfly reintroductions is an ongoing project at the River
Wey, England, headed by Cyril Bennett of The John Spedan Lewis Trust for the Advance of
Natural Sciences. This and several other organizations began a collaborative effort in 2003 to reintroduce eggs and larvae of the burrowing mayfly Ephemera danica to the South Wey
(Bennett 2007). The initial viability of the transplanted larvae had those involved optimistic, and
recently, Bennett and Gilchrist (2010) wrote: “…the mayfly, Ephemera danica , was successfully reintroduced to the River Wey in Surrey and the River Wye in Derbyshire.”
67 International Union for the Conservation of Nature and Natural Resources (IUCN)
These reintroductions would not likely be possible were it not for the International Union for the Conservation of Nature and Natural Resources (IUCN)—an organization created in 1948 by the United Nations Educational, Scientific and Cultural Organization (UNESCO), for the purposes of promoting solidarity between nations (and thus preventing another World War):
“since wars begin in the minds of men, it is in the minds of men that the defences of peace must be constructed”
- Preamble to the Constitution of UNESCO
In 1951, the IUCN published “The State of the Protection of Nature in the World in
1950”—an ambitious compilation of 70 reports from 70 different countries. In 1988, the
increasing number of species reintroduction projects, and the concern with the lack of quality
control, led the IUCN to form a subgroup: the Reintroduction Specialist Group (RSG). The RSG
developed and published reintroduction “Guidelines” in 1998, and has since been the leading
authority, and watchdog, of scientifically-based reintroduction projects (IUCN 1998; Seddon et al
2007). The IUCN-RSG “Guidelines for Re-Introduction” provided the template for this project,
unofficially named the “Green Drake Restoration Project”.
68 Strategy and Process
Preliminary stage of reintroduction
Results of preliminary studies
“Chemistry” – Via the analyses of substrate and water samples, it was determined that the restoration of Spring Creek’s “eastern green drake” mayfly population to its historic range would
not likely be hindered by OC pesticides or other pollutants.
“Toxicity” – Via the bioassessments, substrate collected from The Rock site of Spring
Creek—an area within the historic range of the “eastern green drake” –was not found to be
especially toxic (both lethally and sub-lethally) to E. guttulata larvae collected from the Coburn
region of Penns Creek.
“Community” – Via macroinvertebrate community sampling, the presence of “pollution- intolerant” taxa within the historic range of the “eastern green drake” demonstrated that macroinvertebrates with similar Tolerance Values to E. guttulata can survive and persist there, and does not rule out the possibility of a new population being established.
Permissions
Governing agencies - The first and most important step in a reintroduction is to learn of relevant regulations and jurisdictions—what bureaucratic “hoops” must first be jumped before developing a protocol. On July 19, 2006, while in the preliminary stages of research, I began this step by contacting the Pennsylvania Department of Conservation and Natural Resources (PA
69 DCNR), describing my research interest, explaining my intent, and inquiring of said “hoops”. In their response, PA DCNR explained that they oversee state parks and state forests, and it is the
Pennsylvania Fish and Boat Commission (PA F&BC) that has jurisdiction over waterways.
In Pennsylvania, aquatic insects are classified as “fishbait”, and a fishing license is required to collect up to 50 individuals; to collect more than 50 individuals for educational or scientific purposes, the collector must both purchase a fishing license and apply for a Scientific
Collector Permit (PFBC-107). Information required in this application includes the applicant’s goals and objectives, the exact location of study sites, specific equipment to be used, and taxa to be collected. Any assistants must also possess a fishing license, and their information must be included in the application. It is only with the approval of the PA F&BC that the project may move forward.
After purchasing a fishing license and applying for a PFBC-107, I contacted the PA
F&BC, providing the same information I had sent to the PA DCNR. On July 24, 2006, the
Natural Diversity Chief of the PA F&BC replied, stating that I appeared to be “taking a thorough approach” and following the correct steps. He called the project proposal “interesting”, and said that he looked forward to learning of the results. This was essentially a “green light” to proceed with my research—the preliminary work necessary to design and implement a reintroduction protocol.
On May 28, 2009, I contacted the Natural Diversity Chief of the PA F&BC, to update him on the different components of the research project, and to inquire about how to gain the permission of the PA F&BC to begin the implementation phase of reintroduction. My specific justifications for beginning this next phase were the ease and potential efficacy of egg collection, and the preliminary survivorship data collected from pilot vEXP reps. His response was unexpected:
70 “I reviewed your current Scientific Collectors' Permit. Your [sic] believe your permit covers your activities, but thanks for asking and keeping me in the loop. I am looking forward to learning how E. guttulata "take" in Spring Creek.” (28 May 2009, personal communication).
Community - Reintroductions are long-term projects that require (significant) financial
and political support, government involvement, permissions and/or permits, and community
approval. This cooperation will only be possible if the impact of restoring E. guttulata populations is predicted to be either irrelevant or beneficial to the residents of the Spring Creek
watershed. Ideally, this should be accomplished in the initial stages of the project; however, it is
equally important to maintain communication between all involved parties throughout the
duration of the project.
Choice of life stage to reintroduce
When a population is not sufficiently large (both in number of organisms and in genetic
diversity) to overcome the effects of predation, competition, disease, catastrophic events, etc.
Allee et al 1949). Hoover (1978) estimated that one female E. guttulata produces 3000 to 4000
eggs. For the congeneric E. danica , Bennett (1996) counted 3000 to 6000 eggs per female,
depending on body length. Survivorship of eggs and early instars has been found to be poor—
Elliot et al (1988) found that 95% of mortality occurred during these stages—and only 0.5% of a
female’s eggs will produce adults (Bennett 2007). Therefore, a conservative estimate is 15 adults
for every female. At the peak of their mating flights on Penns Creek, a team of five can easily
collect 200 females in one evening. Transplanting 600,000 E. guttulata eggs improves the
chances of overcoming the genetic and reproductive pitfalls of small populations.
71 Site selection for egg transplants
The area of “the Rock” in Benner Township (40 ◦51’03.41”N, 77 ◦49’19.23”W) has been chosen to be the site of egg transplants for several reasons. It is located within the historic range of E. guttulata (Aurelio 1953). It was the site of substrate collection for chemical analysis (see:
Chapter 3) and for the bioassessment (see: Chapter 4); in the former, it was not found to contain significant levels of OC pesticides, and in the latter, it was found to be not significantly more toxic than Penns Creek substrate. Finally, this was a community sampling site (see: Chapter 5), which yielded four genera of macroinvertebrates with equal or lower Tolerance Values than E. guttulata .
Implementation
Egg collection and transplantation
In order for eggs to be collected, we must first estimate the time to subadult emergence.
This is done by Gregory Hoover, who monitors E. guttulata larvae at the collection site. It is possible to predict the timing of subimaginal emergence by observing gut content and the color of
larval wingpads—darkened wing pads and no obvious gut content are indicative of preparation
for its impending subimaginal molt.
Mean temperatures and precipitation are also used to predict emergence dates. Penns
Creek populations of E. guttulata tend to emerge in late May when central Pennsylvania has
experienced above-average temperatures and below-average rain- and snowfall, and tend to
emerge in early-to-mid June with below-average temperatures and above-average precipitation.
72 Coordination of volunteers
Several weeks in advance, volunteer collectors are recruited from Penn State’s
Department of Entomology, as well as from other departments and local NGOs. Outings are usually decided with little notice, so the available volunteer pool must be sufficiently large. In addition to my “Assistant” and myself, three to ten volunteers are needed for each collection date.
Because of the distance between State College and the site of egg collection, travel arrangements need to be made for each volunteer.
Notifications
Twenty-four hours prior to expected collection dates, the PA F&BC are notified of our intended location and planned activities. Failure to abide by this and other PAF&BC regulations could effectively nullify the Scientific Collection Permit that allows this reintroduction project to continue.
Preparation of equipment and materials
The method of egg collection requires the use of stream water (to elicit an ovipositional response). To prevent potential inter-stream contamination, water from the Spring Creek transplant site is collected prior to arriving at the Penns Creek collection site. Spring Creek water and Penns Creek water are never mixed. Wide mouth glass jars to be used in the collection of eggs are steam-disinfected between outings. Each volunteer is encouraged to bring his/her own aerial collecting net, and spare aerial nets are brought to every outing.
73 Collection of females and their eggs
Because mating flights take place at treetop height, prime locations for collecting gravid females are stable (for swinging nets) and elevated above the stream (for intercepting females in their descent to oviposit). Two such locations on Penns Creek have been utilized: a metal suspension bridge near the intersection of SR3002 and Cherry Run Road (Union County), and a wooden footbridge at the end of Tunnel Road at Coburn (Centre County). The latter is the preferred location due to its closer proximity to State College, relative stability, and sufficiently dense E. guttulata populations.
Eggs are collected by capturing females on their streamward descent, and placing them in wide mouth glass jars containing Spring Creek water. Contact with the water elicits oviposition.
When all available jars are full, they are transported directly to the reintroduction site for dispersal.
Egg transplants
At the transplant site, eggs are deposited into the stream by wading to the edge of the depostional zone, then emptying the contents of each jar into the flow. Stream water is used to dislodge any eggs clinging to the bottom of the jar.
Post-implementation stage of reintroduction
Monitoring for E. guttulata larvae is conducted either concurrently with community collections (see Chapter 5 for methodology), or in targeted outings. For targeted outings, the sampling equipment and techniques are those used to collect E. guttulata larvae from Penns
74 Creek for the bioassessments (Chapter 4). Targeted sampling is performed from the transplant site to approximately 1.4 km downstream (the Benner Springs hatchery).
Mark Vinson, formerly of the Utah State University’s Buglab, and the pioneer of their
“salmonfly” reintroduction project, commented on the frustration of post-transplant monitoring:
“It is amazing though that it seems like you are moving lots of insects when you move several thousand, but then you go back a day or two later and they are tough to find. The river never stops trying to move them downstream.” (7 Aug 2007, personal communication).
Since 2010, there have been rumors of sightings of E. guttulata adults at Spring Creek, but no specimens have been collected or photographs taken. Without evidence to confirm these sightings, it was possible that E. guttulata adults observed at Spring Creek could have been unintentionally transported from another stream (eg. an adult is attracted to the interior lights of a vehicle, and becomes trapped inside). Therefore, searching for larvae in the area of the egg transplant site is a more favorable and reliable method of monitoring.
Monitoring concurrent with community collections
-November 5, 2009 – For community collections in riffles and runs, my assistant kicked into a D-frame kick net, while I held a hand net downstream from him, to intercept organisms not captured by the D-frame kick net. For community collections in pools and snags, my assistant sweept the areas with a D-frame kick net. We employed these methods at “the Rock” and at two sites 154 and 347 meters downstream. No larvae were recovered at any of the three sites.
-May 19, 2010 – For spring community collections, my assistant and I employed the same sampling equipment and strategies used for the fall collections. Again, no larvae were recovered at any of the three sites.
75 Targeted monitoring
-June 7, 2010 – Starting from “the Rock” and working downstream for 1.4 km, my assistant and I employed the shovel/hand net collection method (used to collect E. guttulata larvae from Penns Creek for the bioassays) to sample three times at every 50 paces. No larvae were recovered.
-August 4, 2011 – Starting from the Benner Springs hatchery and working upstream to
“the Rock”, my assistant and I used the shovel/hand net collection method to sample one to three times at every 50 paces. Macroinvertebrate abundance and diversity appeared to be very low in areas of dense algal growth; therefore we sampled one time in such areas, and three times in other areas. One female larva was recovered at the egg transplant site (“the Rock”). The larva was active, responded “normally” to tactile stimuli, and exhibited typical photophobic behaviors. The larva’s head capsule was 2.2 mm wide, and its body was sufficiently long to estimate that its subimaginal emergence would have been the following spring. A video of the larva is available online, at http://www.youtube.com/user/GDRP09 .
Discussion
When collecting larvae for bioassessment purposes, the number collected was greater
than needed, to counter any collection-related injuries or deaths, thus ensuring a sufficient
number of healthy larvae were available for each rep of the experiment. All larvae not used in the bioassays were sexed, measured (head capsule width at a 0.1 mm scale), and preserved
individually in 90% ethyl alcohol. These data were intended for use in a “cohort study” that was
abandoned due to time constraints, but three collection dates (two in July, one in August) became
76 useful for making comparisons with the Spring Creek larva recovered from the egg transplant site.
Female larvae collected from Penns Creek on 15 July 2009 had a mean head capsule width of 2.164 mm (s.d. = 0.338 mm, N = 11), and widths ranged from 1.4 mm to 2.6 mm. The head capsule width of the Spring Creek female collected on 4 August 2011 would fall within this range; a one-sample t-test confirmed that it was not significantly different from the mean head capsule width of the 15 July 2009 Penns Creek females (T = -0.36, p = 0.729, 95% CI).
Female larvae collected from Penns Creek on 16 August 2009 had a mean head capsule width of 2.3571 mm (s.d. = 0.3613 mm, N = 21), and widths ranged from 1.4 mm to 2.8 mm. As with the previous month’s collections, the head capsule width of the Spring Creek female collected on 4 August 2011 would fall within this range, and a one-sample t-test confirmed that it was not significantly different from the mean of the 16 August 2009 Penns Creek females (T =
1.99, p = 0.06, 95% CI).
Female larvae collected from Penns Creek on 14 July 2010 had a mean head capsule width of 2.3471 mm (s.d. = 0.1586 mm, N = 17), and widths ranged from 2.0 mm to 2.6 mm. The head capsule width of the Spring Creek female collected on 4 August 2011 would fall within this range; however, it was significantly different from the corresponding mean of the 14 July 2010
Penns Creek females (T = 3.82, p = 0.001, 95% CI).
The execution of a reintroduction project requires a substantial amount of planning, and a balance of adherence to protocols and ability to be flexible procedurally.
Estimates of recolonization
On the evening of June 5, 2011, our team collected 27 jars of females and their eggs. If we use a conservative estimate of 50 females per jar and use Hoover (1978)’s estimate of 4597
77 eggs per Penns Creek E. guttulata female, then that would amount to 229,850 eggs per jar, and an estimated total of 6,205,950 eggs collected that evening.
The mating swarms were considerably less dense on the evening of June 3, 2011, and a total of 9 jars of females and their eggs were collected. Using the previously stated females-per- jar and eggs-per-female estimates, our total of that evening was 2,068,650 eggs, and for all of
2011 was 8,274,600 eggs. Bennett (2007) estimated that 0.5% of an Ephemera female’s eggs will produce adults—if so, then we could conceivably see 41,373 adults from the 2011 transplants.
In 2010, we collected a total of 39 jars of females and their eggs—potentially 8,964,150 eggs yielding 44,820 adults in 2010 alone. There were 49 jars in 2009, and thus potentially
112,626,550 eggs yielding 56,313 adults for that first year of transplants.
Kolar et al (1997) used logistic models to predict Hexagenia sp. recolonization of Lake
Erie:
Hexagenia population: dNh/dt = rhNh (1-atT) – (rh/Kh x Nh2) + I – ahNhNc – NhFh - aoNh
Chironomid population: dNc/dt = rcNc – (rc/Kc x Nc2) – acNhNc – NcFc
Fish predation on Hexagenia : dFh/dt = rfFh – (rf/Pf) x Fh2
Fish predation on Chironomids: dFc/dt = Pf – Fh
Low oxygen effect: If t = j , ao = 0.1 , otherwise ao = 1
Variables for equations:
Fh = fish predation on Hexagenia (they used 13 larvae m-2yr-1) Fc = fish predation on Chironomids I = immigration of Hexagenia (they used 2.6 larvae m-2yr-1) Nh = Hexagenia population Nc = Chironomid population T = toxicity (they used a 50% reduction in number of larvae during 20 years)
78 Coefficients for equations:
ah = Hexagenia competition rate (they used 0.0001) ac = Chironomid competition rate (they used 0.001) ao = dissolved O2 effect (they used 90% mortality within the first 20 years) at = toxic effect rate (random catastrophic event—used 50% reduction during 20 years) Kh = carrying capacity for Hexagenia (they used 350 larvae/m2) Kc = carrying capacity for Chironomids (they used ___ larvae/m2) Pf = total predation rate (they used 13 larvae m-2yr-1) rh = annual population growth for Hexagenia (they used 0.3, 0.6, 0.9 and 1.2) rc = annual population growth for Chironomids rf = increase in predation on Hexagenia
“r” estimates for Hexagenia spp. in lotic populations range from 0.12 to 0.3 (Horst 1976) to 0.6 (Nelson 1970, Hudson and Swanson 1972)
My modification of Kolar et al 1997:
Nt+1 = Nt + (rN - rN2/K) + I – (Pr&α*N)
Where “I” (for “Immigration”) represents an estimated number of first instars (per square
meter) hatched from the transplanted eggs, and where “Pr&α” (for “Predation & Competition”)
represents losses due to predation, competition, catastrophes, etc. Bennett (2007) estimated that
0.5% of an Ephemera sp. female’s eggs will produce adults (99.5% loss). Assuming E.
guttulata spend two years at the larval stage, then annual losses would be 49.75%; therefore,
Pr&α = 0.4975.
The model was run with r = 0.3, 0.6, 0.9, 1.2, and 2. Carrying capacity (K) was set at
either 200 or 250 larvae per square meter, assuming lotic Ephemera are more sensitive to
dissolved oxygen levels than lentic Hexagenia . The number of larvae at t = 0 was set at either 1
(similar to Kolar et al 1997) or at 10 (representing the first year of egg transplants, as there have been no existing populations of E. guttulata found in Spring Creek for decades). Also, if we
79 assume a two-year larval stage, there will be no adults in year one, and thus “r” in year one should be zero.
Figure 6.1 shows the prediction made by this model, if egg transplants continue indefinitely, yielding 50 larvae per year, carrying capacity is 200 larvae/m 2, and annual population growth (r) = 0.3. (The number of Ephemera larvae over time is per square meter.)
Number of Ephemera larvae over time r = 0.3, K = 200, I = 50
100
80
60
40
20 Number of larvae . . larvae of Number
0 1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49 51 Time in years
Figure 6-1 Number of E. guttulata larvae over time, with indefinite egg transplants (r = 0.3)
Figure 6.2 shows the prediction made by this model using the same parameters, but if egg
transplants are discontinued as of year 4:
80
Number of Ephemera larvae over time r = 0.3, K = 200, I = 50 for years 1-3, then I = 0
100
80
60
40
20 Number of larvae . . larvae of Number
0 1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49 51 Time in years
Figure 6-2 Number of E. guttulata larvae over time, if I = 0 as of year 4 (r = 0.3)
If we use the highest value of “r” for Hexagenia sp. (r = 0.6) according to Nelson (1970) and Hudson and Swanson (1972), conservatively increase the carrying capacity to 250 larvae/m 2 , and assume that transplants yield 100 larvae per year, then the population will persist if
“Immigration” is discontinued after year 10 (Figure 6.3):
Number of Ephemera larvae per square meter over time r = 0.6, K = 250, I = 100 for years 1-10, then I = 0
290 240 190 140 90
Number of larvae . . larvae of Number 40 -10 1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49 51 Time in years
Figure 6-3 Number of E. guttulata larvae over time, if I = 0 as of year 11 (r = 0.6)
81 Implications
Figures 6-1 through 6-3 illustrate the long-term nature of reintroduction projects. In other iterations of the model (not shown), when transplants ended after the third year, increasing the number of transplanted larvae by a factor of 10 did not lead to population persistence, but rather a more dramatically rapid extinguishment.
No one knew this better than the river keeper William James Lunn. For 45 years, Lunn
worked on the River Test (UK) for the Houghton Fishing Club (Hills 1936). Lunn is perhaps
most famous for his fly-patterns, but his most impressive accomplishment was the successful
reestablishment of mayfly populations in the Houghton waters. What is especially impressive was
his determination, as it took twenty-six years to accomplish.
Looking forward
Thanks in part to its 83,184 miles of streams and rivers (website: PA-DCNR), sportfishing in Pennsylvania contributes significantly to the state’s revenue. In 2001, more than
$800 million was spent on direct fishing expenditures in Pennsylvania (website: PA-F&BC).
Local economies are reported to benefit as well—a 1994 study of nine central Pennsylvania counties (Bedford, Blair, Cambria, Fayette, Fulton, Huntingdon, Indiana, Somerset, and
Westmoreland) found that direct sales related to fishing and boating generated $10.2 million, and supported 665 employees with a total of $9.5 million in income (website: PAF&BC).
Admittedly, more comprehensive sales, employment, and income data are needed to properly estimate the industry’s worth.
One of the most famous—and surely lucrative—draws to central Pennsylvania’s waterways is the “eastern green drake” mayfly. Their “hatch” (subimaginal emergence) and
82 eventual “spinner-fall” (post-coital mortality) are major events in sportfishing, as hungry trout will gorge themselves, and may not distinguish an angler’s imitation from the actual mayfly. The
Pennsylvania DCNR describes Poe Paddy State Park (through which Penns Creek flows), as “a trout angler's paradise featuring the nationally recognized green drake mayfly hatch” (website:
DCNRb). Reestablishment of Spring Creek’s E. guttulata population could potentially be a boon to the local economy.
On a broader scale, restoring biodiversity may also be an ecological boon. A recent article by Cardinale and his colleagues (2011) argued that greater diversity may actually improve the “health” of a stream. In this study, nitrate uptake was 4.5 times greater in tanks containing eight algal species than in tanks containing one species. It may be possible that such benefits of niche partitioning are not limited to primary producers.
Several organizations in the local community have contributed commendable efforts toward rectifying 55 years of neglect and misuse wrought on Spring Creek. Restoring the
“eastern green drake” mayfly can be seen as analogous to restoring community pride, representing a reward for decades of selfless labor.
I can only hope that it will not take twenty-six years to accomplish.
83
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Appendix A
Pollutants detected in substrate and water collected from Spring and Penns Creeks
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Appendix B
Macroinvertebrate taxa collected from Spring Creek and Penns Creek
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VITA
Hannah L. Stout
EDUCATION:
Ph.D. Entomology Aug 2005 – May 2012 The Pennsylvania State University University Park, PA
BS Biobehavioral Health, Neuroscience Minor Aug 1999 – Dec 2002 The Pennsylvania State University University Park, PA
PROFESSIONAL EXPERIENCE
Founder and Principal Investigator Jan 2009 – present Green Drake Restoration Project State College, PA
Graduate Research and Teaching Assistant Aug 2005 – May 2011 Department of Entomology The Pennsylvania State University University Park, PA
Assistant Director May 2008 – Jun 2008 Bug Camp for Kids The Pennsylvania State University University Park, PA
Docent Coordinator Apr 2006 – Jun 2008 Frost Entomological Museum The Pennsylvania State University University Park, PA
Insect Identification Specialist Jun 2006 – Aug 2006 Invertebrate Inventory of Gettysburg National Military Park Center for BioDiversity Research The Pennsylvania State University University Park, PA
AWARDS
Graduate Student Conservation Research Award May 2009 Society for Freshwater Science
College of Agricultural Sciences Graduate Student Travel Award Oct 2007 The Pennsylvania State University University Park, PA
William Yendol Memorial Research Award Oct 2007 The Pennsylvania State University University Park, PA